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Different Smad2 partners bind a common hydrophobic pocket in Smad2 via a defined proline-rich motif

2002; Springer Nature; Volume: 21; Issue: 1 Linguagem: Inglês

10.1093/emboj/21.1.145

1460-2075

Rebecca A. Randall, Stéphane Germain, Gareth J. Inman, Paul A. Bates, Caroline S. Hill,

Renal and related cancers

Article15 January 2002free access Different Smad2 partners bind a common hydrophobic pocket in Smad2 via a defined proline-rich motif Rebecca A. Randall Rebecca A. Randall Laboratory of Developmental Signalling, Imperial Cancer Research Fund, 44 Lincoln's Inn Fields, London, WC2A 3PX UK Search for more papers by this author Stéphane Germain Stéphane Germain Present address: INSERM U36, Collège de France, 11 Place Marcelin Berthelot, 75231 Paris, France Search for more papers by this author Gareth J. Inman Gareth J. Inman Laboratory of Developmental Signalling, Imperial Cancer Research Fund, 44 Lincoln's Inn Fields, London, WC2A 3PX UK Search for more papers by this author Paul A. Bates Paul A. Bates Biomolecular Modelling Laboratory, Imperial Cancer Research Fund, 44 Lincoln's Inn Fields, London, WC2A 3PX UK Search for more papers by this author Caroline S. Hill Corresponding Author Caroline S. Hill Laboratory of Developmental Signalling, Imperial Cancer Research Fund, 44 Lincoln's Inn Fields, London, WC2A 3PX UK Search for more papers by this author Rebecca A. Randall Rebecca A. Randall Laboratory of Developmental Signalling, Imperial Cancer Research Fund, 44 Lincoln's Inn Fields, London, WC2A 3PX UK Search for more papers by this author Stéphane Germain Stéphane Germain Present address: INSERM U36, Collège de France, 11 Place Marcelin Berthelot, 75231 Paris, France Search for more papers by this author Gareth J. Inman Gareth J. Inman Laboratory of Developmental Signalling, Imperial Cancer Research Fund, 44 Lincoln's Inn Fields, London, WC2A 3PX UK Search for more papers by this author Paul A. Bates Paul A. Bates Biomolecular Modelling Laboratory, Imperial Cancer Research Fund, 44 Lincoln's Inn Fields, London, WC2A 3PX UK Search for more papers by this author Caroline S. Hill Corresponding Author Caroline S. Hill Laboratory of Developmental Signalling, Imperial Cancer Research Fund, 44 Lincoln's Inn Fields, London, WC2A 3PX UK Search for more papers by this author Author Information Rebecca A. Randall1, Stéphane Germain2, Gareth J. Inman1, Paul A. Bates3 and Caroline S. Hill 1 1Laboratory of Developmental Signalling, Imperial Cancer Research Fund, 44 Lincoln's Inn Fields, London, WC2A 3PX UK 2Present address: INSERM U36, Collège de France, 11 Place Marcelin Berthelot, 75231 Paris, France 3Biomolecular Modelling Laboratory, Imperial Cancer Research Fund, 44 Lincoln's Inn Fields, London, WC2A 3PX UK *Corresponding author. E-mail: [email protected] The EMBO Journal (2002)21:145-156https://doi.org/10.1093/emboj/21.1.145 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Transforming growth factor-β (TGF-β)/activin-induced Smad2/Smad4 complexes are recruited to different promoter elements by transcription factors, such as Fast-1 or the Mix family proteins Mixer and Milk, through a direct interaction between Smad2 and a common Smad interaction motif (SIM) in the transcription factors. Here we identify residues in the SIM critical for Mixer–Smad2 interaction and confirm their functional importance by demonstrating that only Xenopus and zebrafish Mix family members containing a SIM with all the correct critical residues can bind Smad2 and mediate TGF-β-induced transcriptional activation in vivo. We identify significant sequence similarity between the SIM and the Smad-binding domain (SBD) of the membrane-associated protein SARA (Smad anchor for receptor activation). Molecular modelling, supported by mutational analyses of Smad2 and the SIM and the demonstration that the SARA SBD competes directly with the SIM for binding to Smad2, indicates that the SIM binds Smad2 in the same hydrophobic pocket as does the proline-rich rigid coil region of the SARA SBD. Thus, different Smad2 partners, whether cytoplasmic or nuclear, interact with the same binding pocket in Smad2 through a common proline-rich motif. Introduction Signals from transforming growth