- Email: [email protected]
NTF2 monomer-dimer equilibrium1
doi:10.1006/jmbi.2001.5136 available online at http://www.idealibrary.com on
J. Mol. Biol. (2001) 314, 465±477
NTF2 Monomer-Dimer Equilibrium Catherine Chaillan-Huntington1, P. Jonathan G. Butler1 James A. Huntington2, Debra Akin3, Carl Feldherr3 and Murray Stewart1* 1
MRC, Laboratory of Molecular Biology, Hills Road, Cambridge CB2 2QH, England 2
University of Cambridge Department of Haematology Wellcome Trust Centre for Molecular Mechanisms in Disease, Addenbrooke's Hospital Site, Hills Road Cambridge CB2 2XY, England 3
Department of Anatomy and Cell Biology, University of Florida College of Medicine Gainesville, FL 32610, USA
Nuclear transport factor 2 (NTF2) mediates nuclear import of RanGDP, a central component of many nuclear traf®cking pathways. NTF2 is a homodimer and each chain has independent binding sites for RanGDP and nuclear pore proteins (nucleoporins) that contain FxFG sequence repeats. We show here that the monomer-dimer dissociation constant for NTF2 obtained by sedimentation equilibrium ultracentrifugation is in the micromolar range, indicating that a substantial proportion of cellular NTF2 may be monomeric. To investigate the functional signi®cance of NTF2 dimerization, we engineered a series of point mutations at the dimerization interface and one of these (M118E) remained monomeric below concentrations of 150 mM. CD spectra and X-ray crystallography showed that M118E-NTF2 preserved the wild-type NTF2 fold, although its thermal stability was 20 deg. C lower than that of the wild-type. M118E-NTF2 bound both RanGDP and FxFG nucleoporins less strongly, suggesting that dissociation of the NTF2 dimer could facilitate RanGDP release and thus nucleotide exchange after it had been transported into the nucleus. Moreover, colloidal gold coated with M118E-NTF2 showed reduced binding to Xenopus oocyte nuclear pores. Overall, our results indicate that dimer formation is important for NTF2 function and give insight into the formation of heterodimers by mRNA export factors such as TAP1 and NXT1 that contain NTF2-homology domains. # 2001 Academic Press
*Corresponding author
Keywords: nuclear traf®cking; protein structure; crystallography; dimerization
Introduction The nuclear pore complexes (NPCs) of the nuclear envelope mediate bi-directional traf®cking of macromolecules between cytoplasm and nucleus.1 ± 4 NPCs are huge macromolecular assemblies constructed from 50 different proteins collectively called nucleoporins,5,6 many of which interact with soluble components of the nucleocytoplasmic transport machinery.7 Many nucleoporins contain tandem sequence repeats based on Present address: C. Chaillan-Huntington, Cambridge Antibody Technology, Science Park, Melbourn, Cambridgeshire SG8 6JJ, England. Abbreviations used: NPC, nuclear pore complex; NTF2, nuclear transport factor 2; NSP1-FF18, NSP1 protein construct containing 18 FxFG repeats; PBS, phosphate-buffered saline; PBST, PBS containing 0.1% (v/v) Tween 20; tm, melting point temperature ( C). E-mail address of the corresponding author: [email protected] 0022-2836/01/30465±13 $35.00/0
FxFG or GLFG cores joined by hydrophilic linkers of variable sequence and length.7,8 Generally, traf®cking of macromolecules such as proteins and nucleic acids is mediated by carriers that are usually members of the importin-b superfamily.1 ± 4 In addition to the importin-family carriers and nucleoporins, the Ras-family GTPase, Ran, and nuclear transport factor 2 (NTF2) are required for ef®cient transport. Ran is a key component of the nuclear traf®cking machinery and also appears to be involved in cellBecause RanGAP1 cycle progression.1 ± 4,9,10 (RanGTPase activating protein) is located in the cytoplasm, whereas the Ran guanine nucleotide exchange factor (RCC1) is located in the nucleus, it is thought that cytoplasmic Ran is primarily GDPbound (RanGDP), whereas nuclear Ran is in the GTP-bound form (RanGTP).11,12 This putative RanGTP gradient between the cytoplasm and nucleoplasm has been proposed to be the basis for determining the directionality of the transport of macromolecules.12 Cargo proteins carrying nuclear # 2001 Academic Press
466 localization signals bind to carrier molecules of the importin-b family in the cytoplasm and are then translocated into the nucleus through nuclear pore complexes, after which the carrier-cargo complex is dissociated by RanGTP. The carrier is then recycled to the cytoplasm bound to RanGTP where RanGAP generates RanGDP, which dissociates from the carrier, freeing it to interact with another cargo molecule and undergo a further import cycle.1 ± 4 The cycling of carriers between nucleus and cytoplasm, and their binding and release of cargo molecules in the appropriate compartment is thus intimately tied to the Ran nucleotide state. Nuclear protein import mediated by importin-b13 and the NTF2-mediated import of RanGDP14 both require a direct interaction between the carrier and FxFG nucleoporins, and analogous nucleoporin-carrier interactions are thought to be involved in other nuclear traf®cking pathways (e.g. see Damelin & Silver15). The nuclear import of RanGDP is mediated by NTF2.16,17 NTF2 is a small homodimeric protein that binds both RanGDP and FxFG repeatcontaining nucleoporins on separate sites.3,18 NTF2 is localized primarily to the nuclear rim,19,20 with lower concentrations in the nucleus and the cytoplasm. NTF2 is an essential protein in yeast21,22 and Caenorhabditis elegans,9 and dominant negative mutants can disrupt nuclear protein import selectively.23 Studies using both Ran and NTF2 engineered mutants have shown that both the NTF2-RanGDP and NTF2-FxFG interactions are required for ef®cient RanGDP import.14,16,17 The Kd values for the interactions of NTF2 with RanGDP or with FxFG repeat nucleoporins are 100 nM and 1 mM, respectively.14,20 On the basis of these values, we recently proposed a model for Ran import where the Ran/NTF2 complex would remain intact during the translocation across the nuclear envelope, ``hopping'' from one FxFG nucleoporin to the next one along the NPC channel.20 We also showed that there was no cooperativity between the interactions involving RanGDP, NTF2, and FxFG repeats.20 Several proteins involved in mRNA nuclear export contain domains that are homologous to NTF2 and that almost certainly share the same fold.24 Thus, vertebrate TAP1 and its yeast counterpart Mex67 have N-terminal RNA-binding domains, followed by four leucine-rich repeats, a domain homologous to NTF2 and ®nally a C-terminal UBA domain.24 ± 26 TAP1 interacts with the small 15 kDa monomeric protein NXT1 (also called p15) that also has an NTF2 fold,24,26 ± 28 and it has been proposed that the formation of the TAP1/ NXT1 heterodimer is mediated by an interaction analogous to that seen between the two chains in the NTF2 homodimer.24 Moreover, NXT1/p15 binds to Ran (albeit in its GTP-bound form) and both Mex67 and TAP1 bind to nucleoporins.29,30 In the case of Mex67, nucleoporin binding appears to require the formation of a heterodimer with Mtr2, which seems to perform a role analogous to NXT1.
NTF2 Monomer-Dimer Equilibrium
Although NTF2 has been implicitly assumed to be a dimer in vivo, the observations with the NTF2 homologues involved in mRNA nuclear export raise the possibility that NTF2 itself may be at least partially dissociated. We therefore determined Kd for the NTF2 monomer-dimer equilibrium and show here that it is in the micromolar range, which is of the same order as the cellular concentration of NTF2,20 indicating that a substantial proportion of NTF2 may be monomeric in vivo. To investigate the functional signi®cance of NTF2 dissociation, we examined its in¯uence on the interaction with RanGDP and FxFG nucleoporins. To do this, we weakened the interaction between NTF2 chains at the dimer interface by mutating methionine residues 84, 102 and 118 to glutamic acid. Dimer formation was reduced substantially in the M118E NTF2 mutant, which also showed a 20 deg. C decrease in thermal stability compared with wildtype NTF2 but no signi®cant alteration of its overall fold in solution. This mutant showed reduced af®nity for both RanGDP and FxFG nucleoporin repeats, and the binding of M118E-NTF2 to Xenopus oocyte NPCs was reduced. Moreover, the results obtained with other mutations at the dimerization interface showed unanticipated ways in which NTF2 dimerization could be stabilized and gave insight into the putative formation of heterodimers by components of the Tap/Mex67 mRNA export machinery.
Results Monomer-dimer equilibrium of NTF2 The aggregation state of NTF2 was examined by sedimentation equilibrium ultracentrifugation, loading samples with initial A280 values of 0.8, w;app against concentration 0.4 and 0.2. Plots of M (Figure 1(a), upper plot) showed that wild-type NTF2 was primarily dimeric at concentrations above 5 mM, but there was clear evidence for dissociation into monomers at lower concentrations. The data obtained from different loading concentrations were superimposed and suggested that the NTF2 dimers were in a rapidly reversible equilibrium with monomers. To better de®ne the concentration range where monomeric NTF2 dominated, sedimentation equilibrium data were obtained with lower loadings, employing concentrations of NTF2 corresponding to initial A230 values of 0.3, 0.2 and 0.1. The results with lower initial NTF2 concentrations (Figure 1(a), lower plot), were very similar to those obtained previously, and clear evidence for dissociation can be seen at NTF2 concentrations below 4 mM. A model for an ideal monomer/dimer equilibrium gave a good ®t to the data of concentration against radius, and examples of plots of the residuals for the ®tting are shown in (Figure 1(b)). This model gave an estimate of the NTF2 dimer dissociation constant (Kd) of 1.1 (0.05) mM.
NTF2 Monomer-Dimer Equilibrium
467
Figure 1. Sedimentation equilibrium analysis of NTF2 monomer-dimer equilibrium. (a) Plots of Mw;app against concentration, derived from cells loaded at different initial concentrations (see Materials and Methods), together with curve ®tted for a monomer-dimer equilibrium. (b) Plots of residuals for direct ®tting of concentration against radius data with model for monomer/dimer equilibrium. The upper plots correspond to the early loading concentrations, while the lower plots correspond to lower loading concentrations employed to obtain a better estimate of Kd, by working over an appropriate concentration range in the cells. These data indicate a Kd of 1 mM for the wt-NTF2 monomer-dimer dissociation constant.
Point mutants at the NTF2 dimerization interface The dissociation constant (Kd) value of 1.1 (0.05) mM is comparable to the concentration for NTF2 in the nucleoplasm (0.6 mM) and the cytoplasm (0.3 mM),20 which suggests that a substantial proportion of NTF2 may be monomeric in vivo. This raised the possibility that dissociation into monomers might in¯uence the interaction of NTF2 with Ran or FxFG nucleoporins. However, it was technically dif®cult to investigate this possibility experimentally because the dissociation constant of the NTF2 dimer was of the same order as its binding constant for RanGDP and FxFG nucleoporins.20 We therefore attempted to produce primarily monomeric NTF2 by introducing point mutations in the NTF2 dimer interface. The crystal structure of NTF231 shows that the interface between chains in the NTF2 dimer is primarily hydrophobic and is formed by the close juxaposition of extensive bsheets from each molecule. Methionine residues 84,
102 and 118 are located within the dimerization interface (Figure 2(a) and (b)) and are in close apposition with hydrophobic residues from the neighbouring chain. We therefore anticipated that mutating these methionine residues to glutamate would reduce the contribution of the hydrophobic effect to dimerization and so produce primarily monomeric NTF2 in solution at concentrations suf®ciently high to measure the association with RanGDP and FxFG nucleoporins directly. All three mutants were assessed by sedimentation equilibrium ultracentrifugation to determine their dissociation constants (Figure 3). Somewhat surprisingly, the effect of the M84E and M102E mutations was slight and, although they had Kd values of 5 (0.5) and 20 (1) mM, respectively, these mutants remained primarily dimeric at the concentrations needed to assess binding to RanGDP and FxFG nucleoporins. However, the M118E mutant had a much more dramatic effect and appeared monomeric over the entire
468
NTF2 Monomer-Dimer Equilibrium
Figure 2. Location of M84, M102 and M118 at the NTF2 dimerization interface. An illustration of the side-chains protruding into the interaction interface between NTF2 chains in the dimer. Methionine residues 84, 102 and 118 are all important components of the hydrophobic centre of the interaction interface. Hydrophobic residues are in grey, acidic residues in red, basic residues in blue and amphipolar residues in white. Also shown are the corresponding side-chains in the putative structures (see Suyama et al.24) of TAP1 and NXT1. Note that neither TAP1 nor NXT1 contains the equivalent of His100 in NTF2.
