Focused Meeting and Satellite to BioScience2005, held at Western Infirmary, Glasgow, U.K., 21–23 July 2005. Organized and Edited by M. Bushell
(Nottingham, U.K.), S. Newbury (Newcastle upon Tyne, U.K.), G. Pavitt (Manchester, U.K.) and A. Willis (Nottingham, U.K.).
The life and death of translation elongation
factor 2
Translation UK
Translation UK
R. Jørgensen*, A.R. Merrill* and G.R. Andersen†1
*Department of Molecular and Cellular Biology, University of Guelph, Guelph, Ontario, Canada N1G 2W1, and †Centre for Structural Biology, Department of
Molecular Biology, University of Aarhus, Gustav Wieds Vej 10C, DK-8000, Denmark
Abstract
eEF2 (eukaryotic elongation factor 2) occupies an essential role in protein synthesis where it catalyses the
translocation of the two tRNAs and the mRNA after peptidyl transfer on the 80 S ribosome. Recent crystal
structures of eEF2 and the cryo-electron microscopy reconstruction of its 80 S complex now provide a substantial structural framework for dissecting the functional properties of this factor. The factor can be modified
by either phosphorylation or ADP-ribosylation, which results in cessation of translation. We review the
structural and functional properties of eEF2 with particular emphasis on the unique diphthamide residue,
which is ADP-ribosylated by diphtheria toxin from Corynebacterium diphtheriae and exotoxin A from
Pseudomonas aeruginosa.
Introduction
In the elongation cycle of protein synthesis, elongation factor 1A delivers aminoacylated tRNA to the ribosomal A-site,
whereafter the peptidyl transferase centre catalyses formation
of the peptide bond leaving a deacylated tRNA in the ribosomal P-site and the newly formed peptidyl-tRNA in the
A-site. This pre-translocational state of the ribosome is
the substrate of the GTPase eEF2 (eukaryotic elongation factor 2) (reviewed in [1]), which catalyses the co-ordinated
movement of the two tRNA molecules, the mRNA and
conformational changes in the ribosome.
The bacterial factor EF2 (formerly EF-G) and the archaeal aEF2 are structurally and functionally homologous
with eEF2. Both eEF2 and EF2 contain six structural domains (Figures 1 and 2). Domains I–V have folds as first described for the bacterial EF2, although insertions are found
in domains I, II and IV of eEF2 [2,3]. Like other GTP-binding
proteins, domain I contains five conserved sequence elements
(G1–G5), forming the binding pocket for GTP/GDP. The G
domain in eEF2 is somewhat bigger and shares no structural similarity with the corresponding domain in EF2. eEF2
is a highly elongated molecule with interesting dynamic
Key words: ADP-ribosylation, cryo-electron microscopy, diphthamide, diphtheria toxin,
elongation factor 2 (EF2), exotoxin A.
Abbreviations used: AdoMet, S-adenosylmethionine; DT, diphtheria toxin; EF2, elongation
factor 2; aEF2, archaeal EF2; eEF2, eukaryotic EF2; EM, electron microscopy; ETA, exotoxin A.
1
To whom correspondence should be addressed (email [email protected]).
properties [2]. Crystal structures have shown two conformations related by a quite large conformational change
involving domain rotations of up to 75◦ (Figure 2). The apoconformation of eEF2 is rather similar to the conformation
of bacterial EF2 in complex with GDP, while binding of the
fungicide sordarin to eEF2 induces a conformation so far not
observed in EF2 [2]. The molecule contains three structural
blocks (domains I–II, domain III and domain IV–V), which
can move relative to each other. No structure is yet known
of either eEF2 or EF2 in complex with GTP, but, since
the G3 region (also called switch II) is expected to change
significantly in response to GTP hydrolysis and is located
at the interface between the three structural blocks, it
appears likely that GTP hydrolysis will be correlated with
conformational changes in the factor. This is in agreement
with elegant kinetic experiments on EF2 showing that GTP
hydrolysis precedes and accelerates translocation [4], and that
an EF2 molecule cross-linked across the domain I–V interface
can still hydrolyse GTP, but is inactive in both translocation
and turnover [5]. Therefore the energy derived from GTP
hydrolysis is used for the promotion of conformational
changes in the ribosome preceding translocation. A rather
complete kinetic scheme for bacterial translocation has been
deduced [6]. Unfortunately, similar kinetic studies of eukaryotic translocation is currently lacking, but the very high
conservation of both ribosomes, tRNA and EF2 makes it
likely that the mechanism of translocation is quite similar in
all kingdoms.
