Hitting transcription in all the right places - Nature

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NEWS AND VIEWS teins in eukaryotes. Subsequent studies revealed that this interaction depends on selenocysteyltRNA, providing insight as to how recycling of EFsec might occur at the ribosome15. The crystal structure of archaeal SELB provided further mechanistic insights and yet another surprise. The location of archaeal SECIS elements at either 3′ or 5′ of the coding region might have suggested separate factors for SECIS binding and tRNA delivery, as observed in eukaryotes. However, the search for archaeal SBP2 homologs, either by sequence homology or RNA-binding assays, has not been fruitful. The likely explanation for this comes with the identification of a potential RNAbinding domain in the C-terminal extension of archaeal SELB, analogous to the bacterial protein16. When the elongation factor domain was modeled on the ribosome, the putative SECIS-binding domain pointed toward the 3′ mRNA entrance cleft, positioning it to interact with the downstream SECIS element. Thus, archaea seem to exhibit the tethering feature of the prokaryotic selenocysteine incorporation mechanism, while at the same time using distal SECIS elements for recoding, similar to the situation in Figure 1b. Finally, there remains the dilemma brought about by recoding at a distance in eukaryotes. How does a protein bound to a secondary structure in the 3′ untranslated region deliver an aminoacyl-tRNA–elongation factor complex to the ribosome far upstream in the coding

region? An even greater challenge is presented by selenoprotein P, with 18 UGA selenocysteine codons in the amphibian version of this protein. Are the ribosomes reprogrammed early in the translation process, perhaps by recruitment or delivery of a recoding factor? When Copeland et al.12 initially purified and cloned SBP2, they reported sequence similarity to the L30 ribosomal protein, as well as the conservation of two RNA-binding motifs in both proteins. L30 homologs are found in eukaryotes and archaea, but not in prokaryotes. Subsequent studies showed that the SECIS-binding region of SBP2 corresponds to the RNA-binding region of L30 (ref. 17). Does this mean that L30 is also capable of binding SECIS elements? If so, do SBP2 and L30 compete or complement each other? Chavatte et al.2 have now shown that L30 is indeed a SECIS-binding protein, but SBP2 and L30 do not bind the same element at the same time, as no complexes containing both proteins could be identified. Their studies reveal that the SECIS element seems to act as a ‘molecular switch,’ alternating between an open structure and a kink-turn structure, as originally proposed by Walczal et al.18 based on RNA structure mapping. In vitro the switch can be triggered by addition of magnesium, promoting formation of the kink-turn and binding by L30. Intriguingly, L30 is also thought to bind a kink-turn structure in 28S ribosomal RNA, providing a possible mechanism for tethering the SECIS element to the ribosome (Fig. 1c).

Is ribosome recoding the sole function of L30? This is not likely, as the protein is conserved in genomes that do not encode selenoproteins, implying that it serves other functions. Future studies on the fascinating process of selenoprotein synthesis, the factors involved, and the evolutionary similarities and differences will almost certainly reveal additional surprises. 1. Gesteland, R.F. & Atkins, J.F. Annu. Rev. Biochem. 65, 741–768 (1996). 2. Chavatte, L., Brown, B.A. & Driscoll, D.M. Nat. Struct. Mol. Biol. 12, 408–416 (2005). 3. Leinfelder, W., Zehelein, E., Mandrand-Berthelot, M.A. & Bock, A. Nature 331, 723–725 (1988). 4. Lee, B.J. et al. Mol. Cell. Biol. 10, 1940–1949 (1990). 5. Zinoni, F., Birkmann, A., Leinfelder, W. & Bock, A. Proc. Natl. Acad. Sci. USA 84, 3156–3160 (1987). 6. Leinfelder, W. et al. J. Bacteriol. 170, 540–546 (1988). 7. Forchhammer, K., Leinfelder, W. & Bock, A. Nature 342, 453–456 (1989). 8. Zinoni, F., Heider, J. & Bock, A. Proc. Natl. Acad. Sci. USA 87, 4660–4664 (1990). 9. Berry, M.J. et al. Nature 353, 273–276 (1991). 10. Buettner, C., Harney, J.W. & Larsen, P.R. J. Biol. Chem. 273, 33374–33378 (1998). 11. Wilting, R., Schorling, S., Persson, B.C. & Bock, A. J. Mol. Biol. 266, 637–641 (1997). 12. Copeland, P.R., Fletcher, J.E., Carlson, B.A., Hatfield, D.L. & Driscoll, D.M. EMBO J. 19, 306–314 (2000). 13. Tujebajeva, R.M. et al. EMBO Rep. 2, 158–163 (2000). 14. Fagegaltier, D. et al. EMBO J. 19, 4796–4805 (2000). 15. Zavacki, A.M. et al. Mol. Cell 11, 773–781 (2003). 16. Leibundgut, M., Frick, C., Thanbichler, M., Bock, A. & Ban, N. EMBO J. 24, 11–22 (2005). 17. Copeland, P.R., Stepanik, V.A. & Driscoll, D.M. Mol. Cell. Biol. 21, 1491–1498 (2001). 18. Walczak, R., Westhof, E., Carbon, P. & Krol, A. RNA 2, 367–379 (1996).

