Lewin’s Genes XII

26.1 Introduction

The phenotypic differences that distinguish the various kinds of cells in a higher eukaryote are largely due to differences in the expression of genes that code for proteins; that is, those transcribed by RNA polymerase II. In principle, the expression of these genes can be regulated at any one of several stages. FIGURE 26.1 distinguishes (at least) six potential control points, which form the following series:

Activation of gene structure: open chromatin

↓

Initiation of transcription and elongation

↓

Processing the transcript

↓

Transport to the cytoplasm from the nucleus

↓

Translation of mRNA

↓

Degradation and turnover of mRNA

FIGURE 26.1 Gene expression is controlled principally at the initiation of transcription. Control of processing may be used to determine which form of a gene is represented in mRNA. The mRNA may be regulated during transport to the cytoplasm, during translation, and by degradation.

Whether a gene is expressed depends on the structure of chromatin both locally (at the promoter) and in the surrounding domain. Chromatin structure correspondingly can be regulated by individual activation events or by changes that affect a wide chromosomal region. The most localized events concern an individual target gene, where changes in nucleosomal structure and organization occur in the immediate vicinity of the promoter. Many genes have multiple promoters; the choice of the promoter can alter the pattern of regulation and influence how the mRNA is used because it will change the 5′ untranslated region (UTR). More general changes may affect regions as large as a whole chromosome. Activation of a gene requires changes in the state of chromatin. The essential issue is how the transcription factors gain access to the promoter DNA.

Local chromatin structure is an integral part of controlling gene expression. Broadly speaking, genes may exist in either of two basic structural conditions. The first is an inactive gene in closed chromatin. Alternatively, genes are found in an “active” state, or open chromatin, only in the cells in which they are expressed, or potentially expressed. The change of structure precedes the act of transcription and indicates that the gene is able to be transcribed. This suggests that acquisition of the active structure must be the first step in gene expression. Active genes are typically found in domains of euchromatin with a preferential susceptibility to nucleases, and hypersensitive sites are created at promoters before a gene is activated (see the Chromatin chapter). A gene that is in open chromatin may actually be active and be transcribed, or it may be potentially active and waiting for a subsequent signal, a condition called poised.

An intimate and continuing connection exists between initiation of transcription and chromatin structure. Some activators of gene transcription directly modify histones; in particular, acetylation of histones is associated with gene activation. Conversely, some repressors of transcription function by deacetylating histones. Thus, a reversible change in histone structure in the vicinity of the promoter is involved in the control of gene expression. These changes influence the association of histone octamers with DNA and are responsible for controlling the presence and structure of nucleosomes at specific sites. This is an important aspect of the mechanism by which a gene is maintained in an active or inactive state.

The mechanisms by which regions of chromatin are maintained in an inactive (silent) state are related to the means by which an individual promoter is repressed. The proteins involved in the formation of heterochromatin act on chromatin via the histones, and modifications of the histones are an important feature in the interaction. Once established, such changes in chromatin can persist through cell divisions, creating an epigenetic state in which the properties of a gene are determined by the self-perpetuating structure of chromatin. The name epigenetic reflects the fact that a gene may have an inherited condition (it may be active or inactive) that does not depend solely on its sequence (see the chapters titled Epigenetics I and Epigenetics II). Once transcription begins, regulation during the elongation phase of transcription is also possible (see the Eukaryotic Transcription chapter). However, attenuation, such as that in bacteria (see the chapter titled The Operon), cannot occur in eukaryotes because of the separation of chromosomes from the cytoplasm by the nuclear membrane. The primary mRNA transcript is modified by capping at the 5′ end and for most protein-coding genes is also modified by polyadenylation at the 3′ end (see the chapter RNA Splicing and Processing). Many genes also have multiple termination sites, which can alter the 3′ UTR, and thus mRNA function and behavior.

Introns must be excised from the transcripts of interrupted genes. The mature RNA must then be exported from the nucleus to the cytoplasm. Regulation of gene expression at the level of nuclear RNA processing might involve any or all of these stages, but the one that has the most evidence concerns changes in splicing; some genes are expressed by means of alternative splicing patterns whose regulation controls the type of protein product (see the RNA Splicing and Processing chapter).

The translation of an mRNA in the cytoplasm can be specifically controlled, as can the turnover rate of the mRNA. This can also involve the localization of the mRNA to specific sites where it is expressed; in addition, the blocking of initiation of translation by specific protein factors may occur. Different mRNAs may have different intrinsic half-lives determined by specific sequence elements (see the chapter mRNA Stability and Localization).

Regulation of tissue-specific gene transcription lies at the heart of eukaryotic differentiation. It is also important for control of metabolic and catabolic pathways. Gene regulators are typically proteins; however, RNAs can also serve as gene regulators. This raises two questions about gene regulation:

How does a protein transcription factor identify its group of target genes?
How is the activity of the regulator itself regulated in response to intrinsic or extrinsic signals?

26.2 How Is a Gene Turned On?

Multicellular eukaryotes typically begin life through the fertilization of an egg by a sperm. In both of these haploid gametes, but especially the sperm, the chromosomes are in super-condensed modified chromatin. Males of some species use positively charged polyamines, such as spermines and spermidines, to replace the histones in sperm chromatin; others include sperm-specific histone variants. Once the process of fusion of the two haploid nuclei is complete in the egg, genes are then activated in a cascade of regulatory events. The general question of how a gene in closed chromatin is turned on can be broken down into (at least) two parts: How is an individual gene that is wrapped up in condensed chromatin identified and targeted for activation? Furthermore, once histone modification and chromatin remodeling begin, how are those processes prevented from spreading to genes that should not be turned on?

First, imagine that replication is one mechanism by which closed chromatin can be disrupted in order to allow DNA-binding sequences to become accessible. Replication opens higher-order chromatin structure by temporarily displacing histone octamers. The occupation of enhancer DNA sites on daughter strands subsequently can be viewed as competition between nucleosomes and gene regulators. Chromatin can be opened if transcription factors are present in high enough concentration, as shown in FIGURE 26.2. If the transcription factor concentration is low, then nucleosomes can bind and condense the region. This occurs in Xenopus embryos as oocyte-specific 5S ribosomal genes are repressed in the embryo after fertilization.

FIGURE 26.2 When replication disrupts chromatin structure, after the Y fork has passed, either chromatin can reform or transcription factors can bind and prevent chromatin formation.

Second, it is clear that some transcription factors can bind to their DNA target sequence in closed chromatin. The DNA exposed on the surface of the histone octamer is potentially accessible. These transcription factors can then recruit the histone modifiers and chromatin remodelers to begin the process of opening the gene region and clearing the promoter (see the section titled Chromatin Remodeling Is an Active Process later in this chapter). Recently described examples of antisense transcription through a gene region can facilitate this process; these are described in more detail in the Noncoding RNA chapter.

Chromatin modification typically originates from a point source (such as an enhancer) and then spreads, in most cases bidirectionally. (In those cases where modification spreads in a unidirectional fashion, the question becomes why it is not spread bidirectionally.) The next question is, what prevents chromatin modification from spreading into distant gene regions?

Activation (as well as repression) is limited by boundaries called insulators or boundary elements (see the Chromatin chapter). Very few of these insulators have been described in detail, and their mechanisms of action are still poorly understood. In one sense, they are very much like enhancers. They are modular, compact sequence sets that bind specific proteins. Insulators can also function within complex loci to separate multiple temporal and tissue-specific enhancers so that only one can function at a time. Boundary elements are also required to prevent the heterochromatin at regions such as the centromeres and telomeres from spreading into euchromatin.

26.3 Mechanism of Action of Activators and Repressors

Initiation of transcription involves many protein–protein interactions between transcription factors bound at enhancers with the basal apparatus that assembles at the promoter, including RNA polymerase. These transcription factors can be divided into two opposing classes: positive activators and negative repressors.

As discussed in the chapter titled The Operon, positive control in bacteria entails a regulator that aids the RNA polymerase in the transition from the closed complex to the open complex. Transcription factors, such as CRP (catabolite repressor protein), in Escherichia coli, typically bind close to the promoter to allow the C-terminal domain of the α subunit of RNA polymerase to make direct physical contact. This usually occurs in a gene having a poor promoter sequence. The activator functions to overcome the inability of the RNA polymerase to open the promoter. Positive control in eukaryotes is quite different. Three classes of activators can be identified that differ by function.

The first class is the true activators (see the Eukaryotic Transcription chapter). These are the classical transcription factors that function by making direct physical contact with the basal apparatus at the promoter (see the next section titled Independent Domains Bind DNA and Activate Transcription) either directly or indirectly, through a coactivator. These transcription factors function on DNA or chromatin templates.

The activity of a true activator may be regulated in any one of several ways, as illustrated schematically in FIGURE 26.3:

A factor is tissue specific because it is synthesized only in a particular type of cell. This is typical of factors that regulate development, such as homeodomain proteins.
The activity of a factor may be directly controlled by modification. HSF (heat shock transcription factor) is converted to the active form by phosphorylation.
A factor is activated or inactivated by binding a ligand. The steroid receptors are prime examples. Ligand binding may influence the localization of the protein (causing transport from cytoplasm to nucleus), as well as determine its ability to bind to DNA.
Availability of a factor may vary; for example, the factor NF-κB (which activates immunoglobulin κ genes in B lymphocytes) is present in many cell types. It is sequestered or masked in the cytoplasm, however, by the inhibitory protein I-κB. In B lymphocytes, NF-κB is released from I-κB and moves to the nucleus, where it activates transcription.
A dimeric factor may have alternative partners. One partner may cause it to be inactive; synthesis of the active partner may displace the inactive partner. Such situations may be amplified into networks in which various alternative partners pair with one another, especially among the helix-loop-helix (HLH) proteins.
The factor may be cleaved from an inactive precursor. One activator is produced as a protein bound to the nuclear envelope and endoplasmic reticulum. The absence of sterols (such as cholesterol) causes the cytosolic domain to be cleaved; it then translocates to the nucleus and provides the active form of the activator.

FIGURE 26.3 The activity of a positive regulatory transcription factor may be controlled by (a) synthesis of protein, (b) covalent modification of protein, (c) ligand binding, or (d) binding of inhibitors that sequester the protein or affect its ability to bind to DNA (e) by the ability to select the correct binding partner for activation and (f) by cleavage from an inactive precursor.

The second class includes the antirepressors. When one of these activators is bound to its enhancer, it recruits the histone modifier enzymes and/or the chromatin remodeler complexes to convert the chromatin from the closed state to the open state. This class has no activity on a DNA template; it only functions on chromatin templates (described later in the section Chromatin Remodeling Is an Active Process).