factor-β (TGF-β) family members are transduced to the nucleus by members of the Smad family. Upon ligand binding, an active complex is formed comprising activated receptor-regulated Smads (R-Smads), such as Smad2 or Smad3 in the case of the ligands TGF-β and activin, and a co-Smad (Smad4; Massagué and Wotton, 2000). R-Smads are activated directly by phosphorylation at their extreme C-terminus by the type I receptor kinase (Massagué and Wotton, 2000). Smad2 and Smad3 are specifically recruited to the receptor complex by an FYVE domain protein called SARA (Smad anchor for receptor activation; Tsukazaki et al., 1998). After ligand stimulation, active Smad complexes rapidly accumulate in the nucleus, where they are directly involved in gene regulation. Smads bind DNA very weakly with limited specificity and thus cooperate with other transcription factors, which target them to specific DNA-binding sites (Massagué and Wotton, 2000; ten Dijke et al., 2000; Shi, 2001). For example, in early Xenopus embryos, Smad complexes activated by activin-related ligands (consisting of XSmad2 and XSmad4β; Howell et al., 1999; Masuyama et al., 1999) are recruited to DNA by transcription factors, such as the forkhead/winged helix protein, XFast-1 (Chen et al., 1996, 1997), or a subset of paired-like homeodomain transcription factors of the Mix family, Mixer and Milk (Germain et al., 2000). These Smad-interacting transcription factors are key determinants of specificity since they have different DNA-binding domains and thus recruit a common activated Smad complex to different promoter elements to activate distinct sets of target genes. Consistent with this idea, Mix and Fast family members are expressed in different regions of the embryo: XFast-1 is highly expressed in prospective ectoderm and mesoderm, whereas Mixer and Milk are expressed in a ring in deeper layers of the mesoderm and endoderm (Hill, 2001). Many different proteins at all levels of the TGF-β signalling pathways have been reported to interact with different combinations of Smads. For example, proteins such as SARA recruit Smad2 and Smad3 to the receptors for phosphorylation, transcription factors recruit different Smad complexes to DNA, co-activators and co-repressors are involved in modulating transcriptional responses, and E3 ubiquitin ligases, such as the Smurfs, bind Smads, which target them or associated proteins for degradation (ten Dijke et al., 2000). To understand these processes, it is important to define Smad interaction motifs in the partner proteins and to understand how these motifs interact specifically with different Smads. We demonstrated previously that members of the Mix and Fast families that bind active Smad complexes do so through a common 25 amino acid Smad interaction motif (SIM), characterized by a conserved core (P-P-N-K-S/T-I/V), present in their C-terminal regions (Germain et al., 2000). The SIM binds to the C-terminal (MH2) domain of phosphorylated Smad2, which in turn binds Smad4. The SIM displays a high degree of binding specificity, interacting only with the MH2 domains of Smad2 and Smad3, and not with those of the BMP-activated R-Smads or Smad4 (Germain et al., 2000). Here we set out to determine what constitutes a functional SIM, to understand the molecular basis for the interaction of the SIM with Smad2, and to determine whether it is unique to these Smad2/Smad3-interacting transcription factors or whether it may represent a generic Smad interaction motif. Using a combination of in vitro and in vivo binding and transcription assays, we have identified the conserved residues of the Mixer SIM critical for interaction with Smad2. The results demonstrate the importance of the very conserved P-P-N-K-S/T-I/V core and two C-terminal flanking residues, and indicate that the SIM is an extended motif. These preferences are observed in the other Xenopus Mix family members and in zebrafish Mixer [bonnie and clyde (bon); Alexander et al., 1999; Kikuchi et al., 2000], since only those family members that contain a SIM with all the correct critical residues bind Smad2 and mediate TGF-β-induced transcriptional activation in vivo. Of the many reported Smad-interacting proteins, few bind the MH2 domain of Smad2 and show exactly the same specificity