Structural Analysis of the Mitotic Regulator hPin1 in Solution

INSIGHTS INTO DOMAIN ARCHITECTURE AND SUBSTRATE BINDING*

Elena Bayer {ddagger} §, Sandra Goettsch {ddagger} §, Jonathan W. Mueller {ddagger} §, Bernhard Griewel {ddagger} §, Elena Guiberman {ddagger} §, Lorenz M. Mayr ¶ and Peter Bayer {ddagger} § ||

From the {ddagger}Max-Planck-Institute for Molecular Physiology, Otto-Hahn-Strasse 11, 44227 Dortmund, Germany, the §Interdisciplinary Center for Magnetic Resonance, 44227 Dortmund, Germany, and the Novartis Pharma AG, Lead Discovery Center/GSO, WSJ-350.102, CH-4002 Basel, Switzerland

Received for publication, January 22, 2003 , and in revised form, April 22, 2003.


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 REFERENCES
 
The peptidyl-prolyl cis/trans isomerase hPin1 is a phosphorylation-dependent regulatory enzyme whose substrates are proteins involved in regulation of cell cycle, transcription, Alzheimer's disease, and cancer pathogenesis. We have determined the solution structure of the two domain protein hPin1-(1–163) and its separately expressed PPIase domain (50–163) (hPin1PPIase) with an root mean square deviation of <0.5 Å over backbone atoms using NMR. Domain organization of hPin1 differs from that observed in structures solved by x-ray crystallography. Whereas PPIase and WW domain are tightly packed onto each other and share a common binding interface in crystals, our NMR-based data revealed only weak interaction of both domains at their interface in solution. Interaction between the two domains of full-length hPin1 is absent when the protein is dissected into the catalytic and the WW domain. It indicates that the flexible linker, connecting both domains, promotes binding. By evaluation of NOESY spectra we can show that the {alpha}1/{beta}1 loop, which was proposed to undergo a large conformational rearrangement in the absence of sulfate and an Ala-Pro peptide, remained in the closed conformation under these conditions. Dissociation constants of 0.4 and 2.0 mM for sulfate and phosphate ions were measured at 12 °C by fluorescence spectroscopy. Binding of sulfate prevents hPin1 aggregation and changes surface charges across the active center and around the reactive and catalytically essential Cys113. In the absence of sulfate and/or reducing agent this residue seems to promote aggregation, as observed in hPin1 solutions in vitro.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 REFERENCES
 
Human Pin1 (hPin1)1 is a key protein in post-phosphorylation regulatory mechanisms. It was originally identified in 1996 in a two-hybrid screen as protein interacting with and suppressing the toxicity of Aspergillus mitotic kinase never-in-mitosis A (NIMA) (1). hPin1 selectively catalyzes peptidyl-prolyl cis/trans isomerization of phosphorylated Ser/Thr-Pro of proteins after they have been targeted by Ser/Thr-Pro-directed kinases (2, 3).

Peptidyl-prolyl isomerization of phosphorylated moieties provides a mechanism for switching a protein between different conformations, thereby influencing protein activity (4, 5), dephosphorylation (6), and subcellular localization (7). hPin1 substrates are proteins involved in regulation of cell cycle, transcription, Alzheimer's disease, and cancer pathogenesis. Phosphorylation-dependent peptidyl-prolyl isomerization is proven, e.g. for regulation of phosphatase Cdc25 function, where hPin1 binds to p-Thr48-Pro and p-Thr67-Pro motifs in Cdc25C. These motifs are crucial for Cdc25C to activate Cdc2 and to trigger the G2/M transition. Conformational changes on Cdc25C induced by hPin1 were detected as sensitivity to proteases or by MPM-2 (phosphospecific mitosis marker antibody) recognition (6, 8). Furthermore, hPin1 may integrate signals mediated by different kinases, as it was shown for the kinase substrate p53. Phosphorylation of p53 at all three sites (Thr33-Pro, Thr81-Pro, and Ser315-Pro) is required for its efficient binding to hPin1 followed by dissociation from p53-directed E3 ligase, thereby stabilizing the p53 protein and activating its tumor-suppressor function (4, 5).

Functionally, hPin1 has been shown to regulate several phases of the cell cycle, including G1/S and G2/M transitions as well as the DNA replication checkpoint (1, 8, 9). The protein is essential for growth of HeLa cells and arrests them in mitosis when depleted (1, 10). hPin1 overexpression activates the cyclin D1 promoter by regulation of c-Jun following JNK (c-Jun N-terminal kinase) phosphorylation, thereby promoting tumor growth (11). Consequently, hPin1 is of great interest for cancer therapy, and many pharmaceutical companies have chosen this protein as a molecular target for drug discovery.

Human Pin1 is a two domain protein. The N-terminal WW domain mediates protein-protein interaction (8) and targeting of hPin1 to the nucleus. Both events are regulated by phosphorylation of residue Ser16 (12). The C-terminal PPIase domain catalyzes peptidyl-prolyl isomerization of pSer/pThr-Pro moieties of the substrate. In assays with phosphorylated Ser/Thr-Pro peptides, the isolated WW domain (hPin1WW) is completely catalytically inactive, whereas the separated PPIase domain (hPin1PPIase) is 90% as active as full-length protein (6). Although the WW domain has no catalytic activity, it binds the phosphorylated peptides with higher affinity than the PPIase domain does. Binding affinities of full-length hPin1 and its isolated WW domain toward peptide substrates only differ in a factor of two, whereas hPin1PPIase shows moderate affinity or no affinity toward them (13). In binding studies with cellular substrates of hPin1, the WW domain was shown to be responsible for interaction, and hPin1PPIase does not bind any of the protein substrates (8). Interestingly, the optimal binding peptide for the WW domain WFYpSPFLE (pintide) is most similar to the substrate WFYpSPR-pNa, for which the highest PPIase activity of hPin1 (kcat/Km = 20,160 mM-1 s-1, unphosphorylated 170 mM-1 s-1) has been measured (2).

Two crystal structures of full-length hPin1 are published and deposited in the Research Collaboratory for Structural Bioinformatics data bank (accession numbers: 1pin [PDB] and 1f8a [PDB] ) (13, 14). In both structures WW and PPIase domains share a common interface. The WW domain and the catalytically active PPIase domain of hPin1 are connected by a glycine- and serine-rich stretch, which plays a yet unknown role in protein function. In crystal structures this linker seems to be flexible and forms no contacts to residues of the two folded domains.

The crystal structures suggest two possible conformations of hPin1. The first structure of hPin1-(1–163), co-crystallized together with an Ala-Pro peptide (14), exhibits the "closed" active site of PPIase domain, including a phosphate-binding {alpha}1/{beta}1 loop, which is complexed to a sulfate ion and partly shelters the substrate binding motif, a proline ring pocket. The WW domain does not participate in substrate interaction, but together with the PPIase domain binds a PEG molecule close to the interface. In the second structure of hPin-(1–163) a 70° rotation of the phosphate-binding loop leads to an "open" conformation of the PPIase domain active site (13). No sulfate ion is complexed to the structure. In that case, the phosphorylated peptide is bound exclusively to the WW domain, resulting in a twist of its {beta}-sheet. Both structures are recognized as two different stages during the substrate-binding event. Sulfate is hypothesized to act as phosphoryl group mimetica, which upon binding together with Ala-Pro dipeptide induces "closing" of the loop region.