concentration range examined and remained monomeric even with initial concentrations as high as A280 0.8. The data shown in Figure 3(a) (bottom plot) indicated that M118E-NTF2 was monomeric up to concentrations of at least 150 mM, while the residual plot (Figure 3(b), lowest plot) showed a good ®t for a non-ideal monomer, with the osmotic second virial coef®cient (B) estimated as 0.0043 Mÿ1. We can therefore conclude that wild-type NTF2 is primarily dimeric at concentrations above 2-4 mM, whereas mutants within the dimerization interface show varying degrees of dissociation. In particular, M118E-NTF2 was essentially monomeric below 150 mM. Crystal structures of M84E-, M102E- and M118E-NTF2 For all the mutants, the crystal structure shows dimers with a Ca backbone similar to the wild-type molecule. The crystal structure of M84E-NTF2 showed putative salt-bridges between His100 and Glu84 (Figure 4(a)) in both chains, which could help mitigate the introduction of a charged residue into the hydrophobic interface. In M102E- and M118E-NTF2, steric constraints prevented the formation of saltbridges between His100 and Glu102 in different chains, although a putative salt-bridge was formed between these residues in the same chain (data not shown, but can be seen in PDB deposition 1JB4). In addition, Glu118 was placed very close to other acidic side-chains and the resulting repulsion would decrease the stability of the interface (this would be especially true at physiological pH, but was probably diminished in the acidic conditions (pH 4.5) needed to produce the crystals of this mutant). Thus, the crystal structures of the three mutants can account for the different Kd values observed for dimerization. Comparison of the three dimer structures indicated that, although the NTF2 fold had been retained in each case, each of the mutations had also introduced a small rigid-body rotation
between the two chains in the dimer (Figure 5). When RanGDP binds to NTF2 there is also a small rigid-body rotation of one NTF2 chain relative to the other (see Stewart et al.32 and Figure 5), albeit in the direction opposite to that seen with the NTF2 mutants in the present study. Overall, it seems that there is considerable plasticity to the NTF2 dimerization interface and this accounts for how small rearrangements in M84E-NTF2, for example, can enable formation of a putative salt-bridge to His100 and so mitigate the effect of introducing a charged residue into the primarily hydrophobic interface. The observation that M118E-NTF2 crystallized as a dimer indicated that the M118E mutation does not abolish the dimerization completely, at least under the acidic conditions (pH 4.5) used to grow the crystals. The sedimentation equilibrium data indicated that, in contrast to M84E and M102E, the M118E mutation perturbs dimerization so that dimeric NTF2 could not be detected at concentrations up to 150 mM. However, it would appear that dimerization of this mutant was still possible in the unusual solvent conditions and the very high concentrations used for crystallization (of the order of millimolar). One function of the PEG used as precipitant, for example, is to increase the activity of macromolecular species due to the excluded volume effect.33 Moreover, the M118ENTF2 crystals were obtained only at pH 4.5, where the ionization of the introduced Glu would be much less than at physiological pH. At lower concentrations (0.01-0.1 mM) in a more physiological solvent, sedimentation equilibrium data indicated that M118E-NTF2 was monomeric. Under these conditions, circular dichroism (Figure 6(a)) indicated that the secondary structure content of M118E mutant was very similar to that of wildtype NTF2, strongly suggesting that the structure of the individual polypeptide chains had not been altered radically when M118E-NTF2 dissociated into monomers. Circular dichroism (CD) spectroscopy also showed that the thermal stability of the different
469
NTF2 Monomer-Dimer Equilibrium
Figure 3. Sedimentation equilibrium analysis of NTF2 mutants. (a) Plots of Mw;app against concentration for mutants, together with curves ®tted either for a monomer/dimer equilibrium (M84E-NTF2 and M102E-NTF2) or for a non-ideal monomer (M118E-NTF2). (b) Plots of residuals for ®tting of concentration against radius data for mutants with models either for a monomer/dimer equilibrium (M84E-NTF2 and M102E-NTF2) or for a non-ideal monomer equilibrium (M118E-NTF2). These data show that M118E-NTF2 was present as monomers at concentrations up to 150 mM.
NTF2 mutants correlated with the strength of their dimerization. Sedimentation equilibrium ultracentrifugation had indicated that, with the protein concentrations employed (35 mM), wild-type NTF2 was present as a dimer, M84-NTF2 and M102NTF2 were partially dissociated and M118E-NTF2 was monomeric. Figure 6 shows that M118E-NTF2 was considerably less stable than wild-type, with a tm value of 51 C compare to 71 C for wild-type NTF2, whereas both the M84E- and M102Emutants showed an intermediate lowering of tm (Table 1).
Influence of NTF2 dimerization on the interaction with RanGDP and FxFG nucleoporins Previous studies have indicated that binding to RanGDP and FxFG nucleoporins are both crucial for NTF2 function.14,16,18,20 We therefore investigated the in¯uence of dimerization on these properties using bead-binding assays. M118E-NTF2 was tested for its ability to be pulled down by Sagarose beads coated with either S-tagged RanGDP or S-tagged Nsp1-FF18 (a construct that contains
470
NTF2 Monomer-Dimer Equilibrium
Figure 4. Putative Glu84-His100 salt-bridge stabilizes the M84E-NTF2 dimer. A portion of the 2Fo ÿ Fc electron density map of the crystal structure of M84E-NTF2 showing putative interactions between Glu84 and His100 that could contribute to the stability of the dimerization interface. The A chain is yellow and the B chain is blue. The interaction between chains is mediated by a water molecule.
18 FxFG repeats of the yeast nucleoporin NSP114,20,34,35) and showed reduced binding to both RanGDP and FxFG nucleoporins compared with wild-type NTF2 (Figure 7). We also tested the ability of M118E-NTF2 to associate with NPCs in vivo by microinjecting colloidal gold coated with either this mutant or wild-type NTF2 into Xenopus oocytes. Electron micrographs (Figure 8) showed that, as observed previously,14,36 colloidal gold coated with wild-type NTF2 bound to the cytoplasmic and nucleoplasmic faces of NPCs, as well as to their central transport channel. By contrast, colloidal gold particles coated with M118E-NTF2 showed greatly reduced binding to NPCs when injected into Xenopus oocytes: there was only slight staining of the nucleoplasmic and cytoplasmic faces of NPCs and very few particles were seen in the central transport channel. The results of these binding studies imply that either the binding of both RanGDP and FxFG nucleoporins is sensitive to the monomer-dimer
Table 1. Thermal unfolding of NTF2 mutants Protein Wild-type NTF2 M84E-NTF2 M102E-NTF2 M118E-NTF2
tm( C) 71 61 61 51
Figure 5. NTF2 rigid-body rotations. Illustration of the rigid-body rotations observed between (black) wildtype NTF2 alone31 and (blue) bound to RanGDP32 and the M84E mutants (red). In each case, the A chains have been superimposed, and so the difference in positions of the B-chains indicates the relative movement between the chains in the dimer.
state of NTF2 or that we have altered the binding sites for both by our mutation. Met118 is located some distance from the RanGDP binding site on NTF232 and, although the precise binding site for FxFG nucleoporins has not been established directly, it appears to be near Trp7,14 which is also a considerable distance from Met118. Therefore, it appeared more likely that the reduced af®nity seen in our binding assays was a result of the monomeric nature of the M118E-NTF2.
Discussion Previous studies using X-ray crystallography,31,32 gel-®ltration37 and NTF2 covalently coupled to Sepharose beads34,35 had all indicated that NTF2 is a homodimer. However, our ultracentrifugation sedimentation equilibrium data (Figure 1) established a dissociation constant for the NTF2 dimer to be Kd 1.1 (0.05) mM. A dissociation constant of this magnitude implies that a considerable degree of NTF2 monomer could be present at concentrations of the order of micromolar or 0.015 mg/ml. Previous in vitro studies had been performed at concentrations of the order of 1 mg/ ml, where NTF2 would indeed be primarily dimeric. However, cellular concentrations of NTF2 are considerably lower than those used in these in vitro studies. Previous results have shown that NTF2 is localized primarily at the nuclear envelope, where it is bound to FxFG
NTF2 Monomer-Dimer Equilibrium
Figure 6. M118E-NTF2 retains its fold in solution at monomeric concentration, but shows a decrease in thermal stability. (a) The CD spectra of wt-NTF2 (continuous line) and M118E-NTF2 (broken line) in solution were indistinguishable, indicating that the M118E mutant had not introduced a major conformational change. These spectra were recorded at a concentration of 35 mM, where sedimentation equilibrium data (Figure 3) indicated that M118E-NTF2 was dissociated into monomers. (b) Thermal stability of wt-NTF2 and the M84E, M102E and M118E mutants determined by CD. The data represent the change in ellipticity (, mdeg) as a function of temperature. The denaturation temperature was taken at the in¯exion point of the rising portion of the curve.
nucleoporins.14,19,20,38 The concentration of NTF2 at the nuclear envelope is of the order of 20 mM,20 but the cytoplasmic and nucleoplasmic concentrations of NTF2 are only of the order of 0.3 mM and 0.6 mM respectively.20 A micromolar dissociation constant therefore would indicate that, although most of the NTF2 at the nuclear pores would be dimeric, a substantial proportion of NTF2 in the bulk of the nucleoplasm and cytoplasm could be monomeric. Therefore, except at the nuclear envelope, a considerable dissociation of dimeric NTF2 into monomers could occur in vivo. This possibility raises the question of the importance of dimerization for NTF2 function.
471 Because the dissociation constant of NTF2 dimers was of the same order of magnitude as that observed for RanGDP and FxFG nucleoporin binding,14,20 we engineered a point mutant (M118E) at the dimerization interface (Figure 2(a) and (b)) that increased the degree of dissociation so that it was monomeric at concentrations up to 150 mM. Although M118E-NTF2 was still able to bind Ran GDP and FxFG nucleoporin repeats, it did so with reduced af®nity, indicating that the oligomeric state of NTF2 in¯uenced its af®nity for both of its substrates. The af®nity of NTF2 for its substrates is crucial for its function.20 Thus, NTF2 has to bind RanGDP suf®ciently strongly to transport it through nuclear pores, yet suf®ciently weakly to release it in the nucleus to enable it to be recharged with RanGTP by RCC1. Although the concentration of NTF2 at the nuclear pores themselves would result in NTF2 being primarily dimeric, the lower NTF2 concentration in the nucleus would favour its dissociation into monomers. Reducing its af®nity for RanGDP in the nucleus through dissociation of the NTF2 dimer could be important in promoting release of RanGDP once it had been transported through the nuclear pore, especially since the NTF2-RanGDP complex has been shown to inhibit nucleotide exchange by RCC1.19 The oligomeric state of NTF2 also in¯uenced the thermal stability of the molecule (Figure 6), so that monomeric M118E-NTF2 showed a decreased stability (51 C) compared to wild-type (71 C). Surprisingly M84E-NTF2 and M102E-NTF2 were still primarily dimeric in solution, although in each case the mutant had a higher Kd than wild-type (5(0.5) and 20(1) mM, respectively). The crystal structures of the mutants gave a rationale for their unexpected retention of dimerization. In both mutants, local rearrangements, involving rigidbody rotation to the two chains coupled with the formation of salt-bridges or hydrogen bonds between the introduced glutamate residue and His100, offset the introduction of a hydrophilic residue into the hydrophobic dimer interface (Figure 5). Thus, although these mutants produced a small but signi®cant rigid-body rotation of the two NTF2 chains, they did not reduce their af®nity suf®ciently to generate a substantial population of monomers under the conditions used to assay for RanGDP and FxFG nucleoporin binding. However in the M118E-NTF2 structure, Glu118 was less able to form a salt-bridge with His100 from the other chain, and so a greater degree of dissociation was produced by this mutation. Interestingly, a yeast analogue of human M84E-NTF2 (yeast M83TNTF2) has a temperature-sensitive phenotype.21,39 Although the crystal structure of yeast wild-type NTF2 has not been determined, it is likely that its structure is very similar to that of the rat protein.39 A plausible explanation of the temperature-sensitive phenotype of the yeast M83T mutation is that, unlike the mutation to glutamate, threonine would not be able to form a compensatory salt-bridge
472
Figure 7. Interaction of M118E-NTF2 with RanGDP and FxFG nucleoporins. S-tagged Ran-GDP or Nsp1-FF18 attached to S-agarose beads were incubated with wild-type or M118E-NTF2 and, after centrifugation and washing, the amount of material bound was assessed by SDS-PAGE. Although M118E-NTF2 bound to both RanGDP and NSP1-FF18, it did so with lower af®nity than wild-type NTF2. Because the af®nity of NTF2 for FxFG nucleoporins was lower than for RanGDP,20 we included W7A-NTF2 (which does not bind FxFG nucleoporins, Bayliss et al.14) as a negative control.
with His100 and so would increase the level of dissociation of the NTF2 dimer. Our CD studies (Figure 6) indicated that the temperature-stability of NTF2 monomers is substantially less than that of the wild-type and so would be consistent with the observed temperature-sensitive phenotype of yeast M83T-NTF2. The results of our mutagenesis studies of the NTF2 dimerization interface provide a structural context for evaluating the heterodimerization of NTF2 homologues involved in mRNA export. Both
NTF2 Monomer-Dimer Equilibrium
TAP1 and NXT1 show suf®ciently strong sequence homology to NTF2 to make it unlikely that they do not also have the same fold and indeed the formation of an NXT1-TAP1 heterodimer has been proposed to be important for their function.24 It has been proposed that this heterodimer forms by using a hydrophobic interface constructed from the extensive b-sheet of the NTF2 fold (Figure 2) and a number of mutations have been engineered into both TAP1 and NXT1 to attempt to evaluate this hypothesis.24 However, although some of these mutants have interfered with dimer formation, others produced a more marginal effect. Moreover, some of the mutations that reduced the TAP1NXT1 interaction (for example, I95D, F109Y and F133D in p15 and L466H, F479W and F499D in Tap) were not in the putative dimerization interface. Although not always conserved as methionine, the residue corresponding to Met84 in vertebrate NTF2 is invariably hydrophobic in TAP1, Mex67 and NXT1, and mutations here (F465H in TAP1; V94D in NXT1) reduce heterodimer formation. However, both TAP1 and NXT1 lack a histidine residue corresponding to NTF2 His100, and so cannot form a compensating saltbridge to stabilize dimerization. At the position corresponding to NTF2 Met102, TAP1 and NXT1 have polar residues (T, N in NXT1, D in TAP1, D or E in Mex67). At the position corresponding to NTF2 Met118, mutations from the conserved hydrophobic residue to Asp (analogous to our M118E mutant) in either TAP1 (V498D) or NXT1 (C132D) inhibit heterodimerization. Our crystal
Figure 8. Electron micrographs of sections of embedded Xenopus oocytes microinjected with colloidal gold (black dots) coated with (a) wild-type or (b) M118E-NTF2. Whereas wild-type NTF2 accumulated on both nuclear and cytoplasmic faces of the NPCs as well as in their central transport channel, the degree of labeling observed with the M118E mutant was greatly reduced, showing only weak accumulation on the NPC faces and little material present in the central transport channel. The scale bar represents 1000 nm.