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Figure 1 Structural alignment of eEF2 from Saccharomyces cerevisiae (EF2 YEAST) and EF2 from Thermus thermophilus (EFG THETH)
The α-helices (circles) and β-strands (arrows) are shaded according to the domain structure. Secondary-structure elements
from domains I, III and V are shaded light grey, while those of domains G , II, and IV are dark grey. The G1–G5 motifs
are indicated by black boxes, the two threonine residues phosphorylated by the eEF2 kinase are marked with ‘P’, and the
diphthamide (encoded as histidine) is labelled with an asterisk (*).
Cryo-EM (electron microscopy) reconstructions of both
sordarin-stabilized eEF2–80 S [7] and antibiotic/GDPNPstabilized EF2–70 S complexes [8–10] have been conducted,
and support a similar mechanism of translocation in bacteria and eukaryotes. The docking of the sordarin-induced
conformation into the 12 Å (1 Å = 0.1 nm) density of eEF2
required only rigid body rotation of two large structural
blocks, whereas docking of EF2 into cryo-EM reconstruction
maps of EF2–70 S complexes at 11–18 Å resolution required
docking of individual small domains, which may be less reliable at low resolution. In the 80 S complex, eEF2 domains I
and V interact with the 60 S subunit, and, in particular, the
rRNA sarcin–ricin loop contacts the nucleotide binding
pocket in domain I. Domain II interacts with the 40 S subunit,
while domains III and IV are in contact with both ribosomal
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subunits. An important functional role of eEF2/EF2 domain IV is also evident from kinetic experiments with a
domain-IV-deletion mutant of EF2, which still has ribosomestimulated GTPase activity, but only catalyses translocation
slowly and fails to be released from the ribosome after GTP
hydrolysis [4].
The diphthamide
In contrast with bacterial EF2, eEF2 and aEF2 contain a
unique and strictly conserved post-translationally modified
histidine residue located at the tip of domain IV (Figure 2).
Previously, NMR analysis suggested the residue to be a 2-[3carboxyamido-3-(trimethylammonio)-propyl] histidine, also
referred to as diphthamide [11,12]. The biosynthesis of
diphthamide is achieved by a complex stepwise addition to the
Translation UK
Figure 2 Conformations of eEF2 from Saccharomyces cerevisiae
(A) eEF2-apo. (B) ADP-ribosylated eEF2 having both the antifungal inhibitor, sordarin (black), GDP (yellow) and Mg2+
(cyan) bound to the factor. The three N-terminal domains of the two structures are superimposed and then translated
horizontally. GDP and Mg2+ shown with the eEF2-apo structure are modelled from the ADP-ribosylated eEF2 by superposition.
The structural domains are coloured in different blue shades and indicated by numerals I–V, except for G . ADPR-DIPH,
ADP-ribosylated diphthamide.
histidine residue [13–19]. The first step is the transfer of the 3amido-3-carboxypropyl group of AdoMet (S-adenosylmethionine) to the imidazole CE1 of the histidine residue and
requires the action of the four proteins, referred to in yeast as
Dph1–Dph4, of which Dph1–3 seem to associate to form a
multimeric complex. In the next step, Dph5 is responsible for
the trimethylation of the resulting amino group, again using
AdoMet as the methyl donor. Finally, the carboxyl group
of the diphthamide is amidated in an ATP-dependent manner
by a yet unidentified enzyme. In yeast eEF2, the diphthamide
is formed by modification of His699 . Recently, we modelled
the diphthamide in a F o –F o electron density map from the
2.6 Å crystal structure of ADP-ribosylated yeast eEF2 [20]
(Figure 3). Its structure was in agreement with the NMR
analysis of the diphthamide, but furthermore suggested the
chiral centre of the third carbon atom to be an R enantiomer.
