Stopping for FISH and Chips along the Chromatin Fiber Superhighway

Molecular Cell 844

et al. and implicates the balanace between JNKs in the regulation of apoptosis and tumorigenesis. Ze’ev Ronai Department of Oncological Sciences Mount Sinai School of Medicine New York, New York 10029 Selected Reading Fuchs, S.Y., Dolan, L., Davis, R.J., and Ronai, Z. (1996). Oncogene 13, 1531–1535. Fuchs, S.Y., Adler, V., Buschman, T., Yin, Z., Wu, X., Jones, S.N., and Ronai, Z. (1998). Genes Dev. 12, 2658–2663. Gao, M., Labuda, T., Xia, Y., Gallangher, E., Fang, D., Liu, Y.C., and Karin, M. (2004). Science, in press Published online September 9, 2004. 10.1126/science.1099414. Figure 1. Functional Differences among JNK1 and JNK2 Under normal, nonstressed growth conditions, c-Jun is predominantly bound to inactive JNK2, resulting in the targeting of c-Jun for degradation, thereby limiting the availability of c-Jun. Upon exposure to stress or DNA damage, activated JNK1 exhibits greater affinity for c-Jun, thereby displacing JNK2 and resulting in the phosphorylation of c-Jun, rendering it stable and transcriptionally active. Subsequently, JNK2 is expected to capture dephosphorylated c-Jun as a means of limiting the duration of c-Jun signaling.

et al., 2002). Further, JNK1 null mice are prone to development of skin tumors, which is not the case for JNK2 null animals (She et al., 2002). Since JNK1 null fibroblasts exhibit lower levels of Jun expression and activity (the current study) and since c-Jun can efficiently induce apoptosis, it is possible that skin tumor development is prompted by failure of mutant cells to undergo apoptosis. The latter substantiates the findings of Sabapathy

Stopping for FISH and Chips along the Chromatin Fiber Superhighway

Gilbert et al. (2004) report in a recent issue of Cell on the analysis of chromatin fiber structure across the human genome. They show that compact chromatin fibers are composed of heterochromatin but also contain some active genes, while open chromatin fibers correlate with regions of highest gene density, but not with gene expression. Heitz is credited with making the first (in 1928) distinction between euchromatin (less compact) and heterochromatin (highly compacted) based upon their microscopic appearance in interphase nuclei (Heitz, 1928). Subsequently, these global features were correlated with regions of active or inactive genes, respectively (Littau et al., 1964). However, the problem with generalizations is that it’s often only a matter of time before advances in technology prove them to be not totally correct. The

Hirosumi, J., Tuncman, G., Chang, L., Gorgun, C.Z., Uysal, K.T., Maeda, K., Karin, M., and Hotamisligil, G.S. (2002). Nature 420, 333–336. Kallunki, T., Su, B., Tsigelny, I., Sluss, H.K., Derijard, B., Moore, G., Davis, R., and Karin, M. (1994). Genes Dev. 8, 2996–3007. Morrison, D.K., and Davis, R.J. (2003). Annu. Rev. Cell Dev. Biol. 19, 91–118. Musti, A.M., Treier, M., and Bohmann, D. (1997). Science 275, 400–402. Nateri, A.S., Riera-Sans, L., Da Costa, C., and Behrens, A. (2004). Science 303, 1374–1378. Sabapathy, K., Hochedlinger, K., Nam, S.Y., Bauer, A., Karin, M., and Wagner, E.F. (2004). Mol. Cell 15, 713–725. She, Q.B., Chen, N., Bode, A.M., Flavell, R.A., and Dong, Z. (2002). Cancer Res. 62, 1343–1348. Treier, M., Staszewski, L.M., and Bohmann, D. (1994). Cell 78, 787–798. Zhu, X.H., Nguyen, H., Halicka, H.D., Traganos, F., and Koff, A. (2004). Mol. Cell. Biol. 13, 6058–6066.

