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Keywords:

  • cross-linking;
  • PII signalling;
  • PPM phosphatase;
  • protein phosphorylation;
  • signal transduction

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Members of the Mg2+- or Mn2+-dependent protein phosphatases/PP2C-like serine/threonine phosphatases (PPM/PP2C) are abundant and widely distributed in prokaryotes and eukaryotes, where they regulate diverse signal transduction pathways. Despite low sequence conservation, the structure of their catalytic core is highly conserved except for a flexible loop termed the flap subdomain. Bacterial PPM/PP2C members without C- or N-terminal regulatory domains still recognize their substrates. Based on the crystal structure of tPphA (a PPM/PP2C member from the cyanobacterium Thermosynechococcus elongatus), variants of tPphA were generated by site-directed mutagenesis to identify substrate specificity determinants. Furthermore, a PPM/PP2C chimera containing the tPphA catalytic core and the flap subdomain of human PP2Cα was also generated. tPphA variants and the chimera were tested towards different artificial substrates and native phosphorylated PII. A binding assay combining chemical crosslinking and pull-down was designed to analyze the binding of the various phosphatase variants to phosphoprotein PII. Together, these data showed that the metal 1–metal 2 cluster in the catalytic center, but not the catalytically active metal 3, is required for the binding of phosphorylated substrate. Residues outside the catalytic center are pivotal for the recognition and turnover of phosphorylated protein substrate. In particular, a histidine residue (His39) of tPphA was identified to play a specific role in protein substrate dephosphorylation. Furthermore, mutations in the variable flap subdomain can affect enzyme activity as well as substrate specificity.

Structured digital abstract


Abbreviations
IPTG

isopropyl thio-β-d-galactoside

pNPP

p-nitrophenyl phosphate

PII-P

phosphorylated PII protein

PP2C

PP2C-like serine/threonine phosphatase

PPM

Mg2+- or Mn2+-dependent protein phosphatase

Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Reversible protein phosphorylation affects the structure and function of proteins that are responsible for the regulation of nearly all biological processes in living organisms. Protein kinases and protein phosphatases control the state of protein phosphorylation. Protein phosphatases can be divided into three main subclasses based on their substrate specificity and protein sequence: protein tyrosine phosphatase [1]; protein serine/threonine phosphatase [2]; and histidine phosphatase [3]. Protein serine/threonine phosphatase can be further classified into three major subfamilies: phosphoprotein phosphatases, the aspartate-based phosphatases, and metal-dependent protein phosphatases (PPM/PP2C) [2]. The human PPM member PP2Cα [4] has been the defining representative of the PPM family, which is therefore also referred to as the PP2C family. The catalytic domain of these family members comprises eight absolutely conserved residues within 11 conserved motifs [5–7]. PP2C homologs are widely present in eukaryotes and prokaryotes where they regulate diverse signaling pathways [8]. In the human genome, there are 16 distinct PP2C genes that give rise to at least 22 different isoforms [9]. They participate in many biological processes including regulation of mitogen-activated protein kinase cascades, cell cycle progression, ubiquitination and degradation of proteins, and mechanisms for cell death and survival [8]. In plants, there are even more PP2C genes with Arabidopsis thaliana harbouring 82 PP2C genes [10]. Except for six, all the others can be classified into 10 groups (groups A–J) based on sequence similarity [10]. Group A contains genes whose products have been shown to act as negative regulators in the abscisic acid (ABA) signaling pathway, which takes part in the response to different environmental stresses such as drought, cold and salinity [8,11]. In bacteria, many PP2Cs lack potential regulatory domains but nevertheless they still specifically dephosphorylate a variety of substrates [12–15].

The PP2C phosphatases require Mg2+ or Mn2+ ions as ligands, which are coordinated by a universally conserved core of aspartate residues. Structural analysis of bacterial and plant PP2C members revealed that these enzymes contain three metal ions (M1, M2 and M3) embedded in a catalytic core with almost invariant structure [11,16–20] except for the variable region termed the flap subdomain [17]. The flap, which protrudes near the M3 binding site, consists of a flexible loop and α-helical segments (Fig. 1A). In the catalytic center, M1 and M2 coordinate a catalytically active nucleophilic water molecule and, according to novel structures, they could interact with two oxygen atoms of the phosphate group [18]. Recently, we could show by mutational and structural analysis of a cyanobacterial PP2C family member (tPphA from Thermosynechococcus elongatus) that the loosely bound M3, which was occasionally observed in several bacterial and plant PP2C homologs, is required for catalysis, presumably by activating a water molecule as proton donor [21]. tPphA is the homolog of PphA from Synechocystis sp. PCC 6803, which is the phosphatase that dephosphorylates the signaling protein PII. The PII protein has been shown to be phosphorylated/dephosphorylated on a seryl residue (Ser49) located at the apex of a large solvent-exposed loop termed T-loop (Fig. 1B) in response to the binding of signaling metabolites [22,23]. The genome of Synechocystis contains eight putative PP2C type homologs [24,25]. However, only one out these eight genes turned out to be responsible for phospho-PII (PII-P) dephosphorylation [11,26], raising the question of the mechanistic basis for the specificity of PII towards PphA. The alignment of these eight PP2Cs showed the greatest sequence variability in the flap subdomain. The tPphA protein displays a highly similar flap subdomain sequence to PphA and it could specifically dephosphorylate PII-P in an effector-molecule-dependent manner [16].

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Figure 1.  (A) The crystal structure of tPphA (PDB 2J86). The amino acids that were analyzed by site-directed mutagenesis are indicated. Metal 1, Metal 2 and Metal 3 (M1, M2 and M3) constitute the catalytic center of tPphA. The flap subdomain is close to M3. (B) The crystal structure of PII from Synechococcus elongatus PCC 7942 (PDB 1QY7). The T-loop is indicated. (C) The crystal structure of SaSTP (PDB 2PK0). The His41 residue from monomer C could make a salt bridge with the Glu152 residue from monomer A. The homolog residue of SaSTP His41 in tPphA is His39. (D) Overlay of the crystal structures of tPphA (colored cyan, PDB 2J86) and human PP2Cα (colored magenta, PDB 1A6Q). The human PP2Cα flap (termed hflap in the figure) and tPphA flap (termed tflap in the figure) show different conformations.

