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Cell_Biology_Alberts. to the establishment and spread of heterochromatin. More generally, the recruited proteins act with the modified histones to determine how and when genes will be expressed, as well as other chromosome functions. In this way, the precise structure of each domain of chromatin governs the readout of the genetic information that it contains, and thereby the structure and function of the eukaryotic cell.


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Cell_Biology_Alberts. H3.3 gene expression,H2AZ chromosome segregation transcriptional repression,macroH2A X-chromosome inactivation Chromatin Acquires Additional variety Through the Site-Specific Insertion of a Small Set of Histone variants In addition to the four highly conserved standard core histones, eukaryotes also contain a few variant histones that can also assemble into nucleosomes. These histones are present in much smaller amounts than the major histones, and they have been less well conserved over long evolutionary times. Variants are known for each of the core histones with the exception of H4; some examples are shown in Figure 4–35.


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Cell_Biology_Alberts. The major histones are synthesized primarily during the S phase of the cell cycle and assembled into nucleosomes on the daughter DNA helices just behind the replication fork (see Figure 5–32). In contrast, most histone variants are synthesized throughout interphase. They are often inserted into already-formed chromatin, which requires a histone-exchange process catalyzed by the ATP-dependent chromatin remodeling complexes discussed previously. These remodeling complexes contain subunits that cause them to bind both to specific sites on chromatin and to histone chaperones that carry a particular variant. As a result, each histone variant is inserted into chromatin in a highly selective manner (see Figure 4–27). Covalent modifications and Histone variants Act in Concert to Control Chromosome Functions


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Cell_Biology_Alberts. Covalent modifications and Histone variants Act in Concert to Control Chromosome Functions The number of possible distinct markings on an individual nucleosome is in principle enormous, and this potential for diversity is still greater when we allow for nucleosomes that contain histone variants. However, the histone modifications are known to occur in coordinated sets. More than 15 such sets can be identified in mammalian cells. However, it is not yet clear how many different types of chromatin are functionally important in cells.


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Cell_Biology_Alberts. Some combinations are known to have a specific meaning for the cell in the sense that they determine how and when the DNA packaged in the nucleosomes is to be accessed or manipulated—a fact that led to the idea of a “histone code.” For example, one type of marking signals that a stretch of chromatin has been newly replicated, another signals that the DNA in that chromatin has been damaged and needs repair, while others signal when and how gene expression should take place. Various regulatory proteins contain small domains that bind to specific marks, recognizing, for example, a trimethylated lysine 4 on histone H3 (Figure 4–36). These domains are often linked together as modules in a single large


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Cell_Biology_Alberts. Figure 4–35 The structure of some histone variants compared with the major histone that they replace. The histone variants are inserted into nucleosomes at specific sites on chromosomes by ATP-dependent chromatin remodeling enzymes that act in concert with histone chaperones (see Figure 4–27). The CENP-A (Centromere Protein-A) variant of histone H3 is discussed later in this chapter (see Figure 4–42); other variants are discussed in Chapter 7. The sequences in each variant that are colored differently (compared to the major histone above it) denote regions with an amino acid sequence different from this major histone. (Adapted from k. Sarma and D. Reinberg, Nat. Rev. Mol. Cell Biol. 6:139–149, 2005. with permission from macmillan Publishers ltd.)


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Cell_Biology_Alberts. Figure 4–36 How a mark on a nucleosome is read. The figure shows the structure of a protein module (called an INg PHD domain) that specifically recognizes histone H3 trimethylated on lysine 4. (A) A trimethyl group. (b) Space-filling model of an INg PHD domain bound to a histone tail (green, with the trimethyl group highlighted in yellow). (C) A ribbon model showing how the N-terminal six amino acids in the H3 tail are recognized. The red lines represent hydrogen bonds. This is one of a family of PHD domains that recognize methylated lysines on histones; different members of the family bind tightly to lysines located at different positions, and they can discriminate between a mono-, di-, and trimethylated lysine. In a similar way, other small protein modules recognize specific histone side chains that have been marked with acetyl groups, phosphate groups, and so on. (Adapted from P.v. Peña et al., Nature 442:100–103, 2006. with permission from macmillan Publishers ltd.) protein or


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Cell_Biology_Alberts. side chains that have been marked with acetyl groups, phosphate groups, and so on. (Adapted from P.v. Peña et al., Nature 442:100–103, 2006. with permission from macmillan Publishers ltd.) protein or protein complex, which thereby recognizes a specific combination of histone modifications (Figure 4–37). The result is a reader complex that allows particular combinations of markings on chromatin to attract additional proteins, so as to execute an appropriate biological function at the right time (Figure 4–38).


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Cell_Biology_Alberts. The marks on nucleosomes due to covalent additions to histones are dynamic, being constantly removed and added at rates that depend on their chromosomal locations. Because the histone tails extend outward from the nucleosome core and are likely to be accessible even when chromatin is condensed, they would seem to provide an especially suitable format for creating marks that can be readily altered as a cell’s needs change. Although much remains to be learned about the meaning of the different histone modifications, a few well-studied examples of the information that can be encoded in the histone H3 tail are listed in Figure 4–39. A Complex of Reader and writer Proteins Can Spread Specific The phenomenon of position effect variegation described previously requires that some modified forms of chromatin have the ability to spread for substantial distances along a chromosomal DNA molecule (see Figure 4–31). How is this possible?


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Cell_Biology_Alberts. The enzymes that add or remove modifications to histones in nucleosomes are part of multisubunit complexes. They can initially be brought to a particular region of chromatin by one of the sequence-specific DNA-binding proteins (transcription regulators) discussed in Chapters 6 and 7 (for a specific example,


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Cell_Biology_Alberts. Figure 4–37 Recognition of a specific combination of marks on a nucleosome. In the example shown, two adjacent domains that are part of the NURF (Nucleosome Remodeling Factor) chromatin remodeling complex bind to the nucleosome, with the PHD domain (red) recognizing a methylated H3 lysine 4 and another domain (a bromodomain, blue) recognizing an acetylated H4 lysine 16. These two histone marks constitute a unique histone modification pattern that occurs in subsets of nucleosomes in human cells. Here the two histone tails are indicated by green dotted lines, and only half of one nucleosome is shown. (Adapted from A.J. Ruthenburg et al., Cell 145:692–706, 2011. with permission from Elsevier.) protein modules scaffold binding to specifc protein histone modifcations on nucleosome attachment to other components in nucleus, leading to gene expression, gene silencing, or other biological function see Figure 7–20). But after a modifying enzyme “writes” its mark on one or a few neighboring


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Cell_Biology_Alberts. to other components in nucleus, leading to gene expression, gene silencing, or other biological function see Figure 7–20). But after a modifying enzyme “writes” its mark on one or a few neighboring nucleosomes, events that resemble a chain reaction can ensue. In such a case, the “writer enzyme” works in concert with a “reader protein” located in the same protein complex. The reader protein contains a module that recognizes the mark and binds tightly to the newly modified nucleosome (see Figure heterochromatin formation, gene silencingK 9


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Cell_Biology_Alberts. Figure 4–38 Schematic diagram showing how a particular combination of histone modifications can be recognized by a reader complex. A large protein complex that contains a series of protein modules, each of which recognizes a specific histone mark, is schematically illustrated (green). This “reader complex” will bind tightly only to a region of chromatin that contains several of the different histone marks that it recognizes. Therefore, only a specific combination of marks will cause the complex to bind to chromatin and attract the additional protein complexes (purple) needed to catalyze a biological function.


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Cell_Biology_Alberts. Figure 4–39 Some specific meanings of histone modifications. (A) The modifications on the histone H3 N-terminal tail are shown, repeated from Figure 4–34. (b) The H3 tail can be marked by different sets of modifications that act in combination to convey a specific meaning. Only a small number of the meanings are known, including the three examples shown. Not illustrated is the fact that, as just implied (see Figure 4–38), reading a histone mark generally involves the joint recognition of marks at other sites on the nucleosome along with the indicated H3 tail recognition. In addition, specific levels of methylation (mono-, di-, or trimethyl groups) are generally required. Thus, for example, the trimethylation of lysine 9 attracts the heterochromatin-specific protein HP1, which induces a spreading wave of further lysine 9 trimethylation followed by further HP1 binding, according to the general scheme that will be illustrated shortly (see Figure 4–40). Also important in this process,


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Cell_Biology_Alberts. spreading wave of further lysine 9 trimethylation followed by further HP1 binding, according to the general scheme that will be illustrated shortly (see Figure 4–40). Also important in this process, however, is a synergistic trimethylation of the histone H4 N-terminal tail on lysine 20.


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Cell_Biology_Alberts. 4–36), activating an attached writer enzyme and positioning it near an adjacent nucleosome. Through many such read–write cycles, the reader protein can carry the writer enzyme along the DNA—spreading the mark in a hand-over-hand manner along the chromosome (Figure 4–40). In reality, the process is more complicated than the scheme just described. Both readers and writers are part of a protein complex that is likely to contain multiple readers and writers, and to require multiple marks on the nucleosome to spread. Moreover, many of these reader–writer complexes also contain an ATP-dependent chromatin remodeling protein (see Figure 4–26C), and the reader, writer, and remodeling proteins can work in concert to either decondense or condense long stretches of chromatin as the reader moves progressively along the nucleosome-packaged DNA.


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Cell_Biology_Alberts. A similar process is used to remove histone modifications from specific regions of the DNA; in this case, an “eraser enzyme,” such as a histone demethylase or his-tone deacetylase, is recruited to the complex. As for the writer complex in Figure 4–40, sequence-specific DNA-binding proteins (transcription regulators) direct where such modifications occur (discussed in Chapter 7). Some idea of the complexity of the above processes can be derived from the results of genetic screens for genes that either enhance or suppress the spreading and stability of heterochromatin, as manifest in effects on position effect variegation in Drosophila (see Figure 4–32). As pointed out previously, more than 100 such genes are known, and most of them are likely to code for subunits in one or more reader–writer–remodeling protein complexes.