The third class includes architectural proteins, such as Yin-Yang; these proteins function to bend the DNA, either bringing bound proteins together to facilitate forming a cooperative complex or bending the DNA the other way to prevent complex formation, as shown in FIGURE 26.4. Note that a strand of DNA may thus be bent in two different directions depending on whether the regulator binds to the top or to the bottom. This is a difference of one-half of a turn of the helix, which is about 5 bp (10.5 bp per turn).

FIGURE 26.4 Architectural proteins control the structure of DNA and thus control whether bound proteins can contact each other.

Several examples of negative control in bacteria, in the lac operon and in the trp operon, were described in the chapter titled The Operon. Repression can occur in bacteria when the repressor prevents the RNA polymerase from converting the promoter from the closed complex to the open complex, as in the lac operon, or bind to the promoter sequence to prevent RNA polymerase from binding, as in the trp operon. Many more mechanisms have been identified by which repressors act in eukaryotes, some of which are illustrated in FIGURE 26.5:

One mechanism of action by which a eukaryotic repressor can prevent gene expression is to sequester an activator in the cytoplasm. Eukaryotic proteins are synthesized in the cytoplasm. Proteins that function in the nucleus have a domain that directs their transport through the nuclear membrane. A repressor can bind to that domain and mask it.
Several variations of that mechanism are possible. One that takes place in the nucleus occurs when the repressor binds to an activator that is already bound to an enhancer and masks its activation domain, thus preventing it from functioning, such as with the Gal80 repressor (see the section later in this chapter titled Yeast GAL Genes: A Model for Activation and Repression).
Alternatively, the repressor can be masked and held in the cytoplasm until it is released to enter the nucleus.
A fourth mechanism is simple competition for an enhancer, where either the repressor and activator have the same binding site sequence or have overlapping but different binding site sequences. This is a very versatile mechanism for a cell because there are two variables at work here: One is strength of a factor binding to DNA, and the second is factor concentration. By only slightly varying the concentration of a factor, a cell can dramatically alter its developmental path.

FIGURE 26.5 A repressor may control transcription by (a) sequestering an activator in the cytoplasm, (b) by binding an activator and masking its activation domain, (c) by being held in the cytoplasm until it is needed, or (d) by competing with an activator for a binding site.

The transcription factors that recruit the histone modifiers and chromatin remodelers have as their counterparts repressors that recruit the complexes that undo (or change) the modifications and remodeling. The same is true for the architectural proteins, where, in fact, the same protein bound to a different site prevents activator complexes from forming.

26.4 Independent Domains Bind DNA and Activate Transcription

The actions of the activator class of transcription factors are the most well-known. Activators must be able to perform multiple functions:

Activators recognize specific DNA target sequences located in enhancers that affect a particular target gene.
Having bound to DNA, an activator exercises its function by binding to components of the basal transcription apparatus.
Many activators require a dimerization domain to form complexes with other proteins.

Can the domains in the activator that are responsible for these activities be characterized? Often an activator has one domain that binds DNA and another, separate domain that activates transcription. Each domain behaves as a separate module that functions independently when it is linked to a domain of the other type. The geometry of the overall transcription complex must allow the activating domain to contact the basal apparatus irrespective of the exact location and orientation of the DNA-binding domain.

Enhancer elements near the promoter may still be an appreciable distance from the start point, and in many cases may be oriented in either direction. Enhancers may even be farther away and always show orientation independence. This organization has implications for both the DNA and proteins. The DNA may be looped or condensed in some way to allow the formation of the transcription complex, permitting interactions between factors bound at both the enhancer and the promoter. In addition, the domains of the activator may be connected in a flexible way, as illustrated in FIGURE 26.6. The main point here is that the DNA-binding and activating domains are independent and are connected in a way that allows the activating domain to interact with the basal apparatus irrespective of the orientation and exact location of the DNA-binding domain.

FIGURE 26.6 DNA-binding and activating functions in a transcription factor may comprise independent domains of the protein.

Binding to DNA is usually necessary for activating transcription, but some transcription factors function without a DNA-binding domain by virtue of protein–protein interactions. Does activation depend on the particular DNA-binding domain? This question has been answered by making hybrid proteins that consist of the DNA-binding domain of one activator linked to the activation domain of another activator. The hybrid functions in transcription at sites dictated by its DNA-binding domain, but in a way determined by its activation domain.

This result fits the modular view of transcription activators. The function of the DNA-binding domain is to bring the activation domain to the basal apparatus at the promoter. Precisely how or where it is bound to DNA is irrelevant, but once it is there, the activation domain can play its role. This explains why the exact locations of DNA-binding sites can vary. The ability of the two types of modules to function in hybrid proteins suggests that each domain of the protein folds independently into an active structure that is not influenced by the rest of the protein.

26.5 The Two-Hybrid Assay Detects Protein–Protein Interactions

The model of domain independence is the basis for an extremely useful assay for detecting protein interactions. The principle is illustrated in FIGURE 26.7. One of the proteins to be tested is fused to a DNA-binding domain. The other protein is then fused to a transcription-activating domain. This is accomplished by linking the appropriate coding sequences in each case and making chimeric proteins by expressing each hybrid gene.

FIGURE 26.7 The two-hybrid technique tests the ability of two proteins to interact by incorporating them into hybrid proteins, where one has a DNA-binding domain and the other has a transcription-activating domain.

If the two proteins that are being tested can interact with one another, the two hybrid proteins will interact. This is reflected in the name of the technique: the two-hybrid assay. The protein with the DNA-binding domain binds to a reporter gene that has a simple promoter containing its target site. It cannot, however, activate the gene by itself. Activation occurs only if the second hybrid binds to the first hybrid to bring the activation domain to the promoter. Any reporter gene can be used where the product is readily assayed, and this technique has given rise to several automated procedures for rapidly testing protein–protein interactions.

The effectiveness of the technique dramatically illustrates the modular nature of proteins. Even when fused to another protein, the DNA-binding domain can bind to DNA, and the transcription-activating domain can activate transcription. Correspondingly, the interaction ability of the two proteins being tested is not inhibited by the attachment of the DNA-binding or transcription-activating domains. (Of course, there are some exceptions for which these simple rules do not apply, and interference between the domains of the hybrid protein prevents the technique from working.)

The power of this assay is that it requires only that the two proteins being tested can interact with each other. They need not have anything to do with transcription (in fact, if the proteins being tested themselves are involved in transcription, it can frequently lead to false positives, as a single hybrid may work as an activator). As a result of the independence of the DNA-binding and transcription-activating domains, all that is required is that they are brought together. This will happen so long as the two proteins being tested can interact in the environment of the nucleus.

26.6 Activators Interact with the Basal Apparatus

The true activator class of transcription factors may work directly when it consists of a DNA-binding domain linked to a transcription-activating domain, as illustrated earlier in Figure 26.5. In other cases, the activator does not itself have a transcription-activating domain (or contains only a weak activation domain), but binds another protein—a coactivator—that has the transcription-activating activity. FIGURE 26.8 shows the action of such an activator. Coactivators can be regarded as transcription factors whose specificity is conferred by the ability to bind to proteins that bind to DNA instead of directly to DNA. A particular activator may require a specific coactivator.

FIGURE 26.8 An activator may bind a coactivator that contacts the basal apparatus.

Although the protein components are organized differently, the mechanism is the same. An activator that contacts the basal apparatus directly has an activation domain covalently connected to the DNA-binding domain. When an activator works through a coactivator, the connections involve noncovalent binding between protein subunits (compare Figures 26.5 and 26.6). The same interactions are responsible for activation, irrespective of whether the various domains are present in the same protein subunit or divided into multiple protein subunits. In addition, many coactivators also contain additional enzymatic activities that promote transcription activation, such as activities that modify chromatin structure (see the section later in this chapter titled Histone Acetylation Is Associated with Transcription Activation).

An activation domain works by making protein–protein contacts with general transcription factors that promote assembly of the basal apparatus. Contact with the basal apparatus may be made with any one of several basal factors, but typically occurs with TF_IID, TF_IIB, or TF_IIA. All of these factors participate in early stages of assembly of the basal apparatus (see the Eukaryotic Transcription chapter). FIGURE 26.9 illustrates the situation in which such a contact is made. The major effect of the activators is to influence the assembly of the basal apparatus.

FIGURE 26.9 Activators may work at different stages of initiation by contacting the TAFs of TF_IID or by contacting TF_IIB.

TF_IID may be the most common target for activators, which may contact any one of several TAFs. In fact, a major role of the TAFs is to provide the connection from the basal apparatus to activators. This explains why the TATA-binding protein (TBP) alone can support basal-level transcription, whereas the TAFs of TF_IID are required for the higher levels of transcription that are stimulated by activators. Different TAFs in TF_IID may provide surfaces that interact with different activators. Some activators interact only with individual TAFs; others interact with multiple TAFs. We assume that the interaction assists the binding of TF_IID to the TATA box, assists the binding of other basal apparatus components around the TF_IID-TATA box complex, or controls the phosphorylation of the C-terminal domain (CTD). In any case, the interaction stabilizes the basal transcription complex, speeds the process of initiation, and thereby increases use of the promoter.

The activating domains of the yeast activator Gal4 (see the section later in this chapter titled Yeast GAL Genes: A Model for Activation and Repression) and others have multiple negative charges, giving rise to their description as “acidic activators.” Acidic activators function by enhancing the ability of TF_IIB to join the basal initiation complex. Experiments in vitro show that binding of TF_IIB to an initiation complex at an adenovirus promoter is stimulated by the presence of Gal4 or other acid activators, and that the activator can bind directly to TF_IIB. Assembly of TF_IIB into the complex at this promoter is therefore a rate-limiting step that is stimulated by the presence of an acidic activator.

The resilience of an RNA polymerase II promoter to the rearrangement of elements, and its indifference even to the particular elements present, suggests that the events by which it is activated are relatively general in nature. Any activators whose activating region is brought within range of the basal initiation complex may be able to stimulate its formation. Some striking illustrations of such versatility have been accomplished by constructing promoters consisting of new combinations of elements.

How does an activator stimulate transcription? Two general types of models can be considered:

The recruitment model argues that the activator’s sole effect is to increase the binding of RNA polymerase to the promoter.
An alternative model is to suppose that the activator induces some change in the transcriptional complex; for example, in the conformation of enzymes such as protein kinases, which increases its efficiency.