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A clue to understanding the specific interaction between PP2C phosphatases and their phosphoprotein substrates came from the crystallographic analysis of the Streptococcus agalactiae PP2C phosphatase SaSTP [19]. An SaSTP crystal with four monomers (A, B, C and D) in the asymmetric unit revealed monomer C in a conformation which interacted with the flap subdomain of the adjacent monomer A (Fig. 1C). The interactions include two salt bridges (Arg12(C)–Glu151(A) and His41(C)–Glu152(A)) and three direct hydrogen bonds (Ser14(C)–Glu152(A), Arg13(C)–Ser155(A) and Ile162(C)–Pro157(A)). Whether this interaction mimicked an enzyme–product complex or was a serendipitous crystallographic artifact was an open question [19]. However, SaSTP His41 is highly conserved among bacterial PP2C phosphatases. The corresponding residue in tPphA is His39, which is located at the tip of a small loop (Fig. 1A). His39 seems to be flexible, since this residue is invisible in one space group (C2221) of the wild-type tPphA crystal [16] and in the crystal of the D193A variant [21]. Whether the residue is indeed involved in substrate recognition or product interaction needs to be resolved. A further clue for understanding substrate recognition and regulation was provided by the atomic structures of A. thaliana PP2Cs in complex with ABA-regulated PP2C inhibitor proteins [11,27]. The structures revealed a highly conserved tryptophan residue in the flap subdomain of ABI1 (ABA-insensitive1) and HAB1 (hypersensitive to ABA1), which directly docked into a cavity of the inhibitor proteins leading to inhibition of ABI1/HAB1 activity, a fact that highlights an important regulatory role of the flap subdomain [11,27–29]. In support of a role of the flap subdomain for substrate recognition is the observation that enzymatically active recombinant human PP2Cα (amino acids 1–297, without the C-regulatory domain) could not dephosphorylate PII-P although a synthetic T-loop phosphopeptide could be readily dephosphorylated [21]. The structure of the catalytic core of tPphA and human PP2Cα is highly similar, but their flap subdomains differ substantially in structure (Fig. 1D) and primary sequence [16].

Since the structures of tPphA and of PII are solved (Fig. 1A, B) [16,30] and the conformational changes involved in PII signaling are well studied [31], these proteins represent a well suited example for studying the fundamental properties of PP2C substrate interaction. In order to find key residues/domains in tPphA for substrate recognition, residues surrounding the catalytic center including the flap segment were analyzed by site-directed mutagenesis (Fig. 1A) and a new PP2C chimera protein (tPphA catalytic center + human PP2Cα flap) was generated (Fig. 2B). Substrate-specific activities of the variants were assayed and a phosphatase binding assay, based on chemical crosslinking under enzymatically arrested conditions, was developed to assay the direct interaction between the enzymes and the PII protein substrate.

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Figure 2.  (A) The motif nomenclature of tPphA according to Bork et al. 1996 [5]. tPphA contains all 11 conserved motifs and eight highly conserved residues (colored red). Motif 5b and motif 6 constitute the flap subdomain. The amino acids, which were analyzed by site-directed mutagenesis, are indicated by blue numbers. (B) The strategy of exchanging the flap subdomains of tPphA and human PP2Cα to generate two chimeric proteins. The tPphA flap subdomain (tflap) is labeled in blue, the human PP2Cα flap subdomain (hflap) is labeled in green. Only the catalytic domain (1–297) of human PP2Cα was used, whereas the C-terminal segment (298–382, grey α helices in the figure) was truncated.

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Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

In this study, several new tPphA variants (R13K, S15A, Q17E, Q17K, M36A, H39A, R160A, T138A and T138E) were created. The previously created variants D18A, D34A, D119A, D193A, D231A [21], H161A and R169A [16] were also included in this research. Figure 1A shows the location of the mutated residues in the 3D structure of tPphA. According to the classic PP2C motif nomenclature [5], the primary sequence of tPphA contains all 11 conserved motifs and eight highly conserved residues (shown in Fig. 2A). Motif 5b and motif 6 constitute the flap subdomain. The positions of mutated residues in tPphA primary sequence are indicated (Fig. 2A). Furthermore, the flap subdomains of tPphA and of human PP2Cα were mutually exchanged to generate two chimeric proteins (Fig. 2B). In chimera A, the human PP2Cα flap subdomain was fused to the tPphA catalytic core. In chimera B, the tPphA flap subdomain was fused to the human PP2Cα catalytic core. Whereas construction of chimera A resulted in soluble and active protein (see below), chimera B was insoluble and could not be re-solubilized under various conditions (details on the manipulation of chimera B are given in the section entitled ‘Cloning, overexpression and purification of human PP2Cα and the chimeras’). Further experiments were therefore only performed with chimera A.

Enzymatic assays of tPphA variants, human PP2Cα and chimera A towards artificial substrates and PII-P