Hitting transcription in all the right places Erwan Lejeune & Andreas G Ladurner A new study shows that CtBP, a transcription corepressor, may mediate its effect by blocking histone acetylation, a mark of active transcription. This activity is modulated by NADH binding, thereby supporting a link between cellular metabolism and gene expression.

Pick a really active gene, one that the cell cannot transcribe into RNA often enough, a location in the genome where RNA polymerase keeps coming back to. Now suppose the cell has to reverse this expression pattern because outside conditions have changed, and it no longer needs to transcribe that gene. How can this be achieved? The authors are in the Gene Expression Unit and Structural & Computational Biology Unit, European Molecular Biology Laboratory, Meyerhofstrasse 1, 69117 Heidelberg, Germany e-mail: [email protected]

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There are several ways that transcription at a particular gene can be silenced in eukaryotic cells. For example, repressors could displace and/or replace transcriptional activators at their binding sites; the core transcription machinery and its accomplices, such as coactivators, could be prevented from doing their job; or seemingly small though specific chemical changes to the chromatin template could signal to stop transcription. One particular transcriptional repressor protein called CtBP can bring about all of these mechanisms, but perhaps not all at the same time (Fig. 1). A new study reported on page 423 of this issue now adds

another repressive activity to CtBP’s bag of tricks. Kim et al.1 show that CtBP can inhibit histone acetylation, a mark of active transcription, by blocking access of nuclear histone acetyltransferases such as p300 to their target. This inhibition is reversed by high NADH levels. CtBP could thus sense the metabolic state of the cell and link it with gene expression. CtBP is a 48-kDa cellular protein that interacts with the C-terminal fragment of the E1A oncoprotein, which contains a PxDLS peptide motif (and thus the name C-terminal binding protein). CtBP reduces the transcriptional activation activity of E1A2, and E1A mutants

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CtBP complex:

Lys4 demethylase Lys9/Lys29 methylase Histone deacetylase

CtBP CtBP CtBP

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NADHdependent inhibition p300 of histone binding by bromodomains

LS PxD

NADH

Mitochondria Chromatin

Figure 1 The transcriptional corepressor CtBP associates with a variety of repressive activities and is regulated by cellular NADH levels. CtBP purifies as a large protein complex11. The subunits of this complex encode a histone H3 Lys4 demethylase12, a histone deacetylase and a histone H3 Lys9/Lys27 methyltransferase. A new study shows that CtBP inhibits PxDLS motif–containing bromodomains1, such as those of the histone acetyltransferase p300. When cellular NADH concentrations are high, CtBP dimerizes and no longer inhibits bromodomain proteins.