The function of the diphthamide is still unknown. Interestingly, mouse and human DPH1 genes were identified previously as the tumour-suppressor gene, OVCA1 [21]. Knockout of one allele of DPH1 in mice results in increased tumour
development, whereas loss of both alleles leads to death
during embryonic development [22]. It is possible that the
effects of Dph1 deficiency result from the lack of diphthamide on eEF2, and therefore the diphthamide may have an
Figure 3 ADP-ribosylated diphthamide of eEF2 with interactions
between the ADP-ribose (ADPR) and eEF2
Hydrogen bonds are shown in yellow, while potential interactions are
shown in light blue.
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important function in translation. On the other hand, several
mutagenesis experiments resulting in strains that lacked
the ability to form diphthamide or mutations of the diphthamide indicate that it is not strictly required for eEF2
function [16,19,23–25]. The cryo-EM studies of eEF2 in complex with the yeast 80 S ribosome show that the area around
the diphthamide at the tip of domain IV is close enough
to interact with the ribosomal decoding site and, in particular,
two universally conserved adenines in helix 44 of 18 S rRNA
[7], which serve to distinguish cognate from near-cognate
codon–anticodon duplexes. This suggests that this area of
eEF2, including the diphthamide, may somehow stabilize the
codon–anticodon pairing during translocation, thereby preventing frame shifts. A cytoplasmic ADP-ribosyltransferase
capable of ADP-ribosylating eEF2 in a wide variety of
eukaryotic cell types has been identified [26–29], suggesting
a possible regulatory mechanism of eEF2. The in vivo significance of this transferase activity, however, is still unclear,
and the protein responsible for ribosylation is yet to be
identified.
Inactivation of eEF2 by kinases and
bacterial toxins
Both mammalian [30] and yeast [31] eEF2 can be phosphorylated by endogenous kinases which leads to translational
down-regulation owing to a reversible inactivation of the
factor. In rabbit eEF2, two threonine residues in the G2
region (also known as switch I) are modified [32], which
correlates well with the lower affinity of phosphorylated rat
eEF2 for GTP, while the affinity for GDP is unaltered [33].
The G2 region is likely to play an important role for binding/hydrolysis of GTP in the ribosomal context, but the
region is disordered/unfolded and solvent-exposed in all
known structures of eEF2 [2,20] and thereby likely to be an
excellent substrate for the kinase. In mammals, the eEF2
kinase belongs to the α-kinase family, and the phosphorylation of eEF2 is regulated by insulin, glutamate and
β-adrenergic agonists (reviewed in [34]). Interestingly, phosphorylation of eEF2 is important in hibernating ground
squirrels as part of their systematic down-regulation of
energy-consuming cellular activities [35]. In yeast, phosphorylation of eEF2 is mediated by the Rck2 kinase and can
be induced by osmotic shock [36].
The diphthamide is the exclusive cellular substrate for
irreversible inactivation by DT (diphtheria toxin) from
Corynebacterium diphtheriae and ETA (exotoxin A) from the
otherwise unrelated Pseudomonas aeruginosa [37,38]. Besides
sharing the eEF2 diphthamide as substrate, the two catalytic
domains of toxins can be aligned to a root mean square
deviation of 2.1 Å and, in particular, have a common core fold
around the NAD-binding site [39]. Both catalyse the transfer
of ADP-ribose from NAD+ to the NE2 atom of the diphthamide imidazole ring in eEF2 [11,12]. ETA is one of several
P. aeruginosa virulence factors, and infections with P. aeruginosa are often observed in immunocompromised patients,
where pronounced multidrug resistance makes antibiotic
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treatment difficult (reviewed in [40]). C. diphtheriae was one
of the first toxin-producing bacteria to be discovered, and
infection was previously considered as one of the most serious
childhood diseases. Although most industrialized countries
today have widespread immunization against diphtheria,
some developing countries still experience outbreaks [41].
