The Role of Chromatin in Molecular Mechanisms of Toxicity

Jonathan G. Moggs1 and George Orphanides

Syngenta CTL, Cheshire SK10 4TJ, United Kingdom

Received March 17, 2004; accepted April 29, 2004


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 CHANGES IN CHROMATIN DURING...
 CHANGES IN GENE EXPRESSION...
 THE ROLE OF CHROMATIN...
 THE ROLE OF CHROMATIN...
 HISTONE MODIFICATIONS SERVE AS...
 DO XENOBIOTICS TARGET CHROMATIN...
 SUMMARY
 REFERENCES
 
Eukaryotic cells store their genetic information in the form of a highly organized nucleoprotein complex termed chromatin. The high degree of compaction of DNA within chromatin places severe constraints on proteins that require access to the DNA template to facilitate gene transcription, DNA replication, and DNA repair. As a consequence, eukaryotic cells have developed sophisticated mechanisms to allow chromatin to be rapidly decompacted locally for access by DNA-binding proteins. Once thought to play only a structural role, it now appears that chromatin plays a key regulatory role by marshalling access to the DNA template. We have reviewed the role played by chromatin in the cellular response to physiological and toxicological stimuli and described how changes in chromatin structure may in the future be used as markers of toxicity. We also review the evidence that chromatin itself is the direct target of certain toxicants and that toxicant-induced perturbations in chromatin structure may precipitate adverse effects.

Key Words: chromatin; histone; toxicology; genome; epigenetic.


    PACKAGING OF DNA INTO CHROMATIN
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 THE ROLE OF CHROMATIN...
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 DO XENOBIOTICS TARGET CHROMATIN...
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A remarkably high degree of compaction is needed to accommodate the mammalian genome in the nucleus of a cell. This is achieved through the formation of a highly organized nucleoprotein structure known as chromatin (Van Holde, 1997Go). The basic repeating subunit of chromatin is the nucleosome, which consists of ~146 bp of DNA wrapped in almost two complete turns around a core octamer of histone proteins comprising two molecules each of histones H2A, H2B, H3, and H4 (Luger et al., 1997Go). Nucleosome core particles are separated from each other by short linker DNA segments (~30 to 40 bp in mammalian cells) whose length can vary among different cell types and species. Nucleosomes then undergo further folding and compaction to form the mature chromatin fiber. The architectural organization of these additional levels of compaction have not been defined, but are known to involve the linker histone H1 and additional nonhistone chromosomal proteins (Vaquero et al., 2003Go). The DNA compaction achieved through chromatin folding occurs at the expense of accessibility. Proteins that need access to DNA to carry out processes such as DNA transcription, replication, recombination, and repair are unable to reach sequences buried inside a chromatin fiber. For this reason, eukaryotes have developed sophisticated mechanisms that allow chromatin to be unravelled locally to facilitate access to DNA (Orphanides and Reinberg, 2002Go).


    CHANGES IN CHROMATIN DURING THE RESPONSE TO ENVIRONMENTAL STIMULI
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Once thought to play a largely passive, structural role, it is now clear that chromatin plays a pivotal role in the regulation of gene expression, chromosome replication, cell cycle progression, and the maintenance of genome integrity by marshalling access to the DNA template. The observation that the structure of chromatin around transcribed genes was more sensitive to attack by nucleases provided an early insight into the dynamic nature of chromatin (Van Holde, 1997Go). The molecular mechanisms that drive these rapid changes in chromatin organization have only recently begun to be defined. In this report, we have described the role played by chromatin in mediating cellular responses to certain classes of toxicant.


    CHANGES IN GENE EXPRESSION IN RESPONSE TO XENOBIOTICS
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One of the principle ways in which a eukaryotic cell responds to changes in its environment is by altering gene expression to change the complement of expressed proteins and, thereby, maintain homeostasis. Consequently, practically all toxic events will result in changes in gene expression. The acetylation of lysine residues in the N-terminal tails of the histone proteins has long been associated with actively transcribed genes (Hebbes et al., 1988Go). However, the role of histone acetylation in the regulation of gene expression awaited discovery of the enzymes responsible for this post-translational modification. The acetylation and deacetylation of histone lysine residues is catalyzed by histone acetyltransferases (HATs; reviewed by Carrozza et al., 2003Go) and histone deacetylases (HDACs; reviewed by de Ruijter et al., 2003Go), respectively. The identification of the first HATs underscored the importance of chromatin in regulating gene expression. Remarkably, many of the HATs were revealed to be previously identified transcriptional coactivators that are recruited to active genes by sequence-specific transcriptional activators. This suggested that the primary role of transcriptional activator proteins is to recruit activities that can alter chromatin structure to increase DNA accessibility. In support of this assertion, ATP-dependent, chromatin-remodeling enzymes, which increase DNA accessibility by using energy derived from the hydrolysis of ATP to alter the structure and position of nucleosomes, are also recruited to transcribed genes by transcription factors (reviewed by Becker and Horz, 2002Go).

