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With the current access to the whole genomes of various organisms and the completion of the first draft of the human genome, there is a strong need for a structure-function classification of protein families as an initial step in moving from DNA databases to a comprehensive understanding of human biology. As a result of the explosion in nucleic acid sequence information and the concurrent development of methods for high-throughput functional characterization of gene products, the genomic revolution also promises to provide a new paradigm for drug discovery, enabling the identification of molecular drug targets in a significant number of human diseases. This molecular view of diseases has contributed to the importance of combining primary sequence data with three-dimensional structure and has increased the awareness of computational homology modeling and its potential to elucidate protein function. In particular, when important proteins or novel therapeutic targets are identifiedlike the family of protein tyrosine phosphatases (PTPs) (reviewed in reference 53)a structure-function classification of such protein families becomes an invaluable framework for further advances in biomedical science. Here, we present a comparative analysis of the structural relationships among vertebrate PTP domains and provide a comprehensive resource for sequence analysis of phosphotyrosine-specific PTPs.

PTPs are a key group of signal transduction enzymes which, together with protein tyrosine kinases, control the levels of cellular protein tyrosine phosphorylation. Protein tyrosine kinases phosphorylate cellular substrates on tyrosine residues, and much progress has been made over the last 20 years in elucidating their significance in signal transduction (for reviews, see references 26, 30, 31, 33, 71, and 72). However, it is only recently that the complexities of the PTPs have been appreciated. Thus, today it is recognized that the capacity of PTPs to dephosphorylate phosphotyrosine residues selectively on their substrates plays a pivotal role in initiating, sustaining and terminating cellular signaling (for reviews, see references 1, 4, 19, 32, 35, 46, 55, and 83). It has been shown that both the catalytic domain and noncatalytic segments of the PTPs contribute to the definition of substrate specificity in vivo. Whereas noncatalytic domains may target the PTPs to specific intracellular compartments in which the effective local concentration of substrate is high (3, 19, 51), the PTP catalytic domains themselves confer site-selective protein dephosphorylation by recognizing both the phosphotyrosine residue to be dephosphorylated and its flanking amino acids in the substrate. The combination of structural studies, kinetic analysis of PTP domains (37, 74, 76, 90, 91, 96), and studies involving substrate-trapping mutants (20, 23, 89) as well as PTP chimeras (60, 82) has convincingly demonstrated that isolated PTP domains may exhibit exquisite substrate selectivity.

The structurally conserved PTP domain defines membership of the PTP family, and three groups of enzymes are capable of dephosphorylating tyrosine-phosphorylated residues (57): (i) classical PTPs, (ii) dual-specificity PTPs, and (iii) low-molecular-weight PTPs. The dual-specificity PTPs and low-molecular-weight PTPs will not be considered further but have been reviewed (43, 70). The classical PTPs, which are the focus of the present study, encompass both transmembrane receptor-like and nontransmembrane enzymes, and the wide spectrum of protein domains present within this family highlight their diverse cellular functions. Most transmembrane receptor-like PTPs (RPTPs) contain two cytoplasmic PTP domains, a membrane proximal domain (D1) and a membrane distal domain (D2), and in addition have a single transmembrane segment and an extracellular domain.

As the study of PTPs has developed, the availability of an impressive number of X-ray crystal structures (for an updated list, see reference 53) and phylogenetically divergent cDNAs has permitted a detailed structural analysis of the evolutionary relationships among vertebrate PTP domains. Together with numerous enzymological studies revealing insights into the mechanism of PTP catalysis (12, 14, 25, 34, 47, 68, 77, 80, 97-99, 101), this development has permitted us to combine an extensive set of amino acid sequences with representative three-dimensional protein structures to derive new and refined information regarding PTP structure, substrate recognition, and evolutionary conservation. Because perturbed levels of tyrosine phosphorylation are associated with diseases such as cancer, autoimmunity, and allergy, we hope that this comprehensive analysis of the PTP family may assist in providing the structural basis for novel therapeutic strategies involving the development of selective PTP inhibitors.