473
NTF2 Monomer-Dimer Equilibrium
structures of the three Met mutants of NTF2 show how rigid-body rotations of the interface can occur to accommodate changes in the dimerization interface and so may account for the failure of some mutations at the putative TAP1-NXT1 interface24 to inhibit heterodimer formation. In summary, we have demonstrated that the NTF2 dimer has a micromolar dissociation constant that is of the order of the cellular concentration of NTF2, raising the possibility that NTF2 may be partially dissociated in vivo. We have engineered point mutants at the NTF2 dimerization interface that increase the degree of dissociation, albeit with two of these (M84E and M102E), compensating salt-bridge formation with His100 partially mitigated the effect of the mutation on interface hydrophobicity and so generated only a small change in Kd. However, M118E-NTF2 remained monomeric below 150 mM, because similar compensating salt-bridges could not form. The heat-denaturation pro®les of the mutants show that the dimerization of NTF2 is important for thermal stability. Moreover, the af®nity of the M118E mutant was reduced for both RanGDP and nucleoporin FxFG repeats. In the context of the concentration of NTF2 in the nucleus, this suggested that dissociation of the NTF2 dimer could speed the release of RanGDP once it had been transported into the nucleus and thus facilitate nucleotide exchange by RCC1. Overall, our results indicate that NTF2 dimerization is important for its function in the nuclear import of RanGDP and are consistent with an analogous TAP1-NXT1 heterodimerization being important for binding to nucleoporins during mRNA export.
successive scans (for each cell) could not be distinguished when overlaid and the later scan was then taken as being operationally at equilibrium. Baselines were measured by overspeeding (to 60,000 rev/min), until the sample had pulled away from the meniscus, and then slowing the centrifuge to 18,000 rev/min before recording a ®nal scan for the baseline. Optical absorbance values were converted to molarities, using the molar extinction coef®cients e230 4.39 104 Mÿ1 and e280 9.98 103 Mÿ1 and allowing for the 12 mm path-length of the cells. The solvent density (r) and partial speci®c volume (n) of NTF2 were calculated from their compositions, as described,41 using the program SEDNTERP (courtesy of John Philo from the RASMB archive), which gave values of 1.0062 g/ml and 0.7345 ml/g, respectively. The data were analyzed by calculating the weight average, apparent molecular mass (Mw;app )) for overlapping sets of 41 consecutive datum points, with the concentration estimated as that of the mid-point, by non-linear regression ®tting (with proFit v.5.1.2, from QuantumSoft, ZuÈrich, Switzerland) to the equation: 2RT w;app d ln cr M dr2 1 ÿ n rw2
where cr is the concentration at radius r, o is the rotational speed (in rad/second), R is the gas constant and T is the temperature (in K). Plots of Mw;app against c were prepared (using proFit) and examined to predict the best model for the dependence. Direct ®tting to the data for c against r, giving a closer approximation to random errors and therefore a better estimation of the parameters,42 was again by non-linear regression (with proFit as above), weighted with the standard deviation of each datum point. For both the model of an ideal monomer/dimer aggregation equilibrium and a non-ideal monomer, the following simpli®cation of the equations was employed:
Experimental Procedures Mutagenesis, protein expression and purification Site-directed mutagenesis of the methionine residues at positions 84, 102 and 118 (ATG codon) to glutamic acid (GAA codon) was performed using the Quick Change site-directed mutagenesis kit (Stratagene, La Jolla, Ca) following the manufacturer's protocol and using oligonucleotides 30 to 37 bases in length. Rat NTF2 (wild-type and mutants), N-terminally S-tagged canine Ran and the 18 FxFG repeat-containing fragment of nucleoporin Nsp1p (NSP1-FF18) were expressed and puri®ed as described.16,20,34 NTF2 concentrations refer to the amount of monomeric chain. Sedimentation equilibrium analysis Sedimentation equilibrium runs were carried out in standard double-sector cells in an An-60 Ti rotor and a Beckman XL-A analytical ultracentrifuge. Samples, in PBS, were analyzed by long column equilibrium at 18,000 rev/min and 5.0 C, with optical absorbance recorded at 230 nm or 280 nm, taking the mean (and standard deviation) of 100 readings for the ®nal data at each radius, using the standard Beckman software. Runs were initially over-speeded, at 25,000 rev/min for six hours, to speed attainment of equilibrium.40 Subsequently, scans were taken at intervals of 24 hours until
1
s
M1 1 ÿ n rw2 2RT
2
where M1 is the monomer concentration and s is a scaling constant. For the monomer/dimer equilibrium, the reference monomer concentration (c1,0), at the reference radius (r0) was calculated from the equation:
c1;0
ÿKd
q Kd2 8Kd A0 ÿ dA=e1 4
3
where Kd is the dissociation coef®cient, A0 is the absorbance at the reference radius (r0), e1 is the monomer extinction coef®cient (and it is assumed that that for the dimer will be twice this) and dA is the baseline absorbance. The absorbance (Ar) at every radius (r) was then calculated from the equation: Ar e1 c1;0 exp s r2 ÿ r20 2 c1;0 exp s r2 ÿ r20 2 Kd
4
dA
For the non-ideal monomer model, the reference concentration (c0) was taken at a reference radius close
474
NTF2 Monomer-Dimer Equilibrium
to the meniscus, so that it had a small value. The absorbance at every radius was then calculated from the equation: Ar e c0 exp s r2 ÿ r20 ÿ 2Bc0 exp s r2 ÿ r20 dA 5 ÿ1
where B is the osmotic second virial coef®cient (in M ). Parameters ®tted included Kd or B, depending upon the model, and a correction factor for M1 (to allow for possible error in n or r) and an error in dA (to allow for errors in measurement). In practice, the ®rst factor was between 0.95 and 1.05, while the second was <0.05 absorbance units. The calculated optical absorbances and residuals for each radius were tabulated at the end of each ®t. The former could be used to calculate and plot the ®tted curve w;app against concentration, for that model, on the of M plot of the experimental values, while the latter was plotted against radius, to con®rm the validity of the ®t to the model. Circular dichroism spectroscopy CD spectra were recorded on a JASCO 810 spectropolarimeter. Spectra in the 200-260 nm range were recorded as an average of three scans. The effect of temperature on changes in protein secondary structure was followed by recording the ellipticity at 222 nm every 0.5 deg. C in a 0.1 cm temperature-controlled cell using a temperature change of 60 deg. C/hour. Before each temperature scan, a spectrum in the range of 200-260 nm was recorded. Protein samples in 100 mM Tris-HCl (pH 7.4), 100 mM NaCl, 2 mM MgCl2, 5 mM DTT, 0.1 mM PMSF were microfuged and degassed prior to use.
ation on beamline 7.2 at the Daresbury Synchrotron Radiation Source (Table 2). Data were reduced using MOSFLM and SCALA.43 R-free ¯ags in the M102E and M118E data sets were assigned to the same re¯ections as in the M84E data set. For M84E-NTF2 molecular replacement and re®nement used the CNS package:44 5 % of the data was excluded during all stages of re®nement for calculation of R-free.45 We located two NTF2 chains in the asymmetric unit using residues 2-127 of chain A of the structure of the P212121 crystal form native NTF2 (PDB accession code 1oun31) as a model. A model with two NTF2 chains in the asymmetric unit would have an expected solvent content of 45 % (v/v), which was within the range commonly observed for protein crystals.46 After rigid-body re®nement and group B-factor re®nement, the R-factor of the molecular replacement solution reduced to 36.6 % (free-R factor 39.7 %). Positional re®nement was preformed with CNS44 using conjugate gradient minimization, Cartesian slow cooling and torsion-angle simulated annealing with strong noncrystallographic symmetry constraints, alternating with model building using O47 and produced a ®nal model for the two NTF2 chains in the unit cell in which the R-factor was reduced to 20.6 % (free-R factor 24.8 %). The ®nal structural model contained residues 4-126 of both NTF2 chains and 117 water molecules (Table 2). Because the crystals of M102E- and M118E-NTF2 had the same symmetry and essentially the same lattice parameters, we were able to use the M84E-NTF2 structure as a starting point and, after rigid-body re®nement, positional re®nement and model building obtained structures with R-factors of 20.4 % and 20.2 %, respectively (R-free 24.1 % and 29.1 %, respectively). Data collection and re®nement statistics for all three crystals are given in Table 2. Drawings of molecular models were produced using Bobscript.48
X-ray crystallography Diffraction-quality crystals of M84E-NTF2 were obtained by vapour diffusion using 4 ml hanging drops composed of 2 ml of 2.5 mg/ml M84E-NTF2 in100 mM Tris-HCl (pH 7.4), 100 mM NaCl, 2 mM MgCl2, 5 mM DTT, 0.1 mM PMSF and 2 ml of well buffer (100 mM sodium acetate (pH 4.5), 50 mM MgCl2, 12 % (w/v) PEG 8000). A crystal of dimensions 50 mm 50 mm 25 mm was transferred to cryoprotectant (well buffer containing 20 % (v/v) glycerol) for less than one minute and was Ê resolution native data set ¯ash-frozen at 100 K. A 2.0 A was collected at 100 K using Cu Ka radiation on a Rigaku generator equipped with Osmic mirrors and a MAR345 detector (Table 2). Diffraction quality crystals of M102E-NTF2 were obtained by vapour diffusion using hanging drops composed of 2 ml of drop buffer containing 2.7 mg/ml of M102E-NTF2 and 2 ml of 100 mM sodium acetate (pH 5.3), 50 mM MgCl2, 10 % PEG 8000 well buffer. A crystal was transferred to cryoprotectant (well buffer containing 20 % glycerol and 20 % PEG 8000) for less than one minute and was ¯ash-frozen at 100 K Ê resolution data set collected in-house and a 2.2 A (Table 2). Diffraction quality crystals of M118E-NTF2 were obtained by vapour diffusion using hanging drops composed of 2 ml of 2.5 mg/ml M118E-NTF2 in drop buffer and 2 ml of 100 mM sodium acetate (pH 4.5), 50 mM MgCl2, 26 % PEG 4000 well buffer. A crystal was transferred to cryoprotectant (well buffer containing 12 % glycerol and 35 % PEG 4000) for less than one minÊ resolution ute and was ¯ash-frozen at 100 K and a 2.3 A Ê wavelength radinative data set collected using 1.488 A
Binding to RanGDP and FxFG nucleoporins M118E-NTF2 was tested for its potential to bind RanGDP or FxFG nucleoporins by pull-down experiments with S-tagged Ran or S-tagged NSP1-FF18 sepharose beads. The beads were obtained by incubating S-agarose matrix (Novagen) with a saturating amount of Escherichia coli lysate expressing S-tagged Ran or S-tagged NSP1-FF18, for one hour at 4 C. To remove non-speci®cally bound proteins from the lysate, beads were washed with PBST and subsequently incubated with wild-type NTF2 or M118E-NTF2 in PBST buffer on a rotating wheel for one hour at 4 C. After two washes with PBST buffer, the NTF2 molecules bound to the Stagged Ran or NSP1-FF18 on the beads were recovered in SDS loading buffer after boiling, and visualized by running the samples on a 10 % to 20 % (w/v) acrylamide SDS/polyacrylamide gradient gel.49 The relative binding of the M118E-NTF2 for S-tagged RanGDP or NSP1-FF18 was then determined by comparing the level of WTNTF2 and M118E-NTF2 recovered under the same conditions. Because the binding to nucleoporins was weaker than to RanGDP,20 we employed W7A-NTF2 as a negative control. Electron microscopy Xenopus laevis were purchased from Xenopus I (Ann Arbor, MI, USA) and maintained in ®ltered arti®cial pond water. Stage VI oocytes, approximately 1.2 mm in
475
NTF2 Monomer-Dimer Equilibrium Table 2. Statistics for P21 crystals of NTF2 mutants NTF2 mutant A. Cell dimensions Ê) a (A Ê) b (A Ê) c (A b (deg.) B. Data collection and reduction Source Ê) Wavelength (A Ê) Resolution range (A Ê) Highest resolution bin (A Total/unique reflectionsa hIi/shIia Completeness (%)a Multiplicitya Rmerge (%)a,b C. Structure refinement Working/free reflections Protein/water non-H atoms Working/freec R-factor (%)d Ê 2) Average B-factor (A rms deviation from ideality Ê) Bonds (A Angles (deg.) Residues in Ramachandran region (%) Most favoured Additionally allowed Generously allowed Forbidden
M84E
M102E
M118E
34.29 79.44 41.75 103.80
35.15 79.07 41.65 104.06
34.81 78.23 41.34 104.45
CuKa 1.5418 22.3-2.0 2.11-2.0 36,984/14,697 10.5 (3.5) 99.7 (99.8) 2.5 (2.5) 5.4 (18.2)
CuKa 1.5418 30-2.23 2.37-2.23 81,646/10,835 11.4 (2.9) 99.7 (99.2) 3.2 (3.1) 8.7 (42.9)
SRS 7.2 1.488 30-2.30 2.38-2.30 73,003/9197 9.6 (2.4) 96.2 (96.2) 2.5 (2.0) 9.8 (27.2)
13,944/735 1988/117 20.7/25.1 20
10,757/549 1988/183 20.4/24.1 34
8722/475 1988/112 20.3/28.9 28
0.010 1.5
0.009 2.1
0.007 1.3
93.2 5.9 0.5 0.5 (B92)
90.9 7.7 0.9 0.5 (92B)
89.1 8.2 1.8 0.9 (92A,B)
a
Highest resolution shell in parentheses. Rmerge hklijIhkl ÿ hIhklij/hklihIhkli, where hIhkli is the mean of the observations Ihkl,i of re¯ection hkl. c Free-R was computed using 10 % of the data assigned randomly. d R-factor 100(hkl[jFo(hkl)j ÿ jFc(hkl)j]/hkljFo(hkl)j), where Fo and Fc are the observed and calculated structure factors, respectively. b
diameter, were obtained from mature females and had well-delineated animal and vegetal hemispheres. ColÊ in diameter were coated loidal gold particles 20-50 A with wild-type NTF2 or the M118E mutant and microinjected into manually defolliculated stage VI oocytes as described36 using a Narishige micromanipulator and 10-15 mm tip diameter micropipettes. After approximately 60 minutes, the injected oocytes were ®xed with glutaraldehyde followed by OsO4, dehydrated and embedded as described.50 Sections were examined in a JEOL 100CX electron microscope. Protein Data Bank accession numbers Co-ordinates for the M84E-, M102E- and M118E crystal structures have been deposited at the RCSB Protein Data Bank with accession numbers 1JB2, 1JB4, and 1JB5.
