* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project
Download Tubulin folding is altered by mutations in a putative GTP binding motif
Survey
Document related concepts
Protein phosphorylation wikipedia , lookup
Endomembrane system wikipedia , lookup
Extracellular matrix wikipedia , lookup
G protein–coupled receptor wikipedia , lookup
Protein moonlighting wikipedia , lookup
Organ-on-a-chip wikipedia , lookup
Cytokinesis wikipedia , lookup
Nuclear magnetic resonance spectroscopy of proteins wikipedia , lookup
Folding@home wikipedia , lookup
Signal transduction wikipedia , lookup
Intrinsically disordered proteins wikipedia , lookup
Protein folding wikipedia , lookup
Chemical biology wikipedia , lookup
Proteolysis wikipedia , lookup
Transcript
Journal of Cell Science 109, 1471-1478 (1996) Printed in Great Britain © The Company of Biologists Limited 1996 JCS3328 1471 Tubulin folding is altered by mutations in a putative GTP binding motif Juan C. Zabala1,*, Ana Fontalba1 and Jesus Avila2 1Departamento de Biologia Molecular, Facultad de Medicina, Universidad de Cantabria, Spain 2Centro de Biologia Molecular, Facultad de Ciencias, Universidad Autonoma de Madrid (CSIC-UAM), Spain *Author for correspondence (e-mail: [email protected]) SUMMARY Tubulins contain a glycine-rich loop, that has been implicated in microtubule dynamics by means of an intramolecular interaction with the carboxy-terminal region. As a further extension of the analysis of the role of the carboxyterminal region in tubulin folding we have mutated the glycine-rich loop of tubulin subunits. An α-tubulin point mutant with a T150rG substitution (the corresponding residue present in β-tubulin) was able to incorporate into dimers and microtubules. On the other hand, four βtubulin point mutants, including the G148rT substitution, did not incorporate into dimers, did not release monomers, but were able to form C900 and C300 complexes (intermediates in the process of tubulin folding). Three other mutants within this region (which approximately encompasses residues 137-152) were incapable of forming dimers and C300 complexes but gave rise to the formation of C900 complexes. These results suggest that tubulin goes through two sequential folding states during the folding process, first in association with TCP1-complexes (C900) prior to the transfer to C300 complexes. It is this second step that implies binding/hydrolysis of GTP, reinforcing our previous proposed model for tubulin folding and assembly. INTRODUCTION stacked rings of 8 non-identical subunits (Lewis et al., 1992; Kubota et al., 1994, 1995). Recently, TCP1 particles have been implicated in actin and tubulin folding (Frydman et al., 1992; Gao et al., 1992; Melki et al., 1993; Rommelaere et al., 1993; Sternlicht et al., 1993; Yaffe et al., 1992). Additionally, TCP1containing chaperonin forms a binary complex with completely unfolded actin or tubulin expressed in E. coli (Gao et al., 1992, 1993). Folding of tubulins follows a complex pathway in which different multimolecular complexes (termed C900 and C300) have been implicated (Zabala and Cowan, 1992; Yaffe et al., 1992; Fontalba et al., 1993, 1995; Campo et al., 1994; Fig. 1). Soon after tubulin is synthesized it is bound to a TCP-1-containing chaperone particle (C900). Pulse-chase experiments have shown that, in the presence of nucleotides tubulin dimers and release factors, the disappearance of C900 complexes is correlated with the formation of C300 complexes and monomers and ultimately with the formation of tubulin dimers (Campo et al., 1994). Recently, we have purified a molecular chaperone named p14 responsible of the release of monomers from C300 complexes (Campo et al., 1994). This protein, p14 (or a closely related protein, cofactor A) has also been identified as a factor that facilitates release of tubulin monomers from the TCP1 complex by modulating the ATPase activity of its cognate chaperonin (Gao et al., 1994), though the interaction of cofactor A with CCT complexes has not been confirmed. Thus, p14 seems to have a pleiotropic role and is important for tubulin subunit folding, via the TCP1 complex, and via the C300 complex. Protein folding in the cell is an intricate process that often requires the assistance of preexisting proteins collectively known as molecular chaperones (Ellis and van der Vies, 1991; Georgopoulos and Welch, 1993; Craig et al., 1994; Morimoto et al., 1994). Two major groups of heat-shock proteins are established as molecular chaperones: the Hsp70 and the Hsp60 (or chaperonin) families. The chaperonin family is conserved in all organisms. The first discovered chaperonin, more than twenty years ago, was GroEL (Georgopoulos et al., 1973). High-resolution crystal structures of GroEL have recently been obtained (Braig et al., 1994). They show a porous cylinder of 14 subunits made of two heptameric rings. Also, an extensive mutational analysis of GroEL has been reported (Fenton et al., 1994). This analysis identified a putative polypeptide-binding domain that is essential for binding of the co-chaperonin GroES, which is required for productive polypeptide release. GroEL in Escherichia coli, Hsp60 in mitochondria and Rubisco-subunit binding protein (RBP) in chloroplasts having similar primary and quaternary structures have been called group I chaperonins (Willison and Kubota, 1994). Recently a second group (group II) of chaperonins has been discovered in archaebacteria and eukaryotes. They are significantly related to the group I chaperonins (Kubota et al., 1994, 1995). The eukaryotic cytosol contains TCP1-complexes, also called CCT, TRiC or cytosolic chaperonin (Lewis et al., 1992; Frydman et al., 1992; Gao et al., 1992). TCP1 is a 60 kDa subunit of the cytosolic hetero-oligomeric chaperonin consisting of two Key words: Native electrophoresis, Tubulin folding, GTP-binding, Site-directed mutant 1472 J. C. Zabala, A. Fontalba and J. Avila RF? Mg PU UT CCT Mg PT ATP AA AA,AA AA RF (p14,?) 