If all the components required for efficient transcription are added up—basal factors, RNA polymerase, activators, and coactivators—the result is a very large apparatus that consists of ~40 proteins. Is it feasible for this apparatus to assemble step by step at the promoter? Some activators, coactivators, and basal factors may assemble stepwise at the promoter, but then they may be joined by a very large complex consisting of RNA polymerase preassembled with further activators and coactivators, as illustrated in FIGURE 26.10.

FIGURE 26.10 RNA polymerase exists as a holoenzyme containing many activators.

Several forms of RNA polymerase in which the enzyme is associated with various transcription factors have been found. The most prominent “holoenzyme complex” in yeast (defined as being capable of initiating transcription without additional components) consists of RNA polymerase associated with a 20-subunit complex called Mediator. Mediator includes products of several genes in which mutations block transcription, including some SRB loci (so named because many of their genes were originally identified as suppressors of mutations in RNA polymerase B, another name for pol II). The name was suggested by its ability to mediate the effects of activators. Mediator is necessary for transcription of most yeast genes. Homologous complexes are required for the transcription of most genes in multicellular eukaryotes as well. Mediator undergoes a conformational change when it interacts with the CTD of RNA polymerase. It can transmit either activating or repressing effects from upstream components to the RNA polymerase. It is probably released when a polymerase starts elongation. Some transcription factors influence transcription directly by interacting with RNA polymerase or the basal apparatus, whereas others work by manipulating the structure of chromatin (see the section later in this chapter, Chromatin Remodeling Is an Active Process).

Thus far, the discussion of gene regulation has focused solely on protein factors. However, in many cases noncoding RNA and antisense transcripts also participate in gene regulation (see the section later in this chapter, Yeast Gal Genes: A Model for Activation and Repression, and the Regulatory RNA and Noncoding RNA chapters). Another RNA-dependent pathway that has been implicated in gene regulation and chromatin structure is RNA interference (RNAi). Recent data in Drosophila demonstrate the involvement of the processing machinery for RNAi—Dicer and Argonaute—associated with chromatin at actively transcribed heat-shock loci. Furthermore, mutations that inactivate this machinery lead to problems with RNA polymerase II positioning properly at the promoter. Sequencing of RNAs associated with Argonaute show small RNAs originating from both strands of the promoter region.

On a global scale, transcription that takes place in a nucleus is not scattered randomly throughout at sites of individual genes, but rather is seen to occur in large foci sometimes called transcription factories. As discussed in the Chromosomes chapter, individual chromosomes are not scattered randomly throughout the nucleus, but rather reside in chromosomal domains. New imaging techniques, including chromatin interaction analysis by paired-end-tagged sequencing, or ChIA-PET, allow researchers to examine interactions between distal loci, including enhancers and promoters. These interactions, seen in human cells, can be surprisingly long range—intragenic, extragenic, and even intergenic. Enhancer–promoter interactions were described earlier. Also seen now are promoter–promoter interactions between both nearby and distal genes, as shown in FIGURE 26.11. The data suggest the intriguing possibility that perhaps eukaryotes do possess a physical mechanism, the chroperon, to coordinate the expression of multiple genes similar to the operon model in prokaryotes.

FIGURE 26.11 Higher-order chromatin interactions synergistically promote transcription of clustered genes. These interactions indicate a topological, combinatorial mechanism of transcription regulation.

Modified from Cell 148 (2012): 1–7.

26.7 Many Types of DNA-Binding Domains Have Been Identified

It is common for an activator to have a modular structure in which different domains are responsible for binding to DNA and for activating transcription. Factors are often classified according to the type of DNA-binding domain. In general, a relatively short motif in this domain is responsible for binding to DNA:

The zinc finger comprises a DNA-binding domain. It was originally recognized in factor TF_IIIA, which is required for RNA polymerase III to transcribe 5S rRNA genes. The consensus sequence of a single finger is:

Cys-X_2–4-Cys-X₃-Phe-X₅-Leu-X₂-His-X₃-His

The zinc-finger motif takes its name from the loop of approximately 23 amino acids that protrudes from the zinc-binding site and is described as the Cys₂/His₂ finger. The zinc is held in a tetrahedral structure formed by the conserved Cys and His residues. This motif has since been identified in numerous other transcription factors (and presumed transcription factors). Proteins often contain multiple zinc fingers, such as the three shown in FIGURE 26.12. Some zinc-finger proteins can bind to RNA.

Steroid receptors (and some other proteins) have another type of zinc finger that is different from the Cys₂/His₂ finger. Its structure is based on a sequence with the zinc-binding consensus:

Cys-X₂-Cys-X₁₃-Cys-X₂-Cys

These sequences are called Cys₂/Cys₂ fingers. The steroid receptors are defined as a group by a functional relationship: Each receptor is activated by binding a particular steroid, such as glucocorticoid binding to the glucocorticoid receptor. Together with other receptors, such as the thyroid hormone receptor or the retinoic acid receptor, the steroid receptors are members of the superfamily of ligand-activated activators with the same general modus operandi: The protein factor is inactive until it binds a small ligand, as shown in FIGURE 26.13. The steroid receptors bind to DNA as dimers—either homodimers or heterodimers. Each monomer of the dimer binds to a half-site that may be palindromic or directly repeated.

The helix-turn-helix motif was originally identified as the DNA-binding domain of phage repressors. The C-terminal α-helix lies in the major groove of DNA and is the recognition helix; the middle α-helix lies at an angle across DNA. The N-terminal arm lies in the minor groove and makes additional contacts. A related form of the motif is present in the homeodomain, a sequence first characterized in several proteins encoded by Homeobox genes involved in developmental regulation in Drosophila, and by the comparable human Hox genes shown in FIGURE 26.14. Homeodomain proteins can be activators or repressors.
The amphipathic helix-loop-helix (HLH) motif has been identified in some developmental regulators and in genes coding for eukaryotic DNA-binding proteins. Each amphipathic helix presents a face of hydrophobic residues on one side and charged residues on the other side. The length of the connecting loop varies from 12 to 28 amino acids. The motif enables proteins to dimerize, either homodimers or heterodimers, and a basic region near this motif contacts DNA, as shown in FIGURE 26.15. Not all of the HLH proteins contain a DNA-binding domain, but rather rely on their partner for sequence specificity. Partners may change during development to provide additional combinations.
Leucine zippers consist of an amphipathic α-helix with a leucine residue in every seventh position. The hydrophobic groups, including leucine, face one side while the charged groups face the other side. A leucine-zipper domain in one polypeptide interacts with a leucine-zipper domain in another polypeptide to form a protein dimer. Rules govern which zippers may dimerize. Adjacent to each zipper is another domain containing positively charged residues that is involved in binding to DNA; this is known as the bZIP (basic zipper) structural motif shown in FIGURE 26.16.

FIGURE 26.12 Zinc fingers may form α-helices that insert into the major groove, which is associated with β-sheets on the other side.

FIGURE 26.13 The first finger of a steroid receptor controls which DNA sequence is bound (positions shown in purple); the second finger controls spacing between the sequences (positions shown in blue).

FIGURE 26.14 Helix 3 of the homeodomain binds in the major groove of DNA, with helices 1 and 2 lying outside the double helix. Helix 3 contacts both the phosphate backbone and specific bases. The N-terminal arm lies in the minor groove and makes additional contacts.

FIGURE 26.15 A helix-loop-helix (HLH) dimer in which both subunits are of the bHLH type can bind DNA, but a dimer in which one subunit lacks the basic region cannot bind DNA.

FIGURE 26.16 The basic regions of the bZIP motif are held together by the dimerization at the adjacent zipper region when the hydrophobic faces of two leucine zippers interact in parallel orientation.

26.8 Chromatin Remodeling Is an Active Process

Transcriptional activators face a challenge when trying to bind to their recognition sites in eukaryotic chromatin. FIGURE 26.17 illustrates two general states that can exist at a eukaryotic promoter. In the inactive state, nucleosomes are present, and they prevent basal factors and RNA polymerase from binding. In the active state, the basal apparatus occupies the promoter, and histone octamers cannot bind to it. Each type of state is stable. In order to convert a promoter from the inactive state to the active state, the chromatin structure must be perturbed in order to allow binding of the basal factors.

FIGURE 26.17 If nucleosomes form at a promoter, transcription factors (and RNA polymerase) cannot bind. If transcription factors (and RNA polymerase) bind to the promoter to establish a stable complex for initiation, histones are excluded.

The general process of inducing changes in chromatin structure is called chromatin remodeling. This consists of mechanisms for repositioning or displacing histones that depend on the input of energy. Many protein–protein and protein–DNA contacts need to be disrupted to release histones from chromatin. There is no free ride: Energy must be provided to disrupt these contacts. FIGURE 26.18 illustrates the principle of dynamic remodeling by a factor that hydrolyzes ATP. When the histone octamer is released from DNA, other proteins (in this case transcription factors and RNA polymerase) can bind.

FIGURE 26.18 The dynamic model for transcription of chromatin relies on factors that can use energy provided by hydrolysis of ATP to displace nucleosomes from specific DNA sequences.

Chromatin remodeling results in several alternative outcomes, as shown in FIGURE 26.19:

Histone octamers may slide along DNA, changing the relationship between the nucleic acid and the protein. This can alter both the rotational and the translational position of a particular sequence on the nucleosome.
The spacing between histone octamers may be changed, again with the result that the positions of individual sequences are altered relative to the histone octamer.
The most extensive change is that an octamer(s) may be displaced entirely from DNA to generate a nucleosome-free gap. Alternatively, one or both H2A-H2B dimers can be displaced, leaving an H2A-H2B-H3-H4 hexamer, or an H3-H4 tetramer, on the DNA.

FIGURE 26.19 Remodeling complexes can cause nucleosomes to slide along DNA, displace nucleosomes from DNA, or reorganize the spacing between nucleosomes.

A major role of chromatin remodeling is to change the organization of nucleosomes at the promoter of a gene that is to be transcribed. This is required to allow the transcription apparatus to gain access to the promoter. Remodeling can also act to prevent transcription by moving nucleosomes onto, rather than away from, essential promoter sequences. Remodeling is also required to enable other manipulations of chromatin, such as repair of damaged DNA (see the Repair Systems chapter).

Remodeling often takes the form of displacing one or more histone octamers. This can result in the creation of a site that is hypersensitive to cleavage with DNase I (see the Chromatin chapter). Sometimes less dramatic changes are observed, such as alteration of the rotational positioning of a single nucleosome, detectable by loss or change of the DNase I 10-bp ladder. Thus, changes in chromatin structure can extend from subtly altering the positions of nucleosomes to removing them altogether.