To shed some light onto hPin1 domain interaction in solution and to investigate the proposed induced fit mechanism upon sulfate binding, we solved the structure of full-length hPin1-(1–163) and of its PPIase-(50–163) domain in complex with sulfate using nuclear magnetic resonance. Weak interaction of domains was detected in the presence of the flexible linker in full-length hPin1, but no complex formation occurred after dissecting the protein into separated domains. NOESY techniques provide information about the integrity of the structure, indicating that the phosphate-binding loop remains in the closed conformation even in the absence of sulfate ions.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 REFERENCES
 
Plasmid Construction—The coding DNA sequence for human Pin1 protein (RefSeq NM_006221 [GenBank] ) was used to search via BLASTN (15) in the dbEST-human data base at NCBI (www.ncbi.nlm.nih.gov) for corresponding EST clones and several of the clones with 5'-sequence identity (National Institutes of Health Image clones 110334, 171663, and 471113) were combined and used to amplify the Pin1 gene by PCR (forward primer: AATAATAATCATATGGCGGACGAGGAGAAGCTG (NdeI site underlined); reverse primer: AATAATAATGAATTCATTACTCAGTGCGGAGGATGATGT (EcoRI site underlined)). This PCR fragment was cloned into the NdeI/EcoRI sites of pET-28a vector to generate a fusion protein with N-terminal poly-His tag and thrombin site for purification of the protein. The DNA encoding the hPin1PPIase-(50–163) protein was constructed and amplified by PCR using the pET-28a_hPin1 gene and the following primers: forward, AATTTAAACATATGCATCACCATCACCATCACGGGGAGCCTGCCAGGGTCCGCTGC (NdeI site underlined) and reverse, AAATTTGAATTCATTACTCAGTGCGGAGGATGATGTGG (EcoRI site underlined). The PCR product was NdeI/EcoRI-ligated into the vector pET-41a (Novagen). The gene for hPinPPIase is preceded by a sequence encoding for the His6 tag. To receive the gene coding for the construct hPin1WW-(6–39) PCR using pET-28a_hPin1 was performed with the following primers: forward, AAATTTAAACCATGGGCAAGCTGCCGCCCGGCTGGGAG (NcoI site underlined) and reverse, AAATTTGAATTCATTAGCCGCTGGGCCGCTCCCACTG (EcoRI site underlined). The PCR product was NcoI/EcoRI-ligated into the vector pET-42_mod_TEV, which was kindly provided by Dr. Axel Scheidig. The gene for hPinWW is preceded by a sequence encoding for glutathione S-transferase protein (GST), followed by a His6 tag and a tobacco etch virus (TEV) protease cleavage site.

Expression and Purification of Human Pin1, Pin1PPIase, and Pin1WW All protein constructs were expressed in Escherichia coli strain BL21(DE3) Codonplus RIL (Novagen). 50-ml overnight cultures of hPin1 or hPin1PPIase were harvested and resuspended into 50 ml of M9 minimal medium enriched with either [15N]ammonium chloride (1 g/liter) for uniformly labeling or with [15N]ammonium chloride (1 g/liter) and [13C]glucose (2.5 g/liter) for uniformly 15N-13C labeling (Cambridge Isotope Laboratory). Expression cultures were grown in isotope enriched minimal medium by inoculation of 1 liter with 50 ml of overnight-adapted cells. For unlabeled protein the 50-ml overnight culture was directly added to 1 liter of 2x YT (yeast extract tryptone) medium. After induction of protein expression at A600 = 0.4 with 1 mM isopropyl-1-thio-{beta}-D-galactopyranoside, cells were shaken for a further 4 h at 37 °C, harvested, and centrifuged at 4 °C for 20 min at 5,000 x g in a Beckman J2-HC centrifuge (Beckman Instruments, Palo Alto, CA). Cell rupture was performed using a Model 110S Microfluidizer (Microfluids, Newton, MA) in 50 mM sodium phosphate or Tris/HCl buffer, pH 8.0, each containing 0.3 mM NaCl, 20 mM imidazole, and 2 mM {beta}-mercapto-ethanol supplemented with CompleteTM, EDTA-free, protease inhibitor mixture (Roche Applied Science, Penzberg, Germany). The cell lysate was ultracentrifuged at 4 °C in a Sorvall Discovery 100 centrifuge at 72,000 x g for 30 min. The supernatant was applied to a nickel-nitrilotriacetic acid Superflow (Qiagen) column (2.5 x 20 cm), equilibrated with either 50 mM sodium phosphate or Tris/HCl buffer, pH 8.0, 0.3 M NaCl, 20 mM imidazole. hPin1 and hPin1PPIase proteins were eluted with an imidazole gradient of 20 to 200 mM in the corresponding buffer, concentrated, and washed in a Microsep microconcentrator (Filtron Technology Corp., Northborough, MA) with an exclusion size of 10,000 Da. The unlabeled GST fusion WW domain construct was expressed in 2x YT medium and purified by affinity chromatography with nickel-nitrilotriacetic acid Superflow. The GST-His6 tags were removed by His-TEV protease (Invitrogen) at 4 °C directly on column. The untagged WW domain was in the flowthrough, whereas the GST-His6 tags and His6-TEV protease remained bound to nickel-nitrilotriacetic acid Superflow. Finally, the WW domain-containing fractions were pooled and concentrated in a Centricon tube with an exclusion size of 1,000 Da.

UV Spectroscopy—UV spectroscopy of the aggregation of hPin1 and all OD measurements were carried out using a CARY 100 Bio UV-visible spectrometer (Varian) equipped with a Peltier temperature control unit.

Fluorescence Titration Experiments—Fluorescence experiments were performed using an SLM Smart 8000 spectrofluorometer (Colora, Lorch, Germany) equipped with a PH-PC9635 photomultiplier. Sample concentration for hPin1PPIase was 1 µM. The buffer contained 50 mM Tris/HCl, 1 mM DTT, pH 6.6, for measurements at 12 °C. For tryptophan fluorescence, samples were excited at 295 nm and the emission intensity was measured at 348 nm. The slit widths for the experiments were 1 and 16 nm. For buffer and volume effects, corrections were done by titration with blank buffer. Data were evaluated using the program Sigmaplot 7.0 by fitting a quadratic equation given by Müller et al. (16).

In equilibrium one-to-one binding of a protein, P, and ion, I, to form a protein-ion complex, PI, can be expressed as in Equation 1.

(Eq. 1)
The equilibrium dissociation constant, Kd, for this reaction is as follows,

(Eq. 2)
where P(0) = [P] + [PI] and I(0) = [I] + [PI], Kd and [PI] can be calculated from Equation 3.

(Eq. 3)
Substitution for [PI] leads to the following equation.

(Eq. 4)
The measured fluorescence intensity, F, is described as,

(Eq. 5)
where F(0) is the fluorescence intensity of the free protein and {Delta}F is the increase in fluorescence when adding sulfate or phosphate ions. Values for the dissociation constant (Kd), the amplitude of fluorescence change, and protein concentration were allowed to vary during the fitting procedure.

NMR Spectra Acquisition—Triple resonance and homonuclear experiments were performed on either a Varian Inova 600 or a Bruker DRX-500 spectrometer, both equipped with shielded Z gradients. The HBHA(CO)NH spectrum was recorded on a Avance 600 spectrometer at Bruker, Rheinstetten, Germany (Dr. Bermel), and 13C HSQC-TOCSY and 13C NOESY spectra were acquired on a Bruker Avance 800 at the PARABIO center in Florence, Italy (Dr. Pieratelli). The temperature for all experiments was set to 300 K. Homonuclear two-dimensional experiments (TOCSY and NOESY) were recorded with unlabeled protein; 15N-heteronuclear HSQC-type spectra were acquired on uniformly 15N-labeled or 13C-15N-labeled samples. The latter protein was used to perform triple resonance spectra. All protein samples (0.6–0.8 mM) were dissolved in phosphate buffer solution (50 mM sodium phosphate, 50 mM NaSO4, 1.0 mM DTT, 5 mM EDTA at pH 6.6) or in Tris/HCl buffer (50 mM, 1 mM DTT at pH 6.6) in H2O:D2O (9:1, v/v). 2,2-Dimethyl-2-silapentane 5-sulfonate, sodium salt was used as an internal standard for calibration of proton resonances. For 15N and 13C calibrations we followed the IUPAC procedure (17). The water resonance was suppressed by applying a WATERGATE sequence (18) or by presaturation. Prior to Fourier transformation, all spectra were multiplied by a sine bell square window function shifted by {pi}/2. Generally, 2048 x 256 data points were used for acquisition of HSQC titration spectra and 1024 x 120 x 80 data points for three-dimensional edited spectra. NMR spectra were processed using the standard Bruker software XWINNMR. Analysis and visual representation of two-dimensional spectra were performed using the NDEE program package (SpinUp Inc., Dortmund, Germany) and three-dimensional spectra were analyzed with the program Aurelia (Bruker) on O2 and Octane workstations (Silicon Graphics Inc.).