These observations have, in recent years, precipitated a renewed interest in chromatin structure. This resulted in the seminal hypothesis that the modification of histone tails by lysine acetylation, lysine and arginine methylation, serine and threonine phosphorylation, and lysine ubiquitination constitutes a histone code that, when translated, dictates the activity of a gene (Strahl and Allis, 2000Go; Turner, 2000Go). The central role of these complex chromatin alterations in the response to toxicants is illustrated by the way in which xenobiotics that target nuclear receptors elicit their effects. The nuclear receptors, of which there are more than 48 in mammals, are a family of ligand-activated transcription factors (Enmark and Gustafsson, 2001Go). A growing number of xenobiotics have been shown to function by directly altering the activity of nuclear receptors. For example, peroxisome proliferators, dioxins, and estrogenic chemicals act via the peroxisome proliferator–activated receptor {alpha} (Johnson et al., 2002Go), aryl hydrocarbon receptor (Denison and Nagy, 2003Go), and estrogen receptors (ER; Singleton and Khan, 2003Go), respectively.

Once activated by ligand (e.g., a toxicant), it appears that the major mechanistic role of nuclear receptors is to induce alterations in chromatin structure to allow RNA polymerase II and its accessory transcription factors access to the gene to be transcribed (Fig. 1). The molecular mechanism through which ligand activation of a nuclear receptor leads to alterations in chromatin structure and gene activation has been studied in detail for ER{alpha} (Moggs and Orphanides, 2001Go). Ligand binding by ER{alpha} induces conformational changes in the receptor that facilitate the successive recruitment and exchange of 46 coregulator proteins during the recognition of specific estrogen response element sequences near to or within ER{alpha} target genes (Metivier et al., 2003Go). These coregulators include ATP-dependent, nucleosome-remodeling enzymes (the BRG1, BRM1, INI1, and BAF170 subunits of the SWI/SNF complex) and histone-modifying enzymes (the HATs CBP, p300, p/CAF, GCN5, and TIP60; HDAC1 and HDAC7; and the histone methyltransferases [HMTs] CARM1 and PRMT1) that together lead to the decompaction of chromatin structure locally.



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FIG. 1. Alterations in chromatin structure during gene activation by xenobiotic-activated nuclear receptors. Once activated by xenobiotics, it appears that the major mechanistic role of nuclear receptors is to induce alterations in chromatin structure (histone protein modifications and chromatin remodeling) to allow RNA polymerase II and its accessory transcription factors access to the gene to be transcribed. The process is initiated by the binding of a ligand (e.g., a toxicant) to a nuclear receptor. This initiates a conformational change in the receptor that facilitates the binding of coactivator proteins. Following binding of the activated nuclear receptor/coactivator complex to gene promoters, the coactivators are able to induce local changes in chromatin structure by modifying histones post-translationally and remodeling chromatin organization. This makes DNA sequences at the transcriptional start site accessible to RNA polymerase II and the general transcription machinery and facilitates the initiation of gene transcription. NR, nuclear receptor; GTFs, general transcription factors; RE, nuclear receptor DNA response element; Ac, histone lysine acetylation; Me, histone lysine or arginine methylation; SWI/SNF, ATP-dependent nucleosome remodeling enzyme; HATs, histone acetyltransferases; HMTs, histone methyltransferases; HDACs, histone deacetylases.

 
The precise way in which histone post-translational modifications lead to alterations in gene activity has not yet been defined in detail. However, additional proteins are likely to recognize these modifications, effectively reading the histone code, leading to an increase in DNA accessibility to facilitate transcription (Fischle et al., 2003aGo; Jenuwein and Allis, 2001Go; Turner, 2002Go). There is a precedent for the recognition of histone modifications by an extensive family of evolutionarily conserved protein modules, chromodomains and bromodomains, found in many proteins associated with chromatin. Chromodomains within the chromatin-silencing proteins polycomb and heterochromatin protein 1 selectively dock onto methylated lysines 9 and 27, respectively, of histone H3 (Bannister et al., 2001Go; Fischle et al., 2003bGo; Min et al, 2003Go), and a similar selective recognition of acetylated histones is exhibited by multiple bromodomain-containing proteins (Kanno et al., 2004Go). In this way, chromatin is a key mediator in the cellular response to toxicants that target nuclear receptors.