The mechanism for the ADP-ribosylation of the diphthamide by the toxin is still unclear. Several studies have shown
that mutations of either the diphthamide or the proteins that
synthesize the diphthamide confer resistance to both DT
and ETA [16,19,23–25]. Kinetic data suggest that the ADPribosylation of eEF2 by ETA or DT is likely to be a SN 1
substitution reaction involving an oxacarbenium cation
[42,43] and results in an inversion of configuration at the
C1 of the ADP-ribose [20]. This indicates that C1 of NAD+
can only be attacked from one side by the nucleophilic
NE2 atom of the diphthamide imidazole. The inversion is
probably caused by the anomeric carbon of the nicotinamide ribose being susceptible to a backside nucleophilic
attack from the NE2 atom in diphthamide after cleavage of the
nicotinamide ribose bond [42]. In addition, kinetic isotope
effect experiments suggest that the complex reaches an
early transition state either by activation of the leaving
nicotinamide group or by increased stabilization of the oxacarbenium ion, which is not due to an increase in the nucleophilic character or participation from NE2 of the diphthamide. The relatively low nucleophilicity of the bulky
diphthamide nucleophile at the proposed transition state
could be a result of steric crowding and activation of transition state formation could result from eEF2 binding, which
then increases the reaction rate [43]. Interestingly, the crystal
structure of the ADP-ribosylated eEF2 showed that the
interaction between Glu553 in ETA (or the corresponding
Glu148 in DT) and the nicotinamide ribose previously determined to be important for catalytic activity [44,45] is
replaced by the highly conserved Asp696 in eEF2 after
ADP-ribosylation (Figure 3) [20]. Near the end of the
catalytic reaction, this interaction of Asp696 in eEF2 with
the 2 -OH of the N-ribose may orient the incoming cation
until the covalent bond with the diphthamide imidazole
is formed. In the structure of the ADP-ribosylated eEF2,
the trimethylammonium group of the diphthamide seems to
interact with the β-phosphate of the ADP-ribose (Figure 3)
and may therefore have an important role in the reaction
mechanism of ADP-ribosylation [20]. This is supported by
previous work showing that the methylation of the diphthamide is very important for the ADP-ribosylating activity of the toxins. The precursor, diphthine, lacking the amide
group, can still be ADP-ribosylated, although at a lower rate
[15,17].
Exactly how ADP-ribosylation of the diphthamide inhibits eEF2 function remains to be determined. Binding
experiments of ADP-ribosylated eEF2 to the ribosome show
a reduction of affinity for the pre-translocational ribosome,
but no changes for the post-translocational ribosome [46].
Other competition and co-sedimentation experiments have
shown that the ADP-ribosylated eEF2 is indeed able to form
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a stable complex with the ribosome [20,47–49]. Another
striking observation was that, while binding of GDP to
wheat germ eEF2 was unaffected by ADP-ribosylation, GTP
binding was significantly reduced, despite being approx. 60 Å
away from the diphthamide [46,50]. These results disagree,
however, with recent fluorescence spectroscopy experiments
of nucleotide binding showing that ADP-ribosylation of
yeast eEF2 has very little, if any, effect in binding both GTP
and GDP [20]. In addition, other binding experiments
have shown that ADP-ribosylated eEF2 still has ribosomedependent GTPase activity and can dissociate from the
ribosome [49].
Future perspectives
Although we have increased our understanding of eEF2
function, the exact mechanism of translocation remains to
be determined, in particular how the energy derived from
GTP hydrolysis is utilized for translocation. If the mechanism
of translocation in eukaryotes and bacteria is conserved,
hydrolysis of GTP by eEF2 is a quite early event and is
used to greatly accelerate translocation. We do not know
what triggers the GTP hydrolysis, but the sarcin–ricin loop is
perhaps the only ribosomal element that comes close enough
to interact with the nucleotide-binding pocket in eEF2 on the
ribosome and therefore could be responsible for triggering
the activity. Advanced kinetic studies similar to those conducted with great success for bacterial translocation are
required urgently for a thorough mechanistic understanding
of the translocation reaction in eukaryotes. In particular, such
experiments may be required to elucidate the exact function of
the diphthamide, which, because of its complicated synthetic
pathway, surely must have an important function in the
cells. The fact that the DPH1 gene has been shown to be
a tumour-suppressor gene, playing an important role in cell
proliferation, indicates that there could be a link between
diphthamide and the control of cell growth.
To obtain a better understanding of how ETA and DT
inactivates the elongation factor, it will be necessary to
carry out further kinetic and structural studies of the interaction between eEF2 and the ribosylating toxins. Finally,
a detailed understanding of how ADP-ribosylated eEF2
inhibits translation may require cryo-EM reconstructions of
the modified factor in complex with the 80 S ribosome.
