NATURE STRUCTURAL & MOLECULAR BIOLOGY VOLUME 12 NUMBER 5 MAY 2005 389

Knowing when not to stopMarla J Berry

A recent study reveals that the eukaryotic ribosomal protein L30 binds to the selenocysteine recoding RNA element and may function to tether the recoding machinery to the translating ribosome.

The author is in the Department of Cell and Molecular Biology, University of Hawaii at Manoa, 1960 East West Road, T514, Honolulu, Hawaii 96822, USA.e-mail: mberry@hawaii.edu

The genetic code was elucidated in the 1960s, revealing the language by which genes are translated into proteins using code words for start, stop and the 20 standard amino acids. In the late 1980s, the code was expanded to add what has been termed the 21st amino acid, seleno cysteine. The position of this amino acid in proteins was shown to correspond to a stop signal, UGA, in the genes, and the term ‘recoding’ was coined to describe this process1. Since that time, unraveling the mystery of how a stop codon is recoded to a sense codon has been the subject of considerable efforts, with a number of surprises arising over the years. The latest of these, reported by Chavatte et al.2 on page 408 of this issue, is the demonstration that eukarya recruit a ribosomal protein, L30, to selenoprotein mRNAs, presumably to assist in recoding at the ribosome.

The UGA code word for selenocysteine is conserved in all three kingdoms of life, as is the presence of unique tRNAs with an anti-codon complementary to UGA3,4. Studies by Bock and colleagues have elucidated the key features of the recoding process by which UGA is read as selenocysteine in bac-teria5–7. In particular, cis-acting RNA struc-tures located immediately downstream of the UGA codons were identified and shown to function in recruiting a complex essential for decoding8. The complex in Escherichia coli consists of the unique selenocysteyl-tRNA, a translation elongation factor termed SELB dedicated to this tRNA, and GTP to provide energy. SELB contains an elongation factor domain similar to those found in the stan-dard elongation factors, EF-Tu and EF-1A, that deliver other tRNAs to the ribosome in prokaryotes and eukaryotes, respectively. In addition to this elongation factor domain, a C-terminal extension of SELB was shown to bind the cis-acting RNA secondary struc-tures, termed selenocysteine incorporation sequence (SECIS) elements, thus recruiting the elongation factor–tRNA complex to the adjacent UGA codon (Fig. 1a).

At the time when recoding components were being characterized in bacteria, only a single selenoprotein had been identified in verte-brates, but the conserved use of UGA in pro-karyotes and eukaryotes raised the possibility that the recoding process might also be similar. For example, it was assumed that in eukaryotes the SECIS RNA element would occupy a similar position and function analogously as observed in prokaryotes. The first surprise came with the demonstration that the structures required for recoding in eukaryotes were found not in the coding region adjacent to the UGA, but rather far downstream in the 3′ untranslated region9. In some mRNAs, these structures are located kilobases away from the UGA codons they serve10. The sequences and structures of the SECIS elements proved to be conserved among eukaryotes but quite distinct from those in pro-karyotes. Thus, the search was on for the pre-sumed eukaryote homolog of bacterial SELB, a putative factor that would recognize eukaryotic SECIS elements and deliver selenocysteyl-tRNA to the upstream UGA codons. Another mecha-nistic difference is that eukaryotes could recode multiple upstream UGAs with a single SECIS

element, whereas in bacteria, the proximity of the SECIS element to the UGA constrains recoding to that position. It was subsequently found that archaea recode UGA from a dis-tance, via structures that differ from those of either prokaryotes or eukaryotes11: although most are found in the 3′ untranslated region, at least one archaeal SECIS is located in the 5′ untranslated region.

The next surprise came with the identifica-tion of two distinct factors in eukaryotes serv-ing the function of the single bacterial protein. Copeland et al.12 identified a SECIS-binding protein termed SBP2 (the first SBP proved to be a false start) that lacked an elongation fac-tor domain. Shortly thereafter, studies in two laboratories reported the identification of the archaeal, nematode, fly and vertebrate seleno-cysteyl-tRNA specific elongation factors13,14. The murine protein, termed EFsec or mSELB, was shown to lack SECIS-specific binding capa-bility, but instead, to interact with SBP2. Thus, the two functions of bacterial SELB—recruit-ment of the elongation factor to the RNA and delivery of selenocysteyl-tRNA to the ribo-some—are carried out by two interacting pro-