    THE ROLE OF CHROMATIN IN THE ACTIVATION OF STRESS RESPONSE GENES
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It is well established that toxicants cause rapid alterations in gene expression by activating protein kinase signaling cascades (reviewed by Tibbles and Woodgett, 1999Go; Yang et al., 2003Go). The resulting rapid, defensive alterations in gene activity require the transmission of a signal directly to the histones present in the chromatin of stress response genes. Within minutes of exposure to toxicants, such as the stress-inducing chemical anisomycin, the phosphorylation of serine 10 of histone H3 and the acetylation of lysines 9 and/or 14 take place (Clayton et al., 2000Go). Additional toxicants that induce rapid phosphorylation of serine 10 on histone H3 include arsenite (He et al., 2003Go) and the DNA-damaging agent cisplatin (Wang and Lippard, 2004Go). Anisomycin-induced histone H3 serine 10 phosphorylation appears to be carried out by the protein kinases MSK1 or RSK2, which themselves are regulated directly by the ERK and p38 MAP kinase signaling cascades (Sassone-Corsi et al., 1999Go; Thomson et al., 1999Go). This has been termed the nucleosomal response and facilitates the rapid alterations in chromatin structure and gene activity required for the proper cellular response to potentially deleterious agents (Clayton and Mahadevan, 2003Go). Moreover, these modifications are believed to be essential for the full transcriptional response to stress-inducing chemicals.


    THE ROLE OF CHROMATIN IN THE RESPONSE TO GENOTOXINS
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Direct damage of DNA by genotoxic agents is the primary mechanism of chemical carcinogenesis. The maintenance of genome integrity in eukaryotes involves a battery of damage surveillance and DNA repair enzymes. Like the gene expression machinery described above, these enzymes must operate within the repressive chromatin environment of the nucleus (reviewed by Green and Almouzni, 2002Go). DNA damage induced by genotoxic agents that covalently modify DNA bases or generate single-strand DNA breaks are primarily recognized and repaired by DNA excision repair pathways (Wood, 1996Go). In an analogous manner to the chromatin decompaction that accompanies the recruitment of RNA polymerase II during gene activation, DNA excision repair is associated with increased histone acetylation and localized chromatin remodeling (Green and Almouzni, 2002Go).

Double-strand breaks (DSBs) are a more severe form of DNA damage and can lead to genomic rearrangements if left unrepaired. Specific changes in chromatin modification state and structure appear to be at the center of the cellular response to DSBs (Fig. 2). The detection of DNA DSBs by the damage surveillance machinery results in the activation of protein kinases (ATM, ATR, and DNA-PK) that phosphorylate serine 139 within a specialized histone variant, histone H2A.X (reviewed by Downs and Jackson, 2003Go). The H2A.X variant makes up between 10 and 15% of total cellular H2A in higher eukaryotes, possesses an extended carboxy terminal tail that extends outwards from the nucleosome, and is incorporated into nucleosomes randomly throughout the genome. The role of histone H2A.X phosphorylation in DSB repair has not been defined in detail. However, recent evidence suggests that H2A.X functions as a tumor suppressor gene that protects cells from the deleterious effects of DNA damage. Transgenic mice lacking a functional H2A.X gene have unstable genomes and are susceptible to tumors, indicating that H2A.X functions to maintain genome integrity (Bassing et al., 2003Go; Celeste et al., 2003Go). Phosphorylation of serine 139 of histone H2A.X also occurs during apoptosis, a process that involves chromatin condensation and DNA fragmentation, the latter of which also generates DSBs.



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FIG. 2. Histone modifications as early markers of DNA damage. The sensing of double-strand DNA breaks (e.g., produced by ionizing radiation or DNA-binding drugs) is accompanied by the activation of protein kinases that rapidly phosphorylate the specialized histone variant H2A.X and also transduce signals to additional signaling and DNA repair proteins. Phosphorylated H2A.X is thought to function by recruiting additional proteins that may enhance the accuracy or efficiency of DNA repair. P, phosphorylation of serine 139 within the specialized histone variant histone H2A.X.

