Title: The structure of mouse HP1 suggests a unique mode of single peptide recognition by the shadow chromo domain dimer
Abstract: Article3 April 2000free access The structure of mouse HP1 suggests a unique mode of single peptide recognition by the shadow chromo domain dimer Sally V. Brasher Sally V. Brasher Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Brian O. Smith Brian O. Smith Present address: Institute of Cell and Molecular Biology, University of Edinburgh, Edinburgh, EH9 9JR UK Search for more papers by this author Rasmus H. Fogh Rasmus H. Fogh Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Daniel Nietlispach Daniel Nietlispach Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Abarna Thiru Abarna Thiru Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Peter R. Nielsen Peter R. Nielsen Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author R. William Broadhurst R. William Broadhurst Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Linda J. Ball Linda J. Ball Present address: Forschungsinstitut fuer Molekulare Pharmakologie, Alfred-Kowalke-Strasse 4, D-10315 Berlin, Germany Search for more papers by this author Natalia V. Murzina Corresponding Author Natalia V. Murzina Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Ernest D. Laue Corresponding Author Ernest D. Laue Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Sally V. Brasher Sally V. Brasher Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Brian O. Smith Brian O. Smith Present address: Institute of Cell and Molecular Biology, University of Edinburgh, Edinburgh, EH9 9JR UK Search for more papers by this author Rasmus H. Fogh Rasmus H. Fogh Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Daniel Nietlispach Daniel Nietlispach Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Abarna Thiru Abarna Thiru Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Peter R. Nielsen Peter R. Nielsen Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author R. William Broadhurst R. William Broadhurst Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Linda J. Ball Linda J. Ball Present address: Forschungsinstitut fuer Molekulare Pharmakologie, Alfred-Kowalke-Strasse 4, D-10315 Berlin, Germany Search for more papers by this author Natalia V. Murzina Corresponding Author Natalia V. Murzina Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Ernest D. Laue Corresponding Author Ernest D. Laue Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK Search for more papers by this author Author Information Sally V. Brasher1, Brian O. Smith2, Rasmus H. Fogh1, Daniel Nietlispach1, Abarna Thiru1, Peter R. Nielsen1, R. William Broadhurst1, Linda J. Ball3, Natalia V. Murzina 1 and Ernest D. Laue 1 1Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge, CB2 1GA UK 2Present address: Institute of Cell and Molecular Biology, University of Edinburgh, Edinburgh, EH9 9JR UK 3Present address: Forschungsinstitut fuer Molekulare Pharmakologie, Alfred-Kowalke-Strasse 4, D-10315 Berlin, Germany ‡S.V.Brasher and B.O.Smith contributed equally to this work *Corresponding authors. E-mail: [email protected] or E-mail: [email protected] The EMBO Journal (2000)19:1587-1597https://doi.org/10.1093/emboj/19.7.1587 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The heterochromatin protein 1 (HP1) family of proteins is involved in gene silencing via the formation of heterochromatic structures. They are composed of two related domains: an N-terminal chromo domain and a C-terminal shadow chromo domain. Present results suggest that chromo domains may function as protein interaction motifs, bringing together different proteins in multi-protein complexes and locating them in heterochromatin. We have previously determined the structure of the chromo domain from the mouse HP1β protein, MOD1. We show here that, in contrast to the chromo domain, the shadow chromo domain is a homodimer. The intact HP1β protein is also dimeric, where the interaction is mediated by the shadow chromo domain, with the chromo domains moving independently of each other at the end of flexible linkers. Mapping studies, with fragments of the CAF1 and TIF1β proteins, show that an intact, dimeric, shadow chromo domain structure is required for complex formation. Introduction The first heterochromatin-associated protein to be characterized was heterochromatin protein 1 (HP1), a suppressor of position effect variegation (PEV) (James and Elgin, 1986; Eissenberg et al., 1990). HP1 is found in complexes with other known heterochromatin proteins, e.g. Su(var)3–7 (Cleard et