2+ GTP ADP C 300 2+ GDP M, D Fig. 1. Folding and incorporation of newly synthesized tubulin into dimers. Newly synthesized (unfolded, UT) tubulin forms a binary complex with CCT particles containing different subunits of TCP-1 related proteins. Tubulin subunits (prefolded, PT) released from CCT particles form complexes of about 300 kDa (C300). p14 chaperone releases β-tubulin monomers (M) from C300 complexes and dimer (D) formation is dependent on GTP hydrolysis, Mg2+ ions and other unidentified release factors (RF). Tubulins are GTP-binding proteins (Jacobs et al., 1974; Weisenberg et al., 1976). β-tubulin is a GTPase, while αtubulin has no enzyme activity (Carlier, 1982). Tubulins have an invariant region rich in glycines that is found in α-, β-, and γ-chains and which is presumed to form a phosphate-binding loop (Burns, 1995). The consensus phosphate binding site (GXXXXGK) common to all non-tubulin GTP-binding proteins does not occur in the forward direction (aminocarboxy) in the sequence of either α- or β-tubulin though it is present in the backward direction (Sternlicht et al., 1987). Davis et al. (1994) have shown that this region encompassing amino acids 103-109, a highly conserved region of β-tubulins, participates in GTP-binding and hydrolysis. A critical feature of the phosphate binding sequence in non-tubulin NTP-binding proteins is that it is contained in a glycine-rich loop that contains a carboxy-terminal lysine residue (Möller and Amons, 1985). Computational analysis of α- and β-tubulin sequences shows both to contain in the forward direction another glycinerich region, commonly observed in NTP-binding proteins (each of which has the highest probability of forming a looped structure); however, in neither case does the sequence contain the lysine residue characteristic of the phosphate loops of nontubulin NTP-binding proteins. It is therefore unclear whether this probable loop constitutes a phosphate binding site in tubulins. The recent observation that a segment of the bacterial FtsZ protein matches very well with the glycine-rich region in tubulins supports its GTP-binding function. FtsZ is an essential cell division protein in E. coli; it is a GTPase and it forms a dynamic structure at the division site under cell cycle control (de Boer et al., 1992; RayChaudhuri and Park, 1992). The systematic mutational analysis carried out on the yeast β-tubulin gene has shown that the lethal substitutions are located in three regions of the protein. One of these regions is the glycine rich loop. Lethal alleles identify regions presumably more critical for β-tubulin function (Reijo et al., 1994). Furthermore, it has been discussed that a mutation in this putative phosphate binding motif both reduces GTP binding and suppresses microtubule dynamics. For these reasons, Sage et al. (1995) concluded that β-tubulin possesses a cryptic GTPase superfamily motif (GXXXXGK) as well a glycine-rich phosphate binding motif (GGGTGSG). These results support the suggestion that this glycine-rich region is involved in the interaction of tubulin with GTP (Burns, 1995). It has previously been shown that incorporation of tubulin subunits into tubulin dimers from C300 complexes and/or monomers requires GTP hydrolysis (Fontalba et al., 1993). Also, GTP is required for the stabilization of the monomeric form once released, but incubation of C300 complexes with GTPγS (a non-hydrolyzable GTP analog) prior to the addition of purified p14 protein completely preclude the release of monomers. Thus, it seems that GTP hydrolysis is required to get monomers released from C300 complexes, as suggested by pulse-chase experiments (Campo et al., 1994). These points lead to the intriguing hypothesis that the hydrolysis required for the folding of the native β-tubulin is an intrinsic property of the β-tubulin protein. Since α-tubulin is a GTP-binding protein that probably does not hydrolyze GTP (Carlier, 1982), it would be possible to test this hypothesis by comparing the effects on the folding process of equivalent mutations in α- and β-tubulins. Padilla et al. (1993) have proposed that in β-tubulin an intramolecular interaction between the C terminus and the putative GTP-binding loop may regulate the dynamic instability of microtubules. Deletion of the carboxy terminus of βtubulin slows down their dimerization rates. The C terminus is important for the folding process itself facilitating the folding of the nascent polypeptide, although proper folding takes place in molecules lacking the C terminus (Fontalba et al., 1995). Thus, we have proposed that the interaction of the GTP binding region with the C terminus could stabilize the folded GTPbinding site during the whole process that ends with the formation of the tubulin dimer. To test these hypotheses, different mutated forms in the putative GTP-binding motif were studied in relation to their coassembly properties and the ability to form the different molecular forms that are found during the folding and dimerization processes. MATERIALS AND METHODS Materials Reticulocyte extracts were