Figure 1 Models for selenocysteine incorporation. (a,b) Models for prokaryotic (a) and (b) archaeal recoding machinery. mRNAs are blue, ribosomes purple, tRNAs yellow, nascent peptide pink, and codons black. SELB from prokaryotes and archae are shown as red and blue ovals (blue, elongation factor domain; red, SECIS-binding domain). (c) Current model of eukaryotic recoding machinery. Eukaryotic EFsec is blue and dark purple (blue, elongation factor domain; dark purple, SBP2-interaction domain). SBP2 is red and L30 green. The kink-turn in the SECIS element is depicted in the right panel.

N E W S A N D V I E W S©

2005

Nat

ure

Pub

lishi

ng G

roup

ht

tp://

ww

w.n

atur

e.co

m/n

smb

390 VOLUME 12 NUMBER 5 MAY 2005 NATURE STRUCTURAL & MOLECULAR BIOLOGY

teins in eukaryotes. Subsequent studies revealed that this interaction depends on selenocysteyl-tRNA, providing insight as to how recycling of EFsec might occur at the ribosome15.

The crystal structure of archaeal SELB pro-vided further mechanistic insights and yet another surprise. The location of archaeal SECIS elements at either 3′ or 5′ of the cod-ing region might have suggested separate fac-tors 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 RNA-binding 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 inter-act with the downstream SECIS element. Thus, archaea seem to exhibit the tethering feature of the prokaryotic selenocysteine incorpora-tion 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 com-plex 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 ribo-somal 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 capa-ble 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 struc-ture 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, promot-ing 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 seleno-protein 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 placesErwan 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?

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 coact-ivators, 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 transcrip-tional 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 his-tone acetylation, a mark of active transcription, by blocking access of nuclear histone acetyl-transferases 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 inter-acts with the C-terminal fragment of the E1A oncoprotein, which contains a PxDLS peptide motif (and thus the name C-terminal bind-ing protein). CtBP reduces the transcriptional activation activity of E1A2, and E1A mutants

The authors are in the Gene Expression Unit and Structural & Computational Biology Unit, European Molecular Biology Laboratory, Meyerhofstrasse 1, 69117 Heidelberg, Germanye-mail: ladurner@embl.de

N E W S A N D V I E W S©

2005

Nat

ure

Pub

lishi

ng G

roup

ht

tp://

ww

w.n

atur

e.co

m/n

smb

NATURE STRUCTURAL & MOLECULAR BIOLOGY VOLUME 12 NUMBER 5 MAY 2005 391

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 pattern-ing defects attributed to a loss of gene repressionby 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 dehydroge-nases3,8. 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 there-fore serve as a metabolic sensor for transcrip-tional regulation.

Inside the cell, CtBP is a component of a large protein complex11 with distinct repres-sive 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 acetyl-transferase p300 (refs. 13,14). Bromodomains normally mediate binding of acetyltransfer-ases 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 concen-trations are raised. Consistently, an NADH-insensitive 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 acetyl-ated 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 bromo domain function17. The data suggest that CtBP and p300 may antagonize each other functionally. Consistently, Kim et al.1 show that the over-expression of p300 leads to an increase in acetyl -ation 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

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 bromo-domains) 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 demethyl-ation 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 tran-scription. 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 modi-fication 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 func-tions and gene repression. Recently, there has been increasing interest in the role of metabo-lism 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 regu-lates another gene repressor, the Sir2 fam-ily 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 acetyl-ated 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 restric-tion) alter their NAD+ levels, and this seems to affect lifespan through the regula-tion 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-

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.

NADH

CtBPCtBP

CtBP

Mitochondria

CtBP complex:

NADH-dependent

inhibition of histone binding by

bromodomains

p300. . . . .. . . . .

Chromatin

Lys4 demethylaseLys9/Lys29 methylase

Histone deacetylase

PxDLS

N E W S A N D V I E W S©

2005

Nat

ure

Pub

lishi

ng G

roup

ht

tp://

ww

w.n

atur

e.co

m/n

smb

392 VOLUME 12 NUMBER 5 MAY 2005 NATURE STRUCTURAL & MOLECULAR BIOLOGY

lism28–31 and also by repression of transcrip-tion 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 under-stand 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 feed-back 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.