To test the effect of amino acid substitutions on tPphA activity, the various phosphatase variants were assayed with the following substrates: the artificial substrate p-nitrophenyl phosphate (pNPP) (Table 1), three different phosphopeptides, of which one corresponds to the phosphorylated T-loop of PII (Table 2), and the physiological substrate PII-P (Fig. 3 and Fig. S1A–C). As shown previously, the alanine variants of the aspartate residues D18, D34, D119, D193 and D231, which coordinate the metals in the catalytic center, are catalytically inactive [21]. The newly generated variants displayed a large heterogeneity with respect to their catalytic properties, as described in more detail in the following. Figure 4A shows a comprehensive comparison of the variants, which were differentially affected in substrate specificity. To facilitate the comparison, the enzyme activities towards the five different substrates were normalized to the respective activity of wild-type tPphA, which was set to 100%. Among all newly created variants, variant Q17K was most strongly affected in catalytic activity, with only about 10% residual activity towards pNPP and the threonyl peptide and no detectable activity towards seryl peptides and PII-P. The tPphA crystal structure (PDB file 2J82 [16]) reveals that Gln17 indirectly coordinates M2 by a bridging water molecule. In many other PP2C homologs, the corresponding position is occupied by a glutamyl residue (see Supplementary file S1 in [20]). Interestingly, the Gln17 to Glu mutation (Q17E) resulted in increased catalytic efficiency towards an artificial phosphoseryl peptide (pS-peptide), whereas the activity towards the other substrates was very similar to wild-type tPphA. Of the other novel variants, variant S15A was not significantly affected in catalytic properties. Several variants (R13K, M36A, T138A, R160A, H161A and R169A) showed a general reduction in catalytic activity, dephosphorylating all tested substrates more slowly than wild-type tPphA. Apparently, these point mutations affected the catalytic activity of the corresponding tPphA variants to different degrees, showing effects on Kcat and Km as presented in Table 1. Residues Arg13 and Met36 are close to the catalytic center of tPphA. The conservative replacement of Arg13 by Lys resulted in a moderate reduction of activity. By contrast, the M36A variant showed strongly reduced catalytic activity towards all substrates, suggesting that the bulky methionyl side chain could be required to stabilize the structure of the catalytic center or to exclude water from the catalytic center. Alanine replacement of residues Thr138, Arg160, His161 and Arg169, all belonging to the flap subdomain (Figs 1A and 2A), moderately reduced enzyme activity towards all tested substrates. This effect on catalysis could be due to subtle flap subdomain interactions with the catalytically active third metal (M3) as shown in previous structures [20,21]. Finally, two variants, H39A and T138E, were differentially affected in substrate reactivity (Fig. 4A). These two variants could dephosphorylate artificial substrates but showed almost no activity towards PII-P. Alanine replacement of residue His39, which is highly conserved in bacterial PP2C family members (H39A), did not at all impair the reactivity towards pNPP, but, intriguingly, this variant displayed reduced activity towards artificial phosphopeptides and was almost completely unable to dephosphorylate PII-P. The undisturbed activity towards pNPP clearly indicates that this peripheral part of the enzyme does not directly take part in catalysis. The impaired reactivity towards phosphopeptides and phosphoprotein implies a role of His39 in substrate specificity. Mutation of Thr138 to Glu (T138E) resulted in a surprisingly high disturbance of activity. This was unexpected, since in numerous bacterial PP2C members the corresponding position is in fact occupied by Asp or Glu residues (see Supplementary file S1 in [20]). The methyl group of Thr138 projects into the basis of the flap subdomain and introduction of a negative charge at this place may distort the conformation of the entire flap subdomain region. A closer look at the data shows that the enzyme lost about 80% of activity when assayed with pNPP; it was even less active towards the artificial pS- and pT-phosphopeptides and was almost completely inactive towards phospho-PII. Strikingly, approximately 50% activity was regained with the T-loop peptide as substrate. Therefore, in addition to general damage of the catalytic activity, the T138E mutation seems to affect substrate specificity.

Table 1. Kinetic parameters of tPphA and its variants towards pNPP. pNPP assays were carried out as described in Experimental procedures. From the apparent reaction velocities of three independent repetitions, the kinetic parameters were calculated by linear fitting using the program graphpad prism 4. Standard errors are given. The results were obtained from single preparations of tPphA variants, human PP2Cα and chimera A purifications and the data come from single experiments.
  Km (mm) pNPP Kcat (s−1) pNPP Kcat/Km (s−1·m−1) Km (mm) Mn2+
WT0.47 ± 0.0700.85 ± 0.0551809 ± 1170.57 ± 0.088
R13K0.88 ± 0.0750.60 ± 0.026682 ± 300.96 ± 0.052
S15A0.42 ± 0.0380.83 ± 0.0201976 ± 480.57 ± 0.042
Q17E0.66 ± 0.0551.28 ± 0.0501939 ± 760.74 ± 0.062
Q17K1.02 ± 0.0900.17 ± 0.008167 ± 82.03 ± 0.187
M36A0.81 ± 0.0590.26 ± 0.009321 ± 110.78 ± 0.049
H39A0.62 ± 0.0441.10 ± 0.0361774 ± 580.68 ± 0.071
T138A0.45 ± 0.0520.52 ± 0.0251156 ± 560.90 ± 0.072
T138E0.74 ± 0.0650.26 ± 0.011351 ± 154.10 ± 0.33
R160A0.83 ± 0.0710.26 ± 0.011313 ± 131.34 ± 0.120
H161A0.57 ± 0.0410.57 ± 0.0181000 ± 320.82 ± 0.053
R169A0.61 ± 0.0620.49 ± 0.023803 ± 381.29 ± 0.118
Human PP2Cα0.45 ± 0.1921.37 ± 0.2593044 ± 5760.48 ± 0.15
Chimera A0.51 ± 0.1700.16 ± 0.024314 ± 470.63 ± 0.093
Table 2. The activity of tPphA and its variants towards three phosphopeptides. Reactions were performed in the buffer as described in Experimental procedures. Triplicate assays were used. Standard errors are given. The results were obtained from single preparations of tPphA variants, human PP2Cα and chimera A purifications and the data come from single experiments. −, activity is below 0.01 nmol·min−1·μg−1.
 pT-peptide (nmol·min−1·μg−1)pS-peptide (nmol·min−1·μg−1)T-loop peptide (nmol·min−1·μg−1)
WT7.73 ± 1.103.06 ± 0.214.22 ± 0.36
R13K4.61 ± 0.030.61 ± 0.131.02 ± 0.10
S15A8.96 ± 0.063.11 ± 0.094.34 ± 0.21
Q17E7.50 ± 0.065.22 ± 0.294.50 ± 0.25
Q17K0.68 ± 0.01 – –
M36A3.77 ± 0.010.52 ± 0.010.61 ± 0.02
H39A2.65 ± 0.170.47 ± 0.070.89 ± 0.02
T138A4.13 ± 0.151.90 ± 0.094.20 ± 0.37
T138E0.59 ± 0.050.098 ± 0.081.83 ± 0.03
R160A3.51 ± 0.061.72 ± 0.212.33 ± 0.12
H161A0.87 ± 0.010.21 ± 0.041.61 ± 0.05
R169A4.26 ± 0.090.70 ± 0.042.17 ± 0.13
Human PP2Cα16.2 ± 0.965.81 ± 0.0810.55 ± 0.53
Chimera A – –
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Figure 3.  (A) Graphical representation of PII-P dephosphorylation assays over a period of 45 min analyzed by non-denaturing gels, shown in Fig. S1A, for tPphA wild-type (WT) and variants S15A, Q17E, T138A, R13K, R169A and H161A. (B) Graphical representation of PII-P dephosphorylation assays over a period of 90 min analyzed by non-denaturing gels, shown in Fig. S1B, for tPphA variants H161A, R160A, H39A, M36A, T138E, Q17K and D193A.