NMR Titration Experiments—For domain interaction, titration experiments HSQC spectra were acquired on 13C-15N double-labeled PPIase domain and unlabeled WW domain. Equivalent amounts of hPin1WW were added to a 100 µM sample of hPin1PPIase to yield 1:0.15, 1:0.30, 1:0.45, 1:0.60, 1:0.75, 1:0.9, and 1:1.05 stoichiometries of protein/protein concentration. For sulfate binding, HSQC spectra were acquired on uniformly 15N-labeled hPin1PPIase (100 µM). 1 M sodium sulfate solution was titrated to the protein to reach a final 1000:1 ratio of sulfate to protein.

NMR Titration Analysis—Titration analysis was done by fitting chemical shift data to a quadratic equation as described in detail in a previous study (19). Chemical shift differences were calculated using Equation 6 (20).

(Eq. 6)

NMR Assignment and Constraint Collection—Assignments of hPin1 (21, 22) (BioMagResBank (BMRB) accession numbers 5305 [BMRB] and 4882) were used. If necessary, additional assignments were generated using the spectra HNCA, HNCACB, CBCA(CO)NH, HC(CO)NH, C(CO)NH, 13C HSQC TOCSY, 15N HSQC-TOCSY, 13C HSQC-NOESY, and 15N HSQC-NOESY recorded on hPin1 or hPin1PPIase samples. Distance constraints were extracted from a 1H homonuclear NOESY spectrum (mixing time, 150 ms) and 15N and 13C HSQC-NOESY spectra. {varphi} angles were calculated on the basis of HNHA spectra and {psi} angles as described previously (23, 24). Hydrogen bond donors were extracted from 15N HSQC spectra recorded on protein samples in D2O. After structure calculation without hydrogen bonds, additional donors and acceptors were introduced using geometrical selection criteria (distance A-H, <3Å in 50% of the structures). 30% of backbone hydrogen bonds and all side-chain hydrogen bonds were added when corresponding bonds were present in the x-ray structures, and those bonds had favored geometry and were not excluded by neighboring distance (NOEs) and dihedral angle constraints. Based on the x-ray structure (14) and on chemical shift changes upon titration, 10 distance constraints were used to fix the sulfate ion to the side chains of residues Lys63, Arg68, and Arg69.

Structure Calculation—Structure calculation for hPin1 and the hPin1PPIase·sulfate complex was performed using the program CNS 1.0 (A. Brünger). High temperature torsion angle dynamics was performed at 50,000 K for 15 ps (1,000 steps) followed by a 15-ps cooling phase. In each case an ensemble of 100 structures was calculated from a random coil template. Ten models were selected on the basis of energetic criteria (low total energy, distance violations (NOE) < 0.2 Å and dihedral angle violations < 5° using the accept.inp routine) to form a representative ensemble of the calculated structures. An average structure for each ensemble was generated, and in the case of hPin1 subsequent energy minimization was applied. All calculations were done on a Silicon Graphics Inc. Octane workstation. The program Sybyl (Tripos Associates, St. Louis, MO) was used for visualization, and the programs Rasmol 2.6 (25), Molscript 2.2.1 (26), and Raster3D (27) were used for figure production. The structures of hPin1 and hPin1PPIase·sulfate have been deposited in the Protein Data Bank (accession numbers 1NMV and 1NMW).

Error Determination—Errors of fitted Kd values and geometrical and energetical data were calculated by commercial programs (Sigmaplot 7.0) and are defined as root mean square deviation,

(Eq. 7)
where is the mean, xi a certain measured value, and n the total number of measured points.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 REFERENCES
 
The WW and PPIase Domains of hPin1 Interact Only Weakly in Solution—We determined the structure of full-length hPin1-(1–163) and its PPIase domain (hPin1PPIase-(50–163)) in phosphate buffer solution (50 mM Na2SO4, 1 mM DTT, 5 mM EDTA), pH 6.6, and 27 °C using multidimensional nuclear magnetic resonance techniques. 1277 (1044) distance constraints, 78 (64) hydrogen bonds as well as 82 (74) {varphi} and 64 (52) {psi} angle constraints were extracted from homo- and 13C-15N heteronuclear NOESY spectra, 15N HSQC spectra, and HNHA spectra of hPin1 and hPin1PPIase. 100 structures were calculated by torsion angle dynamics (CNS 1.0), and, on the basis of a set of exclusion parameters, an ensemble of 10 structures (Table I) was chosen to represent the final fold. Fig. 1 shows an overlay of the structures of full-length hPin1 fitted on either the WW domain (Fig. 1A) or the PPIase domain (Fig. 1B). Both domains could be determined with high precision resulting in low r.m.s.d. values over backbone atoms of 0.43 Å (WW) and 0.48 Å (PPIase), respectively. The sequence of hPin1 and the number of experimentally determined contacts, each residue is involved in, are presented in Fig. 1 (C and D). Although the WW and PPIase domain share a common interface in crystal structures, no distance constraints (NOEs) could be observed in any of the NOESY spectra between residues Ile28, Thr29, Asn30, and Ala31 and residues Asp136, Ala137, Ala140, Leu141, Ser147, and Gly148, all of which are proposed to be involved in domain interaction (13, 14).


View this table:
[in this window]
[in a new window]
 
TABLE I
Coordinate precision and structural statistics of the calculated structures of hPin1 and hPin1PPIase Vdw energies were calculated with the CHARMM force field on the basis of a repel function using hard Vdw spheres of atoms.

 


View larger version (44K):
[in this window]
[in a new window]
 
FIG. 1.
Solution structure of hPin1. Ensemble of ten energetically favorable structures calculated with the program CNS1.0 were fitted and superimposed on either the WW domain (A) or the PPIase domain (B). C, sequence of full-length hPin1. D, the number of NOE contacts, each residue involved is plotted against the corresponding residue number. Secondary structure elements are drawn at the top of the plot. Black arrows represent {beta}-strands, and gray vertical bars are {alpha}-helices. A double arrow indicates the linker region.

 

A prerequisite for the presence of NOEs between atoms of these two domains is a relatively tight binding of the interacting partners. Lifetime of the complex should be long enough for NOEs to build-up. Consequently, distance constraints between binding partners are not observed in NOESY spectra of weakly bound and short lived complexes. A characteristic feature for weak interaction is the observation of high koff values. In such a case information on the interaction of binding partners can be obtained from chemical shift perturbation experiments. Binding can be monitored by changes in chemical shifts of resonances in NMR spectra. Spectra are collected in a way that the concentration of one of the proteins is stationary while concentration of the second protein is stepwise increased.

To investigate whether both domains of hPin1 interact in solution, we cloned and expressed the PPIase domain comprising residues Gly50 to Glu163 (hPin1PPIase) and compared the chemical shifts of HN resonances in the 15N HSQC spectrum of hPinPPIase to the corresponding resonances in the 15N HSQC spectrum of full-length hPin1. As shown in Fig. 2, large chemical shift differences were observed of residues thought to form the proposed interface (black) and of neighboring amino acids.