obtained from Promega, [35S]methionine (>1,000 Ci/mmol) from Amersham, GTP and m7G(5′)ppp(5′) from Boehringer Mannheim or Serva, and some of the oligonucleotides from Isogen Bioscience (Amsterdam). Generation of expression plasmids containing sitedirected mutations To generate site-directed mutants of β1- or α4-tubulin, different restriction fragments containing sequences encoding wild-type β1 and α4 tubulin (Wang et al., 1986; Villasante et al., 1986) were isolated and cloned into M13 vectors. Briefly, a 0.84 kb BalI-SacI fragment encoding internal sequences of β1 was isolated and cloned into bacteriophage M13mp19 DNA digested with SmaI and SacI. Oligonucleotide-directed changes (Table 1) were introduced using the method of Kunkel (1985). Subcloned 0.75 kb AatII-SacI fragments containing the site-directed changes were substituted for wild-type β1-tubulin sequences contained in the β1-pSV expression plasmid (Fig. 2). The correct substitution of fragments containing a single amino acid change was confirmed by restriction analysis using the appropriate enzyme for which a site was created or destroyed and sequence analysis of the final construct. In the case of double mutants β1M147K/G148T and β1M147F/G148T (Table 1), the 0.75 kb AatII-SacI fragments were substituted for β1K147 tubulin sequences contained in the β1K147-pSV expression plasmid. The correct substitution of fragments containing two amino acid changes was confirmed by restriction analysis with AvaI and Site-directed mutagenesis of tubulins 1473 Table 1. Single and double amino acid substitutions introduced into β1- and α4-tubulins Oligonucleotide primers (5′-3′) *CCATGC(C/A)TGAG(C/G)CCGTGC(C/G)T(C/T)CACCCA ” GAGCAGGGTGCCCTTGCCCGAGCCCGTGCC GAGCAGGGTGCCAAAGCCCGAGCCCGTGCC GAGCAGGGTG(G/C)TCATGCCCGAGCCCGTGCC ” CATGAGCAGGCTGCCCATGCCCGAGCCCGTGCC CATGAGCAGGGTGGTCTTGCCTGAGCCCGTGCC CATGAGCAGGGTGGTAAAGCCTGAGCCCGTGCC CAGCAGAGAGC(T/C)GAAGCCAGAGCC ” CAGCAGAGAGGTC(T/A)TGCCAGAGCCGGT ” Complementary sequence (nt) Mutations 416-442 ” 424-453 ” ” ” 424-456 ” ” 436-459 ” 433-459 ” β1G141E β1G146C β1M147K β1M147F β1G148S β1G148T β1T149S β1M147K/G148T β1M147F/G148T α4T150G α4T150S α4F149K α4F149M Nucleotide changes were introduced into the cDNAs encoding murine wild-type β1- and α4 tubulin isotypes by site-directed mutagenesis (see Materials and Methods). Oligonucleotides used and their complementary coding sequences are shown. Several oligonucleotides were synthesized with ambiguities to obtain several mutants from each. Nucleotides in brackets mean 50% each. The first oligonucleotide (*) was used to produce several mutants from which only β1G141E and β1G146C were used in this work. Nucleotide changes introduced to produce amino acid substitutions appear in bold. Nucleotide changes that created or destroyed a restriction site without an amino acid substitution were introduced in some cases and appear underlined (see Materials and Methods). Fig. 2. Construction of mutant β1 cDNA-containing plasmids. The BalI-SacI fragment of β1 was cloned into M13mp19 DNA digested with SmaI and SacI. Oligonucleotide changes (Table 1) were introduced using the method of Kunkel (1985). Subcloned AatII-SacI fragments were substituted for wild-type β1sequences previously cloned into the pSV-expression plasmid. These constructs were used for in vivo copolymerization experiments (see Materials and Methods). In the case of double mutants (Table 1) these fragments were substituted for β1K147sequences (see Materials and Methods). In order to carry out the in vitro experiments, mutant sequences contained in the pSV expression plasmid were isolated and cloned into the SP6-plasmid pSP64. Restriction sites: L (BalI), A (AatII), S (SacI), H (HindIII), E (EcoRI) and M (SmaI). The arrowhead indicates the position at which nucleotide changes were introduced. 1474 J. C. Zabala, A. Fontalba and J. Avila sequence analysis of the final construct. To generate equivalent constructs under the control of the SP6 promoter, the 1.3 kb HindIIIEcoRV fragments with either single or double amino acids changes were substituted for wild-type or β1K147 tubulin sequences contained in the SP6-plasmid pSP64. To create equivalent constructs of α4 mutants, 0.5 kb BamHI-PstI and 0.6 kb EcoRI-SphI fragments encoding internal sequences of α4 were isolated and cloned into bacteriophage M13mp18 DNA digested with BamHI-PstI or EcoRI-SphI. Oligonucleotide-directed changes (Table 1) were introduced using the method of Kunkel (1985). Subcloned fragments containing the site-directed changes were substituted for wild-type α4-tubulin sequences contained in the α4-pSP64 expression plasmid. The correct substitution of fragments containing each amino acid change was confirmed by restriction analysis and sequence analysis of the final construct. Non-coupled in vitro transcription and translation cDNAs encoding wild-type or mutant α4- and β1-tubulins (Lewis et al., 1985; Villasante et al., 1986; Wang et al., 1986) cloned into SP6 vectors were used as templates for in vitro transcription (Melton et al., 1984). Plasmids were linearized with ScaI, PvuI or HindIII and transcribed in vitro using SP6 RNA polymerase in the presence of m7G(5′)ppp(5′). Transcribed mRNAs were translated in a rabbit reticulocyte cell-free system (Pelham and Jackson, 1976) in the presence of [35S]methionine (Amersham; >1,000 Ci/mmol) for 1 hour at 30°C. β-TUBULIN E 140 Other techniques Purified brain tubulin was prepared by consecutive passages through phosphocellulose and cation-exchange FPLC as described (Zabala and Cowan, 1992; Fontalba et al., 1993). Copolymerization of in vitro synthesized wild-type and mutant tubulins with microtubule proteins were performed essentially as previously described (Zabala and Cowan, 1992; Fontalba et al., 1995). Native gel electrophoresis was carried out as previously described (Zabala and Cowan, 1992; Fontalba et al., 1993, 1995; Campo et al., 1994). RESULTS Analysis of in vitro synthesized site-directed α- and β-tubulin mutants Fontalba et al. (1993, 1995) proposed a general pathway for tubulin folding and dimer assembly (Fig. 1). In this model, newly synthesized tubulin forms a complex with the chaperonin TCP-1 before it is transferred to C300 complexes as intermedi- GGGTGSGMGT 149 K-T F-T 100 200 300 400 AA AA C N α-TUBULIN MG KS 142 GGGTGSGFT(A)S 151 100 N Transfection of cultured cells and immunofluorescence analyses Adherent HeLa cells or BSC-1 monkey fibroblast cells free of mycoplasma contamination were grown on glass coverslips placed on Petri dishes in Dulbecco’s modified Eagle medium containing 10% defined calf serum. The cells were transfected by calcium phosphate precipitation with pSV vectors (Mulligan and Berg, 1981) containing full-length cDNAs encoding either wild-type β1-tubulin (Wang et al., 1986) or site-directed mutants. Forty-eight hours after transfection, cells were fixed, permeabilized and analyzed by double label indirect immunofluorescence (Osborn and Weber, 1982) using the rabbit β1-specific antibody (Lewis and Cowan, 1988) (to detect the expression of transfected sequences) together with a guinea pig anti-α-tubulin antibody (to detect endogenous α-tubulin isotypes). Visualization was done after incubation with the secondary antibodies, FITC-conjugated goat antirabbit IgG (Boehringer Mannheim) and rhodamine-conjugated goat anti-guinea pig IgG (Cappel). Cells were viewed with an immunofluorescence microscope using a Zeiss Plan-Neofluar ×63 objective and photographed. FT C KS S 200 300 400 AA AA C Fig. 3. Site-directed mutagenesis in the glycine rich loop. Diagrams for murine α- and β-tubulins show the glycine-rich loop as a black box. Hatched regions illustrate the divergent carboxy-terminal domain. Dotted regions represent one of the most conserved regions present in tubulins close to the carboxy terminus. The white box at the amino terminus of β-tubulin denote the position of first four aminoacids implicated in an autoregulatory mechanism (Pachter et al., 1987). Amino acid numbers are shown as superscripts. Sitedirected changes are shown both in α- and β-tubulins (summarized in Table 1). All of them contain single amino acid changes except FT and K-T, which contain double amino acid changes. Shaded symbols represent mutant β-tubulins that stop in the C900 step of the folding process. Outline symbols represent mutant β-tubulins that stop after the C300 step. Unmodified symbols represent mutant proteins that behave as wild-type tubulins. The shadowed symbol in α-tubulin represents the altered mutant described in the text. Amino acid sequences are presented in the one letter code. ates in the assembly of the heterodimer. Tubulin is a GTPbinding protein (Jacobs et al., 1974) that, in the absence of GTP, quickly loses its functional conformation. Thus, tubulin should bind GTP during its folding process. We have used site-directed mutagenesis to study the effect of mutations of the GTP binding site in the formation of the different multimolecular complexes, monomers and dimers, which takes place during the folding process. We constructed several site-directed mutations within the glycine-rich loop, which approximately encompasses residues 137-152, and are indicated in Fig. 3. We first analyzed the products of in vitro translation of αand β-tubulin mutants by electrophoresis in an SDS-polyacrylamide gel. Under these denaturing conditions, each polypeptide has a mobility indistinguishable from the mobility of the corresponding native tubulin, or from in vitro translated wild-type α- or β-tubulins (data not shown). Fig. 4 shows the analysis of nine β-tubulin and four α-tubulin different mutants under non-denaturing conditions and in the presence of GTP. All of the α4-tubulin mutants except one (α4F149K) gave a Site-directed mutagenesis of tubulins 1475 Fig. 4. Analysis of altered α- and β-tubulins under non-denaturing electrophoresis. Translation products analyzed by electrophoresis under non-denaturing conditions through 7% (A) and 4.5% (B) polyacrylamide gels. C900, C300, D and M denote the position of different multimolecular complexes, dimers and monomers, respectively. pattern indistinguishable from that of wild-type α4-tubulin. In the case of mutant α4F149K, the dimeric band characteristic of wild-type α4-tubulin (Zabala and Cowan, 1992) was not observed, suggesting that this mutation interferes with the ability to fold and dimerize and thus to incorporate into microtubules. Beta-tubulin mutants β1M147K, β1G148S, β1G148T and β1M147F/G148T result in a similar pattern. In each of these mutants, neither of the two faster migrating bands (monomers and dimers) characteristic of wild-type β-tubulin was observed. The bands corresponding in mobility to the most prominent bands characteristic of wild-type β1-tubulin (C300) are also evident. β1G141E, β1G146C and β1M147K/G148T mutants gave neither the bands corresponding to monomers and dimers nor the ones corresponding to C300 complexes. Finally, other mutants (β1M147F and β1T149S), gave patterns indistinguishable from those of wildtype β-tubulin. To