al., 1997) and Su(var)3–9 (Aagaard et al., 1999), and its mutation causes recessive embryonic lethality due to defects in chromosome morphology, lengthened prophase and subsequent chromosome segregation (Kellum and Alberts, 1995). Mutations can also result in aberrant association of chromosomes and multiple telomeric fusions (Fanti et al., 1998). HP1 homologues have been found in many other organisms from Schizosaccharomyces pombe (Lorentz et al., 1994; Ekwall et al., 1995) to mammals (Singh et al., 1991; Saunders et al., 1993). There are three HP1 protein family members in mammals, HP1α, HP1β (MOD1) and HP1γ (MOD2), and different patterns of phosphorylation and localization point to possible differences in their function (Minc et al., 1999). In Drosophila, phosphorylation of HP1 is required for efficient heterochromatin targeting and, possibly, heterochromatin assembly (Eissenberg et al., 1994; Zhao and Eissenberg, 1999). In mammals, HP1α and HP1γ exist in different phosphorylated forms, becoming hyper-phosphorylated at mitosis. In contrast, HP1β remains as a unique isoform throughout the cell cycle (Minc et al., 1999). In Drosophila, HP1 is localized to heterochromatin, to telomeres and to discrete regions of euchromatin (Kellum et al., 1995; Fanti et al., 1998). In mouse and human cells, HP1α is found predominantly in centromeres, HP1β (MOD1) is distributed widely on the chromosome, and HP1γ (MOD2) localizes mostly to euchromatin (Minc et al., 1999); their localization changes during the cell cycle in both Drosophila (Kellum et al., 1995) and mammals (Minc et al., 1999; Murzina et al., 1999). Recently, we have shown that as mitosis is approached some HP1β dissociates from heterochromatin, as histone H3 becomes hyper-phosphorylated, and then reassociates at the end of mitosis when H3 is dephosphorylated (Murzina et al., 1999). The HP1 proteins make up one class of chromo domain proteins (Paro and Hogness, 1991), having an N-terminal chromo domain and a related C-terminal shadow chromo domain (Aasland and Stewart, 1995; Koonin et al., 1995). Many other chromo domain proteins are also involved in the regulation of gene expression resulting from alterations in chromatin structure (Cavalli and Paro, 1998).The polycomb protein (Pc) represents a second important class, in which an N-terminal chromo domain is present, but the shadow domain is not. These generally much larger proteins contain a different conserved sequence motif in the C-terminus (Paro, 1990; Paro and Hogness, 1991). The construction of chimeric proteins, consisting of either the HP1 protein with its chromo domain replaced by that from Pc, or the Pc protein with an HP1 chromo domain, has shown that both the chromo and shadow domains are important for correct chromatin localization. These results suggested that they may interact independently with different targets in heterochromatin (Platero et al., 1995). Although HP1 proteins are located in chromatin, they do not appear to bind to DNA directly (Singh et al., 1991; Ball et al., 1997). Rather, they have been found to interact with a number of different proteins. So far, the only known example of an interaction involving the chromo domain is the interaction of Drosophila HP1 with the origin recognition complex (ORC) that is required for initiation of eukaryotic DNA replication (Pak et al., 1997). All other known interactions are mediated via the shadow chromo domain. The importance of the shadow chromo domain in HP1 function is emphasized by the fact that a truncated HP1 mutant Su(var)2–504, lacking part of the shadow domain, does not localize in either heterochromatin, euchromatin or at telomeres (Fanti et al., 1998). Both the mouse and human HP1 proteins have been shown to interact with the transcriptional intermediary factors (TIFs) α and β (Le Douarin et al., 1996; Nielsen et al., 1999; Ryan et al., 1999). TIF1β (or KAP1) also binds to proteins containing the KRAB domain (Kruppel-associated box), one of the most widely distributed transcriptional repressor domains in mammals (Friedman et al., 1996; Moosmann et al., 1996). It has been suggested that the HP1–TIF–KRAB complex might recruit hetero-chromatin-like complexes to specific loci on the chromosome defined by the DNA binding site of the Kruppel transcription factors (Ryan et al., 1999). TIF1α is a nuclear protein that interacts directly with the ligand-dependent activation domain of certain nuclear hormone receptors to suppress transcription (Le Douarin et al., 1995); it has also beenshown to be located in euchromatin (Remboutsika et al., 1999). Both TIF1α and TIFβ appear to repress transcription via histone deacetylases(Nielsen et al., 1999). Recently, we have demonstrated that the large subunit of chromatin assembly factor 1 (CAF1p150) binds to mouse HP1 proteins. The interaction is required for the association of CAF1 with heterochromatin in non-S-phase cells, and CAF1 also promotes the incorporation of MOD1 into nascent chromatin during DNA replication in vitro. We identified a peptide motif that is conserved in both the TIFs (Le Douarin et al., 1996) and the large subunit of CAF1p150 that interacts with the shadow chromo domain (Murzina et al., 1999). Thus the results so far suggest that the HP1 proteins may function as adaptors, bringing together different proteins in multi-protein complexes, and locating them in heterochromatin via protein–protein interactions with the chromo and shadow chromo domains. In order to understand the mechanisms involved we are studying the structure and interactions of the mouse HP1β protein, MOD1 (Murzina et al., 1999). We show here that the full-length MOD1 protein is a dimer where the interaction is mediated via the C-terminal shadow chromo domain (MOD1C). We present the structure of MOD1C and show that an intact dimer structure is required for the interaction with the CAF1 and TIF1β proteins. Results Shadow domain structure Based on limited proteolysis data (Ball et al., 1997), and a sequence alignment of the different HP1 proteins (Figure 1),residues 104–171 of MOD1 were expressed in Escherichia coli and purified for structural studies. The purified MOD1C protein had the expected amino acid composition and molecular mass (data not shown). Studies of 15N relaxation of the backbone amides suggested that the protein was larger and tumbled more slowly in solution than the homologous chromo domain (Figure 2). Equilibrium sedimentation analysis subsequently confirmed that the shadow chromo domain is dimeric in solution (data not shown). Figure 1.Sequences of the chromo domains (A) and the shadow chromo domains (B) from different HP1 proteins, numbered so that they correspond to MOD1. Secondary structure elements observed in MOD1 are shown above the alignments; cylinders represent 310 (3–10) or α-helices (α1 and α2), arrows represent β-strands and circles indicate β-bulges. For each domain the residues that make up the hydrophobic core of a 'subunit’ are shaded in yellow and other residues considered important for the structure are shown in green (Gly and Pro). Charged residues in the chromo domain, which are replaced by hydrophobic residues in the shadow chromo domain, are coloured blue (basic) and red (acidic). The red boxes enclose the structured parts of the proteins. Residues that form the dimer interface in the shadow chromo domain are boxed and shaded in grey. Mutations described in this paper are indicated below the alignment. The proteins are, from the top, mouse MOD1/human HP1β (residues 1–81 and 103–185), mouse HP1γ (1–80 and 97–173), human HP1γ (1–80 and 97–173), human HP1α (1–80 and 106–191), mouse HP1α (1–80 and 106–191), Drosophila melanogaster HP1 (1–84 and 132–206), Drosophila virilis HP1 (1–84 and 139–213) and Schizosaccharomyces pombe SWI6 (59–145 and 252–328). Download figure Download PowerPoint Figure 2.Backbone 15N T2 relaxation times for full-length MOD1 (black), free N-terminal domain and free C-terminal domain (both white). Amide groups in full-length MOD1 were assigned (where possible) by comparing spectra of the full-length protein with those of the individual domains. The T2s were calculated by non-linear least-squares fitting (Broadhurst et al., 1995). Download figure Download PowerPoint The structure of MOD1C is well defined except for the five N-terminal residues, 104–109, and the C-terminal residue, 171, which are flexible in solution as judged by the 15N relaxation experiments (see Figure 3). The shadow chromo domain structure thus corresponds to residues 110–170 of mouse MOD1. The structures satisfy the experimentally derived distance restraints, with on average less than one violation >0.5 Å per structure, and have good stereochemistry and van der Waals packing (see Table I). Figure 3.The structure of the shadow chromo domain dimer from MOD1. The backbone r.m.s.d. for the structure is 0.63 Å over the monomer and 0.81 Å over the dimer. (A) A stereo plot of the backbone traces of the ensemble of 16 calculated structures; the two monomers are depicted in red and blue. (B) A cartoon representation of the shadow chromo domain dimer (again red and blue) with the chromo domain from MOD1 (yellow) for comparison.