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Figure 4.  Enzymatic assays of tPphA variants, human PP2Cα and chimera A. Relative activities of tPphA variants, human PP2Cα and chimera A towards different substrates are shown in (A) and (B). The activities of wild-type tPphA towards five substrates were set as 100% and the activities of other enzymes towards these five substrates were adjusted accordingly. pNPP indicates the value for Kcat/Km from the pNPP assay (Table 1). pT-peptide indicates the relative activity with RRA(pT)VA as substrate, pS-peptide indicates the activity with RRA(pS)VA peptide as substrate and T-loop peptide indicates the activity towards the CRYRG(pS)EYTV peptide (Table 2). PII-P indicates the initial activity of PII-P dephosphorylation (retrieved from the initial slope as shown in the curves of Fig. 3A, B and Fig. S1C).

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The activities of human PP2Cα and chimera A (tPphA catalytic core + human PP2Cα flap) were also assayed towards the above mentioned five different substrates. Human PP2Cα dephosphorylated artificial substrates faster than wild-type tPphA but was unable to dephosphorylate PII-P (Tables 1 and 2 and Fig. S1C). The inability to dephosphorylate PII-P is not due to an unfavorable amino acid context around phosphoseryl residue 49, since the phosphopeptide corresponding to the phosphorylated T-loop of PII is readily dephosphorylated. To prove that the flap subdomain of human PP2Cα is responsible for the specific exclusion of PII-P as substrate, the flap was grafted on the catalytic core of tPphA (chimera A). The resulting hybrid protein retained indeed some residual activity; however, it could only dephosphorylate pNPP to some extent (17% residual activity) but could not dephosphorylate phosphopeptides and PII-P (Tables 1 and 2 and Fig. 4B), indicative of an injured enzyme. The catalytic properties in the pNPP dephosphorylation assay showed that the hybrid enzyme was strongly affected for Kcat, whereas the Km values for pNPP and Mn2+ were very similar to the respective Km values of wild-type tPphA (Table 1). In agreement with these properties, isothermal titration calorimetry analysis of metal binding by chimera A revealed that chimera A bound Mn2+ with a similar affinity to wild-type tPphA (Fig. S2 and Table S1). These results imply that chimera A is able to form a metal cluster which is partly functional, but access of the substrates to the catalytic site is severely hampered in the hybrid protein. Only the smallest substrate, pNPP, can be processed in the catalytic site, albeit with reduced kinetics.

An assay to study the interaction between PII-P and tPphA

To examine how the various mutations of tPphA residues directly affect the binding of PII-P to the phosphatase, we sought to directly analyze the interaction of the proteins. Initial attempts to pull down PII-P with His-tagged tPphA on Ni-nitrilotriacetic acid (Ni-NTA) beads were unsuccessful, indicating that the interaction is either very weak and/or transient. Next, we tried to stabilize the interaction by using conditions under which the dephosphorylation reaction is arrested. In the structure of tPphA, obtained by crystallization in a 100 mm CaCl2 containing buffer, Mg2+ was found in the M1 position and Ca2+ in M2 and M3 sites [16]. Analyzing the effect of Ca2+ on wild-type tPphA activity showed that Ca2+ did not support any activity of tPphA and partially inhibited the Mg2+-supported activity towards phosphopeptides and PII-P (Fig. S3). Therefore, pull-down assays were performed under inhibitory buffer conditions (Ca2+ or EDTA, see below) without, however, recovering PII protein. Another attempt to stabilize the interaction intermediates was the addition of glutaraldehyde prior to affinity purification of tPphA and subsequent examination of PII and tPphA recovery by immunoblot analysis. The crosslinking reaction by the bis-aldehyde homobifunctional crosslinker glutaraldehyde proceeds through the formation of Schiff bases between aldehyde and lysine residues of crosslinked proteins [32]. Finally, conditions could be found under which PII could be specifically recovered in the tPphA elution fractions after glutaraldehyde treatment. Controls (PII with and without crosslinking reaction in the absence or presence of tPphA) revealed that the recovery of PII specifically depended on tPphA and proper buffer conditions (Fig. S4A). The fact that almost no crosslink products between PII and tPphA were found on SDS gels after heating the samples in SDS sample buffer can be attributed to the instability of the Schiff base formed through glutaraldehyde crosslinking [32,33]. A Ca2+-containing buffer was necessary to efficiently recover PII-P by this procedure, whereas no pull-down occurred in the presence of Mg2+ (see the lane labeled Mg2+ in Fig. 5A) or in the absence of divalent cations (see the lane labeled ‘−’ in Fig. 5A). Examination of the phosphorylation status of PII incubated under these assay conditions revealed that PII-P was dephosphorylated in the Mg2+-containing buffer, whereas almost no dephosphorylation occurred in the Ca2+-containing buffer or in the presence of EDTA (Fig. 5B). Therefore, the low recovery of PII-P in the presence of Mg2+ was probably due to dephosphorylation of PII-P and a weaker interaction of non-phosphorylated PII with tPphA. On the other hand, the lack of pull-down in the absence of any divalent metal may indicate a metal requirement for phosphoprotein binding (see below). To directly compare the recovery of phosphorylated and non-phosphorylated PII by this procedure, the same amount (0.2 μg) of either PII-P or strep-tagged PII (strep-PII) was used in a pull-down experiment in the presence of Ca2+ (Fig. 5C), and indeed much more PII-P was recovered than non-phosphorylated strep-PII.

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Figure 5.  (A) PII-P recovered by co-purification with tPphA after glutaraldehyde crosslinking in buffer containing Ca2+, Mg2+ or no divalent cation (−). 0.5 μg PII-P was used in the binding assay. Afterwards, the samples were analyzed by SDS/PAGE and immunoblot (anti-PII). CL, (PII) × 3, (PII) × 2 and (PII) × 1 are crosslinking products, trimeric PII, dimeric PII and monomeric PII, respectively. (B) Dephosphorylation of PII-P by tPphA under different conditions. 0.5 μg PII-P was incubated with 10 μg tPphA on ice in different buffers for 2 h, corresponding to the incubation time during the PII-P–tPphA binding assay. The reaction volume was 50 μL containing 110 mm NaCl together with either 2.5 mm Ca2+, 2.5 mm Mg2+and 5 mm EDTA, or without additions, as indicated above. After 2 h incubation, the phosphorylation status of PII was determined by non-denaturing PAGE and immunoblot analysis as described previously [38]. The non-phosphorylated form PII0 and the phosphorylated forms containing one, two or three phosphorylated subunits (PII1, PII2 and PII3) are indicated. (C) Comparison of PII-P (0.2 μg) and strep-PII (0.2 μg) binding to tPphA (10 μg) in Ca2+-containing buffer. PII-P and strep-PII was recovered by co-purification with tPphA after glutaraldehyde crosslinking in buffer containing Ca2+. Afterwards, the samples were analyzed by SDS/PAGE and immunoblot (anti-PII).

