From the
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.
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ABSTRACT |
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INTRODUCTION |
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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-(1163),
co-crystallized together with an Ala-Pro peptide
(14), exhibits the
"closed" active site of PPIase domain, including a
phosphate-binding 1/
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-(1163) 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
-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-(1163) and of its PPIase-(50163) 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.
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MATERIALS AND METHODS |
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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--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
-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 SpectroscopyUV 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 ExperimentsFluorescence 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) |
![]() | (Eq. 2) |
![]() | (Eq. 3) |
![]() | (Eq. 4) |
![]() | (Eq. 5) |
NMR Spectra AcquisitionTriple 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 (0.60.8
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 /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 ExperimentsFor 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 AnalysisTitration 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 CollectionAssignments 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. angles were calculated on the basis of HNHA spectra and
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 CalculationStructure 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 DeterminationErrors 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) |
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RESULTS |
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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.
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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 InteractionA 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, AC). 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.
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Structural Comparison of hPin1 in Solution and in the Crystal
StateBoth 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 H1 protons face each other (data not shown), the ring
of either His157
(14) or His59
(13) are rotated by 180°
around the preceding C
C
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 (6.510.4) and low r.m.s.d. values
(<2.1 Å) along the C 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
-sheet twist. A data base search using the
program DALI
(www2.ebi.ac.uk/dali/)
revealed C
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
1/
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
angles of the superimposed
structures are plotted to indicate differences in conformation.
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The Loop Region Responsible for Binding the Substrate Phosphoryl Group
Is in a Closed ConformationThe major differences between both
crystal structures are found in the phosphate-binding
1/
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 50100 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 of residue Ala116 and the N
proton of
Trp73 are shown in Fig.
6, where parts of NOESY spectra, recorded under different solvent
conditions, are plotted.
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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 CenterNext, 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.
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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 6070% 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 78 mM.
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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.
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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 1.52 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.
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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 1/
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
1/
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
16) and linker region (amino acids 3949) 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
528 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 -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
-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 50100 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 1/
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 -carbon or
-proton resonances. We could not
follow the resonance shift of the Cys113 beta protons; however, we
found an upfield shift in the corresponding
-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 1012 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 (78 mM at 27 °C). From these experiments we can estimate that the dissociation constants for both ions in cells at 37 °C are around 2050 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 25 mM (total concentration, 5060 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.
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FOOTNOTES |
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* 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.
|| 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.
2 R. Kinne, personal communication.
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ACKNOWLEDGMENTS |
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REFERENCES |
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