In the present study, we have compiled a total of 319 vertebrate PTP sequences, including splice variants and partially overlapping sequences. Subsequent analysis narrowed these 319 GenBank entries down to 113 distinct PTP catalytic domain sequences (including non-transmembrane PTPs and domains D1 of RPTPs) and 38 domain D2 sequences from human and other vertebrate species. From this collection of 151 PTP domain sequences, we identified 37 distinct human PTP genes, which we aligned to assist in the identification of conserved regions. This sequence comparison allowed the classification of the vertebrate PTP family into 17 principal subtypes. The motifs identified from our amino acid sequence alignment are reviewed in terms of their location in the tertiary structure and, where relevant, their catalytic function. As a low-resolution and automated homology modeling approach, we applied the methodology of C-regiovariation score analysis (10, 11) to identify foci within the PTP domain tertiary structure, where amino acid conservation extended in three dimensions. The conserved foci identified by this approach are discussed including a previously unrecognized conserved cluster of residues located on the face of the molecule opposite the active site.

Collection of unique vertebrate PTP domains. Following the identification of the first PTP in 1988 (84), intense efforts in the application of PCR and low-stringency screening led to the rapid discovery of a wide variety of other PTP family members. As often happens in rapidly developing research fields, several identical PTPs were independently cloned by different research groups and hence given different names and accession numbers. Consequently, the first step in our structural study was to compile a database of unique PTP domains. A BLAST search (2) of the National Center for Biotechnology Information (NCBI) GenBank database was performed using nucleotide sequences encoding several divergent PTP catalytic domains (PTP1B, SHP1, MEG2, PEST, PTPH1, PTPD1, CD45, RPTPµ, LAR, RPTP, RPTP, RPTP, STEP, and the PTP-like protein IA2). This sequence similarity search generated over 3,500 database hits that, following the exclusion of expressed sequence tags and sequences encoding dual specificity and low-molecular-weight phosphatases, identified 319 vertebrate PTP entries. Alignment of the 5' untranslated regions and the amino acid sequences of these 319 entries revealed a large number of different splice variants, partially overlapping sequences, and duplicate database entries. In total, 113 distinct PTP catalytic domains and 38 domain D2 sequences from tandem domain RPTPs were uncovered. This collection of PTP domains contains 37 human PTP genes and ortholog sequences from vertebrate species (Table 1). Our compilation of PTP-related sequences illustrates the redundancies often observed among GenBank database entries. Moreover, since many of the deposited sequences lack structural or functional annotation, there is a strong requirement for grouping these entries in order to gain access to the combined body of biochemical, structural, and/or functional information known for any given PTP. To this end, we have grouped the entries in Table 1 based on PTP domain sequence similarity (by subtype) and have identified the most likely human orthologs. In addition, to facilitate access to MEDLINE literature for any given PTP of interest, an electronic version of Table 1 can be retrieved (http://science.novonordisk.com/ptp), in which the accession numbers are hyperlinked to the NCBI website and PubMed literature database (http://www.ncbi.nlm.nih.gov). We have also mapped the chromosomal locations of the 37 human PTPs described in this study, allowing a detailed description of their intron and exon structure. Genomic clones, EMBL accession numbers, and the position of these PTPs in the human genome are summarized in Table 2. In addition, we acknowledge that the draft of the human genome contains additional sequences that conform to the PTP consensus motifs, but the expression of these hypothetical proteins has not yet been verified. Only transcripts that have been confirmed from in vitro or in vivo studies are considered in the present structure-function analysis of PTP domains.

Primary sequence alignment of PTP domains. To provide a platform for classification of members of the PTP family and for the identification of conserved residues, a multiple-sequence alignment was constructed using the entire set of vertebrate PTPs identified above (Table 1). In Fig. 1, we have reduced the alignment to a sequence comparison between the 37 human PTP domains, but the extended version used in our analysis can be retrieved from the World Wide Web (http: //science.novonordisk.com/ptp) and includes all 113 vertebrate PTP catalytic domains. To enable the assessment of both the level of conservation and the degree of sequence variation, the alignment is color coded according to amino acid identity (see legend to Fig. 1). The N- and C-terminal boundaries for this alignment correspond to residues 1 to 279 in PTP1B and encompass all invariant residues and structurally conserved elements. In the past, the PTP domain has been described to consist of ~250 amino acids, but the extensive set of PTPs included in this multiple-sequence alignment, combined with structural knowledge and secondary-structure prediction algorithms, has permitted us to identify conservation at the N terminus of the PTP domain comprising the 1' and 2' helices of PTP1B (37) (Fig. 1). We now define the PTP domain as comprising ~280 residues. In Fig. 1, an alignment of domain D2 of RPTPs is included, but these domains were not used in the definition of the PTP consensus sequence.