After submission of this paper, the structure of a
mutant bacterial EF2 in complex with GDPNP has been
published, and shows only very limited conformational
changes compared with the structure of the EF2–GDP
complex [51]. The recent structure of eEF2 in complex with
exotoxin A explains the requirement of the diphthamide for
ADP-ribosylation of eEF2 [52].
References
1 Merrick, W.C. and Nyborg, J. (2000) in Translational control of gene
expression (Sonenberg, N., Hershey, J.W.B. and Mathews, M.B.,
eds.), pp. 89–126, Cold Spring Harbor Laboratory Press,
Cold Spring Harbor
2 Jorgensen, R., Ortiz, P.A., Carr-Schmid, A., Nissen, P., Kinzy, T.G. and
Andersen, G.R. (2003) Nat. Struct. Biol. 10, 379–385
3 Laurberg, M., Kristensen, O., Martemyanov, K., Gudkov, A.T., Nagaev, I.,
Hughes, D. and Liljas, A. (2000) J. Mol. Biol. 303, 593–603
4 Rodnina, M.V., Savelsbergh, A., Katunin, V.I. and Wintermeyer, W. (1997)
Nature (London) 385, 37–41
5 Peske, F., Matassova, N.B., Savelsbergh, A., Rodnina, M.V. and
Wintermeyer, W. (2000) Mol. Cell 6, 501–505
6 Savelsbergh, A., Katunin, V.I., Mohr, D., Peske, F., Rodnina, M.V. and
Wintermeyer, W. (2003) Mol. Cell 11, 1517–1523
7 Spahn, C.M., Gomez-Lorenzo, M.G., Grassucci, R.A., Jorgensen, R.,
Andersen, G.R., Beckmann, R., Penczek, P.A., Ballesta, J.P. and Frank, J.