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Interaction of tPphA variants, human PP2Cα and chimera A with PII-P

The interaction of tPphA variants with PII-P was investigated using the glutaraldehyde crosslinking and pull-down assay described above. Figure 6A shows the immunoblot analysis of PII recovered from the pull-down assays with tPphA variants. The control for tPphA recovery is shown in Fig. S5. In many cases, there was a clear correlation between reactivity towards and binding of PII-P: variants Q17K and T138E which displayed very low activity towards PII-P could not pull-down PII-P, whereas the same position variants Q17E and T138A, which were only slightly impaired in reactivity towards PII-P, showed apparently equal or more binding of PII-P than wild-type tPphA. Furthermore, tPphA variants D18A, D34A and D193A, coordinating the M1–M2 cluster and being completely inactive [21], could not bind any PII-P (Fig. 6A). By contrast, the variants of the loosely M1-coordinating Asp231 residue (D231A) and of the M3-coordinating Asp119 residue (D119A), which are also unable to dephosphorylate PII-P, could still bind this substrate. The flap subdomain variants (R160A, H161A as well as R169A), which are partially impaired in catalytic activity, were also partially impaired in PII-P binding. Surprisingly, the variant H39A was able to recover PII-P although the enzyme was severely impaired in dephosphorylating PII-P. The human PP2Cα and chimera A, both having an active metal center (see above), could not recover PII-P (Fig. 6B), in agreement with the lack of PII-P dephosphorylation.

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Figure 6.  (A) Assay of PII-P binding to wild-type tPphA (WT) and the variants according to the crosslinking and pull-down protocol (see Fig. 5A). The recovery of PII (in monomeric, dimeric, trimeric and tPphA crosslinked forms, see indications in Fig. 5A) is shown by anti-PII immunoblotting. (B) Assay of PII-P binding to tPphA, chimera A and human PP2Cα. Recovery of PII was visualized by anti-PII immunoblotting. These assays were performed three times with similar results. All assays were performed as described in Experimental procedures.

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Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

As deduced from this study, the selection of the phosphoprotein substrate, here the PII-P protein, requires a concerted interaction of the phosphatase with the substrate. The M1–M2 catalytic core is the central region for binding of PII-P. Structures of the PP2C member MspP (from Mycobacterium smegmatis) in various complexes with phosphate, sulfate or the phosphate analog cacodylate showed that two oxygen atoms of the substrate phosphate group directly interacted with M1 and M2 by bidentate coordination [18]. The MspP–cacodylate complex was suggested to mimic the competent enzyme–substrate complex [18]. Modeling of the cacodylate molecule from the MspP structure into the tPphA structure in such a way that the distance between cacodylate and M1 and M2 remains constant showed that the cacodylate oxygen atoms would replace the oxygen atoms from water molecules coordinated by M1 and M2 in tPphA [21], which indicates that M1–M2 of tPphA could indeed directly bind the substrate phosphate. In agreement, the variants, which cannot correctly assemble the M1–M2 core, were impaired in recovering PII-P in pull-down experiments. D18A and D34A variants were shown to be highly impaired in metal binding [21] and the crystal structure of the D193A showed that the position of M1 was shifted by 1.8 Å in this variant [21]. By contrast, the third metal (M3) appears to be less important for substrate binding, since the D119A variant recovered PII-P in the binding assay. The recently solved crystal structure of the D119A variant [21] showed that the M1–M2 cluster and the entire catalytic core were not affected compared with the wild-type structure. However, M3 was specifically missing and the segment of the flap subdomain near M3 was moved by 1.8 Å towards the center. All these data together with previous investigations [18,21] suggest that the positively charged M1–M2 core of tPphA is critical for trapping the substrate phosphate group, whereas M3 is involved in the subsequent dephosphorylation reaction.

According to the enzymatic properties of the H39A variant, the His39 residue appears to play an important role for the specificity towards substrates, since reactivity towards pNPP was unaffected but the activity towards phosphopeptides was substantially impaired and the reactivity towards PII-P almost vanished (Fig. 4A). The catalytic center of the H39A variant is apparently not impaired, as expected from the distant location of the His39 residue relative to the catalytic center. His39 belongs to a small loop (at the beginning of motif 3, Figs 1A and 2A) which is located on the distal end of the catalytic face (on the opposite site of the flap subdomain) flanking a cleft, which protrudes towards M2 (Fig. 1A). The other site of this cleft is flanked by Ser15, which can be replaced by Ala without loss of substrate specificity. His39 is invisible in one space group (C2221) of the crystal structures of tPphA [16] and the variant D193A [21], indicating that this part of the protein is flexible. The present work provides the first evidence by mutational analysis that the conserved His39 residue is indeed involved in substrate recognition, as already suspected from the crystallographic analysis of the S. agalactiae PP2C phosphatase SaSTP (see Introduction) [19]. According to the pull-down data, it appears that His39 is not essential for the coarse binding of PII-P. Instead, His39 might be involved in fine-tuning the formation of a productive substrate–enzyme complex that allows the subsequent dephosphorylation reaction.

In the cleft flanked by His39 and Ser15, near M2, residue Gln17 plays another crucial role in tPphA function. Gln17 is the homolog residue of HAB1 Glu203. In the co-crystal structure of PYR1 (pyrabactin resistance1) with HAB1 from A. thaliana, HAB1 Glu203 directly formed a hydrogen bond with PYR1 Ser85. This interaction was suggested to mimic the interaction of a serine-containing peptide with the catalytic center of a PP2C phosphatase [27]. The study showed that Gln17 can be replaced by Glu without loss of function. The Q17E tPphA variant recovered even higher amounts of PII-P than the wild-type enzyme. Similar to the interaction of HAB1 Glu203 with PYR1 Ser85, tPphA Glu17 may take part in substrate binding, explaining the increased recovery of PII-P by Q17E. By contrast, the Q17K variant was almost completely inactive. The positive charge of Lys17 could impair the binding of M2 and thus compromise enzyme function. In agreement with that conclusion, the variant could not recover PII-P in the pull-down assay (Fig. 6A).