View larger version (318K): [in this window] [in a new window]   FIG. 1.   Sequence comparison of human PTP domains. Shown is an amino acid sequence alignment of 37 human PTP domains (from nontransmembrane PTP and RPTP domains D1) (above) and comparison with domain D2 sequences of RPTPs (below). Amino acids are numbered according to the residue position in human PTP1B. The locations of -helices and -strands (based on the X-ray crystal structure of PTP1B [7]) are shown at the top of the alignment. Twenty-two invariant residues (underscored) and 42 highly conserved residues (>80% identity) are indicated at the bottom of the alignment. The PTP consensus motifs (M1 to M10) are detailed in Table 2. Amino acids are color coded according to their degree of conservation, as indicated below the alignment. Nonconserved residues involved in the definition of substrate selectivity-determining regions are boxed with black lines (see text and Fig. 9). The four-residue conserved linker in tandem RPTP enzymes is boxed in yellow (above) and corresponds to encircled area 1 in Fig. 8. Sequences were aligned using the Clustalw algorithm and the Genetics Computer Group PileUp software (version 8.1) by applying the BLOSUM 62 scoring matrix together with default gap creation and extension penalty. Alignment of the N termini of the PTP domains was guided by crystallographic structural data and secondary structure predictions (nnpredict at http://www.cmpharm.ucsf.edu). The complete alignment of all vertebrate PTP domains can be retrieved (http://science.novonordisk.com/ptp) in several standard GCG formats, including MSF, TFA, and ALN.

PTP family can be classified into 17 subtypes. Since phylogenetic analysis of sequence alignments serves as a useful tool for the classification of homologous proteins, we derived a phylogenetic tree from our alignment of 113 PTP catalytic domains (Fig. 2). The clustering of sequences into divergent branches of this tree provided a basis for subdivision of PTP family members (see figure legend for details). In total, 17 principal PTP subtypes were identified as indicated in Fig. 2. In addition, all PTP domain D1 sequences from tandem domain RPTPs clustered into one major trunk of the phylogenetic tree, allowing the definition of a PTP supertype encompassing these five RPTP subtypes (R1/R6, R2A, R2B, R4, and R5). The high intrasubtype sequence identity of 60 to 80% among domain D1 of these RPTPs, compared to 45 to 60% among PTP domains of the RPTP-like subtype (R3), which contain only one PTP domain, supports earlier suggestions that during evolution, intragenic catalytic domain duplication (i.e., duplication of the PTP domain within an ancestral PTP gene) preceded gene duplication (88). Consistent with this concept, all domain D2 sequences also clustered into one separate branch of the phylogenetic tree (data available at http://science.novonordisk.com/ptp), suggesting structural, and perhaps functional, conservation among these PTP domains. The result of the present classification system, together with a diagram of the overall domain structure of a representative member of each PTP subtype, is presented in Fig. 3.

View larger version (46K): [in this window] [in a new window]   FIG. 2.    Classification of family of PTPs into 17 subtypes. Shown is an unrooted tree derived from the alignment of 113 vertebrate PTP domain sequences (residue positions 1 to 279 in human PTP1B). The tree was drawn by the neighbor-joining method (73). The horizontal distance indicates the degree of sequence divergence, and the scale at the top corner represents the number of substitution events (10 per 100 amino acids). Seventeen PTP domain subtypes were identified from the phylogram: nine nontransmembrane subtypes (NT1 to NT9), five tandem receptor-like subtypes (R1/R6, R2A, R2B, R4, and R5), and three single-domain RPTP subtypes (R3, R7, and R8 [subtype R8 is believed to be catalytically inactive]). As a statistical test of the significance of sequence similarity within PTP subtypes, bootstrap values were calculated (values are at the dendogram node). With the exception of the RPTP-like subtype (R3) and the tandem PTP domain supertype, all subdivisions were assigned based on maximal bootstrap values (1,000). (A tree including the PTP domain D2 sequences can be viewed [http://science.novonordisk.com/ptp], and the raw data files can also be retrieved in several standard GCG formats).