(C) A close up stereo view of the inter-monomer interface with the side chains of interfacial residues shown; key residues are labelled. These plots were produced using MOLSCRIPT and Raster3D (Kraulis, 1991; Merritt and Bacon, 1997). Download figure Download PowerPoint Table 1. Structural statistics for the final ensemble of 16 refined structures of the HP1β shadow chromo domain All structures Closest to mean NOE restraints no. of restraints used 2090 2090 average restraint violation (Å) 0.0061 ± 0.0006a 0.0055 over one monomer (Å) 0.63 0.42 over both monomers (Å) 0.81 0.62 all heavy atoms over one monomer (Å) 1.08 0.85 over both monomers (Å) 1.17 0.95 Parameter r.m.s.d. from idealized geometry bonds (Å) 0.002 ± 9×10−5b 0.002 angles (°) 0.333 ± 0.010b 0.329 improper dihedrals (°) 0.225 ± 0.018b 0.201 Final energy EL–Jc (kJ/mol) −441 ± 44 −360 Ramachandran plot quality (location of non-Gly and non-Pro residues) most favoured region (%) 69.9 69.8 additionally allowed region (%) 21.8 22.6 disallowed region (%) 2.8 5.7 a The sum of NOE violations divided by the total number of restraints and averaged over the ensemble. b The r.m.s.d. for each structure averaged over the ensemble. c The Lennard–Jones potential was not used at any stage in the refinement. As expected from their homology, each monomer of MOD1C forms a compact fold very similar to that of the chromo domain (see Figure 3). MOD1C, however, forms a symmetrical dimer, burying 687 ± 47 A2 of surface area, in which the interface principally involves the C-terminal α-helices of each monomer. Residues that form the dimer interface in the shadow chromo domain are indicated in Figure 1; the main contacts involve I161, Y164 and L168 (see Figure 3C). There are also significant contacts between residue 153 in the α1 helix of one monomer and residue 161 in the α2 helix of the other, between residue 158 and the peptide backbone of residue 154, and between the side chain of W170 and A125, L132 and Y164. Mutations disrupting the dimer structure The importance of particular residues for dimer formation was investigated by mutating residues either to those found in the monomeric chromo domains or to glutamate, thereby introducing a negative charge into the hydrophobic dimer interface. We then determined the relative size of the different mutant proteins (I161 to A or E, Y164 to L or E and W170 to A or E) using gel-filtration chromatography. As shown in Figure 4, wild-type MOD1C and the Y164L, W170A and W170E mutants eluted at the same column volume, suggesting that the size and shape of each mutant protein are similar to those of the wild type. Changing Y164 to L does not disrupt dimer formation, but did decrease the stability of the protein, which showed a tendency to aggregate and precipitate from solution. In contrast, the Y164E, I161A and I161E mutants appear to have a lower molecular weight than the wild-type protein (see Figure 4). We confirmed that each of these proteins was intact by SDS–PAGE and that the Y164E and I161E mutants were similar in structure to the wild-type protein by circular dichroism (data not shown). Sedimentation analysis of these mutants showed that the I161E mutant is monomeric, whereas the Y164E mutant showed weak association with a Kd of ∼500–600 μM (data not shown). Taken together, the data show that mutation of either residue 161 or 164 results in a protein that is structured and essentially monomeric. The results also confirm the importance of these residues for formation of the dimer interface. Figure 4.MOD1C mutations disrupting the dimer structure. Wild-type MOD1C and the mutants were expressed as His-tagged fusions in E.coli, purified with Ni–NTA spin columns (Qiagen) and loaded directly onto a Superdex S75 gel-filtration column (24 ml bed volume) to assess the size of the proteins. Gel-filtration elution profiles are presented for wild-type MOD1C (wt) and the mutants W170A, W170E, Y164L, Y164E, I161E and I161A. The arrow indicates aggregates that eluted in the void volume of the column. Download figure Download PowerPoint The structure of intact MOD1 We have shown previously that the recombinant chromo domain exists as a monomer in solution (Ball et al., 1997), and in this work we have determined that the shadow chromo domain is a dimer. Sedimentation analysis of the intact protein showed that the molecular weight in solution is 42 kDa, approximately twice that calculated from the sequence (21.5 kDa). These results indicate that