characteristics of these two classes of chromatin have been the subject of much interest in recent years based on the identification of specific histone modifications that are associated with “active” or “inactive” chromatin, collectively referred to as the “histone code” (for a review see Fischle et al., 2003). Chromatin structure represents an important mechanism by which genes can be regulated by providing the proper environment and binding sites for regulatory proteins to act. This is important for establishing and maintaining the proper gene expression program for a particular cell type, and also alterations in chromatin structure can result in different disease phenotypes, including cancer. Most notably, aberrant expression of polycomb group proteins have been shown to result in changes in chromatin structure and the gene expression program in breast and prostate cancer cells (Kleer et al., 2003; Varambally et al., 2002). Numerous studies have examined the organization of chromatin fibers in the active and inactive state with regard to specific genes (for a review see Spector, 2003). However, global chromatin fiber structure has never previously been investigated at the genomic level. In a recent issue of Cell, Bickmore’s group fractionated human lymphoblastoid chromatin into open, bulk, and closed chromatin fibers and assayed the distribution of these

Previews 845

fibers by hybridization to metaphase chromosomes and to genomic microarrays with respect to gene density and activity (Gilbert et al., 2004). Chromatin was digested to yield a fragment size of 10–30 kb average length and was sedimented through an isokinetic sucrose gradient. Each fraction was subsequently separated by size into open, bulk, and compact chromatin fractions. In situ hybridization to metaphase chromosomes with probes derived from compact chromatin resulted in hybridization to G bands that are regions of low gene density. Based upon this the authors concluded that such euchromatic regions of the human genome with a very low gene density are packaged into chromatin fibers with a similar level of compaction as heterochromatin. Consistent with this result, Fakan and colleagues have previously shown by immunoelectron microscopy that polycomb group proteins are associated with some euchromatic regions of the human genome (Cmarko et al., 2003). Based upon this they have suggested that active and PcG-silenced loci occur within the same spatially limited nuclear domain. To identify sequences enriched in open chromatin, probes were derived from open chromatin fragments, and their hybridization pattern to metaphase chromosomes was compared to probes derived from bulk chromatin. Not surprisingly, open chromatin probes hybridized to the most gene-rich chromosomes 16, 17, 19, and 22, and these same probes poorly hybridized to genepoor chromosomes 13 and 18. To assess the hybridization of these probes to the human genome at higher resolution and to directly relate the findings to genomic sequence, the probes were hybridized to genomic DNA microarrays. Interestingly, clones with enriched hybridization to open chromatin tended to be clustered in contiguous regions, as did clones from compact chromatin. Transitions between open and closed chromatin regions were sharp, consistent with the possibility that boundaries may exist between these regions. More interestingly, Bickmore’s group did not observe a tight correlation between gene expression and enrichment of open chromatin fibers (Gilbert et al., 2004). For example, only one of eight genes in the 22q11.21 region (UFD1L) is transcriptionally active and, surprisingly, the inactive genes in this region were also enriched in open chromatin fibers. Since genes can be found in open chromatin in the absence of transcription and transcription can occur in regions of compact chromatin (i.e., PITPNB in 22q12.1), the authors suggest that open chromatin is not an absolute requirement for transcription and it is not transcription per se that is necessary to open the chromatin. In light of these findings it would be interesting to examine the epigenetic marks associated with these genes in order to determine the molecular basis for the observed results and whether an inactive gene that is present in open chromatin is modified in a similar or different way than an inactive gene that is present in compact chromatin (i.e., facultative versus constitutive heterochromatin). Future studies examining the compaction level versus the transcriptional status, using expression microarrays, at specific cell cycle phases should provide a more complete understanding of the observed findings. Of particular interest in this regard is whether inactive genes within regions of open