 

    HISTONE MODIFICATIONS SERVE AS MARKERS OF TOXICITY
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The observation that rapid and specific post-translational histone modifications accompany defined cellular responses to certain toxicants suggests that they may serve as useful markers of toxicity. For example, phosphorylation of histone H2A.X occurs very rapidly upon the formation of DSBs and, therefore, may act as a sensitive and specific marker for this type of cellular insult. Antibodies that recognize specifically histone H2A.X phosphorylated at position 139 are available commercially and have been used to detect DSBs resulting from DNA damage (Banath and Olive, 2003Go; Olive and Banath, 2004Go; Taneja et al., 2004Go). Moreover, the same antibodies can be used to detect apoptosis. During apoptosis, H2A.X phosphorylation occurs long before morphological changes such as internucleosomal fragmentation or phosphotidylserine externalization (Rogakou et al., 2000Go). Therefore, immunostaining for H2A.X phosphorylation is a sensitive marker for the onset of apoptosis. Another histone modification that occurs during apoptosis is the phosphorylation of histone H2B on serine 14 (Cheung et al., 2003Go). In contrast to H2A.X phosphorylation, H2B phosphorylation occurs later in the apoptotic pathway during the chromatin condensation step that precedes genome fragmentation; therefore, it marks cells that are committed to apoptosis. The use of chromatin changes as biomarkers is not limited to alterations in histone post-translational modifications. Recent data reveal that a linker histone variant, histone H1.2, is used as a signaling molecule during DNA damage–induced apoptosis (Konishi et al., 2003Go). DNA DSBs induce translocation of nuclear H1.2 to the cytoplasm, where it promotes the release of cytochrome c from mitochondria by activating the Bcl-2 family protein BAK. Thus, the appearance of histone H1.2 in the cytoplasm will mark DNA damage–induced apoptosis.


    DO XENOBIOTICS TARGET CHROMATIN STRUCTURE DIRECTLY?
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Evidence is accumulating that toxicants may elicit their effects by targeting chromatin structure directly. Proof that direct perturbation of chromatin integrity can lead to adverse effects is provided by the observations that mutations, in components of the machinery that regulates chromatin structure, directly precipitate human disease (Table 1; Ausio et al., 2003Go; Huang et al., 2003Go). For example, chromosomal translocations of the genes encoding the HAT enzyme CBP and the HMT enzyme MLL are frequently associated with leukemias (Schneider et al., 2002Go; Timmermann et al., 2001Go). Also, mutations in the gene encoding the histone H3 kinase RSK2 are found in patients with Coffin-Lowry syndrome (Sassone-Corsi et al., 1999Go). Therefore, it is possible that direct disruption by toxicants of the enzymes that maintain proper chromatin organization leads to adverse effects. In support of this hypothesis, the toxic metal nickel can inhibit the GCN5 HAT enzyme in vitro (Broday et al. 2000Go); and the inhibition of histone acetylation by nickel compounds in vivo has been associated with the silencing of gene expression (Yan et al., 2003Go). Furthermore, the transformation of cultured cells to anchorage- independent growth by nickel compounds can be inhibited and reversed by the HDAC inhibitor trichostatin A (Zhang et al., 2003Go). Thus, concomitant alterations in histone acetylation and gene expression appear to be important for nickel-induced cellular transformation.


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TABLE 1 Association of Human Disease with Defects in Chromatin Structure

 
It will be important to determine the specificity of toxicants such as nickel for the numerous enzymes involved in regulating histone acetylation, particularly in light of the observations that nickel exposure can also alter chromatin structure through the truncation of histone H2A via cleavage of an octapeptide from its C-terminus (Karaczyn et al., 2003Go). Interestingly, another carcinogenic metal, chromium, has recently been shown to perturb HAT and HDAC enzyme occupancy on polycyclic aromatic hydrocarbon–inducible genes, resulting in the inhibition of their transcription (Wei et al., 2004Go). Chromium is known to form DNA protein cross-links, and these might be responsible for blocking the normal function of chromatin-modifying enzymes during aryl hydrocarbon receptor–mediated gene activation.