of Smad binding as the SIM. The cytoplasmic protein SARA is the best understood Smad partner with these characteristics. We find significant sequence similarity between the SIM and the proline-rich rigid coil region of the Smad-binding domain (SBD) of SARA. We demonstrate that the SIM binds to a region of the Smad2 MH2 domain, which also binds the rigid coil region of the SARA SBD (Wu et al., 2000), and that the SARA SBD competes with the SIM for binding to Smad2. We propose a molecular model in which a shallow hydrophobic groove on the surface of the MH2 domain of Smad2 is responsible for recruiting different Smad2 partners, such as SARA in the cytoplasm or SIM-containing transcription factors in the nucleus, via a defined proline-rich motif. Results Definition of the residues in the SIM critical for interaction with Smad2 Alignment of the known functional SIMs in Fast and Mix family members reveals the characteristic conserved core P-P-N-K-S/T-I/V, flanked by other highly conserved residues (Germain et al., 2000). To understand how the SIM interacts with the Smad2 MH2 domain, we determined which of the conserved residues are absolutely required for interaction with Smad2. Selected amino acids of Mixer were mutated to alanine, either alone or in pairs (Figure 1A), and the resulting mutants were assayed for their ability to interact with Smad2. Figure 1.Identification of residues critical for SIM/Smad2 interaction. (A) The sequence of the Mixer SIM indicating the residues that have been mutated to alanine, either singly or in pairs. The conserved core of the SIM is underlined (Germain et al., 2000). (B) Analysis of the ability of in vitro translated Mixer mutants to interact with GSTSmad2C by bandshift assay using the MBS from the DE as probe. GSTSmad2C was titrated over the range 20, 10, 5, 2.5 and 1.25 ng. Mixer derivatives bound to DNA are indicated, as are ternary complexes containing GSTSmad2C (arrow). (C) Analysis of the ability of 35S-labelled in vitro translated Mixer mutants to interact with GSTSmad2C in a GST pull-down assay. Bound protein was visualized by SDS–PAGE and autoradiography. Lane 1 corresponds to the interaction between wild-type Mixer and GST to estimate non-specific binding. Twenty per cent input protein is shown. The amount of Mixer derivative bound is quantitated as a percentage of input and these values were normalized such that the binding of wild-type Mixer was 100. Three independent experiments gave similar results. (D) Analysis of the ability of Mixer mutants to interact with phosphorylated Smad2 in an IP western blot. Extracts were prepared from uninduced or TGF-β-induced NIH 3T3 cells that had been transfected with the different Flag-tagged Mixer mutants with Myc-tagged Smad2 and Smad4. Extracts were assayed by IP of complexes with anti-Flag antibody followed by western blotting with anti-phospho-Smad2 antibody (P-Smad2, top blot) or by western blotting the whole extract with anti-Myc antibody (middle blot) or with anti-Flag antibody (bottom blot). Download figure Download PowerPoint First, in vitro synthesized Mixer mutants were assayed by bandshift for their ability to interact with a glutathione S-transferase (GST) fusion protein of the Smad2 MH2 domain (GSTSmad2C), using a probe corresponding to the Mixer-binding site (MBS) from the goosecoid DE (Figure 1B; Germain et al., 2000). The assay was made semi-quantitative by titrating GSTSmad2C over a range of concentrations. The single mutations that had the greatest effect on the ability of Mixer to interact with GSTSmad2C were P291A, P292A and N293A in the core, and M300A and P305A in the C-terminal flanking sequence. In all cases, either no supershift was seen with GSTSmad2C at any input concentration (N293A) or efficient supershifts were only detected at the higher concentrations of GSTSmad2C (P291A, P292A, M300A and P305A). The other single mutations either had no effect (F287A, F290A and D299A) or small effects (K294A, T295A and I296A). Mutants with double mutations in core residues (P291A+K294A and T295A+I296A) were almost completely defective for binding to GSTSmad2C, underlining the importance of the core residues for Smad2C binding (Figure 1B). A double mutation in the N-terminal flanking sequence (D286A+F287A) had no effect on GSTSmad2C binding, suggesting that these flanking residues do not play a role in the interaction with Smad2C. All Mixer derivatives bound DNA as well as wild-type Mixer, indicating that mutating the SIM had no impact on DNA binding. The binding of the Mixer mutants to Smad2C was also assayed in the absence of DNA in a GST pull-down assay (Figure 1C). On the whole, the data agreed well with the bandshift assays. Mutation of N293 alone completely blocked Mixer interaction with GSTSmad2C. Other mutations that severely inhibited binding to GSTSmad2C were P292A, M300A, P305A and the combined mutations of P291A+K294A and T295A+I296A in agreement with the bandshift assays (Figure 1C). The P291A mutant was the only one that behaved significantly differently in the two assays; it interacted with GSTSmad2C more efficiently in the pull-down assay than would be expected from the bandshift assay (see below). We then confirmed the relative importance of the core residues of the SIM and the C-terminal flanking residues, M300 and P305, for Smad2 interaction in vivo in an immunoprecipitation (IP) western assay, measuring the ability of the Mixer mutants to interact with phosphorylated Smad2 upon TGF-β induction in NIH 3T3 cells (Figure 1D). The results generally agreed with the in vitro binding analyses, and again indicated that P292 and N293 in the core and M300 and P305 in the C-terminal flanking region are absolutely required in vivo for Mixer to interact with phosphorylated Smad2. In addition, mutation of I296 also had a severe effect on the ability of Mixer to interact with phosphorylated Smad2. The Mixer mutants were then assayed for their ability to recruit active endogenous Smad2/Smad4 complexes to DNA by assessing their ability to mediate TGF-β-induced transcriptional activation via the goosecoid DE in NIH 3T3 cells (Figure 2). Wild-type Mixer mediated a 10-fold increase in transcriptional activation upon TGF-β stimulation due to recruitment of active endogenous Smad complexes (Figure 2; Germain et al., 2000). Mutants severely defective for interaction with Smad2 in some or all of the binding assays were either completely incapable or very poor at mediating TGF-β-induced transcriptional activation in vivo (Figure 2; P292A, N293A, I296A, M300A and P305A and the double mutants P291A+K294A and T295A+I296A). Control bandshifts and western blots of whole-cell extracts made from cells transfected with Flag-tagged wild-type or mutant Mixer derivatives demonstrated they were all efficiently expressed and bound DNA (Figure 1D and data not shown). Figure 2.The ability of Mixer derivatives to interact with Smad2C correlates with their ability to mediate TGF-β-induced transcription via the DE. NIH 3T3 cells were transfected with the (DE)4–luciferase reporter and plasmids expressing Flag-tagged wild-type Mixer or mutants. Cells were incubated for 8 h in the presence or absence of TGF-β. Luciferase was quantitated relative to β-Gal activity and the value for TGF-β-induced transcription using wild-type Mixer was set at 100. The data are the mean and standard deviation of three independent experiments. Download figure Download PowerPoint We can now define residues in the SIM required for Smad2 binding. The conserved core residues P-P-N-K-T-I are important for interacting with Smad2 and recruiting activated Smad2/Smad4 complexes to DNA for transcriptional activation. Of these residues, P292 and N293 are the most critical for Smad2 interaction, since mutation of either one greatly reduced Smad2 binding. The importance of the other core residues only became apparent when double mutations were tested. The residues N-terminal to the core do not appear to play an important role in Smad2 interaction, but two flanking residues C-terminal to the core are critical for the function of the SIM; these are M300 and P305. This indicates that the SIM extends beyond the conserved core. The Mixer SIM is necessary and sufficient for mediating TGF-β-inducible transcription The Mixer SIM is required for mediating TGF-β-inducible transcription (Figure 2; Germain et al., 2000) but is it sufficient or are other sequences in Mixer also required? A fusion of the Mixer SIM (amino acids 283–307) with the Gal4 DNA-binding domain [Gal4(1–95)–SIM] was tested for its ability to confer TGF-β-inducible transcription in NIH 3T3 cells on (Gal4-OP)5–luciferase. The reporter alone was inactive and non-responsive to TGF-β (Figure 3A). Co-transfection of Gal4(1–95)–SIM conferred a low level of transcriptional activity on the reporter in the absence of TGF-β, which was increased ∼6-fold upon TGF-β stimulation (Figure 3A). A mutant SIM fusion protein, Gal4(1–95)–SIM(PP mut), which does not interact with Smad2, was completely inactive in the presence or absence of TGF-β (Figure 3A). Both Gal4 fusion proteins were equally well expressed and bound DNA efficiently (Figure 3B). Figure 3.The SIM is sufficient to confer TGF-β inducibility in vivo. (A) Schematics of the Gal4(1–95) fusion of the SIM (residues 283–307 of Mixer) or mutant SIM in which P291 and P292 are mutated to alanine. NIH 3T3 cells were transfected with (Gal4-OP)5–luciferase together with plasmids expressing Gal4(1–95), Gal4(1–95)–SIM or Gal4(1–95)–SIM(PP mut) as indicated; TGF-β inductions and luciferase assays were as above. The value for TGF-β-induced transcription using Gal4(1–95)–SIM was set at 100. (B) The Gal4(1–95) derivatives are equally expressed. Whole-cell extracts from NIH 3T3 cells transfected with the Gal4(1–95) derivatives as indicated were assayed by bandshift using a Gal4-binding site as a probe. The complexes were confirmed as Gal4(1–95) derivatives bound to DNA since they were all abolished by addition of the anti-Gal4 antibody, which recognizes the DNA-binding domain. (C) TGF-β-inducible transcription mediated by Gal4(1–95)–SIM results from recruitment of active Smad complexes. NIH 3T3 cells were transfected with (Gal4-OP)5–luciferase together with plasmids expressing Gal4(1–95)–SIM and plasmids expressing increasing amounts of Mixer, or Mixer(PP mut) which cannot bind Smad2. TGF-β inductions and luciferase assays were as above. The value for TGF-β-induced transcription using Gal4(1–95)–SIM was set at 100. In (A) and (C), the data are the mean and standard deviation of three independent experiments. Download figure Download PowerPoint Overexpression of Mixer could compete with Gal4(1–95)–SIM for binding to endogenous Smads, confirming that the TGF-β-induced transcriptional activation was due to recruitment of active Smad complexes (Figure 3C). Competition with Mixer(PP mut) had no effect on the ability of the Gal4(1–95)–SIM fusion to confer TGF-β inducibility on the reporter. Thus, the Mixer SIM is both necessary and sufficient to mediate TGF-β-induced transcription in vivo through its interaction with active Smad complexes. Identification of Smad-interacting members of the Xenopus and zebrafish Mix families We have demonstrated the importance of the core residues of the SIM (in particular P292 and N293) and also two critical C-terminal flanking residues (M300 and P305) for interaction with Smad2. To determine whether these preferences are observed in other naturally occurring SIMs, we analysed other Mix family members to see which have functional SIMs. The SIM of Xenopus Mixer was aligned with the equivalent regions of the other Xenopus Mix family members, Mix.1, Bix1, Milk (Bix2), Bix3 and Bix4 (Rosa, 1989; Ecochard et al., 1998; Henry and Melton, 1998), and the zebrafish orthologue of Mixer, bon (Figure 4A; Alexander et al., 1999; Kikuchi et al., 2000). The Xenopus and zebrafish Mixers, Milk and Bix3 all have a recognizable SIM with the P-P-N-K-T-I core and the methionine and proline residues at positions equivalent to 300 and 305 in Xenopus Mixer, respectively. Based on our analysis of the SIM, we would predict that these would all interact with Smad2. Although Mix.1, Bix1 and Bix4 contain a subset of these residues, they do not contain all of them and we would predict that they would not interact with Smad2. Figure 4.Mix family members that contain a SIM interact with Smad2 and mediate TGF-β-induced transcriptional activation in vivo. (A) Alignment of the SIM regions of six Xenopus Mix family members and zebrafish Mixer. Above, family members that contain the P-P-N-K-T-I core and the C-terminal M and P known to be critical for Smad2 interaction (black shading). Below, family members that have no predicted SIM (open boxes). (B) Equal amounts of 35S-labelled