Chromatin remodeling is undertaken by ATP-dependent chromatin remodeling complexes, which use ATP hydrolysis to provide the energy for remodeling. The heart of the remodeling complex is its ATPase subunit. The ATPase subunits of all remodeling complexes are related members of a large superfamily of proteins, which is divided into subfamilies of more closely related members. Remodeling complexes are classified according to the subfamily of ATPase that they contain as their catalytic subunit. There are many subfamilies; four major ones (SWI/SNF, ISWI, CHD, and INO80/SWR1) are shown in TABLE 26.1. The first remodeling complex described was the SWI/SNF (“switch sniff”) complex in yeast, which has homologs in all eukaryotes. The chromatin remodeling superfamily is large and diverse, and most species have multiple complexes in different subfamilies. Budding yeast have two SWI/SNF-related complexes and three ISWI complexes. At least four different ISWI complexes have been characterized in mammals. Remodeling complexes range from small heterodimeric complexes (the ATPase subunit plus a single partner) to massive complexes of 10 or more subunits. Each type of complex may undertake a different range of remodeling activities.

TABLE 26.1 Remodeling complexes can be classified by their ATPase subunits.

Type of Complex	SWI/SNF	ISWI	CHD	INO80/SWRI
Yeast	SWI/SNFRSC	ISW1aISW1bISW2	CHDI	INO80/SWR1
Fly	dSWI/SNF (brahma)	NURFCHRACACF	JMIZ	Tip60
Human	hSWI/SNF	RSFhACF/WCFRhCHRACWICH	NuRD	INO80 SRCAP
Frog		WICHCHRACACF	Mi-2

SWI/SNF is the prototypic remodeling complex. Its name reflects the fact that many of its subunits are encoded by genes originally identified by swi or snf mutations in Saccharomyces cerevisiae. (swi mutants cannot switch mating type, and snf—sucrose nonfermenting—mutants cannot use sucrose as a carbon source.) Mutations in these loci are pleiotropic, and the range of defects is similar to those shown by mutants that have lost part of the CTD of RNA polymerase II. Early hints that these genes might be linked to chromatin came from evidence that these mutations show genetic interactions with mutations in genes that code for components of chromatin: SIN1, which encodes a nonhistone chromatin protein, and SIN2, which encodes histone H3. The SWI and SNF genes are required for expression of a variety of individual loci. Approximately 120 S. cerevisiae genes require SWI/SNF for normal expression, which is about 2% of the total number of genes. Expression of these loci may require the SWI/SNF complex to remodel chromatin at their promoters. Each yeast cell has only about 150 complexes of SWI/SNF. The related RSC (remodels the structure of chromatin) complex is more abundant and is essential for viability. It acts at approximately 700 target loci.

Different subfamilies of remodeling complexes have distinct modes of remodeling, reflecting differences in their ATPase subunits, as well as effects of other proteins in individual remodeling complexes. SWI/SNF complexes can remodel chromatin in vitro without overall loss of histones or can displace histone octamers. These reactions likely pass through the same intermediate in which the structure of the target nucleosome is altered, leading either to reformation of a (remodeled) nucleosome on the original DNA or to displacement of the histone octamer to a different DNA molecule. In contrast, the ISWI family primarily affects nucleosome positioning without displacing octamers, in a sliding reaction in which the octamer moves along DNA. The activity of ISWI requires the histone H4 tail as well as binding to linker DNA.

The DNA and histone octamer have many contact points; 14 have been identified in the crystal structure. All of these contacts must be broken for an octamer to be released or for it to move to a new position. How is this achieved? The ATPase subunits are distantly related to helicases (enzymes that unwind double-stranded nucleic acids), but remodeling complexes do not have any unwinding activity. Present thinking is that remodeling complexes in the SWI/SNF and ISWI classes use the hydrolysis of ATP to translocate DNA on the nucleosomal surface, essentially by creating a twisting motion. This twisting creates a mechanical force that allows a small region of DNA to be released from the surface and then repositioned. This mechanism creates transient loops of DNA on the surface of the octamer; these loops are themselves accessible to interact with other factors, or they can propagate along the nucleosome, ultimately resulting in nucleosome sliding. In the case of SWI/SNF complexes, this activity can also result in nucleosome disassembly, first by displacement of the H2A/H2B dimers, then of the H3/H4 tetramer.

Different remodeling complexes have different roles in the cell. SWI/SNF complexes are frequently involved in transcriptional activation, whereas some ISWI complexes act as repressors, using their remodeling activity to slide nucleosomes onto promoter regions to prevent transcription. Members of the CHD (chromodomain helicase DNA-binding) family have also been implicated in repression, particularly the Mi-2/NuRD complexes, which contain both chromatin remodeling and histone deacetylase activities. Remodelers in the SWR1/INO80 class have a unique activity: In addition to their normal remodeling capabilities, some members of this class also have histone exchange capability, in which individual histones (usually H2A/H2B dimers) can be replaced in a nucleosome, typically with the H2AZ histone variant (see the Chromatin chapter).

26.9 Nucleosome Organization or Content Can Be Changed at the Promoter

How are remodeling complexes targeted to specific sites on chromatin? Most remodelers do not contain subunits that bind specific DNA sequences, though there are a few exceptions. This suggests the model shown in FIGURE 26.20, in which remodelers are recruited by activators or repressors.

FIGURE 26.20 A remodeling complex binds to chromatin via an activator (or repressor).

The interaction between transcription factors and remodeling complexes gives a key insight into their modus operandi. The transcription factor Swi5 activates the HO gene in yeast, a gene involved in mating-type switching. (Note that despite its name Swi5 is not a member of the SWI/SNF complex.) Swi5 enters the nucleus near the end of mitosis and binds to the HO promoter. It then recruits SWI/SNF to the promoter. Swi5 is then released, leaving SWI/SNF at the promoter. This means that a transcription factor can activate a promoter by a “hit and run” mechanism, in which its function is fulfilled once the remodeling complex has bound. This is more likely to occur with genes that are cell-cycle regulated or otherwise transiently activated; it is equally common at many genes for transcription factors to remain associated with target genes for long periods.

The involvement of remodeling complexes in gene activation was discovered because the complexes are necessary to enable certain transcription factors to activate their target genes. One of the first examples was the GAGA factor, which activates the Drosophila hsp70 promoter. Binding of GAGA to four (CT)_n-rich sites near the promoter disrupts the nucleosomes, creates a hypersensitive region, and causes the adjacent nucleosomes to be rearranged so that they occupy preferential instead of random positions. Disruption is an energy-dependent process that requires the NURF remodeling complex, a complex in the ISWI subfamily. The organization of nucleosomes is altered so as to create a boundary that determines the positions of the adjacent nucleosomes. During this process, GAGA binds to its target sites in DNA, and its presence fixes the remodeled state.

The PHO system was one of the first in which it was shown that a change in nucleosome organization is involved in gene activation. At the PHO5 promoter, the bHLH activator Pho4 responds to phosphate starvation by inducing the disruption of four precisely positioned nucleosomes, as depicted in FIGURE 26.21. This event is independent of transcription (it occurs in a TATA^– mutant) and independent of replication. The promoter has two binding sites for Pho4 (and another activator, Pho2). One is located between nucleosomes, which can be bound by the isolated DNA-binding domain of Pho4; the other lies within a nucleosome, which cannot be recognized. Disruption of the nucleosome to allow DNA binding at the second site is necessary for gene activation. This action requires the presence of the transcription-activating domain and appears to involve at least two remodelers: SWI/SNF and INO80. In addition, chromatin disassembly at PHO5 also requires a histone chaperone, Asf1, which may assist in nucleosome removal or act as a recipient of displaced histones.

FIGURE 26.21 Nucleosomes are displaced from promoters during activation. The PHO5 promoter contains nucleosomes positioned over the TATA box and one of the binding sites for the Pho4 and Pho2 activators. When PHO5 is induced by phosphate starvation (–Pi), promoter nucleosomes are displaced.

A survey of nucleosome positions in a large region of the yeast genome shows that most sites that bind transcription factors are free of nucleosomes. Promoters for RNA polymerase II typically have a nucleosome-free region (NFR) approximately 200 bp upstream of the start point, which is flanked by positioned nucleosomes on either side. These positioned nucleosomes typically contain the histone variant H2AZ (called Htz1 in yeast); the deposition of H2AZ requires the SWR1 remodeling complex. This organization appears to be present in many human promoters as well. It has been suggested that H2AZ-containing nucleosomes are more easily evicted during transcription activation, thus poising promoters for activation; however, the actual effects of H2AZ on nucleosome stability in vivo are controversial.

It is not always the case, though, that nucleosomes must be excluded in order to permit initiation of transcription. Some activators can bind to DNA on a nucleosomal surface. Nucleosomes appear to be precisely positioned at some steroid-hormone response elements in such a way that receptors can bind. Receptor binding may alter the interaction of DNA with histones and may even lead to exposure of new binding sites. The exact positioning of nucleosomes could be required either because the nucleosome “presents” DNA in a particular rotational phase or because there are protein–protein interactions between the activators and histones or other components of chromatin. Thus, researchers have moved some way from viewing chromatin exclusively as a repressive structure to considering which interactions between activators and chromatin can be required for activation.

The MMTV promoter presents an example of the need for specific nucleosomal organization. It contains an array of six partly palindromic sites that constitute the hormone response element (HRE). Each site is bound by one dimer of hormone receptor (HR). The MMTV promoter also has a single binding site for the factor NF1 and two adjacent sites for the factor OTF. HR and NF1 cannot bind simultaneously to their sites in free DNA. FIGURE 26.22 shows how the nucleosomal structure controls binding of the factors.

FIGURE 26.22 Hormone receptor and NF1 cannot bind simultaneously to the MMTV promoter in the form of linear DNA, but can bind when the DNA is presented on a nucleosomal surface.

The HR protects its binding sites at the promoter when hormone is added, but does not affect the micrococcal nuclease-sensitive sites that mark either side of the nucleosome. This suggests that HR is binding to the DNA on the nucleosomal surface; however, the rotational positioning of DNA on the nucleosome prior to hormone addition allows access to only two of the four sites. Binding to the other two sites requires a change in rotational positioning on the nucleosome. This can be detected by the appearance of a sensitive site at the axis of dyad symmetry (which is in the center of the binding sites that constitute the HRE). NF1 can be detected on the nucleosome after hormone induction, so these structural changes may be necessary to allow NF1 to bind, perhaps because they expose DNA and abolish the steric hindrance by which HR blocks NF1 binding to free DNA.