the full-length protein is also a dimer, but left open the question as towhat inter-domain interactions occur in native MOD1. The mobility of the individual domains can be estimated from the backbone 15N T2 relaxation times, which depend on the rotational correlation time of the molecule—larger and more slowly tumbling molecules having shorter T2 values. The average 15N T2 is 99 ms for the free N-terminal domain and 62 ms for the free C-terminal domain. These data show that the free N-terminal domain tumbles significantly faster in solution than the free C-terminal domain, consistent with the former being a monomer and the latter a dimer. For the full-length protein, the average 15N T2 is 61 ms for the N-terminal domain and 26 ms for the C-terminal domain. (The individual values and the actual residues compared are shown in Figure 2). In the full-length protein both domains have shortened T2 values, consistent with a larger overall structure. Significantly, however, the T2 values in the C-terminal domain remain lower than those in the N-terminal domain. The difference in 15N T2 values between the two domains suggests strongly that they have different mobility in the full-length protein and must therefore be moving largely independently of each other. The consistently longer T2 values for the N-terminal domain suggest that it remains unassociated with other parts of the molecule in the full-length protein. These conclusions are also supported by the fact that the linker region between the two domains is unstructured (see Figure 2), consistent with its high level of accessibility to proteases (Ball et al., 1997). In summary, the results suggest a structure for full-length MOD1 where it dimerizes through the C-terminal domain alone, with the two N-terminal domains moving independently of each other at the end of flexible linkers. The shadow chromo domain dimer binds to a single CAF MIR peptide Both CAF1p150 (Murzina et al., 1999) and TIF1β (Le Douarin et al., 1996; Nielsen et al., 1999; Ryan et al., 1999) share a conserved peptide motif named MIR, for MOD1 interacting region, which is essential for their interaction with MOD1. To investigate the interaction between MOD1C and CAF1 further, we expressed a 66 amino acid peptide from mouse CAF1p150 (amino acids 204–269) as a His-tagged fusion protein in E.coli. This peptide comprises the overlapping sequence found in all of the fragments of CAF1 that were originally isolated in the two-hybrid screen with MOD1 (Murzina et al., 1999). The stoichiometry of binding was studied using gel-filtration chromatography. Samples were analysed on a Superdex 75 column after mixing varying amounts of CAF MIR with a constant amount of MOD1C (Figure 5). A single peak, corresponding to the complex, was detected during gel filtration of a 1:1 mixture of CAF MIR to MOD1C dimer (Figure 5, trace d). Mixtures with lower CAF MIR:MOD1C dimer ratios showed an additional peak corresponding to free MOD1 (e.g. Figure 5, trace c), whilst mixtures with higher CAF MIR:MOD1C dimer ratios showed an additional peak corresponding to free CAF1 (e.g. trace e). At no ratio could we see all three peaks at the same time, indicating that the complex is stable during gel filtration. It is particularly noteworthy that, at a CAF MIR:MOD1C dimer ratio of 2:1 (trace e), we do see free CAF1. This would not be expected if two molecules of CAF1 bound to one molecule of MOD1C dimer, i.e. if one molecule of CAF1 bound to each subunit of the MOD1C dimer. Figure 5.Titration of MOD1C with the CAF1 MIR shows that one CAF1 peptide binds to one MOD1C dimer. The figure shows traces (A280) from gel filtration on a Superdex S75 column (2.4 ml) of MOD1C only (a) and different mixtures containing MOD1C dimer and CAF MIR in the ratios 1:0.33 (b), 1:0.8 (c), 1:1 (d), 1:2 (e) and 1:4 (f). The purity of the MOD1C and CAF MIR samples was checked by SDS–PAGE prior to mixing the proteins in the appropriate ratios. The arrows mark the positions at which free MOD1C, free CAF MIR and their complex eluted from the column. A small peak was observed eluting in the void volume of the column, suggesting that a small amount of high-molecular-weight aggregates was present in the mixtures. (Note that CAF MIR absorbs less strongly than MOD1C at 280 nm.) Download figure Download PowerPoint The results of the gel filtration suggest that one molecule of CAF MIR binds to one MOD1C dimer. To confirm this we performed equilibrium sedimentation analysis of a 25 residue MIR-containing peptide from CAFp150 (amino acids 211–235, Mr 