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Cell_Biology_Alberts. Figure 4–40 How the recruitment of a reader–writer complex can spread chromatin changes along a chromosome. The writer is an enzyme that creates a specific modification on one or more of the four nucleosomal histones. After its recruitment to a specific site on a chromosome by a transcription regulatory protein, the writer collaborates with a reader protein to spread its mark from nucleosome to nucleosome by means of the indicated reader–writer complex. For this mechanism to work, the reader must recognize the same histone modification mark that the writer produces; its binding to that mark can be shown to activate the writer. In this schematic example, a spreading wave of chromatin condensation is thereby induced. Not shown are the additional proteins involved, including an ATP-dependent chromatin remodeling complex required to reposition the modified nucleosomes. barrier DNA Sequences block the Spread of Reader–writer Complexes and thereby Separate Neighboring Chromatin Domains


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Cell_Biology_Alberts. barrier DNA Sequences block the Spread of Reader–writer Complexes and thereby Separate Neighboring Chromatin Domains The above mechanism for spreading chromatin structures raises a potential problem. Inasmuch as each chromosome contains one continuous, very long DNA molecule, what prevents a cacophony of confusing cross-talk between adjacent chromatin domains of different structure and function? Early studies of position effect variegation had suggested an answer: certain DNA sequences mark the boundaries of chromatin domains and separate one such domain from another (see Figure 4–31). Several such barrier sequences have now been identified and characterized through the use of genetic engineering techniques that allow specific DNA segments to be deleted from, or inserted in, chromosomes.


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Cell_Biology_Alberts. For example, in cells that are destined to give rise to red blood cells, a sequence called HS4 normally separates the active chromatin domain that contains the human β-globin locus from an adjacent region of silenced, condensed chromatin. If this sequence is deleted, the β-globin locus is invaded by condensed chromatin. This chromatin silences the genes it covers, and it spreads to a different extent in different cells, causing position effect variegation similar to that observed in Drosophila. As described in Chapter 7, the consequences are dire: the globin genes are poorly expressed, and individuals who carry such a deletion have a severe form of anemia.


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Cell_Biology_Alberts. In genetic engineering experiments, the HS4 sequence is often added to both ends of a gene that is to be inserted into a mammalian genome, in order to protect that gene from the silencing caused by spreading heterochromatin. Analysis of this barrier sequence reveals that it contains a cluster of binding sites for histone acetylase enzymes. Since the acetylation of a lysine side chain is incompatible with the methylation of the same side chain, and specific lysine methylations are required to spread heterochromatin, histone acetylases are logical candidates for the formation of DNA barriers to spreading (Figure 4–41). However, several other types of chromatin modifications are known that can also protect genes from silencing. Figure 4–41 Some mechanisms of barrier action. These models are derived from experimental analyses of barrier action, and a combination of several of them may function at any one site.


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Cell_Biology_Alberts. Figure 4–41 Some mechanisms of barrier action. These models are derived from experimental analyses of barrier action, and a combination of several of them may function at any one site. The tethering of a region of chromatin to a large fixed site, such as the nuclear pore complex illustrated here, can form a barrier that stops the spread of heterochromatin. The tight binding of barrier proteins to a group of nucleosomes can make this chromatin resistant to heterochromatin spreading. (C) by recruiting a group of highly active histone-modifying enzymes, barriers can erase the histone marks that are required for heterochromatin to spread. For example, a potent acetylation of lysine 9 on histone H3 will compete with lysine 9 methylation, thereby preventing the binding of the HP1 protein needed to form a major form of heterochromatin. (based on A.g. west and P. Fraser, Hum. Mol. Genet. 14:R101–R111, 2005. with permission from Oxford University Press.)


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Cell_Biology_Alberts. A.g. west and P. Fraser, Hum. Mol. Genet. 14:R101–R111, 2005. with permission from Oxford University Press.) The Chromatin in Centromeres Reveals How Histone variants Can Create Special Structures


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Cell_Biology_Alberts. Nucleosomes carrying histone variants have a distinctive character and are thought to be able to produce marks in chromatin that are unusually long-lasting. An important example is seen in the formation and inheritance of the specialized chromatin structure at the centromere, the region of each chromosome required for attachment to the mitotic spindle and orderly segregation of the duplicated copies of the genome into daughter cells each time a cell divides. In many complex organisms, including humans, each centromere is embedded in a stretch of special centromeric chromatin that persists throughout interphase, even though the centromere-mediated attachment to the spindle and movement of DNA occur only during mitosis. This chromatin contains a centromere-specific variant H3 histone, known as CENP-A (Centromere Protein-A; see Figure 4–35), plus additional proteins that pack the nucleosomes into particularly dense arrangements and form the kinetochore, the special structure required for


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Cell_Biology_Alberts. as CENP-A (Centromere Protein-A; see Figure 4–35), plus additional proteins that pack the nucleosomes into particularly dense arrangements and form the kinetochore, the special structure required for attachment of the mitotic spindle (see Figure 4–19).


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Cell_Biology_Alberts. A specific DNA sequence of approximately 125 nucleotide pairs is sufficient to serve as a centromere in the yeast S. cerevisiae. Despite its small size, more than a dozen different proteins assemble on this DNA sequence; the proteins include the CENP-A histone H3 variant, which, along with the three other core histones, forms a centromere-specific nucleosome. The additional proteins at the yeast centromere attach this nucleosome to a single microtubule from the yeast mitotic spindle (Figure 4–42).


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Cell_Biology_Alberts. The centromeres in more complex organisms are considerably larger than those in budding yeasts. For example, fly and human centromeres extend over hundreds of thousands of nucleotide pairs and, while they contain CENP-A, they do not seem to contain a centromere-specific DNA sequence. These centromeres largely consist of short, repeated DNA sequences, known as alpha satellite DNA in humans. But the same repeat sequences are also found at other (non-centromeric) positions on chromosomes, indicating that they are not sufficient to direct centromere formation. Most strikingly, in some unusual cases, new human centromeres (called neocentromeres) have been observed to form spontaneously on fragmented chromosomes. Some of these new positions were originally euchromatic and lack alpha satellite DNA altogether (Figure 4–43). It seems that centromeres in complex organisms are defined by an assembly of proteins, rather than by a specific DNA sequence.


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Cell_Biology_Alberts. Inactivation of some centromeres and genesis of others de novo seem to have played an essential part in evolution. Different species, even when quite closely Figure 4–42 A model for the structure of a simple centromere. (A) In the yeast Saccharomyces cerevisiae, a special centromeric DNA sequence assembles a single nucleosome in which two copies of an H3 variant histone (called CENP-A in most organisms) replace the normal H3. (b) How peptide sequences unique to this variant histone (see Figure 4–35) help to assemble additional proteins, some of which form a kinetochore. The yeast kinetochore is unusual in capturing only a single microtubule; humans have much larger centromeres and form kinetochores that can capture 20 or more microtubules (see Figure 4–43). The kinetochore is discussed in detail in Chapter 17. (Adapted from A. Joglekar et al., Nat. Cell Biol. 8:581–585, 2006. with permission from macmillan Publishers ltd.)


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Cell_Biology_Alberts. Figure 4–43 evidence for the plasticity of human centromere formation. (A) A series of A-T-rich alpha satellite DNA sequences is repeated many thousands of times at each human centromere (red), and is surrounded by pericentric heterochromatin (brown). However, due to an ancient chromosome breakage-and-rejoining event, some human chromosomes contain two blocks of alpha satellite DNA, each of which presumably functioned as a centromere in its original chromosome. Usually, chromosomes with two functional centromeres are not stably propagated because they attach improperly to the spindle and are broken apart during mitosis. In chromosomes that do survive, however, one of the centromeres has somehow become inactivated, even though it contains all the necessary DNA sequences. This allows the chromosome to be stably propagated. (b) In a small fraction (1/2000) of human births, extra chromosomes are observed in cells of the offspring. Some of these extra chromosomes, which have formed from a


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Cell_Biology_Alberts. to be stably propagated. (b) In a small fraction (1/2000) of human births, extra chromosomes are observed in cells of the offspring. Some of these extra chromosomes, which have formed from a breakage event, lack alpha satellite DNA altogether, yet new centromeres (neocentromeres) have arisen from what was originally euchromatic DNA.


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Cell_Biology_Alberts. The complexity of centromeric chromatin is not illustrated in these diagrams. The alpha satellite DNA that forms centromeric chromatin in humans is packaged into alternating blocks of chromatin. One block is formed from a long string of nucleosomes containing the CENP-A H3 variant histone; the other block contains nucleosomes that are specially marked with dimethyl lysine 4 on the normal H3 histone. Each block is more than a thousand nucleosomes long. This centromeric chromatin is flanked by pericentric heterochromatin, as shown. The pericentric chromatin contains methylated lysine 9 on its H3 histones, along with HP1 protein, and it is an example of “classical” heterochromatin (see Figure 4–39).


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Cell_Biology_Alberts. related, often have different numbers of chromosomes; see Figure 4–14 for an extreme example. As we shall discuss below, detailed genome comparisons show that in many cases the changes in chromosome numbers have arisen through chromosome breakage-and-rejoining events, creating novel chromosomes, some of which must initially have contained abnormal numbers of centromeres—either more than one, or none at all. Yet stable inheritance requires that each chromosome should contain one centromere, and one only. It seems that surplus centromeres must have been inactivated, and/or new centromeres created, so as to allow the rearranged chromosome sets to be stably maintained. The changes in centromere activity just discussed, once established, need to be perpetuated through subsequent cell generations. What could be the mechanism of this type of epigenetic inheritance?