26.10 Histone Acetylation Is Associated with Transcription Activation

All of the core histones are subject to multiple covalent modifications, as discussed in the Chromatin chapter. Different modifications result in different functional outcomes. One of the most extensively studied modifications (and the first to be characterized in detail) is lysine acetylation. All core histones are dynamically acetylated on lysine residues in the tails (and occasionally within the globular core). As described in the Chromatin chapter, certain patterns of acetylation are associated with newly synthesized histones that are deposited during DNA synthesis in S phase. This specific acetylation pattern is then erased after histones are incorporated into nucleosomes.

Outside of S phase, acetylation of histones in chromatin is generally correlated with the state of gene expression. The correlation was first noticed because histone acetylation is increased in a domain containing active genes, and acetylated chromatin is more sensitive to DNase I. This occurs largely because of acetylation of the nucleosomes (on specific lysines) in the vicinity of the promoter when a gene is activated.

The range of nucleosomes targeted for modification can vary. Modification can be a local event—for example, restricted to nucleosomes at a promoter. It can also be a general event, extending over large domains or even to an entire chromosome. Global changes in acetylation occur on sex chromosomes. This is part of the mechanism by which the activities of genes on sex chromosomes are altered to compensate for the presence of two X chromosomes in one sex but only one X chromosome in the other sex (see the chapter titled Epigenetics II). The inactive X chromosome in female mammals has underacetylated histones. The superactive X chromosome in Drosophila males has increased acetylation of H4. This suggests that the presence of acetyl groups may be a prerequisite for a less condensed, active structure. In male Drosophila, the X chromosome is acetylated specifically at K16 of histone H4. The enzyme responsible for this acetylation is called MOF; MOF is recruited to the chromosome as part of a large protein complex. This “dosage compensation” complex is responsible for introducing general changes in the X chromosome that enable it to be more highly expressed. The increased acetylation is only one of its activities.

Acetylation is reversible. Each direction of the reaction is catalyzed by a specific type of enzyme. Enzymes that can acetylate lysine residues in proteins are called histone acetyltransferases (HATs); when these enzymes target lysines in nonhistones, they are also known more generically as lysine (K) acetyltransferases (KATs). The acetyl groups are removed by histone deacetylases (HDACs). HAT enzymes are categorized into two groups: Those in group A act on histones in chromatin and are involved with the control of transcription; those in group B act on newly synthesized histones in the cytosol and are involved with nucleosome assembly.

Two inhibitors have been useful in analyzing acetylation. Trichostatin and butyric acid inhibit histone deacetylases and cause acetylated nucleosomes to accumulate. The use of these inhibitors has supported the general view that acetylation is associated with gene expression; in fact, the ability of butyric acid to cause changes in chromatin resembling those found upon gene activation was one of the first indications of the connection between acetylation and gene activity.

The breakthrough in analyzing the role of histone acetylation was provided by the characterization of the acetylating and deacetylating enzymes and their association with other proteins that are involved in specific events of activation and repression. A basic change in the view of histone acetylation was caused by the discovery that previously identified activators of transcription turned out to also have HAT activity.

The connection was established when the catalytic subunit of a group A HAT was identified as a homolog of the yeast regulator protein Gcn5. It then was shown that yeast Gcn5 itself has HAT activity, with histones H3 and H2B as its preferred substrates in vivo. Gcn5 had previously been identified as part of an adaptor complex required for the function of certain enhancers and their target promoters. It is now known that Gcn5’s HAT activity is required for activation of a number of target genes.

Gcn5 was the prototypic HAT that opened the way to the identification of a large family of related acetyltransferase complexes conserved from yeast to mammals. In yeast, Gcn5 is the catalytic subunit of several HAT complexes, including the 1.8-MDa Spt-Ada-Gcn5-acetyltransferase (SAGA) complex, which contains several proteins that are involved in transcription. Among these proteins are several TAF_IIs. In addition, the Taf1 subunit of TF_IID is itself an acetyltransferase. Some functional overlap exists between TF_IID and SAGA, most notably that yeast can survive the loss of either Taf1 or Gcn5 but cannot tolerate the deletion of both. This might suggest that an acetyltransferase activity is essential for gene expression, and that it can be provided by either TF_IID or SAGA. As might be expected from the size of the SAGA complex, acetylation is only one of its functions. The SAGA complex has histone H2B deubiquitylation activity (dynamic H2B ubiquitylation/deubiquitylation is also associated with transcription), and also contains subunits possessing bromodomains and chromodomains, allowing this complex to interact with acetylated and methylated histones.

One of the first general activators to be characterized as HAT was p300/CREB-binding protein (CBP). (Actually, p300 and CBP are different proteins, but they are so closely related that they are often referred to as a single type of activity.) p300/CBP is a coactivator that links an activator to the basal apparatus (see Figure 26.8). p300/CBP interacts with various activators, including the hormone receptors AP-1 (c-Jun and c-Fos) and MyoD. p300/CBP acetylates multiple histone targets, with a preference for the H4 tail. p300/CBP interacts with another coactivator, PCAF, which is related to Gcn5 and preferentially acetylates H3 in nucleosomes. p300/CBP and PCAF form a complex that functions in transcriptional activation. In some cases yet another HAT can be involved, such as the hormone receptor coactivator ACTR, which is itself a HAT that acts on H3 and H4. One explanation for the presence of multiple HAT activities in a coactivating complex is that each HAT has a different specificity, and that multiple, different acetylation events are required for activation. This enables the picture for the action of coactivators to be redrawn, as shown in FIGURE 26.23, where RNA polymerase II is bound at a hypersensitive site and coactivators are acetylating histones in the nucleosomes in the vicinity.

FIGURE 26.23 Coactivators may have HAT activities that acetylate the tails of nucleosomal histones.

Group A HATs, like ATP-dependent remodeling enzymes, are typically found in large complexes. FIGURE 26.24 shows a simplified model for their behavior. HAT complexes can be targeted to DNA by interactions with DNA-binding factors. The complex also contains effector subunits that affect chromatin structure or act directly on transcription. It is likely that at least some of the effectors require the acetylation event in order to act (such as the deubiquitylation activity of SAGA).

FIGURE 26.24 Complexes that control acetylation levels have targeting subunits that determine their sites of action (usually subunits that interact with site-specific DNA-binding proteins), HAT or HDAC enzymes that acetylate or deacetylate histones, and effector subunits that have other actions on chromatin or DNA.

The effect of acetylation may be both quantitative and qualitative. In cases where the effect of charge neutralization on chromatin structure is key, a certain minimal number of acetyl groups should be required to have an effect, and the exact positions at which they occur are largely irrelevant. In the case where the role of acetylation is primarily in the creation of a binding site (for a bromodomain-containing factor, for example), the specific position of the acetylation event will be critical. The existence of complexes containing multiple HAT activities might be interpreted either way—if individual enzymes have different specificities, multiple activities might be needed either to acetylate a sufficient number of different positions or because the individual events are necessary for different effects upon transcription. At replication, it appears (at least with respect to histone H4) that acetylation at any two of three particular positions is adequate, favoring a quantitative model in this case. Where chromatin structure is changed to affect transcription, acetylation at specific positions is important (see the chapter titled Epigenetics I).

As acetylation is linked to activation, deacetylation is linked to transcriptional repression. Whereas site-specific activators recruit coactivators with HAT activity, site-specific repressor proteins can recruit corepressor complexes, which often contain HDAC activity.

In yeast, mutations in SIN3 and RPD3 result in increased expression of a variety of genes, indicating that Sin3 and Rpd3 proteins act as repressors of transcription. Sin3 and Rpd3 are recruited to a number of genes by interacting with the DNA-binding protein Ume6, which binds to the URS1 (upstream repressive sequence) element. The complex represses transcription at the promoters containing URS1, as illustrated in FIGURE 26.25. Rpd3 is a histone deacetylase, and its recruitment leads to deacetylation of nucleosomes at the promoter. Rpd3 and its homologs are present in multiple HDAC complexes found in eukaryotes from yeast to humans; these large complexes are typically built around Sin3 and its homologs.

FIGURE 26.25 A repressor complex contains three components: a DNA-binding subunit, a corepressor, and a histone deacetylase.

In mammalian cells, Sin3 is part of a repressive complex that includes histone-binding proteins and the Rpd3 homologs HDAC1 and HDAC2. This corepressor complex can be recruited by a variety of repressors to specific gene targets. The bHLH family of transcription regulators includes activators that function as heterodimers, including MyoD. This family also includes repressors, in particular the heterodimer Mad–Max, where Mad can be any one of a group of closely related proteins. The Mad–Max heterodimer (which binds to specific DNA sites) interacts with Sin3–HDAC1/2 complex and requires the deacetylase activity of this complex for repression. Similarly, the SMRT corepressor (which enables retinoid hormone receptors to repress certain target genes) binds mSin3, which, in turn, brings the HDAC activities to the site. Another means of bringing HDAC activities to a DNA site can be an interaction with MeCP2, a protein that binds to methylated cytosines, a mark of transcriptional silencing (see the Eukaryotic Transcription and Epigenetics I chapters).

Absence of histone acetylation is also a feature of heterochromatin. This is true of both constitutive heterochromatin (typically involving regions of centromeres or telomeres) and facultative heterochromatin (regions that are inactivated in one cell although they may be active in another). Typically the N-terminal tails of histones H3 and H4 are not acetylated in heterochromatic regions (see the chapter titled Epigenetics I).

26.11 Methylation of Histones and DNA Is Connected

DNA methylation is associated with transcriptional inactivity, whereas histone methylation can be linked to either active or inactive regions, depending on the specific site of methylation. Numerous sites of lysine methylation are present in the tail and core of histone H3 (a few of which occur only in some species), and a single lysine in the tail of H4 is methylated. In addition, three arginines in H3 and one in H4 are also methylated. Because lysines can be mono-, di-, or trimethylated, and arginines can be mono- or dimethylated (see the Chromatin chapter), the number of potential functional methylation marks is large.

For example, di- or trimethylation of H3K4 is associated with transcriptional activation, and trimethylated H3K4 occurs around the start sites of active genes. In contrast, H3 methylated at K9 or K27 is a feature of transcriptionally silent regions of chromatin, including heterochromatin and smaller regions containing one or more silent genes. Whole-genome studies can help to uncover general patterns of modifications linked to different transcriptional states, as shown in FIGURE 26.26.