A further region of the enzyme that specifies substrate recognition could be the flap subdomain. Two major lines of experimental evidence, both with some limitations, are in favor of this conclusion. The first line comes from the comparison of human PP2Cα and tPphA: both enzymes are highly active towards artificial substrates and human PP2Cα is even very active towards the peptide corresponding to the phosphorylated T-loop of PII; however, it is completely inactive towards the entire phosphoprotein PII-P. The major structural difference concerns the flap subdomain (Fig. 1D), which we therefore suspected plays an important role in hindering the dephosphorylation of PII-P. Nevertheless, other differences exist, which could be important as well. For example, His39, highly conserved in bacterial PP2C members, is not conserved in human PP2Cα. The attempt to graft the flap of human PP2Cα on tPphA and vice versa had only limited success: only the combination of tPphA core and human PP2Cα flap resulted in a soluble protein, but this enzyme was only partially active. Nevertheless, some conclusions can be drawn from studying this chimeric protein. The fact that the enzyme bound metals with similar affinity to wild-type tPphA and could dephosphorylate pNPP, albeit with reduced velocity, indicates that chimera A can form at least a partially active catalytic center. The large flap subdomain may collapse on top of the catalytic site and thereby limit its accessibility, which would explain why only the smallest substrate, pNPP, can be dephosphorylated. In future studies, the catalytic core and flap subdomain of enzymes which are more closely related will be combined, expecting that these chimeras will be catalytically less impaired. The second experimental approach to narrow down the function of the flap subdomain was by introducing point mutations. Almost every mutation in the flap region (T138E, R160A, H161A and R169A) reduced the activity of the respective variant, indicating a general role of the flap subdomain in catalytic efficiency. Subtle interactions with the nearby M3 metal could play a role, as suspected earlier [20,21]. Whereas most flap subdomain variants did not lead to conclusive results, the T138E mutation is particularly intriguing. In many other PP2C homologs, this position is naturally occupied by an Asp or Glu residue (see Supplementary file S1 in [20]). In tPphA, the Thr to Glu replacement heavily affected enzyme activity and strongly reduced binding to PII-P. Activity could be partially restored (to 50% of wild-type) by using the T-loop peptide as substrate, whereas the PII-P substrate is neither turned over nor bound. The fact that a single amino acid substitution in the flap subdomain region is sufficient to completely alter the substrate specificity agrees with the assumption that the flap subdomain is involved in substrate selection.

The phosphatase must be able to precisely recognize its substrate protein, and therefore fine-tuned interactions between phosphatase and substrate are necessary. Here, we have shown that the His39 residue, M1–M2 core and probably the flap subdomain are involved in this recognition process. Only when a productive substrate–enzyme complex is formed is the enzyme able to dephosphorylate its substrate. This mechanism prevents dephosphorylation of non-cognate phosphoproteins. For many bacterial PP2Cs, this recognition process seems to be sufficient, since they lack further substrate binding domains and solely depend on the specificity provided by the interaction between substrate and enzyme on the face of the phosphatase. The flap subdomain is the region with the highest sequence variability in bacterial PP2C members. Further experiments are required to clearly establish whether the flap subdomain is indeed the clue to distinguishing between the different protein substrate specificities of the various PP2C members.

Experimental procedures

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Cloning, overexpression and purification of tPphA variants

The tPphA gene (tlr2243) encoding tPphA was amplified from plasmid pET32a + tPphA [16] by using the T7 primers (Table S2), which amplify genes between the T7 promoter and T7 terminator in the pET (Novagen, Darmstadt, Germany) series plasmids. The resulting PCR product, which contains an NdeI site at the tPphA start codon and a 3′ terminal XhoI restriction site was first cloned using the pGEM-T easy vector system (Promega, Madison, WI, USA). Following restriction of the resulting plasmid with NdeI and XhoI, the tPphA fragment was isolated and cloned between the NdeI and XhoI sites of His-tag expression vector pET15b-vector (Novagen), generating an N-terminal fusion to the 6× His-tag sequence from the vector. The procedure of site-directed mutagenesis of tPphA was carried out with the QuickChange XL site-directed mutagenesis kit (Stratagene, La Jolla, CA, USA). The primers for site-directed mutagenesis are shown in Table S2. All constructs were checked by DNA sequencing. For overproduction of the recombinant enzymes, the constructs were transformed into Escherichia coli BL21 (DE3) cells (New England Biolabs GmbH, Frankfurt am Main, Germany) and grown in LB medium supplemented with ampicillin (100 μg·mL−1). When the optical density (D600) of the culture reached a value of 1–1.5, isopropyl thio-β-d-galactoside (IPTG) was added to a final concentration of 0.5 mm to induce protein overproduction. After 4–5 h induction at 25 °C, the cells were harvested by centrifugation and lysed by sonification (Branson Sonifier 250, Danbury, CT, USA) on ice in a buffer consisting of 20 mm Tris/HCl, pH 7.8, 75 mm KCl, 500 mm NaCl, 5 mm MgCl2, 0.5 mm EDTA, 0.2 mm phenylmethylsulfonyl fluoride. The lysate was centrifuged for 30 min at 16 000 g. The supernatant was centrifuged again for 30 min at 16 000 g. The clarified cell extract was used for protein purification. HIS-Select Cartridges (Sigma, St Louis, MO, USA) were used to purify His-tag proteins. After purification, the proteins were dialyzed in a buffer (20 mm Hepes, pH 7.4, 75 mm KCl, 500 mm NaCl, 2 mm MgCl2, 0.5 mm EDTA, 1 mm dithiothreitol, 50% glycerol) and analyzed by SDS/PAGE. The concentrations of the purified proteins were determined by the Bradford method (Rotiquant; Carl Roth, Karlsruhe, Germany) and from UV spectra by calculation with an absorption coefficient at 280 nm of 27 960 m−1·cm−1 (calculated from http://www.expasy.ch/tools/protparam.html).