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Cell_Biology_Alberts. It has been proposed that de novo centromere formation requires an initial seeding event, involving the formation of a specialized DNA–protein structure that contains nucleosomes formed with the CENP-A variant of histone H3. In humans, this seeding event happens more readily on arrays of alpha satellite DNA than on other DNA sequences. The H3–H4 tetramers from each nucleosome on the parental DNA helix are directly inherited by the sister DNA helices at a replication fork (see Figure 5–32). Therefore, once a set of CENP-A-containing nucleosomes has been assembled on a stretch of DNA, it is easy to understand how a new centromere could be generated in the same place on both daughter chromosomes following each round of cell division. One need only assume that the presence of the CENP-A histone in an inherited nucleosome selectively recruits more CENP-A histone to its newly formed neighbors.


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Cell_Biology_Alberts. There are some striking similarities between the formation and maintenance of centromeres and the formation and maintenance of some other regions of heterochromatin. In particular, the entire centromere forms as an all-or-none entity, suggesting that the creation of centromeric chromatin is a highly cooperative process, spreading out from an initial seed in a manner reminiscent of the phenomenon of position effect variegation that we discussed earlier. In both cases, a particular chromatin structure, once formed, seems to be directly inherited on the DNA following each round of chromosome replication. A cooperative recruitment of proteins, along with the action of reader–writer complexes, can thus not only account for the spreading of specific forms of chromatin in space along the chromosome, but also for its propagation across cell generations—from parent cell to daughter cell (Figure 4–44).


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Cell_Biology_Alberts. Experiments with Frog Embryos Suggest that both Activating and Repressive Chromatin Structures Can be Inherited Epigenetically Epigenetic inheritance plays a central part in the creation of multicellular organisms. Their differentiated cell types become established during development, and persist thereafter even through repeated cell-division cycles. The daughters of a liver cell persist as liver cells, those of an epidermal cell as epidermal cells, and so on, even though they all contain the same genome; and this is because distinctive patterns of gene expression are passed on faithfully from parent cell to daughter cell. Chromatin structure has a role in this epigenetic transmission of information from one cell generation to the next.


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Cell_Biology_Alberts. One type of evidence comes from studies in which the nucleus of a cell from a frog or tadpole is transplanted into a frog egg whose own nucleus has been removed (an enucleated egg). In a classic set of experiments performed in 1968, it was shown that a nucleus taken from a differentiated donor cell can be reprogrammed in this way to support development of a whole new tadpole (see Figure 7–2). But this reprogramming occurs only with difficulty, and it becomes less and less efficient as nuclei from older animals are used. Thus, for example, less than 2% of the enucleated eggs injected with a nucleus from a tadpole epithelial cell developed to the swimming tadpole stage, compared with 35% when the donor nuclei were taken instead from an early (gastrula-stage) embryo. With new experimental tools, the cause of this resistance to reprogramming can now be traced. It arises, at least in part, because specific chromatin structures in the original differentiated nucleus tend to persist and be


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Cell_Biology_Alberts. the cause of this resistance to reprogramming can now be traced. It arises, at least in part, because specific chromatin structures in the original differentiated nucleus tend to persist and be transmitted through the many cell-division cycles required for embryonic development. In experiments with Xenopus embryos, specific forms of either repressive or active chromatin structures could be demonstrated to persist through as many as 24 cell divisions, causing the misplaced expression of genes. Figure 4–45 briefly describes one such experiment,


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Cell_Biology_Alberts. Figure 4–44 How the packaging of DNA in chromatin can be inherited following chromosome replication. In this model, some of the specialized chromatin components are distributed to each sister chromosome after DNA duplication, along with the specially marked nucleosomes that they bind. After DNA replication, the inherited nucleosomes that are specially modified, acting in concert with the inherited chromatin components, change the pattern of histone modification on the newly formed nucleosomes nearby. This creates new binding sites for the same chromatin components, which then assemble to complete the structure. The latter process is likely to involve reader– writer–remodeling complexes operating in a manner similar to that previously illustrated in Figure 4–40.


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Cell_Biology_Alberts. inject normal no injection inject mutant H3.3 mRNA (control) H3.3 mRNA cells analyzed for MyoD expression and for H3.3 histone on MyoD promoter focused on chromatin containing the histone variant, H3.3. We shall return to these phenomena in the final section of Chapter 22, where we discuss stem cells and the ways in which one cell type can be converted into another. Although a great deal remains to be learned about the functions of different chromatin structures, the packaging of DNA into nucleosomes was probably crucial for the evolution of eukaryotes like ourselves. To form a complex multicellular organism, the cells in different lineages must specialize by changing the accessibility and activity of many hundreds of genes. As described in Chapter 21, this process depends on cell memory: each cell holds a record of its past developmental history in the regulatory circuits that control its many genes. That record, it seems, is partly stored in the structure of the chromatin.


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Cell_Biology_Alberts. Although bacteria also have cell memory mechanisms, the complexity of the memory circuits in higher eukaryotes is unparalleled. Strategies based on local variations in chromatin structure, unique to eukaryotes, can enable individual genes, once they are switched on or switched off, to stay in that state until some new factor acts to reverse it. At one extreme are structures like centromeric chromatin that, once established, are stably inherited from one cell generation to the next. Likewise, the major “classical” type of heterochromatin, which contains long arrays of the HP1 protein (see Figure 4–39), can persist stably throughout life. In contrast, a form of condensed chromatin that is created by the Polycomb group of proteins serves to silence genes that must be kept inactive in some conditions, but are active in others. The latter mechanism governs the expression of a large number of genes that encode transcription regulators important in early embryonic development, as discussed


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Cell_Biology_Alberts. but are active in others. The latter mechanism governs the expression of a large number of genes that encode transcription regulators important in early embryonic development, as discussed in Chapter 21. There are many other variant forms of chromatin, some with much shorter lifetimes, often less than the division time of the cell. We shall say more about the variety of chromatin types in the next section.


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Cell_Biology_Alberts. Figure 4–45 evidence for the inheritance of a gene-activating chromatin state.


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Cell_Biology_Alberts. The well-characterized MyoD gene encodes a master transcription regulatory protein for muscle, myoD (see p. 399). This gene is normally turned on in the indicated region of the young embryo where somites form. when a nucleus from this region is injected into an enucleated egg as shown, many of the progeny cell nuclei abnormally express the myoD protein in non-muscle regions of the “nuclear transplant embryo” that forms. This abnormal expression can be attributed to maintenance of the MyoD promoter region in its active chromatin state through the many cycles of cell division that produce the blastula-stage embryo—a so-called “epigenetic memory” that persists in this case in the absence of transcription. The active chromatin surrounding the MyoD promoter contains the variant histone H3.3 (see Figure 4–35) in a lys4 methylated form. As indicated, an overproduction of this histone caused by injecting excess mRNA encoding the normal H3.3 protein increases both H3.3 occupancy on the MyoD


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Cell_Biology_Alberts. (see Figure 4–35) in a lys4 methylated form. As indicated, an overproduction of this histone caused by injecting excess mRNA encoding the normal H3.3 protein increases both H3.3 occupancy on the MyoD promoter and the epigenetic myoD production, whereas injection of an mRNA producing a mutant form of H3.3 that cannot be methylated at lys4 reduces the epigenetic myoD production. Such experiments provide evidence that an inherited chromatin state underlies the epigenetic memory observed. (Adapted from R.k. Ng and J.b. gurdon, Nat. Cell Biol. 10:102–109, 2008. with permission from macmillan Publishers ltd.)


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Cell_Biology_Alberts. In the chromosomes of eukaryotes, DNA is uniformly assembled into nucleosomes, but a variety of different chromatin structures is possible. This variety is based on a large set of reversible covalent modifications of the four histones in the nucleosome core. These modifications include the mono-, di-, and trimethylation of many different lysine side chains, an important reaction that is incompatible with the acetylation that can occur on the same lysines. Specific combinations of the modifications mark many nucleosomes, governing their interactions with other proteins. These marks are read when protein modules that are part of a larger protein complex bind to the modified nucleosomes in a region of chromatin. These reader proteins then attract additional proteins that perform various functions.


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Cell_Biology_Alberts. Some reader protein complexes contain a histone-modifying enzyme, such as a histone lysine methylase, that “writes” the same mark that the reader recognizes. A reader–writer–remodeling complex of this type can spread a specific form of chromatin along a chromosome. In particular, large regions of condensed heterochromatin are thought to be formed in this way. Heterochromatin is commonly found around centromeres and near telomeres, but it is also present at many other positions in chromosomes. The tight packaging of DNA into heterochromatin usually silences the genes within it.


Cell_Biology_Alberts

Cell_Biology_Alberts. The phenomenon of position effect variegation provides strong evidence for the inheritance of condensed states of chromatin from one cell generation to the next. A similar mechanism appears to be responsible for maintaining the specialized chromatin at centromeres. More generally, the ability to propagate specific chromatin structures across cell generations makes possible an epigenetic cell memory process that plays a role in maintaining the set of different cell states required by complex multicellular organisms.


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Cell_Biology_Alberts. Having discussed the DNA and protein molecules from which the chromatin fiber is made, we now turn to the organization of the chromosome on a more global scale and the way in which its various domains are arranged in space. As a 30-nm fiber, a typical human chromosome would still be 0.1 cm in length and able to span the nucleus more than 100 times. Clearly, there must be a still higher level of folding, even in interphase chromosomes. Although the molecular details are still largely a mystery, this higher-order packaging almost certainly involves the folding of the chromatin into a series of loops and coils. This chromatin packing is fluid, frequently changing in response to the needs of the cell.


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Cell_Biology_Alberts. We begin this section by describing some unusual interphase chromosomes that can be easily visualized. Exceptional though they are, these special cases reveal features that are thought to be representative of all interphase chromosomes. Moreover, they provide ways to investigate some fundamental aspects of chromatin structure that we have touched on in the previous section. Next, we describe how a typical interphase chromosome is arranged in the mammalian cell nucleus. Finally, we shall discuss the additional tenfold compaction that chromosomes undergo in the passage from interphase to mitosis. Chromosomes Are Folded into large loops of Chromatin


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Cell_Biology_Alberts. Chromosomes Are Folded into large loops of Chromatin Insight into the structure of the chromosomes in interphase cells has come from studies of the stiff and enormously extended chromosomes in growing amphibian oocytes (immature eggs). These very unusual lampbrush chromosomes (the largest chromosomes known), paired in preparation for meiosis, are clearly visible even in the light microscope, where they are seen to be organized into a series of large chromatin loops emanating from a linear chromosomal axis (Figure 4–46 and Figure 4–47). In these chromosomes, a given loop always contains the same DNA sequence that remains extended in the same manner as the oocyte grows. These chromosomes are producing large amounts of RNA for the oocyte, and most of the genes Figure 4–46 A model for the chromatin domains in a lampbrush chromosome.