View larger version (16K):
[in this window]
[in a new window]
 
FIG. 2.
Chemical shift differences between amide protons of the PPIase domain of hPin1 and hPin1PPIase. The shift difference in ppm is plotted against the corresponding residue number. Residues involved in the binding interface (crystal structure) are represented by black bars; residues where technically no shift difference could be measured are represented by white negative bars.

 

The differences in chemical shifts between resonances of hPin1PPIase and the PPIase domain of full-length hPin1 indicate that interaction of domains in hPin1 occurs in solution along the interface observed in crystal structures. In contrast to the crystal structures (13, 14) equilibrium exists in solution within the intact hPin1 between a "bound" (complexed) and a "free" state, in which both domains are connected by a flexible linker but do not interact with each other. In solution the main populated conformation is the free state of hPin1.

The Interdomain Linker Plays a Major Role in Promoting Domain Interaction—A short flexible linker comprising Asn40 to Gly49 connects both domains of hPin1. In crystal structures no contacts from amino acids of the linker exist to any other residues from either the WW or PPIase domain. Resonances corresponding to amino acids of this region are very weak or absent in 15N HSQC spectra, indicating that the linker is also flexible in solution. Consequently, no long or medium range NOEs were found for residues Asn40 to Gly49.

To elucidate a possible role of the linker in domain interaction, we dissected the protein into its independent domains, excluding the linker region, to monitor their interaction by NMR. For this purpose we expressed the WW domain from residue Lys6 to Gly39 (hPin1WW) and studied binding to hPin1PPIase. Subsequent amounts of unlabeled hPin1WW were added to a 100 µM solution of 15N-uniformly labeled hPin1PPIase, and 15N HSQC spectra were recorded at each state of titration. In case of changes in chemical shifts, plotting differences versus the concentration of added WW domain allows to estimate the Kd value describing complex formation (Fig. 3, A–C). Surprisingly, no shifts were observed after a 1:1 ratio of both proteins was reached (Fig. 3E). The spectrum shown in Fig. 3C (1:1 ratio) is only the sum of the spectra shown in Figs. 3A (hPin1PPIase) and 3B (hPin1WW). As can be seen from Fig. 3E, the chemical shift differences observed in 15N HSQC spectra between resonances in the PPIase domain of full-length hPin1 (green) and hPin1PPIase (red) cannot be regained in the spectrum, in which the single domains are present in a 1:1 ratio (blue). These results demonstrate that no domain interaction occurs in the absence of the linker but that presence of the linker promotes complex formation.



View larger version (17K):
[in this window]
[in a new window]
 
FIG. 3.
Part of the amide region of one- and two-dimensional NMR spectra recorded at pH 6.6 and 300 K. One-dimensional spectra of a 100 µM solution of 15N-labeled hPin1PPIase (A), unlabeled hPin1WW (B), and a 1:1 mixture of 15N-labeled hPin1PPIase and unlabeled hPin1WW (C) are shown. Heteronuclear spectra were recorded without 15N decoupling. D, one-dimensional spectrum of part of the amide region of full-length hPin1 acquired with 15N decoupling. E, an overlay of HSQC spectra recorded with samples described in A, C, and D is shown. Part of the amide region is presented. Colored letters highlight one-dimensional spectra of samples A, C, and D. Resonances in HSQC spectra of the corresponding samples are plotted with identical colors.

 

Structural Comparison of hPin1 in Solution and in the Crystal State—Both crystal structures were determined in complex with small ligands, where either the active center of the PPIase domain is occupied by an Ala-Pro moiety and a PEG molecule is bound to the composite domain interface (14) or a phosphorylated peptide is bound to the WW domain (13). Minor differences in the active centers of the PPIase domains of crystal and solution structure may be direct consequences of these different binding modes. There is evidence that the two highly conserved histidine ring systems (His157 and His59) have different orientations in the three models. Whereas in the solution structure the rings are fixed by a number of distance constraints in a way that both H{epsilon}1 protons face each other (data not shown), the ring of either His157 (14) or His59 (13) are rotated by 180° around the preceding C{beta}–C{gamma} bond in the crystal structures.

Because domains of hPin1 interact only weakly in solution, we had to compare both folds (WW and PPIase domain) separately to their corresponding domains in the crystal structures and to equivalent structures deposited in the RCSB data bank. The topology of the WW domain of full-length hPin1 is similar to those of many other WW domains solved so far. The program SSM (www.ebi.ac.uk/msd-srv/ssm) was used for a structural similarity search against the RCSB data bank and yielded high Z score values (6.5–10.4) and low r.m.s.d. values (<2.1 Å) along the C{alpha} trace to at least 20 other RCSB entries. Highest identity was found for hPin1WW (22) complexed to a Cdc25 peptide (1i6g [PDB] , r.m.s.d. 1.22 Å) and to the WW domain (14) of the crystal structure of hPin1 (1pin [PDB] , r.m.s.d. 1.39 Å). A lower convergence of 1.68 Å was found to the WW domain of the crystal structure (1f8a [PDB] ), complexed to a peptide isolated from the CTD of RNA polymerase II (13). In this structure the bound ligand induced a {beta}-sheet twist. A data base search using the program DALI (www2.ebi.ac.uk/dali/) revealed C{alpha} trace r.m.s.d. values of 1.5 and 1.4 Å between the PPIase domain of hPin1 in solution and the crystal structures solved previously in Refs. 14 and 13, respectively. In this comparison residues of the {alpha}1/{beta}1 loop region were excluded from the fit procedure. An overlay of the PPIase domains of solution and crystal structures is shown in Fig. 4 (A and B). Differences in {varphi} angles of the superimposed structures are plotted to indicate differences in conformation.



View larger version (22K):
[in this window]
[in a new window]
 
FIG. 4.
Structural comparison of solution and crystal structures of the PPIase domain of hPin1. Top, the C{alpha} trace of the PPIase domain of the solution structure (red) is superimposed with the crystal structure solved by Verdecia et al. (13) (blue) (A) and Ranganathan et al. (14) (cyan) (B). Bottom, differences in {varphi} angles between the solution structure and crystal structures from A and B are plotted.

 

The Loop Region Responsible for Binding the Substrate Phosphoryl Group Is in a Closed Conformation—The major differences between both crystal structures are found in the phosphate-binding {alpha}1/{beta}1 loop region, which either covers the active site of the PPIase domain (14) or sticks out from the core of the protein (13). The structure exhibiting the closed conformation is complexed to a sulfate ion ligated by residues Arg68, Arg69, and Lys63 (basic cluster) of the loop region (Fig. 4A, blue), whereas in the "extended" case, no such ion is present (Fig. 4B, cyan). Because the sulfate ion can mimic the phosphoryl group of a substrate molecule (14, 29), the two crystal structures are regarded as time-resolved snapshots of an induced-fit mechanism that facilitates substrate binding (13). Ranganathan et al. (14) suggest an obligatory interaction of substrate and the basic cluster during catalysis, based on the observation that hPin1 enzymatic activity against a tested set of peptide substrates has decreased upon addition of sulfate or phosphate. They propose that binding of substrates and these multivalent ions must be mutually exclusive. This hypothesis can explain why hPin1 slowly aggregates during spectra acquisition in the absence of sulfate ions within a time scale of hours. The extended loop comprises hydrophobic residues like Ile78 that might destabilize the protein and enhance protein aggregation. When the loop is closed, the side-chain atoms of residue Ile78 are alternatively packed into a core comprising side chain atoms of e.g. residues Pro70 and Lys63.