chromatin are poised for activity in subsequent cell cycle phases. In a recent study examining replication timing in lymphoblastoid cells at 1 Mb resolution a positive correlation was found between DNA replication timing and a range of genome parameters including GC content, gene density, and transcriptional activity (Woodfine et al., 2004). While Bickmore and colleagues find an overall correspondence between replication timing and chromatin structure at 1 Mb resolution, at high resolution this breaks down as many regions depleted in open chromatin were found to replicate early (Gilbert et al., 2004). To bring the story back to the native context of an interphase nucleus the authors asked whether regions enriched in open chromatin fibers were cytologically more decondensed than regions of the same chromosome that are depleted of open chromatin fibers. To do so the interphase distance between probes for two regions of 11p was examined. Probes used for 11p15.5 were enriched in open chromatin while those used for 11p14.1-p13 were depleted of open chromatin. When the mean square interphase distance between signals from pairs of probes was examined, a linear relationship was observed to genomic separations of 0.25–2.0 Mb. However, the slopes of the lines indicated that 11p15.5 is less cytologically condensed than 11p14.1-p13. Support for this type of chromatin organization exists as numerous extremely active regions of chromatin have previously been shown to be highly decondensed and in some cases to loop out and reside outside of the bulk region of their interphase chromosome territory (for a review see Spector, 2003). In summary, while human nature often pushes us to think of situations in “black and white” this is usually not the way they are; it is the levels of gray that show us that there are variations on a theme. In the present study, Bickmore and colleagues reveal a more interesting organization of the genome than the simplistic bimodal separation into euchromatin and heterochromatin. They show that the ability of a gene to be activated is not lost because it is packaged more compactly than its neighbors and conversely an inactive gene residing among active genes in open chromatin can remain silenced. Obviously there are other “factors” that come into play. It is left to future studies to unravel this organization and determine how it is regulated and how it changes in response to genomic insults that lead to diseases such as cancer. David L. Spector Cold Spring Harbor Laboratory Cold Spring Harbor, New York 11724 Selected Reading Cmarko, D., Verschure, P.J., Otte, A.P., Van Driel, R., and Fakan, S. (2003). J. Cell Sci. 116, 335–343. Fischle, W., Wang, Y., and Allis, C.D. (2003). Curr. Opin. Cell Biol. 15, 172–183. Gilbert, N., Boyle, S., Fiegler, H., Woodfine, K., Carter, N.P., and Bickmore, W.A. (2004). Cell 118, 555–566. Heitz, E. (1928). Jahrb. Wiss Bot. 69, 762–818. Kleer, C.G., Cao, Q., Varambally, S., Shen, R., Ota, I., Tomlins, S.A.,

Molecular Cell 846

Ghosh, D., Sewalt, R.G., Otte, A.P., Hayes, D.F., et al. (2003). Proc. Natl. Acad. Sci. USA 100, 11606–11611. Littau, V.C., Allfrey, V.G., Frenster, J.H., and Mirsky, A.E. (1964). Proc. Natl. Acad. Sci. USA 52, 93–100. Spector, D.L. (2003). Annu. Rev. Biochem. 72, 573–608.

Synaptic Complex Revisited: A Homologous Recombinase Flips and Switches Bases

While it is still unclear how RecA and its eukaryotic homologs conduct genome-wide homology searches, Radding and colleagues report in this issue of Molecular Cell (Folta-Stogniew et al., 2004) that the latter stages of homologous recognition or alignment involve base flipping (localized melting) and switching (annealing) at A:T rich regions. The key and unique problem in the biochemistry of homologous recombination is how the proteins involved pair any two arbitrary but homologous DNAs. In all current models, a homologous recombination protein loads onto a single-strand DNA (ssDNA), forms a presynaptic filament, and scans a target duplex. Once homology is recognized, a synaptic (pairing) complex consisting of three strands encased in a protein filament is formed. Eventually, DNA strands are exchanged, one of the original duplex strands is released, and a new duplex (heteroduplex) is created. RecA homologs exist in all three domains of life: Bacteria, Archaea (RadA), and Eukaryota (Rad51 and Dmc1). Still, most of our understanding of the mechanism of homologous recognition stems from studies of RecA. The first two DNA pairing activities identified for RecA were its ability to catalyze the renaturation of singlestrand DNAs and the pairing of single-stranded DNA with a superhelical duplex DNA. How these two pairing activities relate to each other and to the search for homology has been a major pursuit of recombination biochemists for the last 25 years. In the context of bacterial conjugation, the RecA-ssDNA filament can search targets in the bacterial genome at a rate of at least 107 bp in about 1 hr. This implies 1000 steps (discrete on and off events) in 1 s or a first-order rate constant of 103 s⫺1. In mammalian meiosis, similar rates apply (CameriniOtero and Hsieh, 1993). Given that the search carried out by RecA can take place in the absence of an expenditure of energy, these earliest contacts by necessity must involve soft or weak interactions, and the initial search cannot entail an energy cost greater than that of unstacking bases and breaking hydrogen bonds in regions longer than a couple of base pairs. Thus, it is useful to think of the kinetic scheme for synapsis as consisting of several steps: one reversible early step, with fast on and off rates, and one or more much slower steps. One elegant way to avoid costly energy expenditures during the search is for the RecA-ssDNA filament to