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Cell_Biology_Alberts. Figure 4–46 A model for the chromatin domains in a lampbrush chromosome. Shown is a small portion of one pair of sister chromatids. Here, two identical DNA double helices are aligned side by side, packaged into different types of chromatin. The set of lampbrush chromosomes in many amphibians contains a total of about 10,000 loops resembling those shown here. The rest of the DNA in each chromosome (the great majority) remains highly condensed. Four copies of each loop are present in the cell, since each lampbrush chromosome consists of two aligned sets of paired chromatids. This four-stranded structure is characteristic of this stage of development of the oocyte, which has arrested at the diplotene stage of meiosis; see Figure 17–56. present in the DNA loops are being actively expressed. The majority of the DNA, however, is not in loops but remains highly condensed on the chromosome axis, where genes are generally not expressed.


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Cell_Biology_Alberts. It is thought that the interphase chromosomes of all eukaryotes are similarly arranged in loops. Although these loops are normally too small and fragile to be easily observed in a light microscope, other methods can be used to infer their presence. For example, modern DNA technologies have made it possible to assess the frequency with which any two loci along an interphase chromosome are held together, thus revealing likely candidates for the sites on chromatin that form the bases of loop structures (Figure 4–48). These experiments and others suggest that the DNA in human chromosomes is likely to be organized into loops of various lengths. A typical loop might contain between 50,000 and 200,000 nucleotide pairs of DNA, although loops of a million nucleotide pairs have also been suggested (Figure 4–49).


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Cell_Biology_Alberts. Further insight has come from another unusual class of cells—the polytene cells of flies, such as the fruit fly Drosophila. Some types of cells, in many organisms, grow abnormally large through multiple cycles of DNA synthesis without cell division. Such cells, containing increased numbers of standard chromosomes, are said to be polyploid. In the salivary glands of fly larvae, this process is taken to an extreme degree, creating huge cells that contain hundreds or thousands of copies of the Figure 4–47 lampbrush chromosomes.


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Cell_Biology_Alberts. (A) A light micrograph of lampbrush chromosomes in an amphibian oocyte. Early in oocyte differentiation, each chromosome replicates to begin meiosis, and the homologous replicated chromosomes pair to form this highly extended structure containing a total of four replicated DNA double helices, or chromatids. The lampbrush chromosome stage persists for months or years, while the oocyte builds up a supply of materials required for its ultimate development into a new individual. (b) An enlarged region of a similar chromosome, stained with a fluorescent reagent that makes the loops active in RNA synthesis clearly visible. (Courtesy of Joseph g. gall.) genome. Moreover, in this case, all the copies of each chromosome are aligned side by side in exact register, like drinking straws in a box, to create giant polytene chromosomes. These allow features to be detected that are thought to be shared with ordinary interphase chromosomes, but are normally hard to see.


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Cell_Biology_Alberts. When polytene chromosomes from a fly’s salivary glands are viewed in the light microscope, distinct alternating dark bands and light interbands are visible (Figure 4–50), each formed from a thousand identical DNA sequences arranged side by side in register. About 95% of the DNA in polytene chromosomes is in bands, and 5% is in interbands. A very thin band can contain 3000 nucleotide pairs, while a thick band may contain 200,000 nucleotide pairs in each of its chromatin strands. The chromatin in each band appears dark because the DNA is more condensed than the DNA in interbands; it may also contain a higher concentration of proteins (Figure 4–51). This banding pattern seems to reflect the same sort of organization detected in the amphibian lampbrush chromosomes described earlier.


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Cell_Biology_Alberts. There are approximately 3700 bands and 3700 interbands in the complete set of Drosophila polytene chromosomes. The bands can be recognized by their different thicknesses and spacings, and each one has been given a number to generate a chromosome “map” that has been indexed to the finished genome sequence of this fly. The Drosophila polytene chromosomes provide a good starting point for examining how chromatin is organized on a large scale. In the previous section, we saw that there are many forms of chromatin, each of which contains nucleosomes with a different combination of modified histones. Specific sets of non-histone proteins assemble on these nucleosomes to affect biological function in different ways. Recruitment of some of these non-histone proteins can spread for long distances along the DNA, imparting a similar chromatin structure to broad tracts DNA product is obtained only if proteins hold the two DNA sequences close together in the cell


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Cell_Biology_Alberts. Figure 4–48 A method for determining the position of loops in interphase chromosomes. In this technique, known as the chromosome conformation capture (3C) method, cells are treated with formaldehyde to create the indicated covalent DNA–protein and DNA–DNA cross-links. The DNA is then treated with an enzyme (a restriction nuclease) that chops the DNA into many pieces, cutting at strictly defined nucleotide sequences and forming sets of identical “cohesive ends” (see Figure 8–28). The cohesive ends can be made to join through their complementary base-pairing. Importantly, prior to the ligation step shown, the DNA is diluted so that the fragments that have been kept in close proximity to each other (through cross-linking) are the ones most likely to join. Finally, the cross-links are reversed and the newly ligated fragments of DNA are identified and quantified by PCR (the polymerase chain reaction, described in Chapter 8). From the results, combined with DNA sequence information, one can


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Cell_Biology_Alberts. and the newly ligated fragments of DNA are identified and quantified by PCR (the polymerase chain reaction, described in Chapter 8). From the results, combined with DNA sequence information, one can derive models for the interphase conformation of chromosomes.


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Cell_Biology_Alberts. Figure 4–49 A model for the organization of an interphase chromosome. A section of an interphase chromosome is shown folded into a series of looped domains, each containing perhaps 50,000–200,000 or more nucleotide pairs of double-helical DNA condensed into a chromatin fiber. The chromatin in each individual loop is further condensed through poorly understood folding processes that are reversed when the cell requires direct access to the DNA packaged in the loop. Neither the composition of the postulated chromosomal axis nor how the folded chromatin fiber is anchored to it is clear. However, in mitotic chromosomes, the bases of the chromosomal loops are enriched both in condensins (discussed below) and in DNA topoisomerase II enzymes (discussed in Chapter 5), two proteins that may form much of the axis at metaphase.


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Cell_Biology_Alberts. normal mitotic right arm of chromosomes at chromosome 2 same scale region where two homologous chromosomes are separated left arm of chromosome 2 left arm of chromocenter chromosome 3 20 µm right arm of chromosome 3 of the genome (see Figure 4–40). Such regions, where all of the chromatin has a similar structure, are separated from neighboring domains by barrier proteins (see Figure 4–41). At low resolution, the interphase chromosome can therefore be considered as a mosaic of chromatin structures, each containing particular nucleosome modifications associated with a particular set of non-histone proteins. Polytene chromosomes allow us to see details of this mosaic of domains in the light microscope, as well as to observe some of the changes associated with gene expression. There Are multiple Forms of Chromatin


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Cell_Biology_Alberts. There Are multiple Forms of Chromatin By staining Drosophila polytene chromosomes with antibodies, or by using a more recent technique called ChIP (chromatin immunoprecipitation) analysis (see Chapter 8), the locations of the histone and non-histone proteins in chromatin can be mapped across the entire DNA sequence of an organism’s genome. Such an analysis in Drosophila has thus far localized more than 50 different chromatin proteins and histone modifications. The results suggest that three major types of repressive chromatin predominate in this organism, along with two major types of chromatin on actively transcribed genes, and that each type is associated with a different complex of non-histone proteins. Thus, classical heterochromatin contains more than six such proteins, including heterochromatin protein 1 (HP1),


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Cell_Biology_Alberts. Figure 4–50 The entire set of polytene chromosomes in one Drosophila salivary cell. In this drawing of a light micrograph, the giant chromosomes have been spread out for viewing by squashing them against a microscope slide. Drosophila has four chromosomes, and there are four different chromosome pairs present. but each chromosome is tightly paired with its homolog (so that each pair appears as a single structure), which is not true in most nuclei (except in meiosis). Each chromosome has undergone multiple rounds of replication, and the homologs and all their duplicates have remained in exact register with each other, resulting in huge chromatin cables many DNA strands thick.


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Cell_Biology_Alberts. The four polytene chromosomes are normally linked together by heterochromatic regions near their centromeres that aggregate to create a single large chromocenter (pink region). In this preparation, however, the chromocenter has been split into two halves by the squashing procedure used. (Adapted from T.S. Painter, J. Hered. 25:465–476, 1934. with permission from Oxford University Press.) Figure 4–51 Micrographs of polytene chromosomes from Drosophila salivary glands. (A) light micrograph of a portion of a chromosome. The DNA has been stained with a fluorescent dye, but a reverse image is presented here that renders the DNA black rather than white; the bands are clearly seen to be regions of increased DNA concentration. This chromosome has been processed by a high-pressure treatment so as to show its distinct pattern of bands and interbands more clearly.


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Cell_Biology_Alberts. (b) An electron micrograph of a small section of a Drosophila polytene chromosome seen in thin section. bands of very different thickness can be readily distinguished, separated by interbands, which contain less condensed chromatin. (A, adapted from D.v. Novikov, I. kireev and A.S. belmont, Nat. Methods 4:483– 485, 2007. with permission from macmillan Publishers ltd; b, courtesy of veikko Sorsa.) Figure 4–52 RNA synthesis in polytene chromosome puffs.