View larger version (38K): [in this window] [in a new window]   FIG. 3.   Schematic representation of PTP family members. Determination of sequence similarity among PTP catalytic domains (Fig. 2) was used to classify the PTP family of enzymes into nine nontransmembrane PTP subtypes (NT) and eight RPTP subtypes (R). Only the human PTPs are listed, and a representative member of each subtype is shown. Synonyms and classifications of all vertebrate PTPs are given in Table 1. PTPs having closely related catalytic domains also tend to be similar in overall structural topology.

Sequence similarity between PTP domains can be used for overall structural classification of PTP family. A major finding from the phylogenetic analysis of the alignment is the very close relationship between PTP domain sequence similarity and the presence of similar structural and functional domains in the full-length proteins (Fig. 3). Thus, the RPTPs, previously classified by their extracellular domains into nine distinct subtypes (8), are categorized into virtually identical groups based solely on catalytic domain sequence homology (hence the use of the existing nomenclature for the RPTPs) (8). However, one difference lies with chicken PTP. Based on its unique extracellular segment, this PTP was previously assigned to its own subtype (R6). but the present classification system suggests that it is the avian homologue of CD45. Therefore, we have included it within the CD45 subtype and defined it as R1/R6.

Ten conserved motifs define family of PTPs. Another application of the present PTP sequence alignment is the identification of conserved motifs that define this class of signal-transducing enzymes. In particular, the definition of consensus amino acid sequenceseither for the PTP family as a whole or for functional and therapeutically interesting PTP subtypeswill help to probe genomes of other organisms for the presence of PTP orthologs and thereby identify relevant model organisms for rapid genetic analysis of the involvement of PTPs in control of fundamental cellular functions.

Superimposition of PTP domains reveals conserved C-backbone trace that allows evaluation of the multiple sequence alignment in 3D space. To date, X-ray crystallographic structures are available for seven different PTP catalytic domains, including the nontransmembrane enzymes (PTP1B, Yop51, SHP1, and SHP2) (7, 27, 79, 92) and RPTP domains (PTPµ, PTP, and LAR) (29, 50, 56). When the crystal structures of vertebrate PTP domains were superimposed, we observed a conserved fold and a consistent C-backbone trace (Fig. 4). This striking conservation of tertiary structure allowed us to quantify the degree of conservation of each amino acid residue in three dimensional space (i.e., relative to the conservation of neighboring residues). In brief, such low-resolution homology modeling, the so-called C-regiovariation score analysis (10, 11), uses the information in a set of aligned sequences and calculates the average degree of conservation which has occurred within a given "sphere of influence" for each residue position along the folded polypeptide backbone of a representative tertiary structure. The method has previously identified interactive sites for cytochrome c, the pancreatic trypsin inhibitor family of proteinases, and carboxypeptidases A and B (10, 11). To avoid bias towards catalytic domains that are represented by a large number of ortholog sequences, we selected a nonredundant set of 37 aligned human PTP catalytic domains (Fig. 1). In agreement with C-regiovariation score analyses of other protein families (10, 11), we observed that a 6- to 8-Å sphere of influence provides an optimal signal-to-noise ratio and yields consistent results for different PTP templates (not shown). All figures in the present work were produced with a sphere of influence of 7 Å.