To investigate the proposed induced fit mechanism and binding affinities of sulfate and phosphate, we studied binding of hPin1PPIase to these ions by fluorescence titration experiments. Changes in the intrinsic fluorescence intensity of Trp73 of hPin1PPIase upon addition of sulfate and phosphate (Tris/HCl buffer) were used to estimate ion-binding affinity of the protein (Fig. 5). hPin1PPIase binds sulfate and phosphate with Kd values of 0.4 and 2.0 mM, respectively (at 12 °C). After proving ion binding, we recorded homo- and 15N heteronuclear NOESY spectra of hPinPPIase and/or hPin1 in the presence of 50–100 mM sodium sulfate and compared them to spectra acquired in the absence of sulfate ions. Surprisingly, almost all distance constraints observed in the presence of sulfate were also observed, when sulfate is lacking. As an example, most prominent NOEs between protons of the methyl group and of the H{alpha} of residue Ala116 and the N{epsilon} proton of Trp73 are shown in Fig. 6, where parts of NOESY spectra, recorded under different solvent conditions, are plotted.



View larger version (17K):
[in this window]
[in a new window]
 
FIG. 5.
Fluorescence titration curves of hPin1PPIase with phosphate and sulfate ions. Filled squares represent the titration curve for phosphate, and filled circles represent the titration curve for sulfate. Both experiments were recorded in Tris/HCl buffer (1 mM DTT) at 12 °C, pH 6.6, and a protein concentration of 1 µM.

 


View larger version (32K):
[in this window]
[in a new window]
 
FIG. 6.
Part of the amide region of two-dimensional 1H NOESY spectra. A NOESY spectrum of hPin1PPIase, in the presence (A) and absence (B) of sulfate is shown. A black frame marks NOEs between the H{epsilon} proton of the indole ring of Trp73 and the H{alpha} and H{beta} protons of Ala116. C, a Molscript representation of part of hPin1PPIase is shown. Residues Ala116 and Trp73 are drawn as ball-and-stick models. The distance constraints obtained from the NOESY spectra are marked by two arrows.

 

Owing to our data the proposed induced fit mechanism does not occur in the hPin1 PPIase domain upon addition of sulfate or, what can be implied, upon addition of substrate molecules in solution. Although the loop region seems to be in the closed conformation in the absence of sulfate, flexibility of some residues changed on a small scale. Upon addition of sulfate, some resonances, which appeared as weak cross-peaks in 15N HSQC spectra recorded without sulfate, gained intensity and changed chemical shifts (e.g. Ser114).

Sulfate Ions Influence Surface Charges Around Residues of the Active Center—Next, we concentrated on the influence of sulfate ions on the structural integrity of the PPIase domain, because sulfate has a stabilizing effect on protein solubility. Therefore, chemical shift differences were measured between hPin1 in the presence and absence of sulfate. The positions of amide HN and N atoms and side-chain atoms, whose resonances undergo chemical shift changes, are shown in Fig. 7 (A and B). Chemical shift changes are only observed for resonances of atoms in close proximity to the sulfate ion. Reasons for chemical shift changes upon ligand binding can either be structural changes, influencing the topological neighborhood of nuclei, or electrostatic shielding or deshielding effects, when charged groups of the ligand change the electronic environment of nuclei in the protein. In Fig. 7C chemical shift differences are plotted against the distance of amide groups to the sulfur atom of the ion. The black curve is fitted under the assumption that the chemical shift is straight proportional to the electric field strength (projected on the vector connecting the sulfur atom and HN of the corresponding amide group) (28). In this case the data can be approximated by a 1/r2 function (black line). Considering experimental errors, resonance shifts observed upon sulfate binding seem to originate from a pure electrostatic effect and are less connected to a structural rearrangement.



View larger version (34K):
[in this window]
[in a new window]
 
FIG. 7.
The influence of sulfate ions on the structure of hPin1PPIase. A, a Molscript representation of hPin1PPIase is shown. Residues whose side-chain resonances undergo chemical shifts in NOESY spectra or 13C HSQC spectra upon addition of sulfate are represented as ball-and-stick models. Blue spheres mark amide resonances that undergo shifts in 15N HSQC spectra upon addition of sulfate. B, shift differences of amide resonances between 15N HSQC spectra recorded in the presence and absence of sulfate ions are plotted against either the corresponding residue number or the distance of the HN proton to the sulfur atom of the sulfate moiety (C).

 

As a control experiment, response of each amino acid of hPin1PPIase on the addition of sulfate was measured in an NMR titration experiment. Subsequent amounts of sulfate were added step-by-step to a 100 µM solution of the protein to reach a final ion concentration of 100 mM. All resonances of residues changing chemical shifts in the case of the PPIase domain of full-length hPin1, also showed chemical shift changes in the HSQC of hPin1PPIase. However, the absolute shift values were decreased to about 60–70% of the original values. Additionally, we observed shift changes of resonances of amino acids Glu83, Phe104, Glu105, and Thr143. From the titration experiment Kd values for sulfate binding at 27 °C could be obtained by plotting the concentration of sulfate ions against the chemical shift differences and fitting data points to a quadratic equation. In Fig. 8 fits of five amino acids are shown. The mean Kd value is ~7–8 mM.



View larger version (15K):
[in this window]
[in a new window]
 
FIG. 8.
Titration of hPin1PPIase with sulfate ions monitored by nuclear magnetic resonance. A 100 µM protein solution was stepwise titrated with a 1 M solution of Na2SO4 to yield a final concentration of 100 mM sulfate (27 °C, pH 6.6). Titration curves of residues His64 ({blacktriangledown}, Kd = 6.9 mM), Ser72 ({blacktriangleup}, Kd = 18.1 mM), Glu131 (•, Kd = 3.0 mM), Asp153 ({blacksquare}, Kd = 2.6 mM), and Ser154 ({diamond}, Kd = 8.1 mM) are plotted.

 

After it could be ruled out that sulfate does induce structural rearrangement on a large scale, we had to look for a stabilizing effect caused by a change in the electrostatics of the protein. Fig. 9 shows GRASP surface charge representations of the sulfate ion binding loop and the active center of hPin1PPIase calculated either with (Fig. 9A) or without (Fig. 9B) sulfate ion electrostatics. Interestingly, the surface charges around the side chain of residue Cys113, a prerequisite for isomerase activity, change from strong positive (free form) to neutral (ion complexed form) upon ion binding. Because the thiol group of this cysteine (in contrast to Cys57) is involved in catalysis and is surface-accessible (30), the pKa value and, therefore, its reactivity might change, too. To investigate, whether Cys113 is responsible for aggregation we performed an experiment on relatively low concentration compared with our NMR conditions, where we followed aggregation by absorption spectroscopy monitored at 550 nm in a UV spectrometer. The aggregation behavior of 40 µM hPin1PPIase was followed in either Tris/HCl buffer solution or a solution where 50 mM sulfate or 1 mM DTT was added. In Fig. 10 the resulting A550 is plotted against time and fitted to a 1-exp function to trace the time course. Although, relative errors in data points are high, the time course of the sample with DTT closely resembles that of the sample where sulfate was added. Without one of these two substances aggregations seems to be faster. Thus, the stabilizing effects, induced upon sulfate binding, can be achieved by addition of reducing agent in the absence of sulfate.



View larger version (44K):
[in this window]
[in a new window]
 
FIG. 9.
GRASP surface representation of hPin1PPIase. Molecular surfaces are colored according to electric potential. Blue colors represent positive charges, red colors are negative charges, and neutral areas are gray. Calculation of electric potential was done, including the potential of the complexed sulfate ion (A) or excluding it (B). Indicated are charged residues and amino acids of the active center.

 


View larger version (17K):
[in this window]
[in a new window]
 
FIG. 10.
Aggregation of hPin1PPIase under various conditions. Aggregation of a 40 µM solution of protein was monitored by UV spectroscopy at a wavelength of 550 nm. Data are fitted to a 1-exp function to trace the time course. Circles represent the time course (solid line) of protein aggregation in Tris/HCl, squares are the time course upon addition of 50 mM sulfate (dashed line), and triangles are the time course upon addition of 1 mM DTT (dotted line).