deleted of the PxDLS motif show a dramatic increase in Ras-mediated tumorigenesis and transformation2,3. These observations suggest that CtBP exerts its negative regulatory effect through the binding of the PxDLS motif. Over the years, it has become clear that CtBP binds a wide range of transcriptional repressors4. CtBP has thus been described as a corepressor. The CtBP family of proteins is well conserved among invertebrates and vertebrates. Genetic studies in Drosophila melanogaster also identified its role as a transcriptional corepressor in vivo, as dCtBP mutants induce segmentation and patterning defects attributed to a loss of gene repression by the short-range repressors Knirps, Krüppel and Snail5–7, all of which contain a PxDLS motif. A similar function has been reported in mammalian systems, as CtBP interacts with several transcription factors that regulate gene expression patterns during differentiation4. An interesting feature of CtBP is its similarity to a subfamily of NAD+-dependent dehydrogenases3,8. CtBP-dependent protein interactions, in fact, turned out to be regulated by the levels of NAD+ and NADH9. Subsequent biochemical and structural analyses have identified genuine dehydrogenase activity in CtBP10, suggesting that its enzymatic activity may stimulate CtBP binding to its in vivo partners. CtBP may therefore serve as a metabolic sensor for transcriptional regulation. Inside the cell, CtBP is a component of a large protein complex11 with distinct repressive chromatin-modifying factors, including LSD1, a histone H3 Lys4-specific demethylase12, histone deacetylases and a histone H3 Lys9/ Lys27-specific methyltransferase. Kim et al.1 discovered an additional activity for the CtBP corepressor when they closely examined the sequences of many bromodomain

proteins. They identified a PxDLS motif in the bromodomain of the nuclear histone acetyltransferase p300 (refs. 13,14). Bromodomains normally mediate binding of acetyltransferases to acetylated histones15,16, but Kim et al.1 demonstrate that CtBP can also directly bind the bromodomains of p300. Previous studies have shown that an increase of NADH levels leads to CtBP dimerization, which in turn increases interaction between CtBP and proteins containing the PxDLS motif. In contrast, Kim et al.1 observe a decrease in interaction between CtBP and the bromodomains of p300 when NADH concentrations are raised. Consistently, an NADHinsensitive CtBP mutant, which does not dimerize, maintains p300 interactions even at high NADH concentrations. In the structure of the p300 bromodomain the PxDLS interaction motif is close to the acetylated lysine–binding pocket, suggesting that CtBP may interfere with binding to acetylated histones. The authors show that CtBP competes with histones for p300 binding and that this competition is NADH-dependent. Accordingly, CtBP represses p300-mediated transactivation that depends on bromodomain function17. The data suggest that CtBP and p300 may antagonize each other functionally. Consistently, Kim et al.1 show that the overexpression of p300 leads to an increase in acetylation of histones H3 and H4, particularly in CtBP–/– knockout cells. When ectopic CtBP is reintroduced in the knockout cells, H3 and H4 acetylation is reduced again. Overall, these data imply that CtBP is involved in regulating histone acetylation and that this function may be linked to cellular metabolism. The novel findings are interesting from two points of view. The first relates to the known components in the CtBP complex that reduce

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transcription by altering chromatin structure. Here, the ability of CtBP to interfere with the coactivator function of p300 (and probably other chromatin factors that contain bromodomains) seems like a convenient, additional mechanism to ensure gene repression. Kim et al.1 did not address whether CtBP in the context of the CtBP protein complex11 can interact with the p300 bromodomain. If so, the CtBP complex would avail itself of a full range of repressive activities, including the demethylation of Lys4 in H3, deacetylation, methylation of Lys9 and Lys27 of H3 and the now-described blocking of coactivation activity by p300. All of these activities eventually could lead to changes in chromatin structure that reduce gene transcription. It is also possible that CtBP may bind bromodomains outside the context of the known CtBP complex. In this role, CtBP’s function may lie upstream of transcriptional activation, as histone acetylation by p300 may be one of the early events in the histone modification game that leads to high transcription levels18. Either way, the ability of CtBP to target bromodomains directly is likely to represent a useful way to lower gene activity. The second and quite intriguing aspect relates to the role of NADH in regulating nuclear functions and gene repression. Recently, there has been increasing interest in the role of metabolism in gene expression and, in particular, in the role of NAD+ in transcription regulation. There is evidence that during repair of single-strand DNA breaks, for instance, large amounts of ADP-ribose polymers are formed using NAD+ as a substrate. As a result, NAD+ levels can drop rapidly and it has been suggested that cells may sense that they are running‘low in energy’ and thus promote apoptosis, rather than continuing in the attempt to fix DNA that has too much damage19,20. In addition, the NAD+/NADH ratio regulates another gene repressor, the Sir2 family of deacetylases. Unusual for enzymes that carry out the hydrolysis of an amide bond, the transcriptional repressor Sir2 uses NAD+ as a cofactor21–23 and converts acetylated lysines, for example, on histones, back to unmodified lysines. For each cycle of the reaction, one molecule of NAD+ is converted to nicotinamide, which is fed back into the NAD pathway, and O-acetyl-ADP-ribose24,25, a little understood metabolite. Organisms on a restrictive diet (so-called calorie restriction) alter their NAD+ levels, and this seems to affect lifespan through the regulation of Sir2 function26,27. The mammalian Sir2 ortholog SirT1, for example, is thought to mediate some of the benefits of calorie restriction through the regulation of a series of transcription factors with roles in fat metabo-