Varambally, S., Dhanasekaran, S.M., Zhou, M., Barrette, T.R., Kumar-Sinha, C., Sanda, M.G., Ghosh, D., Pienta, K.J., Sewalt, R.G., Otte, A.P., et al. (2002). Nature 419, 624–629. Woodfine, K., Fiegler, H., Beare, D.M., Collins, J.E., McCann, O.T., Young, B.D., Debernardi, S., Mott, R., Dunham, I., and Carter, N.P. (2004). Hum. Mol. Genet. 13, 191–202.

recognize homology on a minimally perturbed duplex. The late Paul Howard-Flanders proposed a triple helix as either a transient, or even stable, intermediate in the reaction (Howard-Flanders et al., 1984). However, all recent efforts have failed to detect such a structure as a stable intermediate. Instead, several groups have described a stable synaptic complex consisting of three strands and RecA, in which strand exchange has already taken place (Roca and Cox, 1997 and references in Folta-Stogniew et al., 2004). In this complex the incoming ssDNA is part of the new duplex and the leaving strand has not yet been released. Leaving aside such an early triplex, one can jump forward and ask what are the steps leading to such a poststrand exchange intermediate. One can envision several slower conformational changes, such as homology recognition via base flipping (melting) and switching (annealing). Charles Radding and colleagues (Folta-Stogniew et al., 2004) employ a biophysical approach to study these slower events (⬍1 s⫺1), taking advantage of the fact that 2-aminopurine, a fluorescent analog of adenine, can form a stable Watson-Crick-like base pair with thymine (2AP:T). Thus, a decrease or increase in fluorescence serves as a sensitive indication of formation or disruption of 2AP:T. Analysis of fluorescence resonance energy transfer (FRET) between fluorescein and rhodamine, placed on the ends of interacting DNA molecules, allows monitoring of both synaptic complex formation and outgoing strand dissociation. In a cleverly designed experiment, Folta-Stogniew et al. (2004) use stop-flow analyses of 2AP fluorescence, along with FRET, to follow all the stages of strand exchange in the presence of ATP and an ATP regenerating system: disruption of base pairing within the original duplex, the establishment of hydrogen bonds between the incoming strand and its complement, formation of the synaptic complex, and strand dissociation. Surprisingly, the rate of 2AP:T formation was equal to that of 2AP:T dissociation, and both were very close to the rate of the assimilation of the incoming strand. The dissociation of the outgoing strand was much slower. The results indicate that the disruption of old A:T base pairs and the formation of new ones occurs simultaneously and that the early synaptic complex is a metastable tristranded structure in which the complementary strand participates in Watson-Crick base pairing with both incoming and outgoing strands. The observation that the formation of the synaptic complex is faster for the substrate with higher A-T content lead Folta-Stogniew et al. to propose that the dynamic structure of the double helix (DNA “breathing”) significantly contributes to the recognition of homology. The extended conformation of single-stranded DNA in the presynaptic filament is proposed to allow rotational mobility of the bases making them available for interactions with spontaneously and transiently opened bases