 

DISCUSSION
We have solved the structure of the two domain peptidyl-prolyl cis/trans isomerase hPin1 in solution and shown that it differs from the structures determined by x-ray crystallography. Although chemical shift mapping was successfully applied to screen amino acids in the PPIase domain involved in binding to the WW domain, interdomain NOE constraints were absent in NOESY spectra. According to this observation the lifetime of the complexed state seems to be very short compared with the lifetime of the unbound state in full-length hPin1. Thus, the dissociation constant Kd is dominated by a high koff value. In the crystal both domains are tightly packed onto each other, whereas in solution this state is in exchange with another one, where no interaction between both domains occurs. Dissection of the molecule and bringing together the separated domains does not lead to any observable complex formation in solution. Thus, the apparent Kd value for interaction of the free domains seems to be in the range of hundreds of micromolar or even millimolar. Domain interaction is promoted by a flexible linker, which does not interact with the rest of the molecule. The linker increases the local concentration of one domain around the other (31). Assuming a linker length of 20 Å, the PPIase domain of hPin1 "senses" a local concentration of the WW domain of ~50 mM (and vice versa) compared with a concentration of 100 µM as used in our experiments for the separated domains.

What is the reason for the differences observed in hPin1 domain interaction in the crystal state and in solution? In crystal structures protein is co-crystallized with either a substrate (13) or a xenobiotic PEG (14) molecule, both of which are complexed to the composite interface region between the WW and PPIase domain. Titration of hPin1 in solution with increasing concentrations of PEG400 (Fig. 11) up to 3% (v/v) induces shifts in resonances of residues, which are at the PEG binding site of the WW in the crystal structure (14). It is very likely that binding of a substrate to the WW domain promotes domain interaction (32), a hypothesis that is summarized in Fig. 12. This hypothesis is in agreement with observations that hPin1WW can bind ligands in the absence of the PPIase domain, but binding is enhanced by a factor of 1.5–2 in the presence of the catalytic domain (13). Similar results were obtained when elucidating the structure of dystrophin (33). The WW domain of dystrophin cannot bind alone the dystroglycan ligand without the adjacent helical EF-hand-like domain. The two domains actually form a composite recognition surface that is critical for the specificity to the substrate molecule.



View larger version (24K):
[in this window]
[in a new window]
 
FIG. 11.
A, Molscript representation of PEG binding site in the crystal structure (14). Residues, whose HN resonances undergo chemical shifts upon PEG addition, and PEG are represented by ball-and-stick models. B, part of a series of hPin1 HSQC spectra recorded with increasing PEG concentration (0, 0.1, 0.5, 1.0, 1.7, and 3.0%). In particular note the shift of Gln33 as marked by an arrow.

 


View larger version (34K):
[in this window]
[in a new window]
 
FIG. 12.
Model for the domain interaction in hPin1. In solution the equilibrium tends to the protein state with non-interacting domains. Upon addition of substrate the equilibrium is shifted toward the complexed form of hPin1.

 

The solution structure of the PPIase domain of hPin1 closely resembles the fold observed by Ranganathan et al. in the crystal state (14). Both structures are very similar and have only minor differences in the {alpha}1/{beta}1 loop region. An extended conformation of this loop element was observed by Verdecia et al. (13) leading to the hypothesis of an induced-fit promoted rearrangement of the loop upon sulfate ion or substrate binding. By solving the structure of hPin1 and the PPIase domain in solution in the presence and absence of sulfate ions, we could demonstrate that no such structural rearrangement occurs, but the loop is in its closed conformation under both conditions. The open conformation observed in one of the crystal structures (13) might have its origin in crystallization conditions or crystal contact formation, but according to our studies, does not represent a snapshot of a substrate-receiving PPIase domain in solution. Nevertheless, some resonances of residues in the {alpha}1/{beta}1-loop and amino acids in topological proximity to it gain intensity upon addition of sulfate. A change in the intensity of resonances in HSQC spectra was also observed for AtPin1 (29), which exhibits a similar phosphoryl-binding loop. The loop region seems to increase its rigidity when a phosphorylated substrate gets bound. This is not surprising taking into account that the sulfate ion or the phosphoryl moiety is trapped by flexible side chains of residues Arg68, Arg69, and Lys63 of hPin1, which thereby undergo a loss of rotational freedom. Additionally, we performed a line width analysis of the HSQC spectra of hPin1 and the PPIase domain with and without sulfate (data not shown). Line width narrowing is observed in the N terminus (amino acids 1–6) and linker region (amino acids 39–49) of hPin1 caused by isotropic motion of the HN vectors. In contrast, only minor differences in the line width of residues of the loop region in the PPIase domain in the presence and absence of sulfate are found. The resonances of residues Arg69, Trp73, and Ser114 gain intensity upon increasing ion concentration. These data are in agreement with results of T2 and HetNOE studies (32). Low HN resonance intensities in the absence of sulfate reflect the local flexibility of the corresponding amino acids. The flexibility of the resonances in the absence of sulfate causes fast motion of the HN vectors around the averaged protein backbone position with about 1- or 2-Å mean deviation, as can be seen in Fig. 1. If no sulfate or phosphorylated substrate is bound, residues Arg69, Trp73, and Ser114 are flexible, because their amide groups are not involved in hydrogen bonds, whereas the motion of the sequential neighboring residues is still restricted. Sulfate binding induces formation of an ion- and water-based hydrogen network (14), including side-chain atoms of Lys63 and backbone atoms of Arg69, Trp73, and Ser114, which subsequently become more rigid. If we expected an opening of the loop, as observed in the crystal structure (13), more than 10 residues would be highly flexible and should undergo structural rearrangements of 5–28 Å. This would cause an isotropic movement of the corresponding HN vectors, and thus very low or negative HetNOEs and line width narrowing of all loop amide resonances should be observed.

The WW domain fold of the solution structure of hPin1 was found to have the highest identity to that of the NMR structure of hPin1WW (1.2 Å, PDB code 1i6g [PDB] ) in complex with a Cdc25 peptide (22) and to the WW domain fold (14) within the crystal structure (1.39 Å, PDB code 1pin [PDB] ) published by Ranganathan et al. These findings might indicate that no substantial conformational changes occur in the small and compact structure upon ligand binding. Similar observations have been made by Wintjens et al. (22) on hPin1WW, where only minor structural changes in the WW domain during substrate binding are reported. This is in contradiction to an observed {beta}-sheet twist in the crystal structure of Verdecia et al. (13) where the WW domain is complexed to a CTD peptide. Because in the crystal structure (14) no substrate but a PEG molecule is complexed to the composite interface of the WW and catalytic domain, it is possible that the presence of both the peptide substrate and PPIase domain is a prerequisite for {beta}-twist induction.

Samples of hPin1 and hPin1PPIase diluted in phosphate or Tris/HCl buffer solutions in concentrations necessary for NMR structure determination showed severe aggregation within several hours at room temperature. Although NOESY spectra could be obtained under such conditions, new isotope-labeled samples had to be prepared for each triple resonance spectrum recorded. Thus, sulfate ions, which were reported to have a stabilizing effect on the PPIase structure of AtPin (29) and hPin1, were used to improve sample quality. Aggregation was not completely abolished under the influence of a 50–100 mM solution of sulfate ions, but it could be slowed down to enable acquisition over a period of several days.

The idea to use sulfate ions is based on two facts. First, sulfate ions mimic the binding properties of phosphoryl moieties of substrate peptides containing phosphorylated serine or threonine residues. Second, the crystal structure of hPin1 solved by Ranganathan et al. (14) is complexed to a sulfate ion, by side chains of arginine and lysine residues of the {alpha}1/{beta}1 loop region. This loop is extended in the crystal structure of Verdecia et al. (13) solved in the absence of sulfate and presents hydrophobic residues to the bulk water. It was hypothesized that upon binding of sulfate large structural rearrangements of the loop region are induced pushing the hydrophobic amino acids toward the core structure of the PPIase domain, thereby, increasing protein stability.