FIGURE 26.26 The distribution of histones and their modifications are mapped on an arbitrary gene relative to its promoter. The curves represent the patterns that are determined via genome-wide approaches. The location of the histone variant H2A.Z is also shown. With the exception of the data on K9 and K27 methylation, most of the data are based on yeast genes.

Reprinted from Cell, vol. 128, B. Li, M. Carey, and J. L. Workman, The Role of Chromatin during Transcription, pp. 707–719. Copyright 2007, with permission from Elsevier [http://www.sciencedirect.com/science/journal/00928674].

Histone lysine methylation is catalyzed by lysine methyltransferases (HMTs or KMTs), most of which contain a conserved region called the SET domain. Like acetylation, methylation is reversible, and two different families of lysine demethylases (KDMs) have been identified: the LSD1 (lysine-specific demethylase 1, also known as KDM1) family and the Jumonji family. Different classes of enzymes demethylate arginines.

In silent or heterochromatic regions, the methylation of H3 at K9 is linked to DNA methylation. The enzyme that targets this lysine is a SET domain–containing enzyme called Suv39h1. Deacetylation of H3K9 by HDACs must occur before this lysine can be methylated. H3K9 methylation then recruits the protein HP1 (heterochromatin protein 1), which binds H3K9me via its chromodomain. HP1 then targets the activity of DNA methyltransferases (DNMTs). Most of the methylation sites in DNA are CpG islands (see the chapter titled Epigenetics I). CpG sequences in heterochromatin are typically methylated. Conversely, it is necessary for the CpG islands located in promoter regions to be unmethylated in order for a gene to be expressed.

Methylation of DNA and methylation of histones are connected in a mutually reinforcing circuit. In addition to the recruitment of DNMTs via HP1 binding to H3K4me, DNA methylation can, in turn, result in histone methylation. Some histone methyltransferase complexes (as well as some HDAC complexes) contain binding domains that recognize the methylated CpG doublet, thus the DNA methylation reinforces the circuit by providing a target for the histone deacetylases and methyltransferases to bind. The important point is that one type of modification can be the trigger for another. These systems are widespread, as can be seen by evidence for these connections in fungi, plants, and animal cells, and for regulating transcription at promoters used by both RNA polymerases I and II, as well as maintaining heterochromatin in an inert state.

26.12 Promoter Activation Involves Multiple Changes to Chromatin

FIGURE 26.27 summarizes three common differences between active chromatin and inactive chromatin:

Active chromatin is acetylated on the tails of histones H3 and H4.
Inactive chromatin is methylated on specific lysines (such as K9) of histone H3.
Inactive chromatin is methylated on cytosines of CpG doublets.

FIGURE 26.27 Acetylation of histones activates chromatin; methylation of DNA and specific sites on histones inactivates chromatin.

The reverse events occur in the activation of a promoter with the generation of heterochromatin. The actions of the enzymes that modify chromatin ensure that activating events are mutually exclusive with inactivating events. For example, the silencing methylation of H3 at K9 and the activating acetylation of H3 at K9 and K14 are mutually antagonistic.

How are histone-modifying enzymes such as acetyltransferases or deacetylases recruited to their specific targets? As with remodeling complexes, the process is likely to be indirect. A sequence-specific activator (or repressor) may interact with a component of the acetyltransferase (or deacetylase) complex to recruit it to a promoter.

Direct interactions also take place between remodeling complexes and histone-modifying complexes. Histone modifications by themselves have little effect on the overall structure or accessibility of chromatin, which instead requires the interactions of chromatin remodelers. Binding by the SWI/SNF remodeling complex may lead, in turn, to binding by the SAGA acetyltransferase complex. Acetylation of histones can then stabilize the association with the SWI/SNF complex (via its bromodomain), making a mutual reinforcement of the changes in the components at the promoter. In fact, the Brg1 ATPase subunit of the human SWI/SNF complex requires H4K8 and K12 acetylation for binding to certain targets in vivo. Some remodeling complexes contain between 4 and 10 bromodomains distributed among different subunits, which may confer different binding specificities for specific acetylated targets.

Histone methylation also results in recruitment of numerous factors that contain methyl-lysine recognition motifs such as chromodomains and plant homeodomain (PHD) fingers. Methylation of histone H3 on K4 recruits the chromodomain-containing remodeler Chd1, which also associates with SAGA. H3K4me also directly recruits another acetyltransferase complex, NuA3, which recognizes H3K4me via a PHD domain in one of its subunits. These are just a few of the interactions that occur during transcription activation, and different genes have different (but often overlapping) complex networks of interactions. A further set of dynamic modifications and interactions serves to facilitate transcriptional elongation and to “reset” the chromatin behind the elongating polymerase.

Many of the events at the promoter can be connected into the series illustrated in FIGURE 26.28. The initiating event is the binding of a sequence-specific component, which is either able to find its target DNA sequence in the context of chromatin or to bind to a site in a nucleosome-free region. This activator recruits remodeling and histone-modifying complexes (only HATs are shown for simplicity). Changes occur in nucleosome structure, and the acetylation or other modification of target histones provides a covalent mark that the locus has been activated. Many of these steps are mutually reinforcing. Initiation complex assembly follows (after any other necessary activators bind), and at some point histones are typically displaced.

FIGURE 26.28 Htz1-containing nucleosomes flank a 200-bp NFR on both sides of a promoter. Upon targeting to the upstream activation sequence (UAS), activators recruit various coactivators (such as Swi/Snf or SAGA). This recruitment further increases the binding of activators, particularly for those bound within nucleosomal regions. More important, histones are acetylated at promoter-proximal regions, and these nucleosomes become much more mobile. In one model (left), a combination of acetylation and chromatin remodeling directly results in the loss of Htz1-containing nucleosome, thereby exposing the entire core promoter to the GTFs and Pol II. SAGA and Mediator then facilitate preinitiation complex (PIC) formation through direct interactions. In the other model (right), which represents the remodeled state, partial PICs could be assembled at the core promoter without loss of Htz1. It is the binding of Pol II and TFIIH that leads to the displacement of Htz1-containing nucleosomes and the full assembly of PIC.

Reprinted from Cell, vol. 128, B. Li, M. Carey, and J. L. Workman, The Role of Chromatin during Transcription, pp. 707–719. Copyright 2007, with permission from Elsevier [http://www.sciencedirect.com/science/journal/00928674].

26.13 Histone Phosphorylation Affects Chromatin Structure

All histones can be phosphorylated in vivo in different contexts. Histones are phosphorylated in three circumstances:

Cyclically during the cell cycle
In association with chromatin remodeling during transcription
During DNA repair

It has long been known that the linker histone H1 is phosphorylated at mitosis, and H1 is an extremely good substrate for the Cdc2 kinase that controls cell division. This led to speculation that the phosphorylation might be connected with the condensation of chromatin, but so far no direct effect of this phosphorylation event has been demonstrated, and it is not known whether it plays a role in cell division. In Tetrahymena, it is possible to delete all the genes for H1 without significantly affecting the overall properties of chromatin, resulting in a relatively small effect on the ability of chromatin to condense at mitosis. Some genes are activated and others are repressed by this change, which suggests that there are alterations in local structure. Mutations that eliminate sites of phosphorylation in H1 have no effect, but mutations that mimic the effects of phosphorylation produce a phenotype that resembles the deletion. This suggests that the effect of phosphorylating H1 is to eliminate its effects on local chromatin structure.

Phosphorylation of serine 10 of histone H3 is linked to transcriptional activation (where it promotes acetylation of K14 in the same tail) and to chromosome condensation and mitotic progression. In Drosophila melanogaster, loss of a kinase that phosphorylates histone H3S10 (JIL-1) has devastating effects on chromatin structure. FIGURE 26.29 compares the usual extended structure of the polytene chromosome (upper photograph) with the structure that is found in a null mutant that has no JIL-1 kinase (lower photograph). The absence of JIL-1 is lethal, but the chromosomes can be visualized in the larvae before they die.

FIGURE 26.29 Flies that have no JIL-1 kinase have abnormal polytene chromosomes that are condensed instead of extended.

Photos courtesy of Jorgen Johansen and Kristen M. Johansen, Iowa State University.

This suggests that H3 phosphorylation is required to generate the more extended chromosome structure of euchromatic regions. JIL-1 also associates with the complex of proteins that binds to the X chromosome to increase its gene expression in males (see the chapter titled Epigenetics II), and JIL-1–dependent H3S10 phosphorylation also antagonizes H3K9 dimethylation, a heterochromatic mark. These results are consistent with a role for JIL-1 in promoting an active chromatin conformation. Interestingly, H3S10 phosphorylation by JIL-1 is itself promoted by acetylation of H4K12 by the ATAC acetyltransferase complex; these complicated interactions make it challenging to determine whether one single modification is key for the transitions in chromatin structure or whether several modifications must occur together. It is also not clear how this role of H3 phosphorylation in promoting transcriptionally active chromatin is related to the requirement for H3 phosphorylation to initiate chromosome condensation in at least some species (including mammals and the ciliate Tetrahymena).

This results in somewhat conflicting impressions of the roles of histone phosphorylation. Where it is important in the cell cycle, it is likely to be as a signal for condensation. Its effect in transcription and repair appears to be the opposite, where it contributes to open chromatin structures compatible with transcription activation and repair processes. (Histone phosphorylation during repair is discussed in the Chromatin and Repair Systems chapters.)

It is possible, of course, that phosphorylation of different histones, or even of different amino acid residues in one histone, has opposite effects on chromatin structure.

26.14 Yeast GAL Genes: A Model for Activation and Repression

Yeast, like bacteria, need to be able to rapidly respond to their environment (see the chapter titled The Operon). In the yeast Saccharomyces cerevisiae, the GAL genes serve a similar function to the lac operon in E. coli. In an emergency, when there is little or no glucose as an energy source and only galactose (or in E. coli, lactose) is available, the cell will survive because it can catabolize the alternate sugar to generate ATP. The GAL system in S. cerevisiae has been a model system to investigate gene regulation in eukaryotes for many years. This section focuses on two of these genes, GAL1 and GAL10, which are shown in FIGURE 26.30. Like most eukaryotic genes, the GAL genes are monocistronic. These two genes are divergently transcribed and regulated from a central control region called the upstream activating sequence (UAS), which is similar to an enhancer. Like the lac operon in E. coli, the GAL genes are induced by their substrate, galactose. For the same reason as in E. coli, the GAL genes are also under another level of control (described shortly)—catabolite repression. They cannot be activated by the substrate galactose when there is a sufficient supply of glucose, the preferred energy source.