Acknowledgements We are most grateful to our colleagues in Cambridge, especially Airlie McCoy, Helen Kent and Richard Bayliss for their many helpful comments and criticisms, and to Trevor Littlewood and Richard Bayliss for assistance with binding studies. C.C.-H. held an EMBO Postdoctor-
al Fellowship (ALTF98-121). This work was supported, in part, by Research grant RG0270/1998 from the Human Frontiers Science Program.
References 1. GoÈrlich, D. & Kutay, U. (1999). Transport between the cell nucleus and the cytoplasm. Annu. Rev. Cell Dev. Biol. 15, 607-660. 2. Nakielny, S. & Dreyfuss, G. (1999). Transport of proteins and RNAs in and out of the nucleus. Cell, 99, 677-690. 3. Bayliss, R., Corbett, A. H. & Stewart, M. (2000). The molecular mechanism of transport of macromolecules through nuclear pore complexes. Traf®c, 1, 448-456. 4. Wente, S. R. (2000). Gatekeepers of the nucleus. Science, 288, 1374-1377. 5. Rout, M. P. & Wente, S. R. (1994). Pores for thought: nuclear pore complex proteins. Trends Cell Biol. 4, 357-365. 6. Rout, M. P., Aitchison, J. D., Suprapto, A., Hjertaas, K., Zhao, Y. & Chait, B. T. (2000). The yeast nuclear pore complex: composition, architecture and transport mechanism. J. Cell Biol. 148, 635-652.
476 7. Ryan, K. J. & Wente, S. (2000). The nuclear pore complex: a protein machine bridging the nucleus and cytoplasm. Curr. Opin. Cell Biol. 12, 361-371. 8. Doye, V. & Hurt, E. (1997). From nucleoporins to nuclear pore complexes. Curr. Opin. Cell Biol. 9, 401411. 9. Quimby, B. B., Lamitina, T., L'Hernault, S. W. & Corbett, A. H. (2000). The mechanism of ran import into the nucleus by nuclear transport factor 2. J. Biol. Chem. 275, 28575-28582. 10. Dasso, M. (2001). Running on ran: nuclear transport and the mitotic spindle. Cell, 104, 321-324. 11. Melchior, F. & Gerace, L. (1998). Two-way traf®cking with Ran. Trends Cell. Biol. 5, 175-179. 12. GoÈrlich, D. (1998). Transport into and out of the cell nucleus. EMBO J. 17, 2721-2727. 13. Bayliss, R., Littlewood, T. D. & Stewart, M. (2000). Structural basis for the interaction between FxFG nucleoporin repeats and importin-b in nuclear traf®cking. Cell, 102, 99-108. 14. Bayliss, R., Ribbeck, K., Akin, D., Kent, H. M., Feldherr, C. M., GoÈrlich, D. & Stewart, M. (1999). Interaction between NTF2 and xFxFG-containing nucleoporins is required to mediate nuclear import of RanGDP. J. Mol. Biol. 293, 579-593. 15. Damelin, M. & Silver, P. A. (2000). Mapping interactions between nuclear transport factors in living cells reveals pathways through the nuclear pore complex. Mol. Cell, 5, 133-140. 16. Ribbeck, K., Lipowsky, G., Kent, H. M., Stewart, M. & GoÈrlich, D. (1998). NTF2 mediates nuclear import of Ran. EMBO J. 17, 6587-6598. 17. Smith, A., Brownawell, A. & Macara, I. G. (1998). Nuclear import of Ran is mediated by the transport factor NTF2. Curr. Biol. 8, 1403-1406. 18. Stewart, M. (2000). Insights into the molecular mechanism of nuclear traf®cking using nuclear transport factor 2 (NTF2). Cell Struct. Funct. 25, 217-225. 19. Yamada, M., Tachibana, T., Imamoto, N. & Yoneda, Y. (1998). Nuclear transport factor p10/NTF2 functions as a Ran-GDP dissociation inhibitor (Ran-GDI). Curr. Biol. 8, 1339-1342. 20. Chaillan-Huntington, C., Villa-Braslavsky, C., Kuhlmann, J. & Stewart, M. (2000). Dissecting the interactions between NTF2, RanGDP and the nucleoporin xFxFG repeats. J. Biol. Chem. 275, 58745879. 21. Corbett, A. H. & Silver, P. A. (1996). The NTF2 gene encodes an essential, highly conserved protein that functions in nuclear transport in vivo. J. Biol. Chem. 271, 18477-18484. 22. Paschal, B. M., Fritze, C., Guan, T. & Gerace, L. (1997). High levels of the GTPase Ran/TC4 relieve the requirement for nuclear protein transport factor 2. J. Biol. Chem. 272, 21534-21539. 23. Lane, C. M., Cushman, I. & Moore, M. S. (2000). Selective disruption of nuclear import by a functional mutant nuclear carrier protein. J. Cell Biol. 151, 321-332. 24. Suyama, M., Doerks, T., Braun, I. C., Sattler, M., Izaurralde, E. & Bork, P. (2000). Prediction of structural domains of TAP reveals details of its interaction with p15 and nucleoporins. EMBO Rep. 1, 53-58. 25. Liker, E., Fernandez, E., Izaurralde, E. & Conti, E. (2000). The structure of the mRNA export factor TAP reveals a cis arrangement of a non-canonical RNP domain and an LRR domain. EMBO J. 19, 5587-5598.
NTF2 Monomer-Dimer Equilibrium 26. Katahira, J., Strasser, K., Podtelejnikov, A., Mann, M., Jung, J. U. & Hurt, E. (1999). The Mex67pmediated nuclear mRNA export pathway is conserved from yeast to human. EMBO J. 18, 2593-2609. 27. Black, B. E., Levesque, L., Holaska, J. M., Wood, T. C. & Paschal, B. M. (1999). Identi®cation of an NTF2-related factor that binds Ran-GTP and regulates nuclear protein export. Mol. Cell. Biol. 19, 86168624. 28. Black, B. E., Holaska, J. M., Levesque, L., OssarehNazari, B., Gwizdek, C., Dargemont, C. & Paschal, B. M. (2001). NXT1 is necessary for the terminal step of Crm1-mediated nuclear export. J. Cell Biol. 152, 141-156. 29. Bachi, A., Braun, I. C., Rodrigues, J. P., Pante, N., Ribbeck, K. & von Kobbe, C. et al. (2000). The Cterminal domain of TAP interacts with the nuclear pore complex and promotes export of speci®c CTEbearing RNA substrates. RNA, 6, 136-158. 30. Strasser, K. & Hurt, E. (2000). Binding of the Mex67p/Mtr2p heterodimer to FXFG, GLFG and FG repeat nucleoporins is essential for nuclear mRNA export. J. Cell Biol. 150, 695-706. 31. Bullock, T. L., Clarkson, W. D., Kent, H. M. & Ê resolution crystal Stewart, M. (1996). The 1.6 A structure of nuclear transport factor 2 (NTF2). J. Mol. Biol. 260, 422-431. 32. Stewart, M., Kent, H. M. & McCoy, A. J. (1998). Structural basis for molecular recognition between nuclear transport factor 2 (NTF2) and the GDPbound form of the Ras-family GTPase Ran. J. Mol. Biol. 277, 635-646. 33. Ellis, R. J. (2001). Macromolecular crowding: an important but neglected aspect of the intracellular environment. Curr. Opin. Struct. Biol. 11, 114-119. 34. Clarkson, W. D., Kent, H. M. & Stewart, M. (1996). Separate binding sites on nuclear transport factor 2 (NTF2) for GDP-Ran and the phenylalanine-rich repeat regions of nucleoporins p62 and Nsp1p. J. Mol. Biol. 263, 517-524. 35. Clarkson, W. D., Corbett, A. H., Paschal, B. M., Kent, H. M., McCoy, A. J. & Gerace, L. et al. (1997). Nuclear protein import is decreased by engineered mutants of nuclear transport factor 2 (NTF2) that do not bind GDP Ran. J. Mol. Biol. 272, 716-730. 36. Feldherr, C. M., Akin, D. & Moore, M. S. (1998). The nuclear import factor p10 regulates the functional size of the nuclear pore complex during oogenesis. J. Cell Sci. 111, 1889-1896. 37. Paschal, B. M. & Gerace, L. (1995). Identi®cation of NTF2, a cytosolic factor for nuclear import that interacts with nuclear pore protein p62. J. Cell Biol. 129, 925-937. 38. Moore, M. S. & Blobel, G. (1994). Puri®cation of a Ran-interacting protein that is required for protein import into the nucleus. Proc. Natl Acad. Sci. USA, 91, 10212-10216. 39. Wong, D. H., Corbett, A. H., Kent, H. M., Stewart, M. & Silver, P. A. (1997). Interaction between the small GTPase Ran/Gsp1p and Ntf2p is required for nuclear transport. Mol. Cell. Biol. 17, 3755-3767. 40. Van Holde, K. E. & Baldwin, R. L. (1958). Rapid attainment of sedimentation equilibrium. J. Phys. Chem. 62, 734-743. 41. Laue, T. M., Shah, B. D., Ridgeway, T. M. & Pelletier, S. L. (1992). Computer-aided interpretation of analytical sedimentation data for proteins. In Analytical Ultracentrifugation in Biochemistry and Polymer Science (Harding, S. E., Rowe, A. J. & Horton, J. C.,
NTF2 Monomer-Dimer Equilibrium
42.
43. 44.
45.
eds), pp. 90-125, Royal Society of Chemistry, Cambridge, UK. Johnson, M. L. & Straume, M. (1994). Comments on the analysis of sedimentation equilibrium experiments. In Modern Analytical Ultracentrifugation (Schuster, T. M. & Laue, T. M., eds), pp. 37-65, Birkhauser, Boston, MA. Collaborative Computational Project Number 4 (1994). The CCP4 suite: programs for protein crystallography. Acta Crystallog. sect. D, 50, 760-763. BruÈnger, A. T., Adams, P. D., Clore, G. M., DeLano, W. L., Gros, P. & Grosse-Kunstleve, R. W. et al. (1998). Crystallography and NMR system: a new software suite for macromolecular structure determination. Acta Crystallog. sect. D, 50, 905-921. BruÈnger, A. T. (1992). Free R value: a novel statistical quality for assessing the accuracy of crystal structures. Nature, 355, 472-474.
477 46. Matthews, B. W. (1968). The solvent content of protein crystals. J. Mol. Biol. 33, 491-497. 47. Jones, T. A., Zou, J.-Y., Cowan, S. W. & Kjeldgaard, M. (1991). Improved methods for building protein models in electron density maps and for locating errors in these models. Acta Crystallog. sect. A, 47, 110-119. 48. Esnouf, R. M. (1997). An extensively modi®ed version of MolScript that includes greatly enhanced colouring capabilities. J. Mol. Graph. Model. 15, 132136. 49. Laemmli, U. K. (1970). Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature, 227, 680-685. 50. Dworetzky, S. I. & Feldherr, C. M. (1988). Translocation of RNA-coated gold particles through the nuclear pores of oocytes. J. Cell Biol. 106, 575-584.