ACKNOWLEDGMENTSE.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 crookMichael 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 recom-bination (HR) or nonhomologous end-joining (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 docu-mented roles for the Mre11 complex in several aspects of DNA metabolism2, most promi-nently in DNA repair. For example, budding yeast mutants lacking Mre11, Rad50 or Xrs2

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 mela-nogaster 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 min-utes of break formation7–9 and recruits ATM/Tel1 via interactions with the Nbs1/Xrs2 sub-unit10,11, activating the kinase12 and thus trig-gering 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 recombina-tion, 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 ABC-type 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 prox-imity. The native complex most likely contains a dimer of heterotrimers, with cross-domain interactions between Rad50 monomers form-ing two ATP-binding sites and, with two Mre11 monomers, a DNA-binding module15 (see Fig. 1a). The intact complex has DNA unwind-ing and endonuclease and exonuclease activi-ties2. 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

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: lichten@helix.nih.gov

N E W S A N D V I E W S©

2005

Nat

ure

Pub

lishi

ng G

roup

ht

tp://

ww

w.n

atur

e.co

m/n

smb

NATURE STRUCTURAL & MOLECULAR BIOLOGY VOLUME 12 NUMBER 5 MAY 2005 393

Cys-X-X-Cys motif in a hook-shaped domain that dimerizes with a second hook domain via cysteine-mediated zinc ion coordination16. Dimerization between two Rad50 protomers in a single complex would form a large loop; dimerization could also occur between two complexes bound to different DNA mol-ecules. Both modes of association are seen by atomic force and electron microscopy18,19, and both could tether broken DNA molecules (see Fig. 1).

Although evidence for Mre11 complex–dependent tethering had been provided in vitro and in vivo20,21, the relevance of teth-ering to function remained to be established. Work in budding yeast showed that replacing the two Rad50 hook cysteines with glycine caused a loss of Mre11 binding and a null mutant phenotype16. However, because the hook is also the turn that allows the anti-parallel coiled coil to form, it remained a distinct possibility that this mutant altered other aspects of Rad50 structure.

Wiltzius et al.1 have now directly exam-ined the role of hook dimerization in the function of Mre11 complex by replacing the hook with a conditional dimerization module. They split budding yeast RAD50 into two genes. Each contained half of the heptad repeat region (Rad50N and Rad50C), and the hook domain was removed in the process. Rad50N and Rad50C coassembled with relatively high efficiency and also associated with Mre11. Despite this appar-ent success in reassembly of the complex, cells containing this ‘hookless’ Mre11 com-

plex behaved as rad50-null mutants. If the hookless complex was otherwise normal, then these results would implicate the Rad50 hook domain, and presumably hook-mediated dimerization, as being important to the function of Mre11 complex.

To prove this, Wiltzius et al.1 restored dimerization to the complex. They attached a human FK506-binding protein (FKBP) derivative at the Rad50N C terminus (Rad50N-FKBP). AP20187, a bivalent ligand containing two FK506 moieties, binds two FKBP domains in a geometry similar to that of zinc-coordinated Rad50 hooks16,22 (see Fig. 1b). AP20187 treatment of cells expressing Rad50N-FKBP and Rad50C caused partial to full suppression of all mutant phenotypes tested, including slow growth, sensitivity to methyl methane sulfonate, and short telomere length. Furthermore, AP20187 partially restored DSB formation to meiotic cells. Although only partial sup-pression was observed, it should be noted that the Mre11 complex had suffered both bisection of a major component and replace-ment of its dimerization domain and ligand with components roughly 20 times larger than the originals. Moreover, function was restored not just in contexts where DNA end-tethering should be important (DNA damage resistance), but also in a context involving a single DNA end (telomere main-tenance), and even a situation (meiotic DSB formation) where DNA ends are absent. These results clearly support the hypothesis that Rad50 hook domain dimerization

impacts all aspects of the function of Mre11 complex.