address if those mutants that behave differently from the wild-type β-tubulin give the band corresponding to the complex with TCP-1 (C900), aliquots of in vitro translation reactions were analyzed on 4.5% polyacrylamide gels (Fontalba et al., 1993). All mutants that gave rise to the formation of C300 complexes form C900 complexes too (data not shown). Fig. 4B shows the analysis of mutants β1G141E, β1G146C, and β1M147K/G148T in which the Fig. 5. Analysis of the coassembly properties of site-directed mutants of β1- and α4-tubulins. (A and B) Coassembly experiments with altered tubulin β1G148T. (C and D) Coassembly experiments with mutant tubulin β1T149S. E and F show coassembly experiments with altered tubulin α4T150G. (G and H) Corresponding coassembly experiments with mutant tubulin α4F149K. Aliquots containing the same amount of total protein from two consecutive and complete cycles of assembly/disassembly were analyzed in two ways: by SDSpolyacrylamide gel electrophoresis (data not shown), and by nondenaturing gel electrophoresis in the presence of GTP (see Materials and Methods). (A,C,E,G) Gels were stained with Coomassie blue; (B,D,F,H) autoradiograms of the same gels as in A, C, E and G, respectively. Tracks: T, aliquots of the in vitro translation reactions; S1 and S2, supernatants after the first and second cycles; P1 and P2, pellets from the first and second cycles; C and D indicate the position of C300 complexes and dimers, respectively, after non-denaturing electrophoresis. presence of the band characteristic of C900 complexes can be observed but not the one corresponding to the other molecular forms. These results support our previous suggestion that the formation of the C300 complexes occurs after tubulin is released from TCP-1-complexes (Fontalba et al., 1993). Coassembly properties of in vitro synthesized βtubulin mutants To assess the ability with which mutant tubulin polypeptides incorporate into microtubules in vitro, we performed two complete cycles of assembly/disassembly with added bovine brain microtubules in vitro (Fig. 5). The products generated were analyzed by electrophoresis under denaturing (data not 1476 J. C. Zabala, A. Fontalba and J. Avila shown) and native conditions. The results found were analogous for all the polypeptides that behave similarly when analyzed by electrophoresis under native conditions. All mutants which gave rise to the formation of the different molecular species were capable of incorporating as dimers into microtubules in vitro (i.e. β1T149S and α4 T150G; Fig. 5D,F); in a similar fashion to that of the wild-type β1 or α4-tubulin isotypes (data not shown). Mutants that cannot form the corresponding molecular forms were not incorporated into microtubules (i.e. β1G148T and α4F149K; Fig. 5B,H). Expression of wild-type and mutant β-tubulin proteins in vivo To assess the ability of tubulin mutants to form heterodimers and to coassemble into microtubules in vivo, each mutation was introduced into sequences encoding a β-tubulin isotype, β1 (Wang et al., 1986). This isotype is not normally expressed in tissue culture cells, but is freely incorporated into all microtubules in these cells without effect on cell growth or viability when expressed from transfected DNA constructs (Lewis et al., 1987). The mutant tubulin cDNAs were cloned into an expression vector, and the expression of transfected DNAs was monitored by indirect double label immunofluorescence using tubulin isotype-specific antisera (Lewis and Cowan, 1988; Gu et al., 1988) that distinguish the expression of introduced DNA from endogenously expressed tubulin genes. Data from such experiments are shown in Fig. 6. Mutants that gave a normal pattern when analyzed by non-denaturing electrophoresis (Fig. 4) or coassembled into microtubules in vitro (Fig. 5) coassembled in vivo to give a microtubule pattern that was essentially identical to that detected using a general α-tubulin antibody (e.g. Fig. 6A,B). On the other hand, several mutants (β1G141E, β1G146C, β1M147K, β1G148S, β1G148T, β1M147F/G148T and β1M147K/G148T) gave either a pattern of diffuse punctate spots (Fig. 6D) or cytoplasmic aggregates (Fig. 6F). In either case, these patterns differ radically from the normal microtubule network present in the transfected cells (Fig. 6C,E). These results are consistent with the data obtained from in vitro experiments, which showed that these mutants behave differently on non-denaturing gels (Fig. 4) and on the in vitro copolymerization experiments (Fig. 5) from wild-type β1 as well as from all other mutants tested. Furthermore, the altered polypeptides are not colocalized with α-tubulin in vivo in the double label experiments (Fig. 6C,E). Therefore, it seems probable that these Fig. 6. Coassembly properties of β-tubulin mutants. (A,C,E) General α-tubulin antibody. (B,D,F) Fields identical to A,C,E, respectively, stained with the isotype-specific anti-β1-tubulin antibody (Lewis et al., 1987). (B) Phenotype (in HeLa cells) of wild-type β1tubulin. Note the presence of an untransfected cell that labels with the general anti-α-tubulin antibody (A, arrowed) but not with the anti-β1-specific antibody (B). (D and F) Phenotypes of mutants β1G141E (in BSC-1 cells) and β1G146C (in HeLa cells) (see Table 1). Cells were analyzed by double label indirect immunofluorescence (Osborn and Weber, 1982) using the rabbit β1-specific antibody (Lewis and Cowan, 1988) (to detect the expression of transfected sequences) and a general guinea pig antiα-tubulin antibody (to detect endogenous α-tubulins). Bar, 20 µm. Site-directed mutagenesis of tubulins 1477 mutant β-tubulins are incapable of forming heterodimers with any endogenous α-tubulin isotype, in vivo. DISCUSSION Tubulin is a GTP-binding protein (Jacobs et al., 1974; Weisenberg et al., 1976) composed of two subunits (α and β). GTP binds to both subunits but whereas the β-subunit could interchange the bound GTP with exogenous nucleotides, a tight nonexchangeable binding was observed for the tubulin α-subunit. Thus, it is conceivable that tubulin subunits bind GTP during their folding process (probably in a different way for each subunit), in a process that occurs by the formation of different multimolecular complexes, in addition to monomers and dimers. The interaction of tubulin with the chaperone particle containing TCP-1 appears to be a fast initial step in the folding process (Yaffe et al., 1992; Campo et al., 1994). This particle seems to contain, in addition to TCP-1 related proteins (Kubota et al., 1994), two Hsp70 proteins (Lewis et al., 1992). Different chaperones assist in the folding of other unrelated proteins along complicated pathways from the unfolded state to the final correctly folded protein (Ellis and van der Vies, 1991). Hsp70 has been involved in binding to microtubules at the carboxy terminus of tubulin, like other microtubule-associated proteins (Sanchez et al., 1994). Moreover, genetic analysis in culture cells suggested the implication of Hsp70 in the folding process of tubulin (Ahmad et al., 1990). Completely denatured tubulins form binary complexes with TCP-1-containing particles in the absence of GTP (Gao et al., 1993), though the release of completely folded tubulin from C300 complexes requires GTP hydrolysis (Fontalba et al., 1993). The purpose of this work was to analyze the different steps in tubulin folding through the study of the effect of single amino acid substitutions in the putative phosphate binding motif of the tubulin molecule. The choice of mutations in the glycine-rich loop for analysis is made difficult because, in the absence of the lysine residue common to the phosphate-binding consensus sequence in nontubulin GTP-binding proteins, it is impossible to confidently align the glycine residues in tubulins with the consensus sequence glycine residues. Therefore, we elected to analyze the behavior of mutations in which one or more of the residues in the β-tubulin glycine-rich loop were changed (Fig. 3). Alpha and β-tubulins differ from one another in their nucleotide-binding properties: while β-tubulins bind GTP and hydrolyze it during microtubule assembly, it is thought that α-tubulins bind GTP in a nonexchangeable manner (Carlier, 1982). Also, β-tubulins have a lower affinity site that accepts ATP as well as GTP, in addition to the very specific GTP site, while α-tubulins have a site more specific for ATP (Jayaram and Haley, 1994). For these reasons, α- and β-tubulins may belong to two different classes of GTPbinding proteins (Dever et al., 1987). Thus, they would differ in some of the regions implicated in GTP hydrolysis. Though αand β-tubulins are very closely related, we decided to introduce those amino acids present in the glycine-rich loop in α-tubulins at the corresponding positions in β-tubulins and vice versa. Two types of mutant tubulins were found: (i) mutant polypeptides able to associate with C900 and C300 complexes; and (ii) mutant polypeptides that only associate with C900 complexes. A glycine substitution at position 148 of β1 for threonine, the corresponding residue present in murine α-tubulins, affected the capacity of the mutant tubulin to be released as monomers and to incorporate into dimers. This mutant is still able to associate with C900 and C300 complexes. On the other hand, a threonine substitution at position 150 in α4 for the corresponding residue in β-tubulins, glycine, did not affect the ability of the mutant αtubulin to incorporate into dimers and microtubules and thus, proper folding. This result is in agreement with the presence of amino acids different from threonine in that location of the αtubulin subunit from other organisms (Burns and Surridge, 1994). Due to the fact that the most prominent difference between αand β-tubulins resides in their GTP-binding and hydrolysis properties and that the glycine-rich loop has been implicated in this phenomenon, the implication of this glycine in the binding and/or hydrolysis of GTP in the case of β-tubulins seems likely. Also, since the GTP bound to β-tubulin subunit could be hydrolyzed and taking into account the results observed for its related protein, FtsZ, it could be suggested that GTP hydrolysis in β-tubulin would be required for dimer formation. Although it is difficult to establish whether a given mutation prevents the proper folding by interfering with GTP binding and/or hydrolysis or by disrupting folding in a GTP-independent manner, the point explained above strengthens our suggestion. Also, more striking substitutions in the glycine-rich loop of both α- and βtubulins, generated mutants (i.e. β1M147K/G148T) able to associate with C900 complexes (to which tubulin associates in a denatured form; Gao et al., 1993) but not with C300 complexes. In these cases, it is more probable that the mutations we introduced disrupted the complete folding of tubulins. In any case, every mutant which was able to associate with C300 complexes gave rise to the formation of C900 complexes, suggesting that tubulin emerges during the folding process