Edited by B. Holland (Received 5 July 2001; received in revised form 24 September 2001; accepted 25 September 2001)
J. Mol. Biol. (2001) 314, 465±477
NTF2 Monomer-Dimer Equilibrium Catherine Chaillan-Huntington1, P. Jonathan G. Butler1 James A. Huntington2, Debra Akin3, Carl Feldherr3 and Murray Stewart1* 1
MRC, Laboratory of Molecular Biology, Hills Road, Cambridge CB2 2QH, England 2
University of Cambridge Department of Haematology Wellcome Trust Centre for Molecular Mechanisms in Disease, Addenbrooke's Hospital Site, Hills Road Cambridge CB2 2XY, England 3
Department of Anatomy and Cell Biology, University of Florida College of Medicine Gainesville, FL 32610, USA
Nuclear transport factor 2 (NTF2) mediates nuclear import of RanGDP, a central component of many nuclear traf®cking pathways. NTF2 is a homodimer and each chain has independent binding sites for RanGDP and nuclear pore proteins (nucleoporins) that contain FxFG sequence repeats. We show here that the monomer-dimer dissociation constant for NTF2 obtained by sedimentation equilibrium ultracentrifugation is in the micromolar range, indicating that a substantial proportion of cellular NTF2 may be monomeric. To investigate the functional signi®cance of NTF2 dimerization, we engineered a series of point mutations at the dimerization interface and one of these (M118E) remained monomeric below concentrations of 150 mM. CD spectra and X-ray crystallography showed that M118E-NTF2 preserved the wild-type NTF2 fold, although its thermal stability was 20 deg. C lower than that of the wild-type. M118E-NTF2 bound both RanGDP and FxFG nucleoporins less strongly, suggesting that dissociation of the NTF2 dimer could facilitate RanGDP release and thus nucleotide exchange after it had been transported into the nucleus. Moreover, colloidal gold coated with M118E-NTF2 showed reduced binding to Xenopus oocyte nuclear pores. Overall, our results indicate that dimer formation is important for NTF2 function and give insight into the formation of heterodimers by mRNA export factors such as TAP1 and NXT1 that contain NTF2-homology domains. # 2001 Academic Press
*Corresponding author
Keywords: nuclear traf®cking; protein structure; crystallography; dimerization
Introduction The nuclear pore complexes (NPCs) of the nuclear envelope mediate bi-directional traf®cking of macromolecules between cytoplasm and nucleus.1 ± 4 NPCs are huge macromolecular assemblies constructed from 50 different proteins collectively called nucleoporins,5,6 many of which interact with soluble components of the nucleocytoplasmic transport machinery.7 Many nucleoporins contain tandem sequence repeats based on Present address: C. Chaillan-Huntington, Cambridge Antibody Technology, Science Park, Melbourn, Cambridgeshire SG8 6JJ, England. Abbreviations used: NPC, nuclear pore complex; NTF2, nuclear transport factor 2; NSP1-FF18, NSP1 protein construct containing 18 FxFG repeats; PBS, phosphate-buffered saline; PBST, PBS containing 0.1% (v/v) Tween 20; tm, melting point temperature ( C). E-mail address of the corresponding author: [email protected] 0022-2836/01/30465±13 $35.00/0
FxFG or GLFG cores joined by hydrophilic linkers of variable sequence and length.7,8 Generally, traf®cking of macromolecules such as proteins and nucleic acids is mediated by carriers that are usually members of the importin-b superfamily.1 ± 4 In addition to the importin-family carriers and nucleoporins, the Ras-family GTPase, Ran, and nuclear transport factor 2 (NTF2) are required for ef®cient transport. Ran is a key component of the nuclear traf®cking machinery and also appears to be involved in cellBecause RanGAP1 cycle progression.1 ± 4,9,10 (RanGTPase activating protein) is located in the cytoplasm, whereas the Ran guanine nucleotide exchange factor (RCC1) is located in the nucleus, it is thought that cytoplasmic Ran is primarily GDPbound (RanGDP), whereas nuclear Ran is in the GTP-bound form (RanGTP).11,12 This putative RanGTP gradient between the cytoplasm and nucleoplasm has been proposed to be the basis for determining the directionality of the transport of macromolecules.12 Cargo proteins carrying nuclear # 2001 Academic Press
466 localization signals bind to carrier molecules of the importin-b family in the cytoplasm and are then translocated into the nucleus through nuclear pore complexes, after which the carrier-cargo complex is dissociated by RanGTP. The carrier is then recycled to the cytoplasm bound to RanGTP where RanGAP generates RanGDP, which dissociates from the carrier, freeing it to interact with another cargo molecule and undergo a further import cycle.1 ± 4 The cycling of carriers between nucleus and cytoplasm, and their binding and release of cargo molecules in the appropriate compartment is thus intimately tied to the Ran nucleotide state. Nuclear protein import mediated by importin-b13 and the NTF2-mediated import of RanGDP14 both require a direct interaction between the carrier and FxFG nucleoporins, and analogous nucleoporin-carrier interactions are thought to be involved in other nuclear traf®cking pathways (e.g. see Damelin & Silver15). The nuclear import of RanGDP is mediated by NTF2.16,17 NTF2 is a small homodimeric protein that binds both RanGDP and FxFG repeatcontaining nucleoporins on separate sites.3,18 NTF2 is localized primarily to the nuclear rim,19,20 with lower concentrations in the nucleus and the cytoplasm. NTF2 is an essential protein in yeast21,22 and Caenorhabditis elegans,9 and dominant negative mutants can disrupt nuclear protein import selectively.23 Studies using both Ran and NTF2 engineered mutants have shown that both the NTF2-RanGDP and NTF2-FxFG interactions are required for ef®cient RanGDP import.14,16,17 The Kd values for the interactions of NTF2 with RanGDP or with FxFG repeat nucleoporins are 100 nM and 1 mM, respectively.14,20 On the basis of these values, we recently proposed a model for Ran import where the Ran/NTF2 complex would remain intact during the translocation across the nuclear envelope, ``hopping'' from one FxFG nucleoporin to the next one along the NPC channel.20 We also showed that there was no cooperativity between the interactions involving RanGDP, NTF2, and FxFG repeats.20 Several proteins involved in mRNA nuclear export contain domains that are homologous to NTF2 and that almost certainly share the same fold.24 Thus, vertebrate TAP1 and its yeast counterpart Mex67 have N-terminal RNA-binding domains, followed by four leucine-rich repeats, a domain homologous to NTF2 and ®nally a C-terminal UBA domain.24 ± 26 TAP1 interacts with the small 15 kDa monomeric protein NXT1 (also called p15) that also has an NTF2 fold,24,26 ± 28 and it has been proposed that the formation of the TAP1/ NXT1 heterodimer is mediated by an interaction analogous to that seen between the two chains in the NTF2 homodimer.24 Moreover, NXT1/p15 binds to Ran (albeit in its GTP-bound form) and both Mex67 and TAP1 bind to nucleoporins.29,30 In the case of Mex67, nucleoporin binding appears to require the formation of a heterodimer with Mtr2, which seems to perform a role analogous to NXT1.
NTF2 Monomer-Dimer Equilibrium
Although NTF2 has been implicitly assumed to be a dimer in vivo, the observations with the NTF2 homologues involved in mRNA nuclear export raise the possibility that NTF2 itself may be at least partially dissociated. We therefore determined Kd for the NTF2 monomer-dimer equilibrium and show here that it is in the micromolar range, which is of the same order as the cellular concentration of NTF2,20 indicating that a substantial proportion of NTF2 may be monomeric in vivo. To investigate the functional signi®cance of NTF2 dissociation, we examined its in¯uence on the interaction with RanGDP and FxFG nucleoporins. To do this, we weakened the interaction between NTF2 chains at the dimer interface by mutating methionine residues 84, 102 and 118 to glutamic acid. Dimer formation was reduced substantially in the M118E NTF2 mutant, which also showed a 20 deg. C decrease in thermal stability compared with wildtype NTF2 but no signi®cant alteration of its overall fold in solution. This mutant showed reduced af®nity for both RanGDP and FxFG nucleoporin repeats, and the binding of M118E-NTF2 to Xenopus oocyte NPCs was reduced. Moreover, the results obtained with other mutations at the dimerization interface showed unanticipated ways in which NTF2 dimerization could be stabilized and gave insight into the putative formation of heterodimers by components of the Tap/Mex67 mRNA export machinery.
Results Monomer-dimer equilibrium of NTF2 The aggregation state of NTF2 was examined by sedimentation equilibrium ultracentrifugation, loading samples with initial A280 values of 0.8, w;app against concentration 0.4 and 0.2. Plots of M (Figure 1(a), upper plot) showed that wild-type NTF2 was primarily dimeric at concentrations above 5 mM, but there was clear evidence for dissociation into monomers at lower concentrations. The data obtained from different loading concentrations were superimposed and suggested that the NTF2 dimers were in a rapidly reversible equilibrium with monomers. To better de®ne the concentration range where monomeric NTF2 dominated, sedimentation equilibrium data were obtained with lower loadings, employing concentrations of NTF2 corresponding to initial A230 values of 0.3, 0.2 and 0.1. The results with lower initial NTF2 concentrations (Figure 1(a), lower plot), were very similar to those obtained previously, and clear evidence for dissociation can be seen at NTF2 concentrations below 4 mM. A model for an ideal monomer/dimer equilibrium gave a good ®t to the data of concentration against radius, and examples of plots of the residuals for the ®tting are shown in (Figure 1(b)). This model gave an estimate of the NTF2 dimer dissociation constant (Kd) of 1.1 (0.05) mM.
NTF2 Monomer-Dimer Equilibrium
467
Figure 1. Sedimentation equilibrium analysis of NTF2 monomer-dimer equilibrium. (a) Plots of Mw;app against concentration, derived from cells loaded at different initial concentrations (see Materials and Methods), together with curve ®tted for a monomer-dimer equilibrium. (b) Plots of residuals for direct ®tting of concentration against radius data with model for monomer/dimer equilibrium. The upper plots correspond to the early loading concentrations, while the lower plots correspond to lower loading concentrations employed to obtain a better estimate of Kd, by working over an appropriate concentration range in the cells. These data indicate a Kd of 1 mM for the wt-NTF2 monomer-dimer dissociation constant.
Point mutants at the NTF2 dimerization interface The dissociation constant (Kd) value of 1.1 (0.05) mM is comparable to the concentration for NTF2 in the nucleoplasm (0.6 mM) and the cytoplasm (0.3 mM),20 which suggests that a substantial proportion of NTF2 may be monomeric in vivo. This raised the possibility that dissociation into monomers might in¯uence the interaction of NTF2 with Ran or FxFG nucleoporins. However, it was technically dif®cult to investigate this possibility experimentally because the dissociation constant of the NTF2 dimer was of the same order as its binding constant for RanGDP and FxFG nucleoporins.20 We therefore attempted to produce primarily monomeric NTF2 by introducing point mutations in the NTF2 dimer interface. The crystal structure of NTF231 shows that the interface between chains in the NTF2 dimer is primarily hydrophobic and is formed by the close juxaposition of extensive bsheets from each molecule. Methionine residues 84,
102 and 118 are located within the dimerization interface (Figure 2(a) and (b)) and are in close apposition with hydrophobic residues from the neighbouring chain. We therefore anticipated that mutating these methionine residues to glutamate would reduce the contribution of the hydrophobic effect to dimerization and so produce primarily monomeric NTF2 in solution at concentrations suf®ciently high to measure the association with RanGDP and FxFG nucleoporins directly. All three mutants were assessed by sedimentation equilibrium ultracentrifugation to determine their dissociation constants (Figure 3). Somewhat surprisingly, the effect of the M84E and M102E mutations was slight and, although they had Kd values of 5 (0.5) and 20 (1) mM, respectively, these mutants remained primarily dimeric at the concentrations needed to assess binding to RanGDP and FxFG nucleoporins. However, the M118E mutant had a much more dramatic effect and appeared monomeric over the entire
468
NTF2 Monomer-Dimer Equilibrium
Figure 2. Location of M84, M102 and M118 at the NTF2 dimerization interface. An illustration of the side-chains protruding into the interaction interface between NTF2 chains in the dimer. Methionine residues 84, 102 and 118 are all important components of the hydrophobic centre of the interaction interface. Hydrophobic residues are in grey, acidic residues in red, basic residues in blue and amphipolar residues in white. Also shown are the corresponding side-chains in the putative structures (see Suyama et al.24) of TAP1 and NXT1. Note that neither TAP1 nor NXT1 contains the equivalent of His100 in NTF2.