Cloning, overexpression and purification of human PP2Cα and the chimeras

His-human PP2Cα was purified as described previously with no modification [21]. Artificial genes for chimera A [tPphA catalytic center (1–137, 174–240) + human PP2Cα flap (165–219)] and chimera B [human PP2Cα catalytic center (1–164, 220–297) + tPphA flap (138–173)] were synthesized and cloned into His-tag pET15b vector by Geneart (Regensburg, Germany). At first, overexpression and purification of His-tagged chimeras were performed as in the previous section. His-tagged chimera A was purified and stored in a buffer (20 mm Hepes, pH 7.4, 75 mm KCl, 500 mm NaCl, 2 mm MgCl2, 0.5 mm EDTA, 1 mm dithiothreitol, 50% glycerol). Unfortunately, most of chimera B became insoluble inclusion bodies in E. coli BL21 (DE3) after being induced by IPTG. Only residual soluble chimera B could be purified by HIS-Select Cartridges (Sigma). After protein purification, the soluble form chimera B could be dialyzed against a buffer (20 mm Hepes, pH 7.4, 75 mm KCl, 500 mm NaCl, 2 mm MgCl2, 0.5 mm EDTA, 1 mm dithiothreitol, 50% glycerol), and it remained soluble. After dialysis, the concentration (0.2 μg·μL−1) of chimera B was determined by the Bradford method. But it precipitated at once when it was added into any other buffer.

In order to obtain soluble chimera B, modified methods for overexpression and purification were applied. In method 1, overexpression of His-tagged chimera B was performed as in the previous section. When the optical density (A600) of the E. coli BL21 (DE3) culture reached a value of 1, IPTG was added to a final concentration of 0.5 mm to induce protein overproduction. After 4 h induction at 25 °C, the cells were harvested by centrifugation and lysed by sonification (Branson Sonifier 250) on ice in a buffer consisting of 20 mm Tris/HCl, pH 7.8, 75 mm KCl, 500 mm NaCl, 5 mm MgCl2, 0.5 mm EDTA, 0.2 mm phenylmethylsulfonyl fluoride. The lysate was centrifuged for 30 min at 16 000 g. The pellet containing most of chimera B in the form of inclusion bodies was redissolved and denatured by a buffer containing 0.1 m NaH2PO4/Na2HPO4, pH 7.4, 6 m guanidine. The solution was centrifuged for 30 min at 16 000 g. After centrifuging, the content of the supernatant was checked by SDS/PAGE. SDS/PAGE showed a strong band at 29 kDa and proved that chimera B could be denatured and solubilized by 6 m guanidine. In order to remove guanidine and refold chimera B, the supernatant was dialyzed against a series of buffers containing different concentrations (6, 4, 2, 1 and 0 m) of guanidine, 40 mm Hepes, pH 7.4, 0.5 m NaCl, 2 mm MgCl2. After the supernatant containing soluble chimera B was exchanged from the dialysis buffer containing 2 m guanidine to the dialysis buffer containing 1 m guanidine, all of chimera B precipitated. Therefore, no chimera B could be refolded.

In method 2, E. coli BL21 (DE3) cells containing plasmid pET15b + chimera B were grown in LB medium supplemented with ampicillin (100 μg·mL−1). When the optical density (A600) of the culture reached 0.4 or 0.6 IPTG was added to a final concentration of 0.5 mm to induce protein overproduction. After 3 h induction at 18 °C, the cells were harvested by centrifugation and lysed by a French pressure cell homogenizer (Emulsiflex-B15; Avestin, Ottawa, Canada) in a buffer consisting of 20 mm Tris/HCl, pH 7.8, 75 mm KCl, 500 mm NaCl, 5 mm MgCl2, 0.5 mm EDTA, 0.2 mm phenylmethylsulfonyl fluoride. The lysate was centrifuged for 30 min at 16 000 g. The supernatant was centrifuged again for 30 min at 16 000 g. The clarified cell extract was used for protein purification. HIS-Select Cartridges (Sigma) were used to purify His-tag chimera B. After purification, the proteins were dialyzed in a buffer (20 mm Hepes, pH 7.4, 75 mm KCl, 500 mm NaCl, 2 mm MgCl2, 0.5 mm EDTA, 1 mm dithiothreitol, 50% glycerol) and analyzed by SDS/PAGE. The concentration (0.3 μg·μL−1) of the purified protein was determined by the Bradford method (Rotiquant; Carl Roth). Chimera B still precipitated when it was exchanged from the first buffer (20 mm Hepes, pH 7.4, 75 mm KCl, 500 mm NaCl, 2 mm MgCl2, 0.5 mm EDTA, 1 mm dithiothreitol, 50% glycerol) to any other buffer. Therefore, no further experiments could be performed with chimera B and the characteristics of chimera B were unknown.

Preparation of phosphorylated PII and strep-PII

Phosphorylated PII protein (PII-P) was prepared from Synechococcus elongatus PCC 7942 as described previously with modifications [34]. To achieve a maximum degree of in vivo PII phosphorylation, the cells were grown in 4.5 L BG-11 medium containing 17.6 mm NaNO3 supplemented with 10 mm NaHCO3 and ferric ammonium citrate was replaced by ferric citrate. When A750 of the cells reached 1.0, the cells were treated with 0.5 mm l-methionine sulfoximine for 4 h and were then harvested by centrifugation. All subsequent steps were carried out at 0–8 °C. The cells were broken by sonification (Branson Sonifier 250) in a buffer consisting of 50 mm Tris/HCl, pH 7.4, 4 mm EDTA, 0.1 mm phenylmethylsulfonyl fluoride. Cell debris was removed by centrifugation for 10 min at 10 000 g and the crude extract was cleared by high speed centrifugation at 100 000 g for 60 min. This crude extract enriched with phosphorylated PII was used for the experiments described in the section ‘The activities of tPphA variants, human PP2Ca and chimera A towards PII-P’ below. The crude extract was applied to an Econo-Pac heparin cartridge (Bio-Rad, Hercules, CA, USA) equilibrated in 10 mm Hepes, pH 7.4. Proteins were eluted with a 100 mL linear gradient of 0–400 mm NaCl in 10 mm Hepes, pH 7.4, at a flow rate of 1 mL·min−1. Fractions (4 mL) were collected in tubes containing 50 μL of 0.4 m EDTA to prevent dephosphorylation of PII-P. The PII-P containing fractions were detected by immunoblot (dot-blot) analysis using PII-specific antibodies. Ammonium sulfate was added to a saturation of 80% to precipitate proteins. The precipitated proteins were collected and resuspended in 10 mm Tris/HCl, pH 7.4, 150 mm NaCl, 5 mm EDTA. The solution was applied to a Superdex 200 10/300 GL column (Amersham Pharmacia, Freiburg, Germany) equilibrated in a buffer (10 mm Tris/HCl, pH 7.4, 150 mm NaCl, 5 mm EDTA) and developed in the same buffer at a flow rate of 0.5 mL·min−1. The PII-P containing fractions were concentrated by ultrafiltration (Microcon YM-10, 10 kDa cut-off; Millipore, Bedford, MA, USA). This purified phospho-PII was used for the experiments described in the sections ‘PII– tPphA binding assay’ and ‘Human PP2Cα and chimera bind to PII-P’ below. Purification of recombinant PII proteins with a C-terminal-fused strep-tag II peptide was performed according to Heinrich et al. [35].