(2004) EMBO J. 23, 1008–1019
8 Valle, M., Zavialov, A., Sengupta, J., Rawat, U., Ehrenberg, M. and
Frank, J. (2003) Cell 114, 123–134
9 Agrawal, R.K., Heagle, A.B., Penczek, P., Grassucci, R.A. and Frank, J.
(1999) Nat. Struct. Biol. 6, 643–647
10 Agrawal, R.K., Penczek, P., Grassucci, R.A. and Frank, J. (1998) Proc. Natl.
Acad. Sci. U.S.A. 95, 6134–6138
11 Van Ness, B.G., Howard, J.B. and Bodley, J.W. (1980) J. Biol. Chem. 255,
10710–10716
12 Van Ness, B.G., Howard, J.B. and Bodley, J.W. (1980) J. Biol. Chem. 255,
10717–10720
13 Moehring, J.M., Moehring, T.J. and Danley, D.E. (1980) Proc. Natl.
Acad. Sci. U.S.A. 77, 1010–1014
14 Dunlop, P.C. and Bodley, J.W. (1983) J. Biol. Chem. 258, 4754–4758
15 Moehring, T.J., Danley, D.E. and Moehring, J.M. (1984) Mol. Cell. Biol. 4,
642–650
16 Chen, J.Y., Bodley, J.W. and Livingston, D.M. (1985) Mol. Cell. Biol. 5,
3357–3360
17 Chen, J.Y. and Bodley, J.W. (1988) J. Biol. Chem. 263, 11692–11696
18 Liu, S., Milne, G.T., Kuremsky, J.G., Fink, G.R. and Leppla, S.H. (2004)
Mol. Cell. Biol. 24, 9487–9497
19 Moehring, J.M. and Moehring, T.J. (1988) J. Biol. Chem. 263,
3840–3844
20 Jorgensen, R., Yates, S.P., Teal, D.J., Nilsson, J., Prentice, G.A., Merrill, A.R.
and Andersen, G.R. (2004) J. Biol. Chem. 279, 45919–45925
21 Schultz, D.C., Vanderveer, L., Berman, D.B., Hamilton, T.C., Wong, A.J. and
Godwin, A.K. (1996) Cancer Res. 56, 1997–2002
22 Chen, C.M. and Behringer, R.R. (2004) Genes Dev. 18, 320–332
23 Foley, B.T., Moehring, J.M. and Moehring, T.J. (1995) J. Biol. Chem. 270,
23218–23225
24 Moehring, T.J., Danley, D.E. and Moehring, J.M. (1979)
Somatic Cell Genet. 5, 469–480
25 Phan, L.D., Perentesis, J.P. and Bodley, J.W. (1993) J. Biol. Chem. 268,
8665–8668
26 Fendrick, J.L. and Iglewski, W.J. (1989) Proc. Natl. Acad. Sci. U.S.A. 86,
554–557
27 Lee, H. and Iglewski, W.J. (1984) Proc. Natl. Acad. Sci. U.S.A. 81,
2703–2707
28 Sayhan, O., Ozdemirli, M., Nurten, R. and Bermek, E. (1986)
Biochem. Biophys. Res. Commun. 139, 1210–1214
29 Sitikov, A.S., Davydova, E.K. and Ovchinnikov, L.P. (1984) FEBS Lett. 176,
261–263
30 Ryazanov, A.G. (1987) FEBS Lett. 214, 331–334
31 Donovan, M.G. and Bodley, J.W. (1991) FEBS Lett. 291, 303–306
32 Price, N.T., Redpath, N.T., Severinov, K.V., Campbell, D.G., Russell, J.M.
and Proud, C.G. (1991) FEBS Lett. 282, 253–258
33 Dumont-Miscopein, A., Lavergne, J.P., Guillot, D., Sontag, B. and Reboud,
J.P. (1994) FEBS Lett. 356, 283–286
34 Browne, G.J. and Proud, C.G. (2002) Eur. J. Biochem. 269, 5360–5368
35 Chen, Y., Matsushita, M., Nairn, A.C., Damuni, Z., Cai, D., Frerichs, K.U.
and Hallenbeck, J.M. (2001) Biochemistry 40, 11565–11570
36 Teige, M., Scheikl, E., Reiser, V., Ruis, H. and Ammerer, G. (2001)
Proc. Natl. Acad. Sci. U.S.A. 98, 5625–5630
37 Iglewski, B.H. and Kabat, D. (1975) Proc. Natl. Acad. Sci. U.S.A. 72,
2284–2288
38 Pappenheimer, Jr, A.M. (1977) Annu. Rev. Biochem. 46, 69–94
39 Bell, C.E. and Eisenberg, D. (1997) Adv. Exp. Med. Biol. 419, 35–43
40 Lyczak, J.B., Cannon, C.L. and Pier, G.B. (2000) Microbes Infect. 2,
1051–1060
41 Mattos-Guaraldi, A.L., Moreira, L.O., Damasco, P.V. and Hirata Junior, R.
(2003) Mem. Inst. Oswaldo Cruz 98, 987–993
42 Beattie, B.K., Prentice, G.A. and Merrill, A.R. (1996) Biochemistry 35,
15134–15142
43 Parikh, S.L. and Schramm, V.L. (2004) Biochemistry 43, 1204–1212
C 2006
Biochemical Society
5
6
Biochemical Society Transactions (2006) Volume 34, part 1
44
45
46
47
48
Carroll, S.F. and Collier, R.J. (1987) J. Biol. Chem. 262, 8707–8711
Douglas, C.M. and Collier, R.J. (1987) J. Bacteriol. 169, 4967–4971
Nygard, O. and Nilsson, L. (1990) J. Biol. Chem. 265, 6030–6034
Bermek, E. (1976) J. Biol. Chem. 251, 6544–6549
Davydova, E.K. and Ovchinnikov, L.P. (1990) FEBS Lett. 261,
350–352
49 Nolan, R.D., Grasmuk, H. and Drews, J. (1976) Eur. J. Biochem. 64,
69–75
C 2006
Biochemical Society
50 Burns, G., Abraham, A.K. and Vedeler, A. (1986) FEBS Lett. 208, 217–220
51 Hansson, S., Singh, R., Gudkov, A.T., Liljas, A. and Logan, D.T. (2005)
FEBS Lett. 579, 4492–4497
52 Jorgensen, R., Merrill, A.R., Yates, S.P., Marquez, V.E., Schwan, A.L.,
Boesen, T. and Andersen, G.R. (2005) Nature (London) 436, 979–984
Received 20 June 2005