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Cell_Biology_Alberts. Figure 4–52 RNA synthesis in polytene chromosome puffs. An autoradiograph of a single puff in a polytene chromosome from the salivary glands of the freshwater midge Chironomus tentans. As outlined in Chapter 1 and described in detail in Chapter 6, the first step in gene expression is the synthesis of an RNA molecule using the DNA as a template. The decondensed portion of the chromosome is undergoing RNA synthesis and has become labeled with 3H-uridine, an RNA precursor molecule that is incorporated into growing RNA chains. (Courtesy of José bonner.) whereas the so-called Polycomb form of heterochromatin contains a similar number of proteins of a different set (PcG proteins). In addition to the five major chromatin types, other more minor forms of chromatin appear to be present, each of which may be differently regulated and have distinct roles in the cell.


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Cell_Biology_Alberts. The set of proteins bound as part of the chromatin at a given locus varies depending on the cell type and its stage of development. These variations make the accessibility of specific genes different in different tissues, helping to generate the cell diversification that accompanies embryonic development (described in Chapter 21). Chromatin loops Decondense when the genes within Them Are Expressed When an insect progresses from one developmental stage to another, distinctive chromosome puffs arise and old puffs recede in its polytene chromosomes as new genes become expressed and old ones are turned off (Figure 4–52). From inspection of each puff when it is relatively small and the banding pattern is still discernible, it seems that most puffs arise from the decondensation of a single chromosome band.


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Cell_Biology_Alberts. The individual chromatin fibers that make up a puff can be visualized with an electron microscope. In favorable cases, loops are seen, much like those observed in amphibian lampbrush chromosomes. When genes in the loop are not expressed, the loop assumes a thickened structure, possibly that of a folded 30-nm fiber, but when gene expression is occurring, the loop becomes more extended. In electron micrographs, the chromatin located on either side of the decondensed loop appears considerably more compact, suggesting that a loop constitutes a distinct functional domain of chromatin structure.


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Cell_Biology_Alberts. Observations in human cells also suggest that highly folded loops of chromatin expand to occupy an increased volume when a gene within them is expressed. For example, quiescent chromosome regions from 0.4 to 2 million nucleotide pairs in length appear as compact dots in an interphase nucleus when visualized by fluorescence microscopy. However, the same DNA is seen to occupy a larger territory when its genes are expressed, with elongated, punctate structures replacing the original dot. New ways of visualizing individual chromosomes have shown that each of the 46 interphase chromosomes in a human cell tends to occupy its own discrete ter- ritory within the nucleus: that is, the chromosomes are not extensively entangled with one another (Figure 4–53). However, pictures such as these present only an average view of the DNA in each chromosome. Experiments that specifically localize the heterochromatic regions of a chromosome reveal that they are often


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Cell_Biology_Alberts. Figure 4–53 Simultaneous visualization of the chromosome territories for all of the human chromosomes in a single interphase nucleus. Here, a mixture of DNA probes for each chromosome has been labeled so as to fluoresce with a different spectra; this allows DNA–DNA hybridization to be used to detect each chromosome, as in Figure 4–10. Three-dimensional reconstructions were then produced. below the micrograph, each chromosome is identified in a schematic of the actual image. Note that homologous chromosomes (e.g., the two copies of chromosome 9) are not in general co-located. (From m.R. Speicher and N.P. Carter, Nat. Rev. Genet. 6:782–792, 2005. with permission from macmillan Publishers ltd.)


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Cell_Biology_Alberts. Figure 4–54 The distribution of gene-rich regions of the human genome in an interphase nucleus. gene-rich regions have been visualized with a fluorescent probe that hybridizes to the Alu interspersed repeat, which is present in more than a million copies in the human genome (see page 292). For unknown reasons, these sequences cluster in chromosomal regions rich in genes. In this representation, regions enriched for the Alu sequence are green, regions depleted for these sequences are red, while the average regions are yellow. The gene-rich regions are seen to be largely absent in the DNA near the nuclear envelope. (From A. bolzer et al., PLoS Biol. 3:826–842, 2005.) closely associated with the nuclear lamina, regardless of the chromosome examined. And DNA probes that preferentially stain gene-rich regions of human chromosomes produce a striking picture of the interphase nucleus that presumably reflects different average positions for active and inactive genes (Figure 4–54).


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Cell_Biology_Alberts. How is most of the chromatin in each interphase chromosome condensed when its genes are not being expressed? A powerful extension of the chromosome conformation capture method described previously (see Figure 4–48), which exploits a high-throughput DNA sequencing technology called massive parallel sequencing (see Panel 8–1, pp. 478–481), allows the connections between all of the different one-megabase (1 Mb) segments of the human genome to be mapped in human interphase chromosomes. The results reveal that most regions of our chromosomes are folded into a conformation referred to as a fractal globule: a knot-free arrangement that facilitates maximally dense packing while, at the same time, preserving the ability of the chromatin fiber to unfold and fold (Figure 4–55). Chromatin Can move to Specific Sites within the Nucleus to Alter gene Expression


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Cell_Biology_Alberts. Chromatin Can move to Specific Sites within the Nucleus to Alter gene Expression A variety of different types of experiments has led to the conclusion that the position of a gene in the interior of the nucleus changes when it becomes highly expressed. Thus, a region that becomes very actively transcribed is sometimes found to extend out of its chromosome territory, as if in an extended loop (Figure 4–56). We will see in Chapter 6 that the initiation of transcription—the first step in gene expression—requires the assembly of over 100 proteins, and it makes sense that this would be facilitated in regions of the nucleus enriched in these proteins. More generally, it is clear that the nucleus is very heterogeneous, with functionally different regions to which portions of chromosomes can move as they are subjected to different biochemical processes—such as when their gene expression changes. It is this issue that we discuss next.


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Cell_Biology_Alberts. Figure 4–55 A fractal globule model for interphase chromatin. An extension of the 3C method in Figure 4–48, called Hi-C, was used to measure the extent to which each of the three thousand 1 mb segments in the human genome was located adjacent to any other of these segments. The results support the type of model shown. In the enlarged fractal globule illustrated, a region of 5 million base pairs is seen to fold in a way that keeps regions that are neighbors along the one-dimensional DNA helix as neighbors in three dimensions; this gives rise to monochromatic blocks in this representation that are obvious both on the surface and in cross section. The fractal globule is a knot-free conformation of the DNA that permits dense packing, yet retains an ability to easily fold and unfold any genomic locus. (Adapted from E. lieberman-Aiden et al., Science 326:289–293, 2009. with permission from AAAS.)


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Cell_Biology_Alberts. E. lieberman-Aiden et al., Science 326:289–293, 2009. with permission from AAAS.) Figure 4–56 An effect of high levels of gene expression on the intranuclear location of chromatin. (A) Fluorescence micrographs of human nuclei showing how the position of a gene changes when it becomes highly transcribed. The region of the chromosome adjacent to the gene (red) is seen to leave its chromosomal territory (green) only when it is highly active. (b) Schematic representation of a large loop of chromatin that expands when the gene is on, and contracts when the gene is off. Other genes that are less actively expressed can be shown by the same methods to remain inside their chromosomal territory when transcribed. (From J.R. Chubb and w.A. bickmore, Cell 112:403–406, 2003. with permission from Elsevier.) Networks of macromolecules Form a Set of Distinct biochemical Environments inside the Nucleus


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Cell_Biology_Alberts. Networks of macromolecules Form a Set of Distinct biochemical Environments inside the Nucleus In Chapter 6, we shall describe the function of a variety of subcompartments that are present within the nucleus. The largest and most obvious of these is the nucleolus, a structure well known to microscopists even in the nineteenth century (see Figure 4–9). The nucleolus is the cell’s site of ribosome subunit formation, as well as the place where many other specialized reactions occur (see Figure 6–42): it consists of a network of RNAs and proteins concentrated around ribosomal RNA genes that are being actively transcribed. In eukaryotes, the genome contains multiple copies of the ribosomal RNA genes, and although they are typically clustered together in a single nucleolus, they are often located on several separate chromosomes.


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Cell_Biology_Alberts. A variety of less obvious organelles are also present inside the nucleus. For example, spherical structures called Cajal bodies and interchromatin granule clusters are present in most plant and animal cells (Figure 4–57). Like the nucleolus, these organelles are composed of selected protein and RNA molecules that bind together to create networks that are highly permeable to other protein and RNA molecules in the surrounding nucleoplasm.


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Cell_Biology_Alberts. Structures such as these can create distinct biochemical environments by immobilizing select groups of macromolecules, as can other networks of proteins and RNA molecules associated with nuclear pores and with the nuclear envelope. In principle, this allows other molecules that enter these spaces to be processed with great efficiency through complex reaction pathways. Highly permeable, 1 µm fibrous networks of this sort can thereby impart many of the kinetic advantages of compartmentalization (see p. 164) to reactions that take place in subregions of the nucleus (Figure 4–58A). However, unlike the membrane-bound compartments in nuclear subcompartments. The large the cytoplasm (discussed in Chapter 12), these nuclear subcompartments—lack sphere here is a Cajal body. The smaller ing a lipid bilayer membrane—can neither concentrate nor exclude specific small darker sphere is an interchromatin granule molecules. cluster, also known as a speckle (see also Figure 6–46). These “subnuclear


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Cell_Biology_Alberts. The cell has a remarkable ability to construct distinct environments to per- organelles” are from the nucleus of a form complex biochemical tasks efficiently. Those that we have mentioned in the Xenopus oocyte. (From k.E. Handwerger nucleus facilitate various aspects of gene expression, and will be further discussed and J.g. gall, Trends Cell Biol. 16:19–26, in Chapter 6. These subcompartments, including the nucleolus, appear to form 2006. with permission from Elsevier.) only as needed, and they create a high local concentration of the many different enzymes and RNA molecules needed for a particular process. In an analogous way, when DNA is damaged by irradiation, the set of enzymes needed to carry out DNA repair are observed to congregate in discrete foci inside the nucleus, creating “repair factories” (see Figure 5–52). And nuclei often contain hundreds of discrete foci representing factories for DNA or RNA synthesis (see Figure 6–47).