concentration range examined and remained monomeric even with initial concentrations as high as A280 0.8. The data shown in Figure 3(a) (bottom plot) indicated that M118E-NTF2 was monomeric up to concentrations of at least 150 mM, while the residual plot (Figure 3(b), lowest plot) showed a good ®t for a non-ideal monomer, with the osmotic second virial coef®cient (B) estimated as 0.0043 Mÿ1. We can therefore conclude that wild-type NTF2 is primarily dimeric at concentrations above 2-4 mM, whereas mutants within the dimerization interface show varying degrees of dissociation. In particular, M118E-NTF2 was essentially monomeric below 150 mM. Crystal structures of M84E-, M102E- and M118E-NTF2 For all the mutants, the crystal structure shows dimers with a Ca backbone similar to the wild-type molecule. The crystal structure of M84E-NTF2 showed putative salt-bridges between His100 and Glu84 (Figure 4(a)) in both chains, which could help mitigate the introduction of a charged residue into the hydrophobic interface. In M102E- and M118E-NTF2, steric constraints prevented the formation of saltbridges between His100 and Glu102 in different chains, although a putative salt-bridge was formed between these residues in the same chain (data not shown, but can be seen in PDB deposition 1JB4). In addition, Glu118 was placed very close to other acidic side-chains and the resulting repulsion would decrease the stability of the interface (this would be especially true at physiological pH, but was probably diminished in the acidic conditions (pH 4.5) needed to produce the crystals of this mutant). Thus, the crystal structures of the three mutants can account for the different Kd values observed for dimerization. Comparison of the three dimer structures indicated that, although the NTF2 fold had been retained in each case, each of the mutations had also introduced a small rigid-body rotation
between the two chains in the dimer (Figure 5). When RanGDP binds to NTF2 there is also a small rigid-body rotation of one NTF2 chain relative to the other (see Stewart et al.32 and Figure 5), albeit in the direction opposite to that seen with the NTF2 mutants in the present study. Overall, it seems that there is considerable plasticity to the NTF2 dimerization interface and this accounts for how small rearrangements in M84E-NTF2, for example, can enable formation of a putative salt-bridge to His100 and so mitigate the effect of introducing a charged residue into the primarily hydrophobic interface. The observation that M118E-NTF2 crystallized as a dimer indicated that the M118E mutation does not abolish the dimerization completely, at least under the acidic conditions (pH 4.5) used to grow the crystals. The sedimentation equilibrium data indicated that, in contrast to M84E and M102E, the M118E mutation perturbs dimerization so that dimeric NTF2 could not be detected at concentrations up to 150 mM. However, it would appear that dimerization of this mutant was still possible in the unusual solvent conditions and the very high concentrations used for crystallization (of the order of millimolar). One function of the PEG used as precipitant, for example, is to increase the activity of macromolecular species due to the excluded volume effect.33 Moreover, the M118ENTF2 crystals were obtained only at pH 4.5, where the ionization of the introduced Glu would be much less than at physiological pH. At lower concentrations (0.01-0.1 mM) in a more physiological solvent, sedimentation equilibrium data indicated that M118E-NTF2 was monomeric. Under these conditions, circular dichroism (Figure 6(a)) indicated that the secondary structure content of M118E mutant was very similar to that of wildtype NTF2, strongly suggesting that the structure of the individual polypeptide chains had not been altered radically when M118E-NTF2 dissociated into monomers. Circular dichroism (CD) spectroscopy also showed that the thermal stability of the different
469
NTF2 Monomer-Dimer Equilibrium
Figure 3. Sedimentation equilibrium analysis of NTF2 mutants. (a) Plots of Mw;app against concentration for mutants, together with curves ®tted either for a monomer/dimer equilibrium (M84E-NTF2 and M102E-NTF2) or for a non-ideal monomer (M118E-NTF2). (b) Plots of residuals for ®tting of concentration against radius data for mutants with models either for a monomer/dimer equilibrium (M84E-NTF2 and M102E-NTF2) or for a non-ideal monomer equilibrium (M118E-NTF2). These data show that M118E-NTF2 was present as monomers at concentrations up to 150 mM.
NTF2 mutants correlated with the strength of their dimerization. Sedimentation equilibrium ultracentrifugation had indicated that, with the protein concentrations employed (35 mM), wild-type NTF2 was present as a dimer, M84-NTF2 and M102NTF2 were partially dissociated and M118E-NTF2 was monomeric. Figure 6 shows that M118E-NTF2 was considerably less stable than wild-type, with a tm value of 51 C compare to 71 C for wild-type NTF2, whereas both the M84E- and M102Emutants showed an intermediate lowering of tm (Table 1).
Influence of NTF2 dimerization on the interaction with RanGDP and FxFG nucleoporins Previous studies have indicated that binding to RanGDP and FxFG nucleoporins are both crucial for NTF2 function.14,16,18,20 We therefore investigated the in¯uence of dimerization on these properties using bead-binding assays. M118E-NTF2 was tested for its ability to be pulled down by Sagarose beads coated with either S-tagged RanGDP or S-tagged Nsp1-FF18 (a construct that contains
470
NTF2 Monomer-Dimer Equilibrium
Figure 4. Putative Glu84-His100 salt-bridge stabilizes the M84E-NTF2 dimer. A portion of the 2Fo ÿ Fc electron density map of the crystal structure of M84E-NTF2 showing putative interactions between Glu84 and His100 that could contribute to the stability of the dimerization interface. The A chain is yellow and the B chain is blue. The interaction between chains is mediated by a water molecule.
18 FxFG repeats of the yeast nucleoporin NSP114,20,34,35) and showed reduced binding to both RanGDP and FxFG nucleoporins compared with wild-type NTF2 (Figure 7). We also tested the ability of M118E-NTF2 to associate with NPCs in vivo by microinjecting colloidal gold coated with either this mutant or wild-type NTF2 into Xenopus oocytes. Electron micrographs (Figure 8) showed that, as observed previously,14,36 colloidal gold coated with wild-type NTF2 bound to the cytoplasmic and nucleoplasmic faces of NPCs, as well as to their central transport channel. By contrast, colloidal gold particles coated with M118E-NTF2 showed greatly reduced binding to NPCs when injected into Xenopus oocytes: there was only slight staining of the nucleoplasmic and cytoplasmic faces of NPCs and very few particles were seen in the central transport channel. The results of these binding studies imply that either the binding of both RanGDP and FxFG nucleoporins is sensitive to the monomer-dimer
Table 1. Thermal unfolding of NTF2 mutants Protein Wild-type NTF2 M84E-NTF2 M102E-NTF2 M118E-NTF2
tm( C) 71 61 61 51
Figure 5. NTF2 rigid-body rotations. Illustration of the rigid-body rotations observed between (black) wildtype NTF2 alone31 and (blue) bound to RanGDP32 and the M84E mutants (red). In each case, the A chains have been superimposed, and so the difference in positions of the B-chains indicates the relative movement between the chains in the dimer.
state of NTF2 or that we have altered the binding sites for both by our mutation. Met118 is located some distance from the RanGDP binding site on NTF232 and, although the precise binding site for FxFG nucleoporins has not been established directly, it appears to be near Trp7,14 which is also a considerable distance from Met118. Therefore, it appeared more likely that the reduced af®nity seen in our binding assays was a result of the monomeric nature of the M118E-NTF2.
Discussion Previous studies using X-ray crystallography,31,32 gel-®ltration37 and NTF2 covalently coupled to Sepharose beads34,35 had all indicated that NTF2 is a homodimer. However, our ultracentrifugation sedimentation equilibrium data (Figure 1) established a dissociation constant for the NTF2 dimer to be Kd 1.1 (0.05) mM. A dissociation constant of this magnitude implies that a considerable degree of NTF2 monomer could be present at concentrations of the order of micromolar or 0.015 mg/ml. Previous in vitro studies had been performed at concentrations of the order of 1 mg/ ml, where NTF2 would indeed be primarily dimeric. However, cellular concentrations of NTF2 are considerably lower than those used in these in vitro studies. Previous results have shown that NTF2 is localized primarily at the nuclear envelope, where it is bound to FxFG
NTF2 Monomer-Dimer Equilibrium
Figure 6. M118E-NTF2 retains its fold in solution at monomeric concentration, but shows a decrease in thermal stability. (a) The CD spectra of wt-NTF2 (continuous line) and M118E-NTF2 (broken line) in solution were indistinguishable, indicating that the M118E mutant had not introduced a major conformational change. These spectra were recorded at a concentration of 35 mM, where sedimentation equilibrium data (Figure 3) indicated that M118E-NTF2 was dissociated into monomers. (b) Thermal stability of wt-NTF2 and the M84E, M102E and M118E mutants determined by CD. The data represent the change in ellipticity (, mdeg) as a function of temperature. The denaturation temperature was taken at the in¯exion point of the rising portion of the curve.
nucleoporins.14,19,20,38 The concentration of NTF2 at the nuclear envelope is of the order of 20 mM,20 but the cytoplasmic and nucleoplasmic concentrations of NTF2 are only of the order of 0.3 mM and 0.6 mM respectively.20 A micromolar dissociation constant therefore would indicate that, although most of the NTF2 at the nuclear pores would be dimeric, a substantial proportion of NTF2 in the bulk of the nucleoplasm and cytoplasm could be monomeric. Therefore, except at the nuclear envelope, a considerable dissociation of dimeric NTF2 into monomers could occur in vivo. This possibility raises the question of the importance of dimerization for NTF2 function.
471 Because the dissociation constant of NTF2 dimers was of the same order of magnitude as that observed for RanGDP and FxFG nucleoporin binding,14,20 we engineered a point mutant (M118E) at the dimerization interface (Figure 2(a) and (b)) that increased the degree of dissociation so that it was monomeric at concentrations up to 150 mM. Although M118E-NTF2 was still able to bind Ran GDP and FxFG nucleoporin repeats, it did so with reduced af®nity, indicating that the oligomeric state of NTF2 in¯uenced its af®nity for both of its substrates. The af®nity of NTF2 for its substrates is crucial for its function.20 Thus, NTF2 has to bind RanGDP suf®ciently strongly to transport it through nuclear pores, yet suf®ciently weakly to release it in the nucleus to enable it to be recharged with RanGTP by RCC1. Although the concentration of NTF2 at the nuclear pores themselves would result in NTF2 being primarily dimeric, the lower NTF2 concentration in the nucleus would favour its dissociation into monomers. Reducing its af®nity for RanGDP in the nucleus through dissociation of the NTF2 dimer could be important in promoting release of RanGDP once it had been transported through the nuclear pore, especially since the NTF2-RanGDP complex has been shown to inhibit nucleotide exchange by RCC1.19 The oligomeric state of NTF2 also in¯uenced the thermal stability of the molecule (Figure 6), so that monomeric M118E-NTF2 showed a decreased stability (51 C) compared to wild-type (71 C). Surprisingly M84E-NTF2 and M102E-NTF2 were still primarily dimeric in solution, although in each case the mutant had a higher Kd than wild-type (5(0.5) and 20(1) mM, respectively). The crystal structures of the mutants gave a rationale for their unexpected retention of dimerization. In both mutants, local rearrangements, involving rigidbody rotation to the two chains coupled with the formation of salt-bridges or hydrogen bonds between the introduced glutamate residue and His100, offset the introduction of a hydrophilic residue into the hydrophobic dimer interface (Figure 5). Thus, although these mutants produced a small but signi®cant rigid-body rotation of the two NTF2 chains, they did not reduce their af®nity suf®ciently to generate a substantial population of monomers under the conditions used to assay for RanGDP and FxFG nucleoporin binding. However in the M118E-NTF2 structure, Glu118 was less able to form a salt-bridge with His100 from the other chain, and so a greater degree of dissociation was produced by this mutation. Interestingly, a yeast analogue of human M84E-NTF2 (yeast M83TNTF2) has a temperature-sensitive phenotype.21,39 Although the crystal structure of yeast wild-type NTF2 has not been determined, it is likely that its structure is very similar to that of the rat protein.39 A plausible explanation of the temperature-sensitive phenotype of the yeast M83T mutation is that, unlike the mutation to glutamate, threonine would not be able to form a compensatory salt-bridge
472
Figure 7. Interaction of M118E-NTF2 with RanGDP and FxFG nucleoporins. S-tagged Ran-GDP or Nsp1-FF18 attached to S-agarose beads were incubated with wild-type or M118E-NTF2 and, after centrifugation and washing, the amount of material bound was assessed by SDS-PAGE. Although M118E-NTF2 bound to both RanGDP and NSP1-FF18, it did so with lower af®nity than wild-type NTF2. Because the af®nity of NTF2 for FxFG nucleoporins was lower than for RanGDP,20 we included W7A-NTF2 (which does not bind FxFG nucleoporins, Bayliss et al.14) as a negative control.
with His100 and so would increase the level of dissociation of the NTF2 dimer. Our CD studies (Figure 6) indicated that the temperature-stability of NTF2 monomers is substantially less than that of the wild-type and so would be consistent with the observed temperature-sensitive phenotype of yeast M83T-NTF2. The results of our mutagenesis studies of the NTF2 dimerization interface provide a structural context for evaluating the heterodimerization of NTF2 homologues involved in mRNA export. Both
NTF2 Monomer-Dimer Equilibrium
TAP1 and NXT1 show suf®ciently strong sequence homology to NTF2 to make it unlikely that they do not also have the same fold and indeed the formation of an NXT1-TAP1 heterodimer has been proposed to be important for their function.24 It has been proposed that this heterodimer forms by using a hydrophobic interface constructed from the extensive b-sheet of the NTF2 fold (Figure 2) and a number of mutations have been engineered into both TAP1 and NXT1 to attempt to evaluate this hypothesis.24 However, although some of these mutants have interfered with dimer formation, others produced a more marginal effect. Moreover, some of the mutations that reduced the TAP1NXT1 interaction (for example, I95D, F109Y and F133D in p15 and L466H, F479W and F499D in Tap) were not in the putative dimerization interface. Although not always conserved as methionine, the residue corresponding to Met84 in vertebrate NTF2 is invariably hydrophobic in TAP1, Mex67 and NXT1, and mutations here (F465H in TAP1; V94D in NXT1) reduce heterodimer formation. However, both TAP1 and NXT1 lack a histidine residue corresponding to NTF2 His100, and so cannot form a compensating saltbridge to stabilize dimerization. At the position corresponding to NTF2 Met102, TAP1 and NXT1 have polar residues (T, N in NXT1, D in TAP1, D or E in Mex67). At the position corresponding to NTF2 Met118, mutations from the conserved hydrophobic residue to Asp (analogous to our M118E mutant) in either TAP1 (V498D) or NXT1 (C132D) inhibit heterodimerization. Our crystal
Figure 8. Electron micrographs of sections of embedded Xenopus oocytes microinjected with colloidal gold (black dots) coated with (a) wild-type or (b) M118E-NTF2. Whereas wild-type NTF2 accumulated on both nuclear and cytoplasmic faces of the NPCs as well as in their central transport channel, the degree of labeling observed with the M118E mutant was greatly reduced, showing only weak accumulation on the NPC faces and little material present in the central transport channel. The scale bar represents 1000 nm.