Another class of chromatin-modifying enzymes that can be directly targeted by toxicants is the HDACs. The promising anticancer agent suberoylanilide hydroxamic acid (Kelly et al., 2003Go) is an HDAC inhibitor that interferes with differentiation, proliferation, and apoptosis in tumor cells. More recently, the short-chain fatty acid and endocrine disruptor methoxyacetic acid has been shown to enhance nuclear receptor activity through a combination of MAP kinase activation and HDAC inhibition (Jansen et al., 2004Go). The disruption of chromatin-modifying enzymes by toxicants might, in principle, be expected to result in a nonspecific dysregulation of genome function and, thus, overt cellular toxicity. However, there is increasing evidence that many chromatin-modifying enzymes target specific subsets of genes. This is exemplified by the distinct transcriptional specificities of the human SWI/SNF BRG1 and BRM nucleosome-remodeling complexes (Kadam and Emerson, 2003Go). Furthermore, there is a precedent for the specific inhibition of distinct members of the HAT family: the HATs p300 and PCAF are inhibited by the peptide CoA conjugates Lys-CoA and H3-CoA-20, respectively (Lau et al., 2000Go).

What would be the consequences of direct chemical perturbation of chromatin structure? In addition to their role in regulating gene expression and DNA repair, chromatin modifications play an important part in the transmission of epigenetic information, epigenetics being the study of heritable alterations in gene expression that occur in the absence of changes in genome sequence (Wolffe and Matzke, 1999Go). Thus, the perturbation of chromatin structure by toxicants may lead to long-term and possibly transgenerational changes in epigenetic programming. The key mechanism that controls the epigenetic regulation of mammalian genomes is the methylation of cytosine bases in DNA (Beck and Olek, 2003Go), which forms the modified base 5-methylcytosine (5-mC). In mammalian DNA, 5-mC is present at a level of 2–5% of all cytosines and is found predominantly on CpG dinucleotides. Clusters of CpG sequences (known as CpG islands) tend to be found near the 5' ends of genes and are usually unmethylated. However, a proportion of these CpGs can be methylated in a developmental stage– and cell type–specific manner, usually resulting in gene silencing. The fact that more than 50 genes have been shown to be abnormally methylated in human tumors suggests that alterations in DNA methylation status may lead to cellular transformation and carcinogenesis (Jones and Baylin, 2002Go). Specifically, it is believed that hypomethylation and hypermethylation of CpG islands can lead to the constitutive activation of oncogenes and the silencing of tumor suppressor genes, respectively.

The influence of chromatin structure on DNA methylation status and the transmission of epigenetic information is just beginning to be revealed (reviewed by Jaenisch and Bird, 2003Go). DNA methylation is controlled by families of DNA methyltransferase (DNMT) enzymes and methyl-cytosine–binding proteins (MBDs). The interrelationship between DNA methylation and chromatin structure was revealed initially by the identification of a protein complex containing both the MBD MeCP2 and the histone deacetylases HDAC1 and HDAC2 (Nan et al, 1998Go). Subsequent studies have revealed numerous interactions among DNMTs, MBDs, and chromatin-modifying enzymes (reviewed by Burgers et al., 2002Go). Regions of DNA that are hypermethylated are associated with chromatin that has been modified by methylation of lysine 9 on histone H3 (Lachner et al., 2003Go). Moreover, this particular histone modification appears to be important for DNA methylation–driven gene silencing. Chromatin-containing histone H3 modified at this position is recognized by the HP1 protein, which binds and promotes compaction of chromatin into the silent heterochromatin state (Bannister et al., 2001Go). Given the role of histone H3 lysine 9 methylation in the silencing of hypermethylated DNA, one may expect that disruption of histone-methylating enzymes (the HMTs) will have adverse consequences. Compelling support for this hypothesis is provided by the observation that stable alterations in the expression of the HMT enzyme EZH2, which methylates histone H3 lysine 27, are associated with numerous prostate and breast cancers (Kleer et al., 2003Go; Varambally et al., 2002Go). Therefore, we speculate that direct inhibition of the machinery that regulates chromatin structure will disrupt epigenetic programming via perturbations in chromatin function and will precipitate adverse effects (Bombail et al., 2004Go).


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It is now clear that chromatin plays an important role in mediating the response to physiological and toxicological stimuli. This is highlighted by the fact that changes in the modification state and structure of chromatin are among the fastest responses to toxicant exposure. The association of a growing number of human diseases with defects in the machinery that controls chromatin structure underlines the importance of chromatin in the maintenance of cellular homeostasis. Consequently, we must consider the possibility that direct perturbation of chromatin structure, perhaps by toxicants that inhibit chromatin- modifying enzymes, may be central to the mechanisms of action of some agents.


    NOTES
 

1 To whom correspondence should be addressed at Syngenta CTL, Alderley Park, Cheshire SK10 4TJ, United Kingdom. Fax: +44 1625 585715. E-mail: jonathan.moggs{at}syngenta.com.


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