By comparing NOEs from the loop region of hPin1PPIase in NOESY spectra recorded in the absence and presence of sulfate, we could show that no significant changes in spectra occur. Only different chemical shifts of resonances were observed, but some resonances of 15N HSQC spectra gained intensity upon addition of sulfate (e.g. Ser114). The conformational rearrangements observed in solution are minor and mainly influence the side chain of the ion chelating residues and the rigidity of amino acids in close proximity. The origin of chemical shift changes on sulfate addition is based on an electrostatic effect. Complexation of sulfate causes changes in surface charges around the active center. These alterations are probably the reason for changes in the intrinsic fluorescence signal of Trp73 observed at 295 nm upon addition of sulfate ions. The increase in signal intensity can either be brought about by a gain in rigidity or a change in polarity of the environment, the indole ring is sensing. As can be seen from Fig. 9 polarity around Ser72 and Trp73 decreases upon sulfate binding, which might explain increase in fluorescence signal intensity.

The most dramatic changes in surface charges can be found at the complexation site (Arg68, Arg69, and Lys63) and at the site of the active center, where amino acids His157, His59, and Cys113 are located. It has been shown by mutation analysis that Cys113 is important for cis/trans isomerase activity (14). Mutation of the cysteine to alanine or serine in hPin1 resulted in a 123- or 20-fold decrease in kcat/Km, respectively. Residue Cys113 is partly accessible by the attack of thiol group-modifying agents. Juglone, a Pin1 inhibitor, specifically attacks the thiol group and leads to a slow loss of structural integrity (30). The second cysteine (Cys57) is buried in the interior and not accessible to, e.g., alkylating agents. Our studies show, that by adding the reducing agent DTT in millimolar quantities to a Tris/HCl buffer solution of hPin1 (after each spectrum acquisition was finished), we could slow down aggregation even in the absence of sulfate ions. One possible explanation for aggregation is that disulfide formation at Cys113 occurs and leads to local unfolding of the protein. This in turn makes the second cysteine accessible for an attack and initiates oligomerization. An SDS-PAGE analysis of precipitated hPin1 without sulfate and DTT (data not shown) reveals the presence of dimers, trimers, and multimers, but only monomers were observed after reducing the sample. DTT prevents aggregation by reduction of disulfide bonds in hPin1.

How can we explain the stabilizing effect of sulfate? The reactivity of the protein thiol group depends on the accessibility of the thiolate group to the solvent, the fraction of thiol present as thiolate, and the intrinsic reactivity of the thiolate (basicity). In our case Cys113 became less reactive upon addition of sulfate (Fig. 10). Cysteine residues in catalytically active sites often have low pKa values (35). One can speculate that the apparent pKa of Cys113 increases (becomes more basic) upon addition of sulfate, making the residue less nucleophilic. Because the pKa defines the extent of ionization and reactivity at the given pH, an increase in the pKa should change the protonation state and induce shifts in the {beta}-carbon or {beta}-proton resonances. We could not follow the resonance shift of the Cys113 beta protons; however, we found an upfield shift in the corresponding {beta}-carbon resonance upon addition of sulfate (pH 6.8). In the absence of sulfate the thiolate may be stabilized by adjacent charged groups. Binding of sulfate changes the surface charges around Cys113 from positive to neutral (Fig. 9), thereby destabilizing the thiolate ion and increasing the pKa of the residue. Most likely, disulfide formation is slowed down by this mechanism.

An interesting conclusion can be drawn from Fig. 7C concerning the electrostatic effect of sulfate. The electrical field introduced by the ion penetrates the protein core and influences amide proton and nitrogen chemical shifts within a radius of about 10–12 Å. A similar effect was observed in the phosphorylated form of the protein hirudin, when the phosphate moiety was titrated from a monoanionic to a dianionic state (34). Here, changes in hydrogen bonds and chemical shifts occurred within a radius of 10 Å. Based on this two observations we can suggest that the cut-off value for electrostatic contribution to energy functions used for the calculation of protein structures, protein dynamics, and molecular modeling procedures has to be at least 10 Å to include the full electric field effect.

Because sulfate is regarded as an analogue for the phosphoryl moiety of a substrate peptide of hPin1 in vitro, we have to elucidate its role in vivo. Do sulfate or phosphate ions in cells play a possible regulatory role in hPin1 function? Competition between phosphate ions and non-phosphorylated tetra-peptides was found in in vitro assays, when the phosphate:substrate ratio was higher than 2000 (14). The dissociation constants obtained by fluorescence spectroscopy at 12 °C are 0.4 mM for sulfate and 2 mM for phosphate, respectively. The Kd values increase upon raising temperature, because it has been measured for sulfate in a NMR titration experiment (7–8 mM at 27 °C). From these experiments we can estimate that the dissociation constants for both ions in cells at 37 °C are around 20–50 mM. The cellular concentrations of free sulfate and free phosphate ions are found to be 1 (total concentration was 10 mM, including bound ions) and 2–5 mM (total concentration, 50–60 mM, including bound ions), respectively.2 Assuming that the concentrations of hPin1 (36) and substrate molecules in cells reach micromolar values and the binding affinities of hPin1 to phosphorylated protein substrates are at least 10 times higher than for non-phosphorylated tetra-peptides (Kd > 500 µM) measured in a previous study (14), competition of multivalent ions for the phosphoryl binding site in hPin1 is negligible. Thus, it is unlikely that sulfate and phosphate ions play a regulatory role in hPin1 function in vivo.

What is the biological and pharmaceutical implication of our studies on hPin1? hPin1 is of great interest for cancer therapy and inhibition of its activity might prevent mitosis and, thus, malignant cell division. Bearing the crystal structure in mind and looking from a drug engineer's view onto this molecular target, one might have two options for efficient development of a new anti-cancer drug. Inhibitors of hPin1 activation can either be addressed against the catalytic center of the PPIase domain or against the phospho-peptide binding site of the WW domain. Our model, describing a more dynamic domain interaction, offers a third strategy for rational drug design. Drugs preventing the formation or untying of a common binding interface might influence hPin1 function, too. Although we do not know yet how essential these dynamics are for hPin1 function in vivo, inhibiting domain interaction might prevent the protein from binding and targeting receptor molecules. Searching for "interface drugs" seems to be very promising, because it has already been shown by co-crystallization with PEG (14) that xenobiotic molecules of low molecular weight might induce interface formation.


    FOOTNOTES
 
The atomic coordinates and structure factors (code 1NMV and 1NMW) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).

* This work was supported by the Deutsche Forschungsgemeinschaft (Grants BA1624/3-2 and BA1624/4-1 (to P. B.)), by the Fonds der Chemischen Industrie Deutschland e.V (to P. B. and J. W. M.), by the Bundesministerium für Bildung und Forschung (BMBF) (to J. W. M.), and by the Max-Planck-Society. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

|| To whom correspondence should be addressed. Tel.: 49-231-133-2222; Fax: 49-231-133-2699; E-mail: peter.bayer{at}mpi-dortmund.mpg.de.

1 The abbreviations used are: hPin1, human Pin1; CTD, C-terminal domain; DTT, dithiothreitol; HSQC, hetero-single-quantum coherence; NOESY, nuclear overhauser enhancement spectroscopy; PPIase, peptidyl-prolyl cis/trans isomerase; TOCSY, total correlated spectroscopy;

PEG, polyethylene glycol; GST, glutathione S-transferase; TEV, tobacco etch virus; r.m.s.d., root mean square deviation. Back

2 R. Kinne, personal communication. Back


    ACKNOWLEDGMENTS
 
We gratefully thank Drs. Roberta Pieratelli at the PARABIO in Florence, Italy, Annalisa Pastore at the National Institute for Medical Research (NIMR) in London, UK, and Wolfgang Bermel at Bruker Rheinstetten, Germany, for help with some of the measurements. We thank Beate Schölermann for excellent technical support.