473
NTF2 Monomer-Dimer Equilibrium
structures of the three Met mutants of NTF2 show how rigid-body rotations of the interface can occur to accommodate changes in the dimerization interface and so may account for the failure of some mutations at the putative TAP1-NXT1 interface24 to inhibit heterodimer formation. In summary, we have demonstrated that the NTF2 dimer has a micromolar dissociation constant that is of the order of the cellular concentration of NTF2, raising the possibility that NTF2 may be partially dissociated in vivo. We have engineered point mutants at the NTF2 dimerization interface that increase the degree of dissociation, albeit with two of these (M84E and M102E), compensating salt-bridge formation with His100 partially mitigated the effect of the mutation on interface hydrophobicity and so generated only a small change in Kd. However, M118E-NTF2 remained monomeric below 150 mM, because similar compensating salt-bridges could not form. The heat-denaturation pro®les of the mutants show that the dimerization of NTF2 is important for thermal stability. Moreover, the af®nity of the M118E mutant was reduced for both RanGDP and nucleoporin FxFG repeats. In the context of the concentration of NTF2 in the nucleus, this suggested that dissociation of the NTF2 dimer could speed the release of RanGDP once it had been transported into the nucleus and thus facilitate nucleotide exchange by RCC1. Overall, our results indicate that NTF2 dimerization is important for its function in the nuclear import of RanGDP and are consistent with an analogous TAP1-NXT1 heterodimerization being important for binding to nucleoporins during mRNA export.
successive scans (for each cell) could not be distinguished when overlaid and the later scan was then taken as being operationally at equilibrium. Baselines were measured by overspeeding (to 60,000 rev/min), until the sample had pulled away from the meniscus, and then slowing the centrifuge to 18,000 rev/min before recording a ®nal scan for the baseline. Optical absorbance values were converted to molarities, using the molar extinction coef®cients e230 4.39 104 Mÿ1 and e280 9.98 103 Mÿ1 and allowing for the 12 mm path-length of the cells. The solvent density (r) and partial speci®c volume (n) of NTF2 were calculated from their compositions, as described,41 using the program SEDNTERP (courtesy of John Philo from the RASMB archive), which gave values of 1.0062 g/ml and 0.7345 ml/g, respectively. The data were analyzed by calculating the weight average, apparent molecular mass (Mw;app )) for overlapping sets of 41 consecutive datum points, with the concentration estimated as that of the mid-point, by non-linear regression ®tting (with proFit v.5.1.2, from QuantumSoft, ZuÈrich, Switzerland) to the equation: 2RT w;app d ln cr M dr2 1 ÿ n rw2
where cr is the concentration at radius r, o is the rotational speed (in rad/second), R is the gas constant and T is the temperature (in K). Plots of Mw;app against c were prepared (using proFit) and examined to predict the best model for the dependence. Direct ®tting to the data for c against r, giving a closer approximation to random errors and therefore a better estimation of the parameters,42 was again by non-linear regression (with proFit as above), weighted with the standard deviation of each datum point. For both the model of an ideal monomer/dimer aggregation equilibrium and a non-ideal monomer, the following simpli®cation of the equations was employed:
Experimental Procedures Mutagenesis, protein expression and purification Site-directed mutagenesis of the methionine residues at positions 84, 102 and 118 (ATG codon) to glutamic acid (GAA codon) was performed using the Quick Change site-directed mutagenesis kit (Stratagene, La Jolla, Ca) following the manufacturer's protocol and using oligonucleotides 30 to 37 bases in length. Rat NTF2 (wild-type and mutants), N-terminally S-tagged canine Ran and the 18 FxFG repeat-containing fragment of nucleoporin Nsp1p (NSP1-FF18) were expressed and puri®ed as described.16,20,34 NTF2 concentrations refer to the amount of monomeric chain. Sedimentation equilibrium analysis Sedimentation equilibrium runs were carried out in standard double-sector cells in an An-60 Ti rotor and a Beckman XL-A analytical ultracentrifuge. Samples, in PBS, were analyzed by long column equilibrium at 18,000 rev/min and 5.0 C, with optical absorbance recorded at 230 nm or 280 nm, taking the mean (and standard deviation) of 100 readings for the ®nal data at each radius, using the standard Beckman software. Runs were initially over-speeded, at 25,000 rev/min for six hours, to speed attainment of equilibrium.40 Subsequently, scans were taken at intervals of 24 hours until
1
s
M1 1 ÿ n rw2 2RT
2
where M1 is the monomer concentration and s is a scaling constant. For the monomer/dimer equilibrium, the reference monomer concentration (c1,0), at the reference radius (r0) was calculated from the equation:
c1;0
ÿKd
q Kd2 8Kd A0 ÿ dA=e1 4
3
where Kd is the dissociation coef®cient, A0 is the absorbance at the reference radius (r0), e1 is the monomer extinction coef®cient (and it is assumed that that for the dimer will be twice this) and dA is the baseline absorbance. The absorbance (Ar) at every radius (r) was then calculated from the equation: Ar e1 c1;0 exp s r2 ÿ r20 2 c1;0 exp s r2 ÿ r20 2 Kd
4
dA
For the non-ideal monomer model, the reference concentration (c0) was taken at a reference radius close
474
NTF2 Monomer-Dimer Equilibrium
to the meniscus, so that it had a small value. The absorbance at every radius was then calculated from the equation: Ar e c0 exp s r2 ÿ r20 ÿ 2Bc0 exp s r2 ÿ r20 dA 5 ÿ1
where B is the osmotic second virial coef®cient (in M ). Parameters ®tted included Kd or B, depending upon the model, and a correction factor for M1 (to allow for possible error in n or r) and an error in dA (to allow for errors in measurement). In practice, the ®rst factor was between 0.95 and 1.05, while the second was <0.05 absorbance units. The calculated optical absorbances and residuals for each radius were tabulated at the end of each ®t. The former could be used to calculate and plot the ®tted curve w;app against concentration, for that model, on the of M plot of the experimental values, while the latter was plotted against radius, to con®rm the validity of the ®t to the model. Circular dichroism spectroscopy CD spectra were recorded on a JASCO 810 spectropolarimeter. Spectra in the 200-260 nm range were recorded as an average of three scans. The effect of temperature on changes in protein secondary structure was followed by recording the ellipticity at 222 nm every 0.5 deg. C in a 0.1 cm temperature-controlled cell using a temperature change of 60 deg. C/hour. Before each temperature scan, a spectrum in the range of 200-260 nm was recorded. Protein samples in 100 mM Tris-HCl (pH 7.4), 100 mM NaCl, 2 mM MgCl2, 5 mM DTT, 0.1 mM PMSF were microfuged and degassed prior to use.
ation on beamline 7.2 at the Daresbury Synchrotron Radiation Source (Table 2). Data were reduced using MOSFLM and SCALA.43 R-free ¯ags in the M102E and M118E data sets were assigned to the same re¯ections as in the M84E data set. For M84E-NTF2 molecular replacement and re®nement used the CNS package:44 5 % of the data was excluded during all stages of re®nement for calculation of R-free.45 We located two NTF2 chains in the asymmetric unit using residues 2-127 of chain A of the structure of the P212121 crystal form native NTF2 (PDB accession code 1oun31) as a model. A model with two NTF2 chains in the asymmetric unit would have an expected solvent content of 45 % (v/v), which was within the range commonly observed for protein crystals.46 After rigid-body re®nement and group B-factor re®nement, the R-factor of the molecular replacement solution reduced to 36.6 % (free-R factor 39.7 %). Positional re®nement was preformed with CNS44 using conjugate gradient minimization, Cartesian slow cooling and torsion-angle simulated annealing with strong noncrystallographic symmetry constraints, alternating with model building using O47 and produced a ®nal model for the two NTF2 chains in the unit cell in which the R-factor was reduced to 20.6 % (free-R factor 24.8 %). The ®nal structural model contained residues 4-126 of both NTF2 chains and 117 water molecules (Table 2). Because the crystals of M102E- and M118E-NTF2 had the same symmetry and essentially the same lattice parameters, we were able to use the M84E-NTF2 structure as a starting point and, after rigid-body re®nement, positional re®nement and model building obtained structures with R-factors of 20.4 % and 20.2 %, respectively (R-free 24.1 % and 29.1 %, respectively). Data collection and re®nement statistics for all three crystals are given in Table 2. Drawings of molecular models were produced using Bobscript.48
X-ray crystallography Diffraction-quality crystals of M84E-NTF2 were obtained by vapour diffusion using 4 ml hanging drops composed of 2 ml of 2.5 mg/ml M84E-NTF2 in100 mM Tris-HCl (pH 7.4), 100 mM NaCl, 2 mM MgCl2, 5 mM DTT, 0.1 mM PMSF and 2 ml of well buffer (100 mM sodium acetate (pH 4.5), 50 mM MgCl2, 12 % (w/v) PEG 8000). A crystal of dimensions 50 mm 50 mm 25 mm was transferred to cryoprotectant (well buffer containing 20 % (v/v) glycerol) for less than one minute and was Ê resolution native data set ¯ash-frozen at 100 K. A 2.0 A was collected at 100 K using Cu Ka radiation on a Rigaku generator equipped with Osmic mirrors and a MAR345 detector (Table 2). Diffraction quality crystals of M102E-NTF2 were obtained by vapour diffusion using hanging drops composed of 2 ml of drop buffer containing 2.7 mg/ml of M102E-NTF2 and 2 ml of 100 mM sodium acetate (pH 5.3), 50 mM MgCl2, 10 % PEG 8000 well buffer. A crystal was transferred to cryoprotectant (well buffer containing 20 % glycerol and 20 % PEG 8000) for less than one minute and was ¯ash-frozen at 100 K Ê resolution data set collected in-house and a 2.2 A (Table 2). Diffraction quality crystals of M118E-NTF2 were obtained by vapour diffusion using hanging drops composed of 2 ml of 2.5 mg/ml M118E-NTF2 in drop buffer and 2 ml of 100 mM sodium acetate (pH 4.5), 50 mM MgCl2, 26 % PEG 4000 well buffer. A crystal was transferred to cryoprotectant (well buffer containing 12 % glycerol and 35 % PEG 4000) for less than one minÊ resolution ute and was ¯ash-frozen at 100 K and a 2.3 A Ê wavelength radinative data set collected using 1.488 A
Binding to RanGDP and FxFG nucleoporins M118E-NTF2 was tested for its potential to bind RanGDP or FxFG nucleoporins by pull-down experiments with S-tagged Ran or S-tagged NSP1-FF18 sepharose beads. The beads were obtained by incubating S-agarose matrix (Novagen) with a saturating amount of Escherichia coli lysate expressing S-tagged Ran or S-tagged NSP1-FF18, for one hour at 4 C. To remove non-speci®cally bound proteins from the lysate, beads were washed with PBST and subsequently incubated with wild-type NTF2 or M118E-NTF2 in PBST buffer on a rotating wheel for one hour at 4 C. After two washes with PBST buffer, the NTF2 molecules bound to the Stagged Ran or NSP1-FF18 on the beads were recovered in SDS loading buffer after boiling, and visualized by running the samples on a 10 % to 20 % (w/v) acrylamide SDS/polyacrylamide gradient gel.49 The relative binding of the M118E-NTF2 for S-tagged RanGDP or NSP1-FF18 was then determined by comparing the level of WTNTF2 and M118E-NTF2 recovered under the same conditions. Because the binding to nucleoporins was weaker than to RanGDP,20 we employed W7A-NTF2 as a negative control. Electron microscopy Xenopus laevis were purchased from Xenopus I (Ann Arbor, MI, USA) and maintained in ®ltered arti®cial pond water. Stage VI oocytes, approximately 1.2 mm in
475
NTF2 Monomer-Dimer Equilibrium Table 2. Statistics for P21 crystals of NTF2 mutants NTF2 mutant A. Cell dimensions Ê) a (A Ê) b (A Ê) c (A b (deg.) B. Data collection and reduction Source Ê) Wavelength (A Ê) Resolution range (A Ê) Highest resolution bin (A Total/unique reflectionsa hIi/shIia Completeness (%)a Multiplicitya Rmerge (%)a,b C. Structure refinement Working/free reflections Protein/water non-H atoms Working/freec R-factor (%)d Ê 2) Average B-factor (A rms deviation from ideality Ê) Bonds (A Angles (deg.) Residues in Ramachandran region (%) Most favoured Additionally allowed Generously allowed Forbidden
M84E
M102E
M118E
34.29 79.44 41.75 103.80
35.15 79.07 41.65 104.06
34.81 78.23 41.34 104.45
CuKa 1.5418 22.3-2.0 2.11-2.0 36,984/14,697 10.5 (3.5) 99.7 (99.8) 2.5 (2.5) 5.4 (18.2)
CuKa 1.5418 30-2.23 2.37-2.23 81,646/10,835 11.4 (2.9) 99.7 (99.2) 3.2 (3.1) 8.7 (42.9)
SRS 7.2 1.488 30-2.30 2.38-2.30 73,003/9197 9.6 (2.4) 96.2 (96.2) 2.5 (2.0) 9.8 (27.2)
13,944/735 1988/117 20.7/25.1 20
10,757/549 1988/183 20.4/24.1 34
8722/475 1988/112 20.3/28.9 28
0.010 1.5
0.009 2.1
0.007 1.3
93.2 5.9 0.5 0.5 (B92)
90.9 7.7 0.9 0.5 (92B)
89.1 8.2 1.8 0.9 (92A,B)
a
Highest resolution shell in parentheses. Rmerge hklijIhkl ÿ hIhklij/hklihIhkli, where hIhkli is the mean of the observations Ihkl,i of re¯ection hkl. c Free-R was computed using 10 % of the data assigned randomly. d R-factor 100(hkl[jFo(hkl)j ÿ jFc(hkl)j]/hkljFo(hkl)j), where Fo and Fc are the observed and calculated structure factors, respectively. b
diameter, were obtained from mature females and had well-delineated animal and vegetal hemispheres. ColÊ in diameter were coated loidal gold particles 20-50 A with wild-type NTF2 or the M118E mutant and microinjected into manually defolliculated stage VI oocytes as described36 using a Narishige micromanipulator and 10-15 mm tip diameter micropipettes. After approximately 60 minutes, the injected oocytes were ®xed with glutaraldehyde followed by OsO4, dehydrated and embedded as described.50 Sections were examined in a JEOL 100CX electron microscope. Protein Data Bank accession numbers Co-ordinates for the M84E-, M102E- and M118E crystal structures have been deposited at the RCSB Protein Data Bank with accession numbers 1JB2, 1JB4, and 1JB5.