View larger version (86K): [in this window] [in a new window]   FIG. 4.   Crystal structures of vertebrate PTP domains show conserved fold and consistent C-backbone trace. PTP1B (magenta), RPTP (gray), RPTPµ (red), LAR (blue), SHP1 (green), and SHP2 (yellow) were aligned and superimposed using Quanta (Molecular Simulations Inc.). For clarity, residues 280 to 298 (C terminal) of PTP1B, 250 to 281 (N terminal) and 522 to 532 (C terminal) of SHP1, and 2 to 218 (N terminal) of SHP2 were omitted from the figure, as well as D2 of LAR. The calculated RMS deviations between all C atoms between PTP1B and other PTPs are as follows: PTP, 1.35 Å; RPTPµ, 2.72 Å; LAR D1, 2.78 Å; SHP1, 3.14 Å; and SHP2, 2.74 Å. For comparison, the RMS deviation between domains D1 and D2 of LAR is 1.3 Å. The X-ray structures are compared in their native open conformation.

Structural motifs make up the most highly conserved regions in PTP structure. The score values for the C-regiovariation analysis are shown in Fig. 5. Conserved residues in conserved surroundings are identified as peaks. Hydrophobic segments in the primary amino acid sequence alignment make up the most highly conserved microenvironments in the PTP structure. Thus, the structural motifs TXXDFWXMXW (M5), IVMXT, (M6) and KCXXYWP (M7) (Table 3) together form a densely packed hydrophobic core with energetically favored T stacking (52) of their aromatic ring systems (Phe95, Trp96, Tyr124, and Trp125). Extensive hydrophobic interactions were also observed between the stretch of amino acids DYINAS (M3) and [F/Y]IAXQGP (M4), which packed together in the PTP crystal structure by arrangement in parallel and anti-parallel -sheets (Fig. 6). Hydrophobic packing is important for protein structures to gain stability (16). In agreement with this concept, thermosensitive variants of LAR, TC-PTP, and PTP1B (54, 86) were found to contain mutations in the hydrophobic motifs above (M2 to M7), indicating a critical role of these residues in stabilizing the secondary structure of the PTP domain. Moreover, projection of the secondary structure of PTP1B onto the alignment (Fig. 1) and C-regiovariation score values (Fig. 5) revealed that the -sheets and -helixes in the structural motifs M2 to M6 are dominated by conservative amino acid substitutions, whereas nonconservation mutations frequently have been accepted in the regions flanking these secondary structures. To visualize the conservation of the core of the PTP structure, conserved residues are indicated on the C-backbone (Fig. 6).

View larger version (26K): [in this window] [in a new window]   FIG. 5.   The HCSAGXGR and IAXQGP motifs reside within the most highly conserved microenvironment of the PTP structure. Residues located within a highly conserved three dimensional space of the PTP structure are identified by peaks. The C-regiovariation score was calculated using the alignment information in Fig. 1 and the tertiary structure of PTP1B as template. Neighboring residues were defined using a three-dimensional 7-Å sphere of influence. Similar results were obtained for a 5- to 8-Å sphere and when using PTP, PTPµ, or SHP2 as templates for C-regiovariation score analysis (results not shown).

View larger version (62K): [in this window] [in a new window]   FIG. 6.   Core structures within the PTP domain are highly conserved and surface loops between secondary structure elements are least conserved. Shown is a ribbon diagram indicating the positions of conserved motifs (M1 to M10) within the tertiary structure. The degree of conservation was determined from C-regiovariation score analysis of 37 aligned human PTP catalytic domains (see Fig. 5). Areas of conservation (blue, most conserved; red, least conserved) are illustrated using the PTP1B catalytic domain as the representative tertiary structure. Shown is the front view of PTP1B looking into the active site. The catalytically essential Cys215 residue is shown in yellow.