    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 REFERENCES
 

  1. Lu, K. P., Hanes, S. D., and Hunter, T. (1996) Nature 380, 544-547[CrossRef][Medline] [Order article via Infotrieve]
  2. Yaffe, M. B., Schutkowski, M., Shen, M., Stukenberg, P. T., Rahfeld, J.-U., Xu, J., Kuang, J., Kirschner, M. W., Fischer, G., Cantley, L. C., and Lu, K. P. (1997) Science 278, 1957-1960[Abstract/Free Full Text]
  3. Zhou, X. Z., Lu, P. J., Wulf, G., and Lu, K. P. (1999) Cell Mol. Life Sci. 56, 788-806[CrossRef][Medline] [Order article via Infotrieve]
  4. Zheng, H., You, H., Zhou, X. Z., Murray, S. A., Uchida, T., Wulf, G., Gu, L., Tang, X., Lu, K. P., and Xiao, Z. X. (2002) Nature 419, 849-853[CrossRef][Medline] [Order article via Infotrieve]
  5. Zacchi, P., Gostissa, M., Uchida, T., Salvagno, C., Avolio, F., Volinia, S., Ronai, Z., Blandino, G., Schneider, C., and Del Sal, G. (2002) Nature 419, 853-857[CrossRef][Medline] [Order article via Infotrieve]
  6. Zhou, X. Z., Kops, O., Werner, A., Lu, P. J., Shen, M., Stoller, G., Kullertz, G., Stark, M., Fischer, G., and Lu, K. P. (2000) Mol. Cell 6, 873-883[Medline] [Order article via Infotrieve]
  7. Ryo, A., Nakamura, M., Wulf, G., Liou, Y. C., and Lu, K. P. (2001) Nat. Cell Biol. 3, 793-801[CrossRef][Medline] [Order article via Infotrieve]
  8. Lu, P. J., Zhou, X. Z., Shen, M., and Lu, K. P. (1999) Science 283, 1325-1328[Abstract/Free Full Text]
  9. Winkler, K. E., Swenson, K. I., Kornbluth, S., and Means, A. R. (2000) Science 287, 1644-1647[Abstract/Free Full Text]
  10. Rippmann, J. F., Hobbie, S., Daiber, C., Guilliard, B., Bauer, M., Birk, J., Nar, H., Garin-Chesa, P., Rettig, W. J., and Schnapp, A. (2000) Cell Growth Differ. 11, 409-416[Abstract/Free Full Text]
  11. Wulf, G. M., Ryo, A., Wulf, G. G., Lee, S. W., Niu, T., Petkova, V., and Lu, K. P. (2001) EMBO J. 20, 3459-3472[Abstract/Free Full Text]
  12. Lu, P. J., Zhou, X. Z., Liou, Y. C., Noel, J. P., and Lu, K. P. (2002) J. Biol. Chem. 277, 2381-2384[Abstract/Free Full Text]
  13. Verdecia, M. A., Bowman, M. E., Lu, K. P., Hunter, T., and Noel, J. P. (2000) Nat. Struct. Biol. 7, 639-643[CrossRef][Medline] [Order article via Infotrieve]
  14. Ranganathan, R., Lu, K. P., Hunter, T., and Noel, J. P. (1997) Cell 89, 875-886[Medline] [Order article via Infotrieve]
  15. Altschul, S. F., Madden, T. L., Schåffer, A. A., Zhang, J., Zhang, Z., Miller, W., and Lipman, D. J. (1997) Nucleic Acids Res. 25, 3389-3402[Abstract/Free Full Text]
  16. Müller, B., Restle, T., Reinstein, J., and Goody, R. S. (1991) Biochemistry 30, 3709-3715[Medline] [Order article via Infotrieve]
  17. Markley, J. L., Bax, A., Arata, Y., Hilbers, C. W., Kaptein, R., Sykes, B. D., Wright, P. E., and Wüthrich, K. (1998) Pure Appl. Chem. 70, 117-142
  18. Piotto, M., Saudek, V., and Sklenar, J. (1992) J. Biomol. NMR 2, 661-665[Medline] [Order article via Infotrieve]
  19. Surmacz, T. A., Bayer, E., Rahfeld, J. U., Fischer, G., and Bayer, P. (2002) J. Mol. Biol. 321, 235-247[CrossRef][Medline] [Order article via Infotrieve]
  20. Ayed, A., Mulder, F. A. A., Yi, G. S., Lu, Y., Kay, L. E., and Arrowsmith, C. H. (2001) Nat. Struct. Biol. 8, 756-760[CrossRef][Medline] [Order article via Infotrieve]
  21. Jacobs, D. M., Saxena, K., Grimme, S., Vogtherr, M., Pescatore, B., Langer, T., Elshorst, B., and Fiebig, K. M. (2002) J. Mol. Biol. 23, 163-164[Medline] [Order article via Infotrieve]
  22. Wintjens, R., Wieruszeski, J. M., Drobecq, H., Rousselot-Pailley, P., Buee, L., Lippens, G., and Landrieu, I. (2001) J. Biol. Chem. 276, 25150-25156[Abstract/Free Full Text]
  23. Gagne, S. M., Tsuda, S., Li, M. X., Chandra, M., Smillie, L. B., and Sykes, B. D. (1994) Protein Sci. 3, 1961-1974[Abstract/Free Full Text]
  24. Bertini, I., Donaire, A., Jimenez, B., Luchinat, C., Parigi, G., Piccioli, M., and Poggi, L. (2001) J. Biomol. NMR 21, 85-98[CrossRef][Medline] [Order article via Infotrieve]
  25. Sayle, R., and Milner-White, E. J. (1995) Trends Biochem. Sci. 20, 374[CrossRef][Medline] [Order article via Infotrieve]
  26. Kraulis, P. J. (1991) J. Appl. Crystallogr. 24, 946-950[CrossRef]
  27. Meritt, E. A., and Bacon, D. J. (1997) Methods Enzymol. 277, 505-524
  28. Williamson, M. P., and Asakura, T. (1993) J. Magn. Res. B101, 63-71[CrossRef]
  29. Landrieu, I., Wieruszeski, J. M., Wintjens, R., Inze, D., and Lippens, G. (2002) J. Mol. Biol. 320, 321-332[CrossRef][Medline] [Order article via Infotrieve]
  30. Henning, L., Christner, C., Kipping, M., Schelbert, B., Rücknagel, K. P., Grabley, S., Küllertz, G., and Fischer, G. (1998) Biochemistry 37, 5953-5960[CrossRef][Medline] [Order article via Infotrieve]
  31. Reinstein, J., Vetter, I. R., Schlichting, I., Rösch, P., Wittinghofer, A., and Goody, R. (1990) Biochemistry 29, 7440-7450[Medline] [Order article via Infotrieve]
  32. Jacobs, D., Saxena, M., Vogtherr, M., Bernado, P., Pons, M., and Fiebig, K. (April 9, 2003) J. Biol Chem. 278, 26174-26182
  33. Huang, X., Poy, F., Zhang, R., Joachimiak, A., Sudol, M., and Eck, M. J. (2000) Nat. Struct. Biol. 7, 634-638[CrossRef][Medline] [Order article via Infotrieve]
  34. Kipping, M., Zarnt, T., Kiessig, S., Reimer, U., Fischer, G., and Bayer, P. (2001) Biochemistry 40, 7957-7963[CrossRef][Medline] [Order article via Infotrieve]
  35. Claiborne, A., Yeh, J. I., Mallett, T. C., Luba, J., Crane, E. J., III, Charrier, V., and Parsonage, D. (1999) Biochemistry 38, 15407-15416[CrossRef][Medline] [Order article via Infotrieve]
  36. Shen, M., Stukenberg, P. T., Kirschner, M. W., and Lu, K. P. (1998) Genes Dev. 12, 706-720[Abstract/Free Full Text]