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Cell_Biology_Alberts. It seems likely that all of these entities make use of the type of tethering illustrated in Figure 4–58B, where long flexible lengths of polypeptide chain and/or long noncoding RNA molecules are interspersed with specific binding sites that concentrate the multiple proteins and other molecules that are needed to catalyze a particular process. Not surprisingly, tethers are similarly used to help to speed biological processes in the cytoplasm, increasing specific reaction rates there (for example, see Figure 16–18).


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Cell_Biology_Alberts. Is there also an intranuclear framework, analogous to the cytoskeleton, on which chromosomes and other components of the nucleus are organized? The nuclear matrix, or scaffold, has been defined as the insoluble material left in the nucleus after a series of biochemical extraction steps. Many of the proteins and RNA molecules that form this insoluble material are likely to be derived from the fibrous subcompartments of the nucleus just discussed, while others may be proteins that help to form the base of chromosomal loops or to attach chromosomes to other structures in the nucleus.


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Cell_Biology_Alberts. Having discussed the dynamic structure of interphase chromosomes, we now turn to mitotic chromosomes. The chromosomes from nearly all eukaryotic cells become readily visible by light microscopy during mitosis, when they coil up to form highly condensed structures. This condensation reduces the length of a typical interphase chromosome only about tenfold, but it produces a dramatic change in chromosome appearance. Figure 4–59 depicts a typical mitotic chromosome at the metaphase stage of mitosis (for the stages of mitosis, see Figure 17–3). The two DNA molecules produced by DNA replication during interphase of the cell-division cycle are separately folded to produce two sister chromosomes, or sister chromatids, held together at their centromeres, as mentioned earlier. These chromosomes are normally covered with a variety of molecules, including large amounts of RNA–protein


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Cell_Biology_Alberts. Figure 4–58 effective compartmentalization without a bilayer membrane. (A) Schematic illustration of the organization of a spherical subnuclear organelle (left) and of a postulated similarly organized subcompartment just beneath the nuclear envelope (right). In both cases, RNAs and/or proteins (gray) associate to form highly porous, gel-like structures that contain binding sites for other specific proteins and RNA molecules (colored objects). (b) How the tethering of a selected set of proteins and RNA molecules to long flexible polymer chains, as in (A), can create “staging areas” that greatly speed the rates of reactions in subcompartments of the nucleus. The reactions catalyzed will depend on the particular macromolecules that are localized by the tethering. The same strategy for accelerating complex sets of reactions is also employed in subcompartments elsewhere in the cell (see also Figure 3–78).


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Cell_Biology_Alberts. Figure 4–59 A typical mitotic chromosome at metaphase. Each sister chromatid contains one of two identical sister DNA molecules generated earlier in the cell cycle by DNA replication (see also Figure 17–21).


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Cell_Biology_Alberts. Figure 4–60 A scanning electron micrograph of a region near one end of a typical mitotic chromosome. Each knoblike projection is believed to represent the tip of a separate looped domain. Note that the two identical paired chromatids (drawn in Figure 4–59) can be clearly distinguished. (From m.P. marsden and U.k. laemmli, Cell 17:849–858, 1979. with permission from Elsevier.) complexes. Once this covering has been stripped away, each chromatid can be seen in electron micrographs to be organized into loops of chromatin emanating from a central scaffolding (Figure 4–60). Experiments using DNA hybridization to detect specific DNA sequences demonstrate that the order of visible features along a mitotic chromosome at least roughly reflects the order of genes along the DNA molecule. Mitotic chromosome condensation can thus be thought of as the final level in the hierarchy of chromosome packaging (Figure 4–61).


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Cell_Biology_Alberts. The compaction of chromosomes during mitosis is a highly organized and dynamic process that serves at least two important purposes. First, when condensation is complete (in metaphase), sister chromatids have been disentangled from each other and lie side by side. Thus, the sister chromatids can easily separate when the mitotic apparatus begins pulling them apart. Second, the compaction of chromosomes protects the relatively fragile DNA molecules from being broken as they are pulled to separate daughter cells.


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Cell_Biology_Alberts. The condensation of interphase chromosomes into mitotic chromosomes begins in early M phase, and it is intimately connected with the progression of the cell cycle. During M phase, gene expression shuts down, and specific modifications are made to histones that help to reorganize the chromatin as it compacts. Two classes of ring-shaped proteins, called cohesins and condensins, aid this compaction. How they help to produce the two separately folded chromatids of a mitotic chromosome will be discussed in Chapter 17, along with the details of the cell cycle. short region of 11 nm form of chromatin “beads-on-a-string” chromatin fber of packed 30 nm NET RESULT: EACH DNA MOLECULE HAS BEEN PACKAGED INTO A MITOTIC CHROMOSOME THAT IS 10,000-FOLD SHORTER THAN ITS FULLY EXTENDED LENGTH 0.1 µm Figure 4–61 Chromatin packing. This model shows some of the many levels of chromatin packing postulated to give rise to the highly condensed mitotic chromosome.


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Cell_Biology_Alberts. Figure 4–61 Chromatin packing. This model shows some of the many levels of chromatin packing postulated to give rise to the highly condensed mitotic chromosome. Chromosomes are generally decondensed during interphase, so that the details of their structure are difficult to visualize. Notable exceptions are the specialized lampbrush chromosomes of vertebrate oocytes and the polytene chromosomes in the giant secretory cells of insects. Studies of these two types of interphase chromosomes suggest that each long DNA molecule in a chromosome is divided into a large number of discrete domains organized as loops of chromatin that are compacted by further folding. When genes contained in a loop are expressed, the loop unfolds and allows the cell’s machinery access to the DNA.


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Cell_Biology_Alberts. Interphase chromosomes occupy discrete territories in the cell nucleus; that is, they are not extensively intertwined. Euchromatin makes up most of interphase chromosomes and, when not being transcribed, it probably exists as tightly folded fibers of compacted nucleosomes. However, euchromatin is interrupted by stretches of heterochromatin, in which the nucleosomes are subjected to additional packing that usually renders the DNA resistant to gene expression. Heterochromatin exists in several forms, some of which are found in large blocks in and around centromeres and near telomeres. But heterochromatin is also present at many other positions on chromosomes, where it can serve to help regulate developmentally important genes.


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Cell_Biology_Alberts. The interior of the nucleus is highly dynamic, with heterochromatin often positioned near the nuclear envelope and loops of chromatin moving away from their chromosome territory when genes are very highly expressed. This reflects the existence of nuclear subcompartments, where different sets of biochemical reactions are facilitated by an increased concentration of selected proteins and RNAs. The components involved in forming a subcompartment can self-assemble into discrete organelles such as nucleoli or Cajal bodies; they can also be tethered to fixed structures such as the nuclear envelope.


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Cell_Biology_Alberts. During mitosis, gene expression shuts down and all chromosomes adopt a highly condensed conformation in a process that begins early in M phase to package the two DNA molecules of each replicated chromosome as two separately folded chromatids. The condensation is accompanied by histone modifications that facilitate chromatin packing, but satisfactory completion of this orderly process, which reduces the end-to-end distance of each DNA molecule from its interphase length by an additional factor of ten, requires additional proteins.


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Cell_Biology_Alberts. In this final section of the chapter, we provide an overview of some of the ways that genes and genomes have evolved over time to produce the vast diversity of modern-day life-forms on our planet. The sequencing of the genomes of thousands of organisms is revolutionizing our view of the process of evolution, uncovering an astonishing wealth of information about not only family relationships among organisms, but also about the molecular mechanisms by which evolution has proceeded.


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Cell_Biology_Alberts. It is perhaps not surprising that genes with similar functions can be found in a diverse range of living things. But the great revelation of the past 30 years has been the extent to which the actual nucleotide sequences of many genes have been conserved. Homologous genes—that is, genes that are similar in both their nucleotide sequence and function because of a common ancestry—can often be recognized across vast phylogenetic distances. Unmistakable homologs of many human genes are present in organisms as diverse as nematode worms, fruit flies, yeasts, and even bacteria. In many cases, the resemblance is so close that, for example, the protein-coding portion of a yeast gene can be substituted with its human homolog—even though humans and yeast are separated by more than a billion years of evolutionary history.


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Cell_Biology_Alberts. As emphasized in Chapter 3, the recognition of sequence similarity has become a major tool for inferring gene and protein function. Although a sequence match does not guarantee similarity in function, it has proved to be an excellent clue. Thus, it is often possible to predict the function of genes in humans for which no biochemical or genetic information is available simply by comparing their nucleotide sequences with the sequences of genes that have been characterized in other more readily studied organisms. In general, the sequences of individual genes are much more tightly conserved than is overall genome structure. Features of genome organization such as genome size, number of chromosomes, order of genes along chromosomes, abundance and size of introns, and amount of repetitive DNA are found to differ greatly when comparing distant organisms, as does the number of genes that each organism contains.


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Cell_Biology_Alberts. A first obstacle in interpreting the sequence of the 3.2 billion nucleotide pairs in the human genome is the fact that much of it is probably functionally unimportant. The regions of the genome that code for the amino acid sequences of proteins (the exons) are typically found in short segments (average size about 145 nucleotide pairs), small islands in a sea of DNA whose exact nucleotide sequence is thought to be mostly of little consequence. This arrangement makes it difficult to identify all the exons in a stretch of DNA, and it is often hard too to determine exactly where a gene begins and ends.