GR[T/S]

PTP signature motif or phosphate-binding loop (motif 9). The active site sequence VHCSXGXGR[T/S]G (residues 213 to 223 in PTP1B) defines the PTP family and is often referred to as the PTP signature motif or the "PTP loop." Residues in this motif (M9) form the phosphate-binding loop, which is located at the base of the active site cleft. The cysteine in the PTP signature motif acts as a nucleophile and accepts phosphate transiently during catalysis (25), and the invariant Arg221 is involved in both substrate binding and in the stabilization of the phosphoenzyme intermediate (99). Our C-regiovariation score analysis identified two conserved polar residues (Glu115 and Arg257) in a microenvironment of the PTP structure which otherwise has accommodated many amino acid substitutions during evolution (Fig. 5). Importantly, these two residues form hydrogen bonds with the PTP loop, with the invariant Glu115 determining the position of Arg221 through a conserved salt bridge between the carboxy and guanidinium groups. Their invariance among human PTPs highlights their principal role in defining the architecture and function of the phosphate-binding loop. The close proximity of the catalytic Cys215 residue to main-chain amide groups of the PTP loop and hydrogen bonding with both the side chain of Arg221 and the hydroxyl group of Ser222 stabilizes the thiolate (deprotonated) form of the cysteine, favoring its function as a nucleophile (7, 94, 97). Moreover, theoretical investigations revealed that Arg257 may also contribute to stabilizing the nucleophilic nature of the active site cysteine (65). Mutation of the catalytic Cys215 to serine or alanine abrogates all enzyme activity while maintaining affinity for substrates in vitro, a feature that has been successfully utilized to obtain structures of PTPs in complex with phosphotyrosine peptide substrates (37, 74, 75, 91).

Phosphotyrosine recognition loop (motif 1). Whereas the phosphate group in the substrate phosphotyrosine residue is surrounded by residues corresponding to the PTP signature motif (37, 99), aromatic (Tyr46 and Phe182) and nonpolar (Val49, Ala217, and lle219) amino acids pack with the phenyl ring of the phosphotyrosine and delineate the boundaries of the active site binding pocket (37). The fact that these five residues are conserved by amino acid similarity suggests that the mechanism for recognition of the phosphotyrosine moiety of the substrate is similar among all tyrosine-specific PTPs. Collectively, the residues KNRY (Lys43 to Tyr46) are known as the phosphotyrosine recognition loop (37) since this element (M1) defines the depth of the active site crevice and hence creates selectivity for phosphotyrosine by excluding the hydrolysis of the shorter phosphoserine or phosphothreonine residues in target proteins (37, 91).

WPD loop (motif 8). The binding of phosphopeptides to the PTP loop promotes a major conformational change in the catalytic site surface loop (residues 179 to 187) that moves several angstroms to close the active site pocket and trap the bound phosphotyrosine (37, 80). The amino acid sequence of this surface loop is quite diverse, except for the WPDXGXP motif that contains a general acid-base catalyst (Asp181) (98). The presence of two proline residues (which do not support hydrogen bonding) and a glycine in the hinge bend region of this segment is critical for the dynamics of the WPD loop motion (63, 64). Comparison of the structures of ligand-free form of the Yersinia PTP (Yop51) and the enzyme complexed with oxyanions (77, 80) has revealed that an interaction between the invariant tryptophan in the WPD loop (Trp354, equivalent to Trp179 in PTP1B) and the above-mentioned arginine in the PTP loop (Arg409, equivalent to Arg221 in PTP1B) plays an important role in closure of the WPD loop. Enzyme kinetic analyses of this PTP has confirmed that mutation of the hinge Trp179 disables catalysis (28, 42). Closure of the WPD loop is critical for phosphoester hydrolysis, since it positions Asp181 close to the scissile oxygen of the tyrosyl substrate, allowing it to donate a proton to the phenolate leaving group (reviewed in references 22 and 95). Consistent with its role as a general acid catalyst, the substitution of Asp181 for an alanine allows phosphorylated substrates to form stable complexes with the enzyme. This "substrate-trapping" mutation has been used to enable isolation and identification of PTP substrates in vitro and in vivo (20, 23, 89, 94). The Asp181-to-alanine mutation creates a more efficient substrate trap than the mutation in which the active site Cys215 is changed to serine or alanine, possibly because the former mutation promotes the hydrophobic properties of the active site cleft and removes the potential for electrostatic repulsion between the Asp181 and the phosphate moiety of the substrate (23, 94).