FIGURE 26.30 The yeast GAL1/GAL10 locus highlighting the UAS and showing the Gal4, Gal80, and Gal3 regulatory proteins and the RSC/nucleosome. Nucleosomes are also positioned at the promoters when the genes are not being transcribed.

Together, the GAL genes are under five different levels of control. The first level is chromatin structure. Mutations in any of the subunits of the chromatin remodeler SWI/SNF and in the acetyltransferase complex SAGA will result in reduced expression of the GAL genes. Second, the UAS has both general enhancer and Mig1 repressor–binding sites. The third level is through a noncoding RNA transcript that assists in maintaining repressed chromatin over the open reading frames. The fourth level is the GAL-specific galactose induction mechanism. The fifth level is catabolite (glucose) repression.

The two GAL genes are unusual in that they lack the typical nucleosome-free region present at the start sites of most yeast genes. Instead, the start sites are contained in well-positioned nucleosomes. The UAS region that controls the GAL genes has an unusual base composition—short-phased AT repeats every 10 base pairs—which causes the DNA to bend. Nucleosomes containing the histone variant H2AZ (Htz1 in yeast) are positioned over the promoters of both GAL1 and GAL10, aided in their positioning in part by the bent DNA.

The GAL10 gene is also an unusual gene in that it has a cryptic promoter in open chromatin at its 3′ end. This promoter transcribes a noncoding RNA that is antisense to GAL10 and extends through and includes GAL1 (see the Regulatory RNA chapter). Transcription is very inefficient and the RNA abundance is extremely low (less than one copy per cell), due, in part, to rapid degradation. Under repressed conditions this promoter is stimulated by the Reb1 transcription factor, usually thought to be an RNA polymerase I transcription factor. The noncoding transcript represses transcription of the GAL1/10 pair of genes by recruiting the Set2 methyltransferase, which leads to H3K36 di- and trimethylation. H3K36me2/me3 recruits an HDAC to deacetylate the chromatin, which, in turn, leads to repressed chromatin structure.

The GAL genes are ultimately controlled by the positive regulator Gal4, which binds as a dimer to four binding sites in the UAS region, as shown in Figure 26.30 and FIGURE 26.31. Its activation domain consists of two acidic patch domains. Gal4, in turn, is regulated by Gal80, a negative regulator that binds to Gal4 and masks its activation domain, preventing it from activating transcription. This is the normal state for the GAL genes: turned off and waiting to be induced. The chromatin architecture of the UAS has been difficult to discern. Recent data from uninduced cells suggest that a partly unwrapped nucleosome is constitutively held in place and positioned by the chromatin-remodeling factor RSC. RSC in yeast, unlike its homologs in higher eukaryotes, has a domain for sequence-specific DNA binding. This complex facilitates the binding of Gal4 by aiding in the phasing of the nucleosomes over the two promoters and prevents them from encroaching on the Gal4 binding sites.

FIGURE 26.31 The yeast GAL1 gene as it is being activated. Gal3 is bound to Gal80 in the nucleus and cytoplasm, preventing it from binding to Gal4 and allowing Gal4 to recruit the transcription machinery and activate transcription.

Gal80, itself is regulated by the negative regulator Gal3, which is controlled by the inducer galactose. Gal80 contains overlapping binding sites for both Gal4 and Gal3. Gal3 is an interesting protein, having very high homology to Gal1, which is a galactokinase enzyme whose function is to phosphorylate galactose. Gal3 has no enzymatic activity, but retains the ability to bind galactose and ATP. This changes the structure of Gal3 to enable it to bind to Gal80 in the presence of NADP. When it does, Gal3 masks the Gal4 binding site of Gal80, preventing it from binding to Gal4. This transition occurs very rapidly, leading to induction of Gal1/10, due primarily to Gal3 binding Gal80 in the nucleus. Gal3 is thus a negative regulator of a negative regulator, which makes it a positive regulator of Gal4. This depletes the nuclear level of Gal80, unmasking Gal4 and allowing activation of the genes. NADP is thought to be a “second messenger” metabolic sensor.

Unmasked Gal4 is now able to begin the process of turning on the GAL1/10 genes through direct contact with a number of proteins at the promoter. During induction, Reb1 no longer binds to the cryptic promoter in GAL10. Gal4 recruits an H2B histone ubiquitylation factor (Rad6), which then stimulates histone di- and trimethylation of histone H3K4 by Set1. Next, the SAGA acetyltransferase complex is recruited by Gal4 and both deubiquitylates H2B and acetylates histone H3, ultimately resulting in the eviction of the poised nucleosomes from the two promoters. The removal is facilitated by the remodeler SWI/SNF and the chaperones Hsp90/70. SWI/SNF is not absolutely required but speeds up the process. This allows the recruitment of TBP/TF_IID, which then recruits RNA polymerase II and the coactivator complex Mediator. Activated Gal4 directly contacts Mediator to ultimately initiate transcription. The elongation control factor TF_IIS is also recruited, which actually plays a role in initiation for at least some genes.

During the elongation phase of transcription, nucleosomes are disrupted (see the Eukaryotic Transcription chapter). In order to prevent spurious transcription from internal cryptic promoters on either strand, histone octamers must re-form as RNA polymerase II passes. A number of histone chaperones and the FACT (facilitating chromatin transcription) complex play a role in the dynamics of octamer disassembly and assembly during elongation.

This system is also poised to rapidly repress transcription when the supply of galactose is used up or glucose becomes available. As Gal4 is activating transcription by RNA polymerase II, protein kinases associated with the activation of the polymerase also phosphorylate Gal4. This phosphorylation then leads to ubiquitination and destruction of Gal4. This turnover may be essential for RNA polymerase clearance and elongation. This is a dynamic system in which there must be a continuous positive signal, the presence of galactose.

Although catabolite repression in eukaryotes is used for the same purpose as in E. coli (which uses cAMP as a positive coregulator), it has a completely different mechanism. Glucose is a preferred sugar source compared to galactose. If the cell has both sugars, it will preferentially use the best source, glucose, and repress the genes for galactose utilization. Glucose repression of the yeast GAL genes is multifaceted. The glucose-dependent switch is the protein kinase Snf1. In low glucose, the GAL genes are transcribed because the general glucose-dependent repressor Mig1 has been inactivated, phosphorylated by Snf1. Glucose repression inactivates Snf1, which allows Mig1 to be active.

A number of other genes involving galactose usage are also downregulated in glucose, including the galactose transporter and Gal4 itself. Glucose inactivates Snf1, which leads to the activation of Mig1 at the GAL locus. Mig1 interacts at the GAL locus with the Cyc8-Tup1 corepressor, which is known to recruit histone deacetylases.

Summary

Transcription factors include basal factors, activators, and coactivators. Basal factors interact with RNA polymerase at the start point within the promoter. Activators bind specific short DNA sequence elements located near promoters or in enhancers. Activators function by making protein–protein interactions with the basal apparatus. Some activators interact directly with the basal apparatus; others require coactivators to mediate the interaction. Activators often have a modular construction in which there are independent domains responsible for binding to DNA and activating transcription. The main function of the DNA-binding domain may be to tether the activating domain in the vicinity of the initiation complex. Some response elements are present in many genes and are recognized by ubiquitous factors; others are present in a few genes and are recognized by tissue-specific factors.

Near the promoters for RNA polymerase II are a variety of short, cis-acting elements, each of which is recognized by a trans-acting factor. The cis-acting elements can be located upstream of the TATA box and may be present in either orientation and at a variety of distances with regard to the start point or downstream within an intron. These elements are recognized by activators or repressors that interact with the basal transcription complex to determine the efficiency with which the promoter is used. Some activators interact directly with components of the basal apparatus; others interact via intermediaries called coactivators. The targets in the basal apparatus are the TAFs of TF_IID, TF_IIB, or TF_IIA. The interaction stimulates assembly of the basal apparatus.

Several groups of transcription factors have been identified by sequence homology. The homeodomain is a sequence of 60 amino acids that regulates development in insects, worms, and humans. It is related to the prokaryotic helix-turn-helix motif and is the DNA-binding motif for these transcription factors.

Another motif involved in DNA binding is the zinc finger, which is found in proteins that bind DNA or RNA (or sometimes both). A zinc finger has cysteine and histidine residues that bind zinc. One type of finger is found in multiple repeats in some transcription factors; another is found in single or double repeats in others.

The leucine zipper contains a stretch of amino acids rich in leucine that are involved in dimerization of transcription factors. An adjacent basic region is responsible for binding to DNA in the bZIP transcription factors.

Steroid receptors were the first members identified of a group of transcription factors in which the protein is activated by binding of a small hydrophobic hormone. The activated factor becomes localized in the nucleus and binds to its specific response element, where it activates transcription. The DNA-binding domain has zinc fingers.

HLH (helix-loop-helix) proteins have amphipathic helices that are responsible for dimerization, which are adjacent to basic regions that bind to DNA. bHLH proteins have a basic region that binds to DNA. They fall into two groups: ubiquitously expressed and tissue specific. An active protein is usually a heterodimer between two subunits, one from each group. When a dimer has one subunit that does not have the basic region, it fails to bind DNA; thus such subunits can prevent gene expression. Combinatorial associations of subunits form regulatory networks.

Many transcription factors function as dimers, and it is common for there to be multiple members of a family that form homodimers and heterodimers. This creates the potential for complex combinations to govern gene expression. In some cases, a family includes inhibitory members whose participation in dimer formation prevents the partner from activating transcription.

Genes whose control regions are organized in nucleosomes usually are not expressed. In the absence of specific regulatory proteins, promoters and other regulatory regions are organized by histone octamers into a state in which they cannot be activated. This may explain the need for nucleosomes to be precisely positioned in the vicinity of a promoter, so that essential regulatory sites are appropriately exposed. Some transcription factors have the capacity to recognize DNA on the nucleosomal surface, and a particular positioning of DNA may be required for initiation of transcription.

Chromatin-remodeling complexes have the ability to slide or displace histone octamers by a mechanism that involves hydrolysis of ATP. Remodeling complexes range from small to extremely large and are classified according to the type of the ATPase subunit. Common types are SWI/SNF, ISWI, CHD, and SWR1/INO80. A typical form of this chromatin remodeling is to displace one or more histone octamers from specific sequences of DNA, creating a boundary that results in the precise or preferential positioning of adjacent nucleosomes. Chromatin remodeling may also involve changes in the positions of nucleosomes, sometimes involving sliding of histone octamers along DNA.