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NEWS AND VIEWS lism28–31 and also by repression of transcription through the deacetylation of histones32. Unlike for Sir2, there is no evidence that CtBP regulates energy homeostasis and physiology through its repressive functions. Nevertheless, it is interesting to note that both Sir2 and CtBP are active when the NAD+/NADH ratio is high. Under this condition, CtBP inhibits p300 coactivator function, based on the results of Kim et al.1, and Sir2 deacetylates histones. These activities together should turn off gene expression. Under conditions where the NAD+/NADH ratio is low, Sir2 is inactive, and CtBP dimerizes and is unable to inhibit p300. Further work will be necessary to understand why the binding of CtBP to proteins containing a PxDLS motif, such as E1A and p300, is differentially regulated by NAD. Clearly, cells could benefit from a feedback loop between their metabolic state and gene expression. It is also clear that metabolic control is more intricate than the examples with Sir2 and CtBP discussed above. Nonetheless, the present study supports the idea that chromatin architecture is under the control of

cellular metabolism. It is mechanistically alluring that such control should occur in the form of direct, regulatory interactions between NAD and distinct chromatin repressors. Surely, further molecular links between cellular metabolites and gene expression will emerge. ACKNOWLEDGMENTS E.L. is supported by an E-STAR fellowship funded by the European Commission’s FP6 Marie Curie Host fellowship for Early Stage Research Training and is a student in the European Molecular Biology Laboratory International PhD program. 1. Kim, J.H., Cho, E.J., Kim, S.T. & Youn, H.D. Nat. Struct. Mol. Biol. 12, 423–428 (2005). 2. Boyd, J.M. et al. EMBO J. 12, 469–478 (1993). 3. Schaeper, U. et al. Proc. Natl. Acad. Sci. USA 92, 10467–10471 (1995). 4. Chinnadurai, G. Mol. Cell 9, 213–224 (2002). 5. Nibu, Y., Zhang, H. & Levine, M. Science 280, 101– 104 (1998). 6. Nibu, Y. & Levine, M.S. Proc. Natl. Acad. Sci. USA 98, 6204–6208 (2001). 7. Poortinga, G., Watanabe, M. & Parkhurst, S.M. EMBO J. 17, 2067–2078 (1998). 8. Turner, J. & Crossley, M. Bioessays 23, 683–690 (2001). 9. Zhang, Q., Piston, D.W. & Goodman, R.H. Science 295, 1895–1897 (2002). 10. Kumar, V. et al. Mol. Cell 10, 857–869 (2002). 11. Shi, Y. et al. Nature 422, 735–738 (2003). 12. Shi, Y. et al. Cell 119, 941–953 (2004).

13. Moran, E. Curr. Opin. Genet. Dev. 3, 63–70 (1993). 14. Arany, Z., Newsome, D., Oldread, E., Livingston, D.M. & Eckner, R. Nature 374, 81–84 (1995). 15. Dhalluin, C. et al. Nature 399, 491–496 (1999). 16. Jacobson, R.H., Ladurner, A.G., King, D.S. & Tjian, R. Science 288, 1422–1425 (2000). 17. Manning, E.T., Ikehara, T., Ito, T., Kadonaga, J.T. & Kraus, W.L. Mol. Cell. Biol. 21, 3876–3887 (2001). 18. Agalioti, T. et al. Cell 103, 667–678 (2000). 19. Pieper, A.A., Verma, A., Zhang, J. & Snyder, S.H. Trends Pharmacol. Sci. 20, 171–181 (1999). 20. Decker, P., Muller, S. Curr. Pharm. Biotechnol. 3, 275–283 (2002). 21. Imai, S., Armstrong, C.M., Kaeberlein, M. & Guarente, L. Nature 403, 795–800 (2000). 22. Landry, J. et al. Proc. Natl. Acad. Sci. USA 97, 5807– 5811 (2000). 23. Smith, J.S. et al. Proc. Natl. Acad. Sci. USA 97, 6658– 6663 (2000). 24. Tanner, K.G., Landry, J., Sternglanz, R. & Denu, J.M. Proc. Natl. Acad. Sci. USA 97, 14178–14182 (2000). 25. Jackson, M.D. & Denu, J.M. J. Biol. Chem. 277, 18535–18544 (2002). 26. Lin, S.J. & Guarente, L. Curr. Opin. Cell Biol. 15, 241–246 (2003). 27. Guarente, L. & Picard, F. Cell 120, 473–482 (2005). 28. Brunet, A. et al. Science 303, 2011–2015 (2004). 29. Picard, F. et al. Nature 429, 771–776 (2004). 30. Yang, Y., Hou, H., Haller, E.M., Nicosia, S.V. & Bai, W. EMBO J. 24, 1021–1032 (2005). 31. Nemoto, S., Fergusson, M.M. & Finkel, T. Science 306, 2105–2108 (2004). 32. Vaquero, A., Scher, M., Lee, D., Erdjument-Bromage, H., Tempst, P. Reinberg, D. Mol. Cell 16, 93–105 (2004).