Catalytic-water motif or Q loop (motif 10). In the QTXXQYXF motif (M10), two glutamine residues (Gln262 and Gln266) and two conserved arginine residues (Arg254 and Arg257) N terminal to this motif form crucial hydrogen bonds with interacting residues of the PTP loop and its amide backbone. In particular, Gln262 positions and activates an active site water molecule involved in the second hydrolysis step of the phosphocysteine enzyme complex (61, 100). Enzyme kinetic analysis of the Yersinia PTP combined with site-directed mutagenesis have revealed that Gln446 (equivalent to Gln262 in PTP1B) and, to a lesser extent, Gln450 (equivalent to Gln266 in PTP1B) is responsible for restricting phosphoryl transfer from the phosphoenzyme intermediate to water and not to other nucleophile acceptors (i.e., preventing the phosphoenzyme intermediate from acting as a kinase phosphorylating undesirable substrates [100]). Notably, within this highly variable area of the PTP structure (Fig. 5), only these arginine and glutamine residues are invariant, consistent with their involvement in catalysis (74, 100) and critical hydrogen bonding with residues of the PTP loop (80).

Conservation of surface-exposed amino acids in vicinity of active site: comparison between cytoplasmic PTPs and RPTP domains D1 and D2. In an attempt to reveal novel structure-function relationships, we performed the C-regiovariation score analysis on three subsets of PTP domains: (i) nontransmembrane PTPs, (ii) receptor-like D1, and (iii) receptor-like D2 (Fig. 7A, B, and C). The crystal structure of PTP1B was used as a template for the intracellular PTPs, whereas the molecular surface conservation among the aligned RPTP domain D1 and D2 sequences was illustrated using the X-ray crystal structure of PTP domain D1 (50). The interchangeable use of the PTP catalytic domains of PTP1B and PTP for the calculation of the C-regiovariation score values is justified by the excellent overlay of their tertiary structure, with a root mean square (RMS) deviation of 1.35 Å between the two domains (the RMS deviations between other PTP domain tertiary structures are given in the legend to Fig. 4). Although conserved residues converged around the active site for the intracellular PTPs and for domain D1 of RPTPs (Fig. 7A and B, respectively), the domain D2 sequences exhibited a much greater variation in the vicinity of the active site (Fig. 7C).

View larger version (60K): [in this window] [in a new window]   FIG. 7.   PTP domains from cytoplasmic PTPs and RPTP domains D1 and D2 show significant differences in their conservation of surface-exposed amino acids. Shown is surface conservation (blue, most conserved; red, least conserved) of PTP domains from nontransmembrane PTPs (A), RPTP domains D1 (B), and RPTP domains D2 (C). Shown is the front view looking into the active site. C-regiovariation score values for the cytoplasmic PTPs are illustrated using the X-ray crystal structure of PTP1B with the catalytically essential Cys215 (yel low) and epidermal growth factor receptor-derived peptide (green) bound within the active site (closed conformation). For ease of comparison, C-regiovariation score values among RPTP domains D1 and D2 sequences are illustrated using the X-ray crystal structure of RPTP domain D1 (50). The EGFR peptide (green) is modeled in the active site of RPTP for orientation using only a closed conformation of the X-ray crystal structures. Amino acids are labeled according to the residue position in human PTP1B with the equivalent residues in RPTP given in brackets (A and B). The conserved four-residue structural linker located at the N terminus of domain D2 (encircled area 1 in panel C), and which constrains the relative orientation of tandem PTP domains in LAR, is compared to the corresponding nonconserved area for the RPTP domain D1 sequences (encircled area 1 in panel B). The amino acid residues defining this conserved linker are boxed and colored yellow in the alignment in Fig. 1.

Identification of conserved interface between domains D1 and D2 of RPTPs. When comparing the RPTP domains D1 and D2 sequences by using the X-ray crystal structure of RPTP domain D1 as template (79), the encircled area 1 in Fig. 7 was found to be highly conserved among RPTP domain D2 sequences (Fig. 7C) but not among RPTP domain D1 sequences (Fig. 7B). Significantly, the residues in this area (Fig. 1, boxed in yellow) were recently identified as the structural linker that constrains the relative orientation of the two PTP domains in LAR (56). Although this is so far the only report describing the X-ray crystal structure of the tandem arrangement of PTP domains, our alignment reveals that this structural linker is highly conserved among all tandem domain-containing RPTPs, suggesting conservation in function. The consensus motif for this four-residue linker is G[D/E]TE (Fig. 1, highlighted in yellow).