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Cell_Biology_Alberts. One very important approach to deciphering our genome is to search for DNA sequences that are closely similar between different species, on the principle that DNA sequences that have a function are much more likely to be conserved than those without a function. For example, humans and mice are thought to have diverged from a common mammalian ancestor about 80 × 106 years ago, which is long enough for the majority of nucleotides in their genomes to have been changed by random mutational events. Consequently, the only regions that will have remained closely similar in the two genomes are those in which mutations would have impaired function and put the animals carrying them at a disadvantage, resulting in their elimination from the population by natural selection. Such closely similar pieces of DNA sequence are known as conserved regions. In addition to revealing those DNA sequences that encode functionally important exons and RNA molecules, these conserved regions will include


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Cell_Biology_Alberts. pieces of DNA sequence are known as conserved regions. In addition to revealing those DNA sequences that encode functionally important exons and RNA molecules, these conserved regions will include regulatory DNA sequences as well as DNA sequences with functions that are not yet known. In contrast, most nonconserved regions will reflect DNA whose sequence is much less likely to be critical for function.


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Cell_Biology_Alberts. The power of this method can be increased by including in such comparisons the genomes of large numbers of species whose genomes have been sequenced, such as rat, chicken, fish, dog, and chimpanzee, as well as mouse and human. By revealing in this way the results of a very long natural “experiment,” lasting for hundreds of millions of years, such comparative DNA sequencing studies have highlighted the most interesting regions in our genome. The comparisons reveal that roughly 5% of the human genome consists of “multispecies conserved sequences.” To our great surprise, only about one-third of these sequences code for proteins (see Table 4–1, p. 184). Many of the remaining conserved sequences consist of DNA containing clusters of protein-binding sites that are involved in gene regulation, while others produce RNA molecules that are not translated into protein but are important for other known purposes. But, even in the most intensively studied species, the function of the majority of


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Cell_Biology_Alberts. while others produce RNA molecules that are not translated into protein but are important for other known purposes. But, even in the most intensively studied species, the function of the majority of these highly conserved sequences remains unknown. This remarkable discovery has led scientists to conclude that we understand much less about the cell biology of vertebrates than we had thought. Certainly, there are enormous opportunities for new discoveries, and we should expect many more surprises ahead.


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Cell_Biology_Alberts. genome Alterations Are Caused by Failures of the Normal mechanisms for Copying and maintaining DNA, as well as by Transposable DNA Elements


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Cell_Biology_Alberts. Evolution depends on accidents and mistakes followed by nonrandom survival. Most of the genetic changes that occur result simply from failures in the normal mechanisms by which genomes are copied or repaired when damaged, although the movement of transposable DNA elements (discussed below) also plays an important part. As we will explain in Chapter 5, the mechanisms that maintain DNA sequences are remarkably precise—but they are not perfect. DNA sequences are inherited with such extraordinary fidelity that typically, along a given line of descent, only about one nucleotide pair in a thousand is randomly changed in the germ line every million years. Even so, in a population of 10,000 diploid individuals, every possible nucleotide substitution will have been “tried out” on about 20 occasions in the course of a million years—a short span of time in relation to the evolution of species.


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Cell_Biology_Alberts. Errors in DNA replication, DNA recombination, or DNA repair can lead either to simple local changes in DNA sequence—so-called point mutations such as the substitution of one base pair for another—or to large-scale genome rearrangements such as deletions, duplications, inversions, and translocations of DNA from one chromosome to another. In addition to these failures of the genetic machinery, genomes contain mobile DNA elements that are an important source of genomic change (see Table 5–3, p. 267). These transposable DNA elements (transposons) are parasitic DNA sequences that can spread within the genomes they colonize. In the process, they often disrupt the function or alter the regulation of existing genes. On occasion, they have created altogether novel genes through fusions between transposon sequences and segments of existing genes. Over long periods of evolutionary time, DNA transposition events have profoundly affected genomes, so much so that nearly half of the DNA in the human


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Cell_Biology_Alberts. sequences and segments of existing genes. Over long periods of evolutionary time, DNA transposition events have profoundly affected genomes, so much so that nearly half of the DNA in the human genome consists of recognizable relics of past transposition events (Figure 4–62). Even more of our genome is thought to have been derived from transpositions that occurred so long ago (>108 years) that the sequences can no longer be traced to transposons.


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Cell_Biology_Alberts. The genome Sequences of Two Species Differ in Proportion to the length of Time Since They Have Separately Evolved The differences between the genomes of species alive today have accumulated over more than 3 billion years. Although we lack a direct record of changes over time, scientists can reconstruct the process of genome evolution from detailed comparisons of the genomes of contemporary organisms. The basic organizing framework for comparative genomics is the phylogenetic tree. A simple example is the tree describing the divergence of humans from the great apes (Figure 4–63). The primary support for this tree comes from comparisons of gene or protein sequences. For example, comparisons between the sequences of human genes or proteins and those of the great apes typically reveal the fewest differences between human and chimpanzee and the most between human and orangutan.


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Cell_Biology_Alberts. For closely related organisms such as humans and chimpanzees, it is relatively easy to reconstruct the gene sequences of the extinct, last common ancestor of the two species (Figure 4–64). The close similarity between human and chimpanzee genes is mainly due to the short time that has been available for the accumulation of mutations in the two diverging lineages, rather than to functional constraints TRANSPOSONS nonrepetitive DNA that is


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Cell_Biology_Alberts. Figure 4–62 A representation of the nucleotide sequence content of the sequenced human genome. The lINEs (long interspersed nuclear elements), SINEs (short interspersed nuclear elements), retroviral-like elements, and DNA-only transposons are mobile genetic elements that have multiplied in our genome by replicating themselves and inserting the new copies in different positions. These mobile genetic elements are discussed in Chapter 5 (see Table 5–3, p. 267). Simple sequence repeats are short nucleotide sequences (less than 14 nucleotide pairs) that are repeated again and again for long stretches. Segmental duplications are large blocks of DNA sequence (1000–200,000 nucleotide pairs) that are present at two or more locations in the genome. The most highly repeated blocks of DNA in heterochromatin have not yet been completely sequenced; therefore about 10% of human DNA sequences are not represented in this diagram. (Data courtesy of E. margulies.) showing the relationship between 1.5


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Cell_Biology_Alberts. have not yet been completely sequenced; therefore about 10% of human DNA sequences are not represented in this diagram. (Data courtesy of E. margulies.) showing the relationship between 1.5 humans and the great apes based on nucleotide sequence data. As indicated, the sequences of the genomes of all four millions of years before present species are estimated to differ from the sequence of the genome of their last common ancestor by a little over 1.5%. because changes occur independently on both diverging lineages, pairwise comparisons reveal twice the sequence divergence from the last common ancestor. For example, human–orangutan comparisons typically show sequence 1.0 0.5 0.0 divergences of a little over 3%, while that have kept the sequences the same. Evidence for this view comes from the observation that the human and chimpanzee genomes are nearly identical even where there is no functional constraint on the nucleotide sequence—such as in the third position of “synonymous” codons


Cell_Biology_Alberts

Cell_Biology_Alberts. the observation that the human and chimpanzee genomes are nearly identical even where there is no functional constraint on the nucleotide sequence—such as in the third position of “synonymous” codons (codons specifying the same amino acid but differing in their third nucleotide).


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Cell_Biology_Alberts. For much less closely related organisms, such as humans and chickens (which have evolved separately for about 300 million years), the sequence conservation found in genes is almost entirely due to purifying selection (that is, selection that eliminates individuals carrying mutations that interfere with important genetic functions), rather than to an inadequate time for mutations to occur. Phylogenetic Trees Constructed from a Comparison of DNA Sequences Trace the Relationships of All Organisms Phylogenetic trees based on molecular sequence data can be compared with the fossil record, and we get our best view of evolution by integrating the two approaches. The fossil record remains essential as a source of absolute dates, human–chimpanzee comparisons show divergences of approximately 1.2%. (modified from F.C. Chen and w.H. li, Am. J. Hum. Genet. 68:444–456, 2001.)


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Cell_Biology_Alberts. Figure 4–64 Tracing the ancestral sequence from a sequence comparison of the coding regions of human and chimpanzee leptin genes. Reading left to right and top to bottom, a continuous gene is illustrated. leptin is a hormone that regulates food intake and energy utilization in response to the adequacy of 300-nucleotide segment of a leptin-coding fat reserves. As indicated by the codons boxed in green, only 5 nucleotides (of 120 441 total) differ between the two species. moreover, in only one of the five positions does the difference in nucleotide lead to a difference in the encoded amino acid. For each of the five variant nucleotide positions, the corresponding sequence in the gorilla is also indicated. In two cases, sequence, while in three cases it agrees with the chimpanzee sequence.


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Cell_Biology_Alberts. the gorilla sequence agrees with the human what was the sequence of the leptin gene in the last common ancestor? The most economical assumption is that evolution minimum number of mutations consistent with the data. Thus, it seems likely that chimp TACCAACAGATCCTCACCAGTATGCCTTCCAGAAACATGATCCAAATATCCAACGACCTG protein Y Q Q I L T S M P S R N M I Q I S N D L the leptin sequence of the last common has followed a pathway requiring the ancestor was the same as the human and chimpanzee sequences when they agree; when they disagree, the gorilla sequence would be used as a tiebreaker. For convenience, only the first 300 nucleotides of the leptin-coding sequences are given. The remaining 141 are identical between humans and chimpanzees.


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Cell_Biology_Alberts. Figure 4–65 The very different rates of evolution of exons and introns, as illustrated by comparing a portion of the mouse and human leptin genes. Positions where the sequences differ by a single nucleotide substitution are boxed in green, and positions that differ by the addition or deletion of nucleotides are boxed in yellow. Note that, thanks to purifying selection, the coding sequence of the exon is much more conserved than is the adjacent intron sequence. based on radioisotope decay in the rock formations in which fossils are found. Because the fossil record has many gaps, however, precise divergence times between species are difficult to establish, even for species that leave good fossils with distinctive morphology.