Acknowledgements We are most grateful to our colleagues in Cambridge, especially Airlie McCoy, Helen Kent and Richard Bayliss for their many helpful comments and criticisms, and to Trevor Littlewood and Richard Bayliss for assistance with binding studies. C.C.-H. held an EMBO Postdoctor-
al Fellowship (ALTF98-121). This work was supported, in part, by Research grant RG0270/1998 from the Human Frontiers Science Program.
References 1. GoÈrlich, D. & Kutay, U. (1999). Transport between the cell nucleus and the cytoplasm. Annu. Rev. Cell Dev. Biol. 15, 607-660. 2. Nakielny, S. & Dreyfuss, G. (1999). Transport of proteins and RNAs in and out of the nucleus. Cell, 99, 677-690. 3. Bayliss, R., Corbett, A. H. & Stewart, M. (2000). The molecular mechanism of transport of macromolecules through nuclear pore complexes. Traf®c, 1, 448-456. 4. Wente, S. R. (2000). Gatekeepers of the nucleus. Science, 288, 1374-1377. 5. Rout, M. P. & Wente, S. R. (1994). Pores for thought: nuclear pore complex proteins. Trends Cell Biol. 4, 357-365. 6. Rout, M. P., Aitchison, J. D., Suprapto, A., Hjertaas, K., Zhao, Y. & Chait, B. T. (2000). The yeast nuclear pore complex: composition, architecture and transport mechanism. J. Cell Biol. 148, 635-652.
476 7. Ryan, K. J. & Wente, S. (2000). The nuclear pore complex: a protein machine bridging the nucleus and cytoplasm. Curr. Opin. Cell Biol. 12, 361-371. 8. Doye, V. & Hurt, E. (1997). From nucleoporins to nuclear pore complexes. Curr. Opin. Cell Biol. 9, 401411. 9. Quimby, B. B., Lamitina, T., L'Hernault, S. W. & Corbett, A. H. (2000). The mechanism of ran import into the nucleus by nuclear transport factor 2. J. Biol. Chem. 275, 28575-28582. 10. Dasso, M. (2001). Running on ran: nuclear transport and the mitotic spindle. Cell, 104, 321-324. 11. Melchior, F. & Gerace, L. (1998). Two-way traf®cking with Ran. Trends Cell. Biol. 5, 175-179. 12. GoÈrlich, D. (1998). Transport into and out of the cell nucleus. EMBO J. 17, 2721-2727. 13. Bayliss, R., Littlewood, T. D. & Stewart, M. (2000). Structural basis for the interaction between FxFG nucleoporin repeats and importin-b in nuclear traf®cking. Cell, 102, 99-108. 14. Bayliss, R., Ribbeck, K., Akin, D., Kent, H. M., Feldherr, C. M., GoÈrlich, D. & Stewart, M. (1999). Interaction between NTF2 and xFxFG-containing nucleoporins is required to mediate nuclear import of RanGDP. J. Mol. Biol. 293, 579-593. 15. Damelin, M. & Silver, P. A. (2000). Mapping interactions between nuclear transport factors in living cells reveals pathways through the nuclear pore complex. Mol. Cell, 5, 133-140. 16. Ribbeck, K., Lipowsky, G., Kent, H. M., Stewart, M. & GoÈrlich, D. (1998). NTF2 mediates nuclear import of Ran. EMBO J. 17, 6587-6598. 17. Smith, A., Brownawell, A. & Macara, I. G. (1998). Nuclear import of Ran is mediated by the transport factor NTF2. Curr. Biol. 8, 1403-1406. 18. Stewart, M. (2000). Insights into the molecular mechanism of nuclear traf®cking using nuclear transport factor 2 (NTF2). Cell Struct. Funct. 25, 217-225. 19. Yamada, M., Tachibana, T., Imamoto, N. & Yoneda, Y. (1998). Nuclear transport factor p10/NTF2 functions as a Ran-GDP dissociation inhibitor (Ran-GDI). Curr. Biol. 8, 1339-1342. 20. Chaillan-Huntington, C., Villa-Braslavsky, C., Kuhlmann, J. & Stewart, M. (2000). Dissecting the interactions between NTF2, RanGDP and the nucleoporin xFxFG repeats. J. Biol. Chem. 275, 58745879. 21. Corbett, A. H. & Silver, P. A. (1996). The NTF2 gene encodes an essential, highly conserved protein that functions in nuclear transport in vivo. J. Biol. Chem. 271, 18477-18484. 22. Paschal, B. M., Fritze, C., Guan, T. & Gerace, L. (1997). High levels of the GTPase Ran/TC4 relieve the requirement for nuclear protein transport factor 2. J. Biol. Chem. 272, 21534-21539. 23. Lane, C. M., Cushman, I. & Moore, M. S. (2000). Selective disruption of nuclear import by a functional mutant nuclear carrier protein. J. Cell Biol. 151, 321-332. 24. Suyama, M., Doerks, T., Braun, I. C., Sattler, M., Izaurralde, E. & Bork, P. (2000). Prediction of structural domains of TAP reveals details of its interaction with p15 and nucleoporins. EMBO Rep. 1, 53-58. 25. Liker, E., Fernandez, E., Izaurralde, E. & Conti, E. (2000). The structure of the mRNA export factor TAP reveals a cis arrangement of a non-canonical RNP domain and an LRR domain. EMBO J. 19, 5587-5598.
NTF2 Monomer-Dimer Equilibrium 26. Katahira, J., Strasser, K., Podtelejnikov, A., Mann, M., Jung, J. U. & Hurt, E. (1999). The Mex67pmediated nuclear mRNA export pathway is conserved from yeast to human. EMBO J. 18, 2593-2609. 27. Black, B. E., Levesque, L., Holaska, J. M., Wood, T. C. & Paschal, B. M. (1999). Identi®cation of an NTF2-related factor that binds Ran-GTP and regulates nuclear protein export. Mol. Cell. Biol. 19, 86168624. 28. Black, B. E., Holaska, J. M., Levesque, L., OssarehNazari, B., Gwizdek, C., Dargemont, C. & Paschal, B. M. (2001). NXT1 is necessary for the terminal step of Crm1-mediated nuclear export. J. Cell Biol. 152, 141-156. 29. Bachi, A., Braun, I. C., Rodrigues, J. P., Pante, N., Ribbeck, K. & von Kobbe, C. et al. (2000). The Cterminal domain of TAP interacts with the nuclear pore complex and promotes export of speci®c CTEbearing RNA substrates. RNA, 6, 136-158. 30. Strasser, K. & Hurt, E. (2000). Binding of the Mex67p/Mtr2p heterodimer to FXFG, GLFG and FG repeat nucleoporins is essential for nuclear mRNA export. J. Cell Biol. 150, 695-706. 31. Bullock, T. L., Clarkson, W. D., Kent, H. M. & Ê resolution crystal Stewart, M. (1996). The 1.6 A structure of nuclear transport factor 2 (NTF2). J. Mol. Biol. 260, 422-431. 32. Stewart, M., Kent, H. M. & McCoy, A. J. (1998). Structural basis for molecular recognition between nuclear transport factor 2 (NTF2) and the GDPbound form of the Ras-family GTPase Ran. J. Mol. Biol. 277, 635-646. 33. Ellis, R. J. (2001). Macromolecular crowding: an important but neglected aspect of the intracellular environment. Curr. Opin. Struct. Biol. 11, 114-119. 34. Clarkson, W. D., Kent, H. M. & Stewart, M. (1996). Separate binding sites on nuclear transport factor 2 (NTF2) for GDP-Ran and the phenylalanine-rich repeat regions of nucleoporins p62 and Nsp1p. J. Mol. Biol. 263, 517-524. 35. Clarkson, W. D., Corbett, A. H., Paschal, B. M., Kent, H. M., McCoy, A. J. & Gerace, L. et al. (1997). Nuclear protein import is decreased by engineered mutants of nuclear transport factor 2 (NTF2) that do not bind GDP Ran. J. Mol. Biol. 272, 716-730. 36. Feldherr, C. M., Akin, D. & Moore, M. S. (1998). The nuclear import factor p10 regulates the functional size of the nuclear pore complex during oogenesis. J. Cell Sci. 111, 1889-1896. 37. Paschal, B. M. & Gerace, L. (1995). Identi®cation of NTF2, a cytosolic factor for nuclear import that interacts with nuclear pore protein p62. J. Cell Biol. 129, 925-937. 38. Moore, M. S. & Blobel, G. (1994). Puri®cation of a Ran-interacting protein that is required for protein import into the nucleus. Proc. Natl Acad. Sci. USA, 91, 10212-10216. 39. Wong, D. H., Corbett, A. H., Kent, H. M., Stewart, M. & Silver, P. A. (1997). Interaction between the small GTPase Ran/Gsp1p and Ntf2p is required for nuclear transport. Mol. Cell. Biol. 17, 3755-3767. 40. Van Holde, K. E. & Baldwin, R. L. (1958). Rapid attainment of sedimentation equilibrium. J. Phys. Chem. 62, 734-743. 41. Laue, T. M., Shah, B. D., Ridgeway, T. M. & Pelletier, S. L. (1992). Computer-aided interpretation of analytical sedimentation data for proteins. In Analytical Ultracentrifugation in Biochemistry and Polymer Science (Harding, S. E., Rowe, A. J. & Horton, J. C.,
NTF2 Monomer-Dimer Equilibrium
42.
43. 44.
45.
eds), pp. 90-125, Royal Society of Chemistry, Cambridge, UK. Johnson, M. L. & Straume, M. (1994). Comments on the analysis of sedimentation equilibrium experiments. In Modern Analytical Ultracentrifugation (Schuster, T. M. & Laue, T. M., eds), pp. 37-65, Birkhauser, Boston, MA. Collaborative Computational Project Number 4 (1994). The CCP4 suite: programs for protein crystallography. Acta Crystallog. sect. D, 50, 760-763. BruÈnger, A. T., Adams, P. D., Clore, G. M., DeLano, W. L., Gros, P. & Grosse-Kunstleve, R. W. et al. (1998). Crystallography and NMR system: a new software suite for macromolecular structure determination. Acta Crystallog. sect. D, 50, 905-921. BruÈnger, A. T. (1992). Free R value: a novel statistical quality for assessing the accuracy of crystal structures. Nature, 355, 472-474.
477 46. Matthews, B. W. (1968). The solvent content of protein crystals. J. Mol. Biol. 33, 491-497. 47. Jones, T. A., Zou, J.-Y., Cowan, S. W. & Kjeldgaard, M. (1991). Improved methods for building protein models in electron density maps and for locating errors in these models. Acta Crystallog. sect. A, 47, 110-119. 48. Esnouf, R. M. (1997). An extensively modi®ed version of MolScript that includes greatly enhanced colouring capabilities. J. Mol. Graph. Model. 15, 132136. 49. Laemmli, U. K. (1970). Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature, 227, 680-685. 50. Dworetzky, S. I. & Feldherr, C. M. (1988). Translocation of RNA-coated gold particles through the nuclear pores of oocytes. J. Cell Biol. 106, 575-584.
Edited by B. Holland (Received 5 July 2001; received in revised form 24 September 2001; accepted 25 September 2001)