View larger version (56K): [in this window] [in a new window]   FIG. 8.   Identification of novel conserved area on surface of PTP domain opposite active site. Shown are surface conservation (C-regiovariation score values) among nontransmembrane PTPs (A), RPTP domains D1 (B), and RPTP domains D2 (C). The tertiary structure is rotated 180° compared to structures in Fig. 7, showing the surface of the molecule opposite the active site. Encircled area II (B and C) corresponds to the interface for domains D1 and D2 as revealed in the X-ray crystal structure of LAR (56). Encircled area III is a novel putative interactive site, which appears to be conserved in all three subsets of PTP domain sequences. Amino acids are labeled according to the residue positions in PTP1B (A) and RPTP (B and C).

Identification of novel conserved pocket on surface of PTP domain opposite active site. In addition to the identification of conserved regions surrounding the active site and at the interface between domains D1 and D2 of RPTPs, the C-regiovariation score analysis identified another focus of conservation that was confined to the area of the molecule opposite the active site (Fig. 8A, B, and C, encircled area III). This surface-accessible area of conservation extends above a shallow hydrophobic pocket formed by residues IIe57, Ala69, IIe82, and Met98. These four residues have adopted a configuration that accommodates extensive van der Waals interactions and only one PTP (the Xenopus, mouse, and human orthologs of MEG2) has accepted nonconservative changes to residues within this pocket. In addition, residues from motif 2 (DXXRVXL) and motif 5 (TXXDFWXMXW) contribute to this conserved microenvironment, which explains why this cluster of conservation can be identified only when the PTP chain fold is considered.

Size and physiochemical nature of conserved pocket is consistent with recognition site for protein-protein interaction. Statistical examination of protein-protein associations suggests a central role for hydrophobic residues at interfacial regions (40, 67). In terms of amino acid composition at protein-protein domain interfaces, it has been noted that there is a preference for larger, nonpolar residues, particularly aromatic amino acids, as well as a few key basic or acidic residues (87). Indeed, from the alignment analysis, we observed that the conserved residues residing on the rim and within the shallow pocket (Fig. 8, encircled area III) are more hydrophobic and aromatic than the remainder of the surface of the PTP molecule, with several charged residues surrounding the hydrophobic pocket. Therefore, the amino acid composition in this region of the PTP molecule is consistent with the identification of a possible novel site of interaction. The size of the solvent-accessible area that appears conserved (i.e., the dark blue area) is ~250 Ų. Usually, the area of protein-protein interaction surfaces are larger (~700 Ų), typically constituting from 7 to 30% of the total surface area of a monomer (39, 87). However, extensive additional surface area can be included in this putative interaction site of the PTP domain if the adjacent less-conserved residues (i.e., the white area in Fig. 8) are considered in the evaluation.

Identification of nonconserved residues surrounding active siteimplications for substrate recognition and inhibitor design. In addition to the identification of three-dimensionally conserved regions, the C-regiovariation score analysis offers a unique opportunity to identify areas in a protein family that are less well conserved and therefore might indicate a specialized function. As an example, analysis of areas in the proximity of the active site of enzymes, here PTPs, may lead to the identification of putative substrate-binding pockets. Furthermore, such analysis in conjunction with primary sequence alignments may allow the identification of unique combinations of amino acid residues that can be addressed in a structure-based design of selective inhibitors.

View larger version (95K): [in this window] [in a new window]   FIG. 9.   Nonconserved amino acids in the proximity of the PTP active site are involved in the recognition of PTP substrates and nonpeptide PTP inhibitors. Shown is the visualization of four selectivity-determining regions on the molecular surface of PTP1B. Areas of conservation (blue, most conserved; red, least conserved) represent the C-regiovariation score values of 37 aligned human PTP catalytic domains (values from Fig. 5). The amino acids involved in defining these four selectivity-determining regions are indicated (boxed) in the alignment in Fig. 1.