In their discussion, Wiltzius et al.1 suggest that the primary function of the Mre11 com-plex might simply be to hold DNA molecules or chromosomes together. However, it should be kept in mind that this complex is also a nuclease and a signal transducer. Structural studies of the archaeal Mre11–Rad50 com-plex indicate that changing the geometry of the globular base can change the trajectory of the coiled coil15, and it seems reasonable to suggest that the converse would be true. As Wiltzius et al.1 point out, changing from an intracomplex tether to an untethered state, or to an intercomplex tether, might alter the geometry of the Mre11 and Nbs1 (Xrs2) proteins bound at the base, with the poten-tial to modulate either the nuclease or ATM/Tel1 activation—and one of these might have played roles in the assays used in this paper. Thus, although this intriguing paper provides solid evidence for a critical contri-bution by Rad50 hook dimerization, it also makes clear the importance of examining its impact on each of Mre11 complex’s sepa-rate activities. Most fortunately, in adapting the FKBP-AP20187 system to Rad50 hook dimerization, Wiltzius et al.1 have also pro-vided the field with the tools to start address-ing these issues.

1. Wiltzius, J.J.W., Hohl, M., Fleming, J.C. & Petrini, J.H. Nat. Struct. Mol. Biol. 12, 403–407 (2005).

2. Assenmacher, N. & Hopfner, K.P. Chromosoma 113, 157–166 (2004).

3. Moore, J.K. & Haber, J.E. Mol. Cell. Biol. 16, 2164–2173 (1996).

4. Bressan, D.A., Baxter, B.K. & Petrini, J.H. Mol. Cell. Biol. 19, 7681–7687 (1999).

5. Lydall, D. J. Cell Sci. 116, 4057–4065 (2003).6. Purdy, A. & Su, T.T. Curr. Biol. 14, R613–R614

(2004).7. Mirzoeva, O.K. & Petrini, J.H. Mol. Cell. Biol. 21,

281–288 (2001).8. Lisby, M., Barlow, J.H., Burgess, R.C. & Rothstein, R.

Cell 118, 699–713 (2004).9. Shroff, R. et al. Curr. Biol. 14, 1703–1711 (2004).10. Nakada, D., Matsumoto, K. & Sugimoto, K. Genes Dev.

17, 1957–1962 (2003).11. Falck, J., Coates, J. & Jackson, S.P. Nature 434, 605–

611 (2005).12. Lee, J.H. & Paull, T.T. Science published online, 24

March 2005 (doi:10.1126/science.1108297).13. Prieler, S., Penkner, A., Borde, V. & Klein, F. Genes Dev.

19, 255–269 (2005).14. Connelly, J.C. & Leach, D.R. Trends Biochem. Sci. 27,

410–418 (2002).15. Hopfner, K.P. et al. Cell 105, 473–485 (2001).16. Hopfner, K.P. et al. Nature 418, 562–566 (2002).17. Moreau, S., Ferguson, J.R. & Symington, L.S. Mol. Cell.

Biol. 19, 556–566 (1999).18. de Jager, M. et al. Mol. Cell 8, 1129–1135 (2001).19. Anderson, D.E., Trujillo, K.M., Sung, P. & Erickson, H.P.

J. Biol. Chem. 276, 37027–37033 (2001).20. Costanzo, V., Paull, T., Gottesman, M. & Gautier, J.

PLoS Biol. 2, e110 (2004).21. Lobachev, K., Vitriol, E., Stemple, J., Resnick, M.A. &

Bloom, K. Curr. Biol. 14, 2107–2112 (2004).22. Rollins, C.T. et al. Proc. Natl. Acad. Sci. USA 97,

7096–7101 (2000).

B A11

N/X

B A11

FKBP

FKBP

FKBP

FKBP

Zn

AP20187

N/X

a b c

Figure 1 Interactions between hook domains in the Mre11 complex. Concepts are derived from refs. 2 and 15–17. (a) Schematic of the Mre11 complex, not to scale, showing Zn-coordinated interactions between coiled-coil domains. Relative geometry of Mre11 (11) and the Walker A (A), Walker B (B) and coiled-coil domains of Rad50 are as suggested15. All other aspects, including geometries of zinc hook (Zn) and Nbs1/Xrs2 subunit (N/X), are imagined. (b) Illustration of how Wiltzius et al.1 replaced the hook domain with an FKBP/AP20187 conditional dimerization module. (c) Possible ways that hook dimerization could tether two DNA molecules. Top, intracomplex dimerization stabilizes the binding of two DNA molecules by a single complex. Bottom left, intracomplex dimerization forms interlocking loops that tether two complexes. Bottom right, intercomplex dimerization tethers two DNA molecules.

N E W S A N D V I E W S©

2005

Nat

ure

Pub

lishi

ng G

roup

ht

tp://

ww

w.n

atur

e.co

m/n

smb