Assay of phosphatase activity with pNPP as artificial substrate

The activities of tPphA variants, human PP2Cα and chimera A towards pNPP were assayed as described previously with modifications [16,21]. Standard assays in a volume of 250 μL contained 0.25–10 μg tPphA variants, 0.25 μg human PP2Cα and 0.25–10 μg chimera A in a buffer consisting of 10 mm Tris/HCl, pH 8.0, 50 mm NaCl, 1 mm dithiothreitol and 2 mm MnCl2. Reactions were started by the addition of 2 mmpNPP at 30 °C and the increase in absorbance at 400 nm was measured in an ELx808 absorbance microplate reader (BioTek, Winooski, VT, USA) against a blank reaction without enzyme. To determine the Km for Mn2+, the pNPP concentration was fixed at 1.5 mm and the Mn2+ concentration was varied from 0.1 to 8 mm. For pNPP catalytic constants, the pNPP concentration was varied from 0.1 to 2 mm. From the linear slope of each reaction, the kinetic parameters Km and Vmax were calculated by nonlinear hyperbolic fitting using the graphpad prism 4 program (GraphPad Software Inc., La Jolla, CA, USA).

Reactivity of tPphA variants, human PP2Cα and chimera A towards phosphopeptides

Three different phosphopeptides were used to assay the activities of tPphA variants and chimera A. The sequences of the three peptides are CRYRG(pS)EYTV, RRA(pT)VA and RRA(pS)VA. CRYRG(pS)EYTV corresponds to the T-loop sequence of the PII protein. In a standard assay, 0.25–10 μg tPphA variants, 0.25 μg human PP2Cα and 0.25–10 μg chimera A were reacted with 100 μm phosphopeptides in a reaction volume of 100 μL containing 50 mm Tris/HCl, pH 8.0, 50 mm NaCl, 2 mm MnCl2 and 1 mm dithiothreitol. Reactions were incubated at 30 °C for 2–10 min, and then stopped by the addition of 50 μL 4.7 m HCl. The released Pi was quantified colorimetrically by the malachite green assay [36,37]. The absorbance of the solution at 630 nm was measured in an ELx808 absorbance microplate reader (BioTek) against a blank reaction, which was stopped at the start point by 50 μL 4.7 m HCl. The activity of all enzymes toward peptides was calculated with a phosphate standard.

The activities of tPphA variants, human PP2Ca and chimera A towards PII-P

Dephosphorylation assays of PII-P were carried out in 10 mm Tris/HCl, pH 8.0, 50 mm NaCl, 1 mm dithiothreitol and 2 mm MgCl2. 30 μL reactions contained 5 μL Synechococcus extract containing 60 ng highly phosphorylated PII and 100 ng enzymes. Reactions were started by the addition of purified phosphatase. After incubation at 30 °C the reactions were stopped by adding 2.5 μL 100 mm EDTA on ice. Subsequently, the phosphorylation state of PII was determined by non-denaturing PAGE and immunoblot analysis as described previously [38].

PII– tPphA binding assay

Each step of the following binding assay was carried out on ice or at 4 °C. tPphA (10 μg wild-type or tPphA variant proteins) was incubated with PII-P (0.2 or 0.5 μg) or strep-PII (0.2 or 5 μg) in 50 μL of buffer I containing 1 mm Tris/HCl, pH 7.4, 2.5 mm CaCl2, 110 mm NaCl, 0.05 mm EDTA and 0.08% (w/w) glutaraldehyde (Carl Roth). After 60 min, 2.5 μL of 1 m imidazole (pH 8.0) was added to the reaction. After further incubating on ice for 15 min, the reaction was diluted with 420 μL buffer II containing 2.5 mm CaCl2, 100 mm NaCl and 0.05% v/v Tween-20. To extract His-tagged tPphA, 30 μL Ni-NTA magnetic agarose beads (Qiagen, Hilden, Germany) were added. The mixture was gently mixed by rotation for 1 h. Thereafter, the Ni-NTA beads were removed with a magnet and washed three times with 0.5 mL washing buffer (10 mm Tris/HCl, pH 8.0, 2.5 mm CaCl2, 100 mm NaCl, 20 mm imidazole). Finally, the bound proteins were eluted by denaturing the sample with 50 μL SDS sample buffer (75 mm Tris/HCl, pH 6.8, 1.5% dithiothreitol, 1% SDS, 10% glycerol, 0.04% bromophenol blue) at 95 °C for 5 min. Subsequently, the eluted proteins were visualized by SDS/PAGE and immunoblot analysis as described previously [39]. In order to reveal the effect of divalent cations on the binding between PII and tPphA, 2.5 mm CaCl2 was either removed or changed to 2.5 mm MgCl2 in buffer I, buffer II and the washing buffer.

Human PP2Cα and chimera A bind to PII-P

The procedures were same as in the previous section.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

We thank Christina Herrmann and Oleksandra Fokina for practical help. This work was supported by DFG grant Fo195 and by GRK 685 ‘infection biology’.

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Supporting Information

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Fig. S1. (A), (B) Time course assay ofPII-P dephosphorylation by tPphA variants. (C) Timecourse assay of PII-P dephosphorylation by tPphA, humanPP2Cα and chimera A.

Fig. S2. Mn2+binding to tPphA,chimera A and human PP2Cα studied by isothermal titration calorimetry.

Fig. S3. (A) Inhibitory effect ofCa2+ on the relative activities of tPphA towardspNPP, pT-peptide, T-loop peptide and PII-P. (B)Inhibitory effect of Ca2+ on the dephosphorylation ofPII-P by wild-type tPphA.

Fig. S4. Glutaraldehyde (GDA) crosslinkingfollowed by Ni-NTA affinity purification (pull-down, PD) usingHis-tPphA (10 μg) with PII-P(0.5 μg).

Fig. S5. Recovery of tPphA from the pull-down assays shown in Fig. 6(A).

Table S1. The affinity and thermodynamicparameters of tPphA, chimera A and human PP2Cα from ITC assay.

Table S2. Primers used for PCR amplification of tPphA and for site-directed mutagenesis.

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