in at least two different folding states, first in association with C900 prior to its transfer to the second, in association with C300 complexes. These results support our previously proposed model for tubulin folding and assembly (Fontalba et al., 1993). The results found with the above mutants suggest that the presence of glycine at position 148 is indeed required for a functional GTP binding site and that β-tubulin GTPase activity is required for β-tubulin monomer release from C300 complexes. It could be instructive to study whether microtubules assembled with the equivalent α-tubulin mutant in yeast might have altered stabilities as well as to demonstrate that these mutations do affect β-tubulin GTPase activity. We thank our colleagues in Santander, Juan M. Garcia-Lobo and Fernando de la Cruz, for helpful discussions, advice and critical comments on the manuscript; Rafael Campo for technical assistance; Maria Lizama for corrections to the English; and Nicholas J. Cowan for providing us with some of the oligonucleotides used in this work and for the opportunity to realize the immunofluorescences with isotype-specific antibodies in his laboratory. We also would like to acknowledge the comments and suggestions of one of the reviewers. This work was supported by DGICYT (to J.C.Z.). REFERENCES Ahmad, S., Ahuja, R., Venner, T. J. and Gupta, R. S. (1990). Identification of a protein altered in mutants resistant to microtubule inhibitors as a member of the major heat shock protein (hsp70) family. Mol. Cell. Biol. 10, 51605165. Braig, K., Otwinowski, Z., Hedge, R., Boisvert, D. C., Joachimiak, A., Horwich, A. and Sigler, P. B. (1994). The crystal structure of the bacterial chaperonin GroEL at 2.8 Å. Nature 371, 578-586. Burns, R. G. and Surridge, C. D. (1994). Tubulin: conservation and structure. 1478 J. C. Zabala, A. Fontalba and J. Avila In Microtubules (ed. J. S. Hyams and C. W. Lloyd), pp. 3-31. Wiley-liss, John Wiley and Sons, Inc., publication, New York. Burns, R. G. (1995). Analysis of the γ-tubulin sequences: implications for the functional properties of γ-tubulin. J. Cell Sci. 108, 2123-2130. Campo, R., Fontalba, A., Sanchez, L. M. and Zabala, J. C. (1994). A 14 kDa release factor is involved in GTP-dependent β-tubulin folding. FEBS Lett. 353, 162-166. Craig, E. A., Weissman, J. S. and Horwich, A. L. (1994). Heat shock proteins and molecular chaperones: mediators of protein conformation and turnover in the cell. Cell 78, 365-372. Carlier, M. F. (1982). Guanosine 5′ triphosphate hydrolysis and tubulin polymerization. Mol. Cell. Biochem. 47, 97-113. Davis, A., Sage, C. R., Dougherty, C. A. and Farrel, K. W. (1994). Microtubule dynamics modulated by guanosine triphosphate hydrolysis activity of β-tubulin. Science 264, 839-841. de Boer, P., Crossley, R. and Rothfield, L. (1992). The essential bacterial celldivision protein FtsZ is a GTPase. Nature 359, 254-256. Dever, T. E., Glynias, M. J. and Merrick, W. C. (1987). GTP-binding domain: three consensus sequence elements with distinct spacing. Proc. Nat. Acad. Sci. USA 84, 1814-1818. Ellis, R. J. and van der Vies, S. M. (1991). Molecular chaperones. Annu. Rev. Biochem. 60, 321-347. Fenton, W. A., Kashi, Y., Furtak, K. and Horwich, A. L. (1994). Residues in chaperonin GroEL required for polypeptide binding and release. Nature 371, 614-619. Fontalba, A., Paciucci, R., Avila, J. and Zabala, J. C. (1993). Incorporation of tubulin subunits into dimers requires GTP hydrolysis. J. Cell Sci. 106, 627-632. Fontalba, A., Avila, J. and Zabala, J. C. (1995). β-tubulin folding is modulated by the isotype-specific carboxy-terminal domain. J. Mol Biol. 246, 628-636. Frydman, J., Nimmesgern, E., Erdjument-Bromage, H., Wall, J. S., Tempst, P. and Hartl, F. U. (1992). Function in protein folding of TRiC, a cytosolic ring complex containing TCP1 and structurally related subunits. EMBO J. 11, 4767-4778. Gao, Y., Thomas, J. O., Chow, R. L., Lee, G.-H. and Cowan, N. J. (1992). A cytoplasmic chaperonin that catalyzes β-actin folding. Cell 69, 1043-1050. Gao, Y., Vainberg, I. E., Chow, R. L. and Cowan, N. J. (1993). Two cofactors and cytoplasmic chaperonin are required for the folding of α- and β-tubulin. Mol. Cell Biol. 13, 2478-2485. Gao, Y., Melki, R., Walden, P. D., Lewis, S. A., Ampe, C., Rommelaere, H., Vandekerckhove, J. and Cowan, N. J. (1994). A novel cochaperonin that modulates the ATPase activity of cytoplasmic chaperonin. J. Cell Biol. 125, 989-996. Georgopoulos, C. P., Hendrix, R. W., Casjens, S. R. and Kaiser, A. D. (1973). Host participation in bacteriophage lambda head assembly. J. Mol. Biol. 76, 45-60. Georgopoulos C. and Welch, W. J. (1993). Role of the major heat shock proteins as molecular chaperones. Annu. Rev. Cell Biol. 9, 601-634. Gu, W., Lewis, S. A. and Cowan N. J. (1988). Generation of antisera that discriminate among mammalian α-tubulins: introduction of specialized isotypes into cultured cell results in their coassembly without disruption of normal microtubule function. J. Cell Biol. 106, 2011-2022. Jacobs, M., Smith, H. and Taylor, E. W. (1974). Tubulin: nucleotide binding and enzymatic activity. J. Mol. Biol. 89, 455-468. Jayaram, B. and Haley, B. E. (1994). Identification of peptides within the base binding domains of the GTP- and ATP-specific binding sites of tubulin. J. Biol. Chem. 269, 3233-3242. Kubota, H., Hynes, G., Carne, A., Ashworth, A. and Willison, K. (1994). Identification of six Tcp-1-related genes encoding divergent subunits of the TCP-1-containing chaperonin. Curr. Biol. 4, 89-99. Kubota, H., Hynes, G. and Willison, K. (1995). The chaperonin containing tcomplex polypeptide 1 (TCP-1). Multisubunit machinery assisting in protein folding and assembly in the eukaryotic cytosol. Eur. J. Biochem. 