Rad50 connects by hook or by crook Michael Lichten The Mre11 protein complex plays important roles in maintaining genome stability. Inter-molecular bridging by the Rad50 protein has now been shown to be critical to this complex’s function. Eukaryotes contain several systems that respond to double-strand DNA breaks (DSBs) and that repair breaks in ways that preserve genome integrity. The Mre11 complex, which contains the mammalian Mre11, Rad50 and Nbs1 proteins (Xrs2 substitutes for Nbs1 in budding yeast), acts in the DSB response, in break repair by either homologous recombination (HR) or nonhomologous endjoining (NHEJ), and in telomere maintenance. A recent study in budding yeast on page 403 of this issue provides insight into how the Mre11 complex accomplishes these tasks1. Studies in a variety of organisms have documented roles for the Mre11 complex in several aspects of DNA metabolism2, most prominently in DNA repair. For example, budding yeast mutants lacking Mre11, Rad50 or Xrs2 The author is in the Laboratory of Biochemistry, Center for Cancer Research, Building 37, Room 6124, National Cancer Institute, Bethesda, Maryland, 20892-4255, USA. e-mail: [email protected]

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show profound sensitivity to DSB-forming agents, a consequence of defects in both NHEJ3 and HR4, and grow slowly and accumulate at metaphase, consistent with unrepaired DNA lesions leading to transient cell cycle arrest in many cells. The Mre11 complex’s functions also extend beyond DNA damage repair. Null mutants in both yeast and in Drosophila melanogaster exhibit telomere metabolism defects, owing at least in part to the complex’s role in mediating DNA end recognition by the ATM (Tel1 in yeast) checkpoint kinase2,5,6. The Mre11 complex localizes to DSBs within minutes of break formation7–9 and recruits ATM/ Tel1 via interactions with the Nbs1/Xrs2 subunit10,11, activating the kinase12 and thus triggering a DNA damage response. Although the activities described above involve end-binding, studies of budding yeast meiosis suggest that the Mre11 complex also may interact with other chromatin features. In this organism, the Mre11 complex is required for formation of the DSBs that initiate meiotic recombination, and a meiosis-specific association of the

complex with potential break sites occurs even in the absence of DSB formation13. Two components of Mre11 complex, Mre11 and Rad50, are found in organisms ranging from archaea and eubacteria to mammals14. Studies of the archaeal proteins15,16 indicate that Mre11 is a two-lobed nuclease. Rad50 is a split ABCtype ATPase; its center contains a long heptad repeat that folds into a 60-nm antiparallel coiled coil, bringing the N-terminal (Walker A) and C-terminal (Walker B) domains in close proximity. The native complex most likely contains a dimer of heterotrimers, with cross-domain interactions between Rad50 monomers forming two ATP-binding sites and, with two Mre11 monomers, a DNA-binding module15 (see Fig. 1a). The intact complex has DNA unwinding and endonuclease and exonuclease activities2. The Mre11 nuclease does not contribute to all of the activities of the eukaryotic complex, as budding yeast mutants in the nuclease active site retain considerable in vivo function17. The apex of the Rad50 coiled coil contains another dimerization interface, a conserved

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