Extensive covalent modifications occur on histone tails, all of which are reversible. Acetylation of histones occurs at both replication and transcription and facilitates formation of a less compact chromatin structure, usually via interactions with ATP-dependent remodelers. Some coactivators, which connect transcription factors to the basal apparatus, have histone acetylase activity. Conversely, repressors may be associated with deacetylases. The modifying enzymes are usually specific for particular amino acids in particular histones. Some histone modifications may be exclusive or synergistic with others.

Large activating (or repressing) complexes often contain several activities that undertake different modifications of chromatin. Some common motifs found in proteins that modify chromatin are the chromodomain (which binds methylated lysine), the bromodomain (which targets acetylated lysine), and the SET domain (which is part of the active sites of histone methyltransferases).

References

26.2 How Is a Gene Turned On?

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26.3 Mechanism of Action of Activators and Repressors

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26.4 Independent Domains Bind DNA and Activate Transcription

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26.5 The Two-Hybrid Assay Detects Protein–Protein Interactions

Research

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26.6 Activators Interact with the Basal Apparatus

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26.7 Many Types of DNA-Binding Domains Have Been Identified

Reviews

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26.8 Chromatin Remodeling Is an Active Process

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26.9 Nucleosome Organization or Content Can Be Changed at the Promoter

Reviews

Lohr, D. (1997). Nucleosome transactions on the promoters of the yeast GAL and PHO genes. J. Biol. Chem. 272, 26795–26798.
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Research

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26.10 Histone Acetylation Is Associated with Transcription Activation

Reviews

Jenuwein, T., and Allis, C. D. (2001). Translating the histone code. Science 293, 1074–1080.
Lee, K. K., and Workman, J. L. (2007). Histone acetyltransferase complexes: one size doesn’t fit all. Nat. Rev. Mol. Cell Biol. 8, 284–295.
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Research

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26.11 Methylation of Histones and DNA Is Connected

Reviews

Bannister, A. J., and Kouzarides, T. (2005). Reversing histone methylation. Nature 436, 1103–1106.
Richards, E. J., Elgin, S. C, and Richards, S. C. (2002). Epigenetic codes for heterochromatin formation and silencing: rounding up the usual suspects. Cell 108, 489–500.
Zhang, Y., and Reinberg, D. (2001). Transcription regulation by histone methylation: interplay between different covalent modifications of the core histone tails. Genes Dev. 15, 2343–2360.

Research

Cuthbert, G. L., Daujat, S., Snowden, A. W., Erdjument-Bromage, H., Hagiwara, T., Yamada, M., Schneider, R., Gregory, P. D., Tempst, P., Bannister, A. J., and Kouzarides, T. (2004). Histone deimination antagonizes arginine methylation. Cell 118, 545–553.
Fuks, F., Hurd, P. J., Wolf, D., Nan, X., Bird, A. P., and Kouzarides, T. (2003). The methyl-CpG-binding protein MeCP2 links DNA methylation to histone methylation. J. Biol. Chem. 278, 4035–4040.
Gendrel, A. V., Lippman, Z., Yordan, C., Colot, V., and Martienssen, R. A. (2002). Dependence of heterochromatic histone H3 methylation patterns on the Arabidopsis gene DDM1. Science 297, 1871–1873.
Johnson, L., Cao, X., and Jacobsen, S. (2002). Interplay between two epigenetic marks: DNA methylation and histone H3 lysine 9 methylation. Curr. Biol. 12, 1360–1367.
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Shi, Y., Lan, F., Matson, C., Mulligan, P., Whetstine, J. R., Cole, P. A., and Casero, R. A. (2004). Histone demethylation mediated by the nuclear amine oxidase homolog LSD1. Cell 119, 941–953.
Tamaru, H., and Selker, E. U. (2001). A histone H3 methyltransferase controls DNA methylation in Neurospora crassa. Nature 414, 277–283.
Tamaru, H., Zhang, X., McMillen, D., Singh, P. B., Nakayama, J., Grewal, S. I., Allis, C. D., Cheng, X., and Selker, E. U. (2003). Trimethylated lysine 9 of histone H3 is a mark for DNA methylation in Neurospora crassa. Nat. Genet. 34, 75–79.
Wang, Y., Wysocka, J., Sayegh, J., Lee, Y. H., Perlin, J. R., Leonelli, L., Sonbuchner, L. S., McDonald, C. H., Cook, R. G., Dou, Y., Roeder, R. G., Clarke, S., Stallcup, M. R., Allis, C. D., and Coonrod, S. A. (2004). Human PAD4 regulates histone arginine methylation levels via demethylimination. Science 306, 279–283.

26.12 Promoter Activation Involves Multiple Changes to Chromatin

Reviews

Li, B., Carey, M., and Workman, J. L. (2007). The role of chromatin during transcription. Cell 128, 707–719.
Orphanides, G., and Reinberg, D. (2000). RNA polymerase II elongation through chromatin. Nature 407, 471–475.

Research

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Hassan, A. H., Neely, K. E., and Workman, J. L. (2001). Histone acetyltransferase complexes stabilize SWI/SNF binding to promoter nucleosomes. Cell 104, 817–827.
Krebs, J. E., Kuo, M. H., Allis, C. D., and Peterson, C. L. (1999). Cell-cycle regulated histone acetylation required for expression of the yeast HO gene. Genes Dev. 13, 1412–1421.
Orphanides, G., LeRoy, G., Chang, C. H., Luse, D. S., and Reinberg, D. (1998). FACT, a factor that facilitates transcript elongation through nucleosomes. Cell 92, 105–116.
Wada, T., Takagi, T., Yamaguchi, Y., Ferdous, A., Imai, T., Hirose, S., Sugimoto, S., Yano, K., Hartzog, G. A., Winston, F., Buratowski, S., and Handa, H. (1998). DSIF, a novel transcription elongation factor that regulates RNA polymerase II processivity, is composed of human Spt4 and Spt5 homologs. Genes Dev. 12, 343–356.

26.13 Histone Phosphorylation Affects Chromatin Structure

Research

Ciurciu, A., Komonyi, O., and Boros, I. M. (2008). Loss of ATAC-specific acetylation of histone H4 at Lys12 reduces binding of JIL-1 to chromatin and phosphorylation of histone H3 and Ser10. J. Cell Sci. 121, 3366–3372.
Wang, Y., Zhang, W., Jin, Y., Johansen, J., and Johansen, K. M. (2001). The JIL-1 tandem kinase mediates histone H3 phosphorylation and is required for maintenance of chromatin structure in Drosophila. Cell 105, 433–443.

26.14 Yeast GAL Genes: A Model for Activation and Repression

Reviews

Armstrong, J. A. (2007). Negotiating the nucleosome: factors that allow RNA polymerase II to elongate through chromatin. Biochem. Cell Biol. 85, 426–434.
Hahn, S., and Young, F. T. (2011). Transcriptional regulation in Saccharomyces cerevisiae: transcription factor regulation and function, mechanisms of initiation, and roles of activators and coactivators. Genetics 189, 705–736.
Peng, G., and Hopper, J. E. (2002). Gene activation by interaction of an inhibitor with a cytoplasmic signaling protein. Proc. Natl. Acad. Sci. USA 99, 8548–8553.
Ptashne, M. (2014). The chemistry of regulation of genes and other things. J. Biol. Chem. 289, 5417–5435.

Research

Ahuatzi, D., Riera, A., Pelaez, R., Herrero, P., and Moreno, F. (2007). Hxk2 regulates the phosphorylation state of Mig1 and therefore its nucleocytoplasmic distribution. J. Biol. Chem. 282, 4485–4493.
Bryant, G. O., Prabhu, V., Floer, M., Wang, X., Spagna, D., Schreiber, D., and Ptashne, M. (2008). Activator control of nucleosome occupancy in activation and repression of transcription. PLoS 6, 2928–2938.
Egriboz, O., Jiang, F., and Hopper, J. E. (2011). Rapid Gal gene switch of Saccharomyces cerevisiae depends on nuclear Gal3, not cytoplasmic trafficking of Gal3 and Gal80. Genetics 189, 825–836.
Floer, M., Bryant, G. O., and Ptashne, M. (2008). HSP90/70 chaperones are required for rapid nucleosome removal upon induction of the GAL genes in yeast. Proc. Natl. Acad. Sci. USA 105, 2975–2980.
Floer, M., Wong, X., Prabhu, V., Berozpe, G., Narayan, S., Spagna, D., Alvarez, D., Kendall, J., Krasnitz, A., Stepansky, A., Hicks, J., Bryant, G. O., and Ptashne, M. (2010). A RSC/nucleosome complex determines chromatin architecture and facilitates activator binding. Cell 141, 407–418.
Henikoff, J. G., Belsky, J. A., Krassovsky, K, MacAlpine, D. M., and Henikoff, S. (2011). Epigenome characterization at single base-pair resolution. Proc. Natl. Acad. Sci. USA 108, 18318–18323.
Houseley, J., Rubbi, L., Grunstein, M., Tollervey, D., and Vogelauer, M. (2008). A ncRNA modulates histone modification and mRNA induction in the yeast GAL gene cluster. Mol. Cell 32, 685–695.
Imbeault, D., Gamar, L., Rufiange, A., Paquet, E., and Nourani, A. (2008). The Rtt106 histone chaperone is functionally linked to transcription elongation and is involved in the regulation of spurious transcription from cryptic promoters in yeast. J. Biol. Chem. 283, 27350–27354.
Ingvarsdottir, K., Edwards, C., Lee, M. G., Lee, J. S., Schultz, D. C., Shilatifard, A., Shiekhattar, R., and Berger, S. (2007). Histone H3 K4 demethylation during activation and attenuation of GAL1 transcription in Saccharomyces cerevisiae. Mol. Cell Biol. 27, 7856–7864.
Kumar, P. R., Yu, Y., Sternglanz, R., Johnston, S. A., and Joshua-Tor, L. (2008). NADP regulates the yeast GAL system. Science 319, 1090–1092.
Muratani, M., Kung, C., Shokat, K. M., and Tansey, W. P. (2005). The F box protein Dsg1/Mdm30 is a transcriptional Coactivator that stimulates Gal4 turnover and cotranscriptional mRNA processing. Cell 120, 887–899.
Westergaard, S. L., Oliveira, A. P., Bro, C., Olsson, L., and Nielsen, J. (2007). A systems approach to study glucose repression in the yeast Saccharomyces cerevisiae. Biotechnol. Bioeng. 96, 134–141.