230, 3-16. Kunkel, T. A. (1985). Rapid and efficient site-specific mutagenesis without phenotypic selection. Proc. Nat. Acad. Sci. USA 82, 488-492. Lewis, S. A., Lee, M. G.-S. and Cowan, N. J. (1985). Five mouse tubulin isotypes and their regulated expression during development. J. Cell Biol. 101, 852-861. Lewis, S. A., Gu, W. and Cowan, N. J. (1987). Free intermingling of mammalian β-tubulin isotypes among functionally distinct microtubules. Cell 49, 539-548. Lewis, S. A. and Cowan, N. J. (1988). Complex regulation and functional versatility of mammalian α- and β-tubulin isotypes during the differentiation of testis and muscle cells. J. Cell Biol. 106, 2023-2033. Lewis, V. A., Hynes, G. M., Zheng, D., Saibil, H. and Willison, K. (1992). Tcomplex polypeptide-1 is a subunit of a heteromeric particle in the eukaryotic cytosol. Nature 358, 249-252. Melki, R., Vainberg, I. E., Chow, R. L. and Cowan, N. J. (1993). Chaperonin-mediated folding of vertebrate actin-related protein and γtubulin. J. Cell Biol. 122, 1301-1310. Melton, D. A., Kreig, P. A., Rebagliati, M. R., Maniatis, T., Zinn, K. and Green, M. R. (1984). Efficient in vitro synthesis of biologically active RNA hybridization probes from plasmids containing a bacteriophage SP6 promoter. Nucl. Acids Res. 12, 7035-7056. Möller, W. and Amons, R. (1985). Phosphate-binding sequences in nucleotide-binding proteins. FEBS Lett. 186, 1-7. Morimoto, R. I., Tissieres, A. and Georgopoulos, C. (1994). Progress and perspectives on the biology of heat shock proteins and molecular chaperones. In The Biology of Heat Shock Proteins and Molecular Chaperones (ed. R. I. Morimoto, A. Tissieres and C. Georgopoulos), pp. 1-30. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Mulligan, R. C. and Berg, P. (1981). Selection for animal cells that express the Escherichia coli gene coding for xanthine-guanine phosphoribosyl transferase. Proc. Nat. Acad. Sci. USA 78, 2072-2076. Osborn, M. and Weber, K. (1982). Immunofluorescence and immunocytochemical procedures with affinity purified antibodies: tubulin containing structures. Meth. Cell Biol. 244, 98-129. Pachter, J. S., Yen T. J. and Cleveland, D. W. (1987). Autoregulation of tubulin expression is achieved through specific degradation of polysomal tubulin mRNAs. Cell 51, 283-292. Padilla, R., Otin, C. L., Serrano, L. and Avila, J. (1993). Role of the carboxyterminal region of β-tubulin on microtubule dynamics through its interaction with the GTP phosphate binding region. FEBS Lett. 325, 173-176. Pelham, H. R. B. and Jackson, R. J. (1976). An efficient mRNA-dependent translation system from reticulocyte lysates. Eur. J. Biochem. 67, 247-256. RayChaudhuri, D. and Park, J. T. (1992). Escherichia coli cell-division gene ftsZ encodes a novel GTP-binding protein. Nature 359, 251-254. Reijo, R. A., Cooper, E. M., Beagle, G. J. and Huffaker, T. C. (1994). Systematic mutational analysis of the yeast β-tubulin gene. Mol. Biol. Cell 5, 29-43. Rommelaere, H., Van Troys, M., Gao, Y., Melki, R., Cowan, N. J., Vandekerckhove, J. and Ampe, C. (1993). Eukaryotic cytosolic chaperonin contains t-complex polypeptide 1 and seven related subunits. Proc. Nat. Acad. Sci. USA 90, 11975-11979. Sage, C. R., Dougherty, C. A., Davis, A. S., Burns, R. G., Wilson, L. and Farrell, K. W. (1995). Site-directed mutagenesis of putative GTP-binding sites of yeast β-tubulin: Evidence that α-, β-, and γ-tubulins are atypical GTPases. Biochemistry 34, 7409-7419. Sanchez, C., Padilla, R., Paciucci, R., Zabala, J. C. and Avila, J. (1994). Binding of heat-shock protein 70 (hsp70) to tubulin. Arch. Biochem. Biophys. 310, 428-432. Sternlicht, H., Yaffe, M. B. and Farr, G. W. (1987). A model of the nucleotide binding site of tubulin. FEBS Lett. 214, 226-235. Sternlicht, H., Farr, G. W., Sternlicht, M. L., Driscoll, J. K., Willison, K. and Yaffe, M. B. (1993). The t-complex polypeptide 1 complex is a chaperonin for tubulin and actin in vivo. Proc. Nat. Acad. Sci. USA 90, 9422-9426. Villasante, A., Wang, D., Dobner, P., Dolph, P., Lewis, S. A. and Cowan, N. J. (1986). Six mouse α-tubulin mRNAs encode five distinct isotypes: testisspecific expression of two sister genes. Mol. Cell Biol. 6, 2409-2419. Wang, D., Villasante, A., Lewis, S. A. and Cowan N. J. (1986). The mammalian β-tubulin repertoire: hematopoietic expression of a novel, heterologous β-tubulin isotype. J. Cell Biol. 103, 1903-1910. Weisenberg, R. C., Deery, W. J. and Dickinson, P. J. (1976). Tubulinnucleotide interactions during the polymerization and the depolymerization of microtubules. Biochemistry 15, 4248-4252. Willison, K. and Kubota, H. (1994). The structure, function and genetics of the chaperonin containing TCP-1 (CCT) in eukaryotic cytosol. In The Biology of Heat Shock Proteins and Molecular Chaperones (ed. R. I. Morimoto, A. Tissieres and C. Georgopoulos), pp. 1-30. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Yaffe, M. B., Farr, G. W., Miklos, D., Horwich, A. L., Sternlicht, M. L. and Sternlicht, H. (1992). TCP1 complex is a molecular chaperone in tubulin biogenesis. Nature 358, 245-248. Zabala, J. C. and Cowan, N. J. (1992). Tubulin dimer formation via the release of α- and β-tubulin monomers from multimolecular complexes. Cell Motil. Cytoskel. 23, 222-230. (Received 11 September 1995 - Accepted 29 February 1996)