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Cell_Biology_Alberts. Phylogenetic trees whose timing has been calibrated according to the fossil record suggest that changes in the sequences of particular genes or proteins tend to occur at a nearly constant rate, although rates that differ from the norm by as much as twofold are observed in particular lineages. This provides us with a molecular clock for evolution—or rather a set of molecular clocks corresponding to different categories of DNA sequence. As in the example in Figure 4–65, the clock runs most rapidly and regularly in sequences that are not subject to purifying selection. These include portions of introns that lack splicing or regulatory signals, the third position in synonymous codons, and genes that have been irreversibly inactivated by mutation (the so-called pseudogenes). The clock runs most slowly for sequences that are subject to strong functional constraints—for example, the amino acid sequences of proteins that engage in specific interactions with large numbers of other proteins and


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Cell_Biology_Alberts. slowly for sequences that are subject to strong functional constraints—for example, the amino acid sequences of proteins that engage in specific interactions with large numbers of other proteins and whose structure is therefore highly constrained, or the nucleotide sequences that encode the RNA subunits of the ribosome, on which all protein synthesis depends.


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Cell_Biology_Alberts. Occasionally, rapid change is seen in a previously highly conserved sequence. As discussed later in this chapter, such episodes are especially interesting because they are thought to reflect periods of strong positive selection for mutations that have conferred a selective advantage in the particular lineage where the rapid change occurred. The pace at which molecular clocks run during evolution is determined not only by the degree of purifying selection, but also by the mutation rate. Most notably, in animals, although not in plants, clocks based on functionally unconstrained mitochondrial DNA sequences run much faster than clocks based on functionally unconstrained nuclear sequences, because the mutation rate in animal mitochondria is exceptionally high.


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Cell_Biology_Alberts. Categories of DNA for which the clock runs fast are most informative for recent evolutionary events; the mitochondrial DNA clock has been used, for example, to chronicle the divergence of the Neanderthal lineage from that of modern Homo sapiens. To study more ancient evolutionary events, one must examine DNA for which the clock runs more slowly; thus the divergence of the major branches of the tree of life—bacteria, archaea, and eukaryotes—has been deduced from study of the sequences specifying ribosomal RNA.


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Cell_Biology_Alberts. In general, molecular clocks, appropriately chosen, have a finer time resolution than the fossil record, and they are a more reliable guide to the detailed structure of phylogenetic trees than are classical methods of tree construction, which are based on family resemblances in anatomy and embryonic development. For example, the precise family tree of great apes and humans was not settled until sufficient molecular sequence data accumulated in the 1980s to produce the pedigree shown previously in Figure 4–63. And with huge amounts of DNA sequence now determined from a wide variety of mammals, much better estimates of our relationship to them are being obtained (Figure 4–66). A Comparison of Human and mouse Chromosomes Shows How the Structures of genomes Diverge


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Cell_Biology_Alberts. A Comparison of Human and mouse Chromosomes Shows How the Structures of genomes Diverge As would be expected, the human and chimpanzee genomes are much more alike than are the human and mouse genomes, even though all three genomes are roughly the same size and contain nearly identical sets of genes. Mouse and human lineages have had approximately 80 million years to diverge through accumulated mutations, versus 6 million years for humans and chimpanzees. In addition, as indicated in Figure 4–66, rodent lineages (represented by the rat and the mouse) have unusually fast molecular clocks, and have diverged from the human lineage more rapidly than otherwise expected.


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Cell_Biology_Alberts. While the way that the genome is organized into chromosomes is almost identical between humans and chimpanzees, this organization has diverged greatly between humans and mice. According to rough estimates, a total of about 180 breakage-and-rejoining events have occurred in the human and mouse lineages since these two species last shared a common ancestor. In the process, although the number of chromosomes is similar in the two species (23 per haploid genome in the human versus 20 in the mouse), their overall structures differ greatly. Nonetheless, even after the extensive genomic shuffling, there are many large blocks of DNA in which the gene order is the same in the human and the mouse. These stretches of conserved gene order in chromosomes are referred to as regions of synteny. Figure 4–67 illustrates how segments of the different mouse chromosomes map onto the human chromosome set. For much more distantly related vertebrates, such as chicken and human, the number of


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Cell_Biology_Alberts. Figure 4–67 illustrates how segments of the different mouse chromosomes map onto the human chromosome set. For much more distantly related vertebrates, such as chicken and human, the number of breakage-and-rejoining events has been much greater and the regions of synteny are much shorter; in addition, they are often hard to discern because of the divergence of the DNA sequences that they contain.


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Cell_Biology_Alberts. An unexpected conclusion from a detailed comparison of the complete mouse and human genome sequences, confirmed by subsequent comparisons between the genomes of other vertebrates, is that small blocks of DNA sequence are being deleted from and added to genomes at a surprisingly rapid rate. Thus, if we assume that our common ancestor had a genome of human size (about 3.2 billion nucleotide pairs), mice would have lost a total of about 45% of that genome from accumulated deletions during the past 80 million years, while humans would have lost about 25%. However, substantial sequence gains from many small chromosome duplications and from the multiplication of transposons have compensated for these deletions. As a result, our genome size is thought to be practically unchanged from that of the last common ancestor of humans and mice, while the mouse genome is smaller by only about 0.3 billion nucleotides.


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Cell_Biology_Alberts. Figure 4–66 A phylogenetic tree showing the evolutionary relationships of some present-day mammals. The length of each line is proportional to the number of “neutral substitutions”—that is, nucleotide changes at sites where there is assumed to be no purifying selection. (Adapted from g.m. Cooper et al., Genome Res. 15:901–913, 2005. with permission from Cold Spring Harbor laboratory Press.) Good evidence for the loss of DNA sequences in small blocks during evolution can be obtained from a detailed comparison of regions of synteny in the human and mouse genomes. The comparative shrinkage of the mouse genome can be clearly seen from such comparisons, with the net loss of sequences scattered throughout the long stretches of DNA that are otherwise homologous (Figure 4–68).


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Cell_Biology_Alberts. DNA is added to genomes both by the spontaneous duplication of chromosomal segments that are typically tens of thousands of nucleotide pairs long (as will be discussed shortly) and by insertion of new copies of active transposons. Most transposition events are duplicative, because the original copy of the transposon stays where it was when a copy inserts at the new site; see, for example, Figure 5–63. Comparison of the DNA sequences derived from transposons in the human and the mouse readily reveals some of the sequence additions (Figure 4–69). It remains a mystery why all mammals have maintained genome sizes of roughly 3 billion nucleotide pairs that contain nearly identical sets of genes, even though only approximately 150 million nucleotide pairs appear to be under sequence-specific functional constraints. The Size of a vertebrate genome Reflects the Relative Rates of DNA Addition and DNA loss in a lineage


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Cell_Biology_Alberts. The Size of a vertebrate genome Reflects the Relative Rates of DNA Addition and DNA loss in a lineage In more distantly related vertebrates, genome size can vary considerably, apparently without a drastic effect on the organism or its number of genes. Thus, the chicken genome, at one billion nucleotide pairs, is only about one-third the size 200,000 bases Figure 4–67 Synteny between human and mouse chromosomes. In this diagram, the human chromosome set is shown above, with each part of each chromosome colored according to the mouse chromosome with which it is syntenic. The color coding used for each mouse chromosome is shown below. Heterochromatic highly repetitive regions (such as centromeres) that are difficult to sequence cannot be mapped in this way; these are colored black. (Adapted from E.E. Eichler and D. Sankoff, Science 301:793–797, 2003. with permission from AAAS.) Figure 4–68 Comparison of a syntenic portion of mouse and human genomes.


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Cell_Biology_Alberts. About 90% of the two genomes can be aligned in this way. Note that while there is an identical order of the matched index sequences (red marks), there has been a net loss of DNA in the mouse lineage that is interspersed throughout the entire region. This type of net loss is typical for all such regions, and it accounts for the fact that the mouse genome contains 14% less DNA than does the human genome. (Adapted from mouse genome Sequencing Consortium, Nature 420:520–562, 2002. with permission from macmillan Publishers ltd.) 10,000 nucleotide pairs of the mammalian genome. An extreme example is the puffer fish, Fugu rubripes (Figure 4–70), which has a tiny genome for a vertebrate (0.4 billion nucleotide pairs compared to 1 billion or more for many other fish). The small size of the Fugu genome is largely due to the small size of its introns. Specifically, Fugu introns, as well as other noncoding segments of the Fugu genome, lack the repetitive DNA that makes up a large portion of the


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Cell_Biology_Alberts. is largely due to the small size of its introns. Specifically, Fugu introns, as well as other noncoding segments of the Fugu genome, lack the repetitive DNA that makes up a large portion of the genomes of most well-studied vertebrates. Nevertheless, the positions of the Fugu introns between the exons of each gene are almost the same as in mammalian genomes (Figure 4–71).


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Cell_Biology_Alberts. While initially a mystery, we now have a simple explanation for such large differences in genome size between similar organisms: because all vertebrates experience a continuous process of DNA loss and DNA addition, the size of a genome merely depends on the balance between these opposing processes acting over millions of years. Suppose, for example, that in the lineage leading to Fugu, the rate of DNA addition happened to slow greatly. Over long periods of time, this would result in a major “cleansing” from this fish genome of those DNA sequences whose loss could be tolerated. The result is an unusually compact genome, relatively free of junk and clutter, but retaining through purifying selection the vertebrate DNA sequences that are functionally important. This makes Fugu, with its 400 million nucleotide pairs of DNA, a valuable resource for genome research aimed at understanding humans. we Can Infer the Sequence of Some Ancient genomes


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Cell_Biology_Alberts. The genomes of ancestral organisms can be inferred, but most can never be directly observed. DNA is very stable compared with most organic molecules, but it is not perfectly stable, and its progressive degradation, even under the best circumstances, means that it is virtually impossible to extract sequence information from fossils that are more than a million years old. Although a modern organism such as the horseshoe crab looks remarkably similar to fossil ancestors that lived 200 million years ago, there is every reason to believe that the horse-shoe-crab genome has been changing during all that time in much the same way as in other evolutionary lineages, and at a similar rate. Selection must have maintained key functional properties of the horseshoe-crab genome to account for the morphological stability of the lineage. However, comparisons between different present-day organisms show that the fraction of the genome subject to purifying selection is small; hence, it is fair to