in vitro translated Mix family members were analysed by SDS–PAGE and autoradiography. Molecular weights of marker proteins are indicated. (C) Mix family members that contain a SIM interact with GSTSmad2C in a bandshift assay. In vitro translated Flag-tagged Mix family members (as in B) were assayed by bandshift assay using the MBS probe in the presence or absence of antibody directed against the tag (upper panel) or with 20 ng of GSTSmad2C (bottom panel). Mix family members bound to DNA are indicated, as are antibody-supershifted complexes in the top panel and ternary complexes with GSTSmad2C in the bottom panel (arrow). (D) SIM-containing Mix family members mediate TGF-β-induced transcriptional activation. NIH 3T3 cells were transfected with (DE)4–luciferase together with plasmids expressing Mix family members. TGF-β inductions and luciferase assays were as above. The value for TGF-β-induced transcription using Mixer was set at 100. The data are the mean and standard deviation of three independent experiments. Download figure Download PowerPoint The ability of the in vitro translated Mix family members (Figure 4B) to interact with GSTSmad2C was tested in a bandshift assay. All the proteins were supershifted by an antibody recognizing the N-terminal Flag tag on the proteins (Figure 4C, top panel), but only Xenopus and zebrafish Mixers, Milk and Bix3 interacted efficiently with GSTSmad2C and formed a ternary complex (bottom panel). This agrees with the presence of a predicted SIM containing all of the residues that we have shown to be critical. Moreover, the Mix family members that contain a SIM could mediate TGF-β-inducible transcription via the DE (6- to 10-fold for Xenopus Mixer, zebrafish Mixer and Bix3; ∼3-fold for Milk; Figure 4D). Mix.1, Bix1 and Bix4, which were not capable of interaction with Smad2, were unable to confer significant TGF-β-inducible transcription onto the DE (Figure 4D). The Mix family members were all synthesized efficiently in vivo (data not shown). The high basal level of transcription seen in Bix1, Milk, Bix4 and to a lesser extent Bix3, may be due to the presence of Q-rich transcriptional activation domains within the sequence of Bix1–4, which are absent from both Mixer and Mix.1 (Tada et al., 1998). Swapping the SIM of Milk into Mix.1 is sufficient to convert Mix.1 into a Smad-interacting transcription factor We next performed a gain-of-function experiment to determine whether it was possible to convert a non-Smad2-interacting Mix family member, Mix.1, into a Smad2-interacting protein. Instead of the PPNK core characteristic of all functional SIMs, the Mix.1 sequence is QTNK (Figure 4A). In initial experiments, we made four point mutations in Mix.1 to mutate Q300 and T301 to proline and N304 and K306 of Mix.1 to threonine, thus creating in Mix.1 the PPNKTIT of Mixer. This mutant did not interact with Smad2, confirming that additional residues in the SIM are required. Consistent with this, creating the PPNKTI motif in Bix1 also failed to generate a functional SIM (data not shown; see Discussion). Mutations in Mix.1 outside the core must also be required to generate a functional SIM, and amino acids 299–314 of Mix.1 were therefore mutated to the corresponding residues of Milk, which contains a functional SIM, to generate Mix.1–Milk(SIM) (Figure 5A). This Mix.1 derivative conferred very strong TGF-β-inducible transcription on the DE (∼9-fold), comparable with that of Mixer. In contrast, Mix.1 was not TGF-β inducible in this assay. Mix.1–Milk(SIM) also interacted with GSTSmad2C in a bandshift assay (Figure 5B). Thus, swapping the functional Milk SIM into Mix.1 is sufficient to convert it into a Smad2-interacting transcription factor. Figure 5.Replacing the inactive SIM region of Mix.1 with the functional Milk SIM is sufficient to enable Mix.1 to mediate TGF-β-induced transcription activation and bind Smad2C. (A) NIH 3T3 cells were transfected with (DE)4–luciferase together with plasmids expressing Mixer, Mix.1 or the Mix.1–Milk(SIM) chimeric protein. TGF-β inductions and luciferase assays were as above. The value for TGF-β-induced transcription using Mixer was set at 100. The data are the mean and standard deviation of three independent experiments. Schematics of wild-type