2736 Da), MOD1C, and the complex of the two. The synthetic peptide was chosen for analytical centrifugation because the larger CAF MIR fragment (residues 204–269) was prone to both degradation and aggregation.MOD1C was mixed with an excess of CAF MIR 25mer and the mixture was eluted through an S75 gel-filtration column to remove excess peptide. Ultracentrifugation gave a molecular weight of 2.93 kDa for the peptide, 16.63 kDa for the MOD1C dimer and 19.17 kDa for the complex. This agrees with the gel-filtration results. [Note that a larger synthetic peptide (25mer) was used here to give a greater molecular weight difference than would have been obtained with the minimal 13mer peptide, see below.] Definition of the MIR peptide binding site on the shadow chromo domain We next sought to identify the MOD1C residues involved in the interaction with mouse CAF1p150. A CAF MIR 13mer (residues 220–232 of mouse CAF1p150), containing only the essential conserved peptide motif found in CAF1 and TIF1β (Murzina et al., 1999), was used in NMR experiments to map its binding site on MOD1C. The 1H and 15N chemical shifts in 2D 1H–15N HSQC spectra of 15N-labelled MOD1C, before and after complex formation with excess unlabelled peptide, were determined to identify residues that are affected by ligand binding or conformational changes. As the majority of cross peaks in the spectrum of the complex were significantly perturbed, a 3D 15N-separated NOESY-HSQC spectrum was used to assign as many of the cross peaks as possible. Of the 58 non-proline residues, whose amide protons do not exchange rapidly with the solvent, 15 had cross peaks that did not change on the addition of peptide. A further 27 gave rise to two HSQC cross peaks of approximately equal intensity, both of which were shifted relative to their position in the isolated protein. For another three residues only one highly perturbed cross peak could be found (the other presumably being too shifted to be easily identified). Finally, the remaining 13 residues could not be identified, their chemical shifts also being too perturbed for analysis with data from this spectrum alone. An estimate of the number of HSQC peaks that remain unassigned suggests, however, that these residues are also likely to give rise to two HSQC cross peaks each. The magnitude of the perturbations in the spectra that are observed upon binding is consistent with the formation of a tight complex and points to some changes in the structure of the protein on complex formation. The residues whose shifts are most strongly perturbed, and therefore most likely to be close to the ligand, form a contiguous region on the surface of the dimer. They lie in the C-terminal end of the second helix (helix α2), the C-terminal tail, the first β-strand and the first half of the second β-strand (see Figure 6). Many of the residues in the protein give rise to two different, perturbed, but identifiable HSQC cross peaks, suggesting that the same residue in the two different MOD1C subunits experiences a different local environment. This is not in itself unexpected, as a complex between a symmetric dimer and a single, non-symmetric peptide must of necessity be asymmetric. It is, however, surprising that the asymmetry is observed over so large a part of the molecule. Neither the NMR nor gel-filtration experiments point to the presence of more than one species in solution, and the presence of free MOD1C can be specifically excluded. There is no sign of free MOD1C in the NMR spectra and its presence is not compatible with a Kd for complex formation of 2 μM, as determined by fluorescence spectroscopy (data not shown). Figure 6.Mapping of the MIR peptide binding site on MOD1C. The molecular surface of the MOD1C dimer is shown (A), together with a cartoon of the structure (B). (The view shown is related to that in Figure 3 by 90° rotation about the vertical axis.) Residues for which there are no data are in white, unperturbed residues are in light blue, residues for which there are two cross peaks are in blue, residues for which only one highly perturbed cross peak could be found are in magenta and the most highly perturbed residues are in red. The position of the Trp170 side chain in the binding site is shown in yellow (see the text for details). The surface plot was made using GRASP (Nicholls et al., 1991). Download figure Download PowerPoint TIF and CAF MIR compete for the MOD1C binding site To investigate whether the TIF and CAF MIRs interact with the same binding site on the shadow chromo domain, we st