Mix.1, Milk and the Mix.1–Milk(SIM) chimera are shown. (B) Mixer, Mix.1 and Mix.1–Milk(SIM) were synthesized in vitro and analysed for their ability to interact with 20 ng of GSTSmad2C in a bandshift assay using the MBS as a probe. Both Mixer and Mix.1–Milk(SIM) interact with GSTSmad2C, as seen by the supershift (arrow). Download figure Download PowerPoint Common residues in the Smad2 MH2 domain are required for interaction with the SIM and with SARA Many proteins have been reported to interact with the different Smad family members, both in the nucleus and cytoplasm (ten Dijke et al., 2000), but other than the SIM-containing transcription factors, few interact with the Smad2 MH2 domain with the same specificity as the SIM. The best characterized is the membrane-bound FYVE domain protein SARA, which like the SIM-containing transcription factors, specifically recognizes Smad2 and Smad3, but not Smad1 or Smad4 (Tsukazaki et al., 1998). The structure of the SARA SBD with the Smad2 MH2 domain has been solved (Wu et al., 2000). The SARA SBD is in an extended conformation comprising a conformationally restrained proline-rich rigid coil, an α-helix and a β-strand. The rigid coil is responsible for the specificity of the interaction with Smad2 and makes contacts with Y366 in α-helix 2 of the Smad2 MH2 domain, with W368, T372 and C374 in the loop joining α-helix 2 to strand β-8, and with residues on strands β-8 and β-9 (Wu et al., 2000). Previous work has demonstrated that residues in α-helix 2 of the Smad2 MH2 domain (including Y366) are also required to interact with Mixer, Milk and Fast-1 via the SIM (Chen et al., 1998; Germain et al., 2000), and dictate the specificity of the interaction such that the transcription factors bind Smad2 and not Smad1 (Chen et al., 1998). This suggests that the SARA SBD rigid coil and the SIM might bind Smad2 in a similar manner. Alignment of the two motifs revealed important similarities (Figure 6A). Strikingly, the residues in the Mixer SIM that we have demonstrated to be critical for Smad2 interaction (P292, N293, M300 and P305) are either identical in the two motifs or, in the case of M300, substituted by a residue with similar properties. Figure 6.The SIM binds to a region of the Smad2 MH2 domain that also binds the SARA SBD. (A) Alignment of the Mixer SIM with the rigid coil region of the SARA SBD (Wu et al., 2000). Identical residues are in red, conservative substitutions are in light green. (B) Model of the Mixer SIM bound to the Smad2 MH2 domain. Ribbon diagram of the region of Smad2 MH2 domain (grey) corresponding to α-helix 2 (α2) and β-strands (β5–β9) that interacts with the Mixer SIM (green). All side chains of the SIM peptide are shown. Only the side chains of Smad2 that make direct contact with the peptide are displayed. Residues of Smad2 coloured cyan have been mutated in this study (see text). Mixer SIM side chains are coloured green and yellow; yellow side chains are important for Smad2 interactions and for conserving the peptide conformation. Some key hydrogen bonds are indicated by dotted red lines (see Table I for a description of these interactions). (C) Molecular surface of Smad2 (grey) and a ribbon representation of the Mixer SIM (green) in the same orientation as in (B). The side chains of the Mixer SIM and Smad2, coloured yellow and cyan, respectively, are as in (B). The binding groove on Smad2 is generally shallow with larger indentations in the areas that contact M300 and P306 of the Mixer SIM. The figure was created using the program GRASP (Nicholls et al., 1991). (D) The region of Smad2 shown in the model is aligned with the analogous region of Smad1 (which does not interact with the SIM or with the SARA SBD). Red residues are identical between the two Smads and black residues are different. The assignment of structural motifs is based on Shi et al. (1997); α2 is α-helix 2 and β5–9 are β-strands. Eight residues are indicated, which are demonstrated from our mutation analysis and/or predicted from the model to be important for SIM–Smad2 interaction (B and C; Table I). Six of these are unique to Smad2. Download figure Download PowerPoint To determine whether the Mixer SIM could form a rigid coil similar to that in the N-terminal region of the SARA SBD