Mercuric ions inhibit mitogen-activated protein kinase dephosphorylation by inducing reactive oxygen species

Mercuric ions inhibit mitogen-activated protein kinase dephosphorylation by inducing reactive oxygen species

Toxicology and Applied Pharmacology 250 (2011) 78–86 Contents lists available at ScienceDirect Toxicology and Applied Pharmacology j o u r n a l h o...

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Toxicology and Applied Pharmacology 250 (2011) 78–86

Contents lists available at ScienceDirect

Toxicology and Applied Pharmacology j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / y t a a p

Mercuric ions inhibit mitogen-activated protein kinase dephosphorylation by inducing reactive oxygen species Hajo Haase, Gabriela Engelhardt, Silke Hebel, Lothar Rink ⁎ Institute of Immunology, Medical Faculty, RWTH Aachen University, Pauwelsstrasse 30, 52074 Aachen, Germany

a r t i c l e

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Article history: Received 6 July 2010 Revised 19 September 2010 Accepted 6 October 2010 Available online 14 October 2010 Keywords: MAPK Mercuric ions ROS Zinc

a b s t r a c t Mercury intoxication profoundly affects the immune system, in particular, signal transduction of immune cells. However, the mechanism of the interaction of mercury with cellular signaling pathways, such as mitogen activated protein kinases (MAPK), remains elusive. Therefore, the objective of this study is to investigate three potential ways in which Hg2+ ions could inhibit MAPK dephosphorylation in the human Tcell line Jurkat: (1) by direct binding to phosphatases; (2) by releasing cellular zinc (Zn2+); and (3) by inducing reactive oxygen species (ROS). Hg2+ causes production of ROS, measured by dihydrorhodamine 123, and triggers ROS-mediated Zn2+ release, detected with FluoZin-3. Yet, phosphatase-inhibition is not mediated by binding of Zn2+ or Hg2+. Rather, phosphatases are inactivated by at least two forms of thiol oxidation; initial inhibition is reversible with reducing agents such as Tris(2-carboxyethyl)phosphine. Prolonged inhibition leads to non-reversible phosphatase oxidation, presumably oxidizing the cysteine thiol to sulfinic- or sulfonic acid. Notably, phosphatases are a particularly sensitive target for Hg2+-induced oxidation, because phosphatase activity is inhibited at concentrations of Hg2+ that have only minor impact on over all thiol oxidation. This phosphatase inhibition results in augmented, ROS-dependent MAPK phosphorylation. MAPK are important regulators of T-cell function, and MAPK-activation by inhibition of phosphatases seems to be one of the molecular mechanisms by which mercury affects the immune system. © 2010 Elsevier Inc. All rights reserved.

Introduction Each year, anthropogenic and natural sources release several thousand metric tons of mercury into the environment (Hammond, 1971). This leads to exposure of humans to methylmercury through the food chain, mostly from fish. In addition, there is uptake of elemental mercury vapor from tooth fillings and ethylmercury from thimerosal, a preservative in vaccines (Clarkson et al., 2007). In the body, these different species are metabolized. Hereby, inorganic mercury is converted from elemental metallic mercury to its higher 2+ oxidation states mercurous (Hg2+ ), 2 ) and mercuric cations (Hg whereby in vivo most inorganic mercury is rapidly oxidized to the divalent form (Clarkson et al., 2007). Mercury is toxic to a wide range of cell types, including lymphocytes and macrophages (Shenker et al., 1992), a process involving reactive oxygen species (ROS) (Kim and Sharma, 2003). Nevertheless, many effects on the immune system are observed at Abbreviations: AA, ascorbic acid; DHR123, dihydrorhodamine 123; ERK, extracellular signal-regulated kinase; IL, interleukin; JNK, c-Jun N-terminal kinase; MAPK, mitogen activated protein kinase; MKP, MAPK phosphatase; N-AC, N-acetylcysteine; MKP, MAPK phosphatase; PHA, phytohaemagglutinin; PTP, protein tyrosine phosphatase; ROS, reactive oxygen species; TCEP, Tris(2-carboxyethyl) phosphine; TNF, tumor necrosis factor; TPEN, N,N,N′,N′-tetrakis-2(pyridyl-methyl)ethylenediamine. ⁎ Corresponding author. Fax: + 49 241 8082613. E-mail address: [email protected] (L. Rink). 0041-008X/$ – see front matter © 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.taap.2010.10.007

doses below cellular or systemic toxicity, leading to impaired immunity at low level exposure (Vas and Monestier, 2008). Mercury can also trigger unwanted immune reactions such as inflammation or autoimmunity (Vas and Monestier, 2008). For example, mercury exposure leads to lymphocytes secreting pro-inflammatory cytokines such as tumor necrosis factor (TNF)-α (Kim et al., 2003; Kim and Sharma, 2003). On the molecular level, inorganic mercury interacts with several components of signal transduction (Rossi et al., 1991). In particular, there are multiple reports describing an activation of the mitogen activated protein kinases (MAPKs) p38, c-Jun N-terminal kinase (JNK), and extracellular signal-regulated kinase (ERK)1/2 by mercury (Barnes and Kircher, 2005; Kim et al., 2002; Kim et al., 2005; Kim and Sharma, 2004; Turney et al., 1999). Hereby, ERK is only weakly activated. Also, ERK activation by other stimulants, e.g. in response to T-cell receptor stimulation, can be disrupted by mercury inhibiting upstream signaling (Mattingly et al., 2001; Ziemba et al., 2006). Not much is known about the mechanism by which mercury affects MAPKs; in the case of p38 MAPK activation, however, it was shown that ROS are involved (Kim et al., 2003; Kim and Sharma, 2003; Kim and Sharma, 2004). MAPKs are an important part of immune cell signal transduction (Dong et al., 2002). Activity of MAPKs is regulated by dual phosphorylation of a Thr-X-Tyr motif (where X denotes any amino acid). Phosphate groups are transferred to the MAPK by upstream

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MAPK kinases (MAPKK) and removed by dual-specificity MAPK phosphatases (MKP), members of the protein tyrosine phosphatase (PTP) family (Junttila et al., 2008). Multiple mechanisms regulate PTP activity, including gene expression, ligand binding, protein degradation, intracellular localization, and phosphorylation (Samet and Tal, 2010). In addition, PTP are inhibited by zinc (Haase and Maret, 2003; Haase and Rink, 2009; Tal et al., 2006; Taylor et al., 2008), presumably by binding to the active site cysteine in the characteristic (H)CX5R signature motif (Haase and Rink, 2009). Furthermore, phosphataseinhibition has also been demonstrated for another group IIb metal, cadmium (Haase et al., 2010).

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Due to its high affinity for thiol groups (Oram et al., 1996), mercury could bind directly to the active site cysteine and interfere with the catalytic reaction. Alternatively, mercuric chloride displaces protein bound zinc, e.g. from metallothionein (Haase et al., 2006), which could result in zinc-mediated phosphatase inhibition. Zinc is a potent inhibitor of protein phosphatases that dephosphorylate MAPK, including MKPs and PP2A (Ho et al., 2008; Zhuo and Dixon, 1997). Another feature of mercury that may be relevant for phosphatase activity is the generation of reactive oxygen species (ROS) (Lund et al., 1993). The catalytic cysteine of phosphatases is highly susceptible to oxidation by ROS, leading to inactivation of these enzymes (Salmeen

Fig. 1. Zinc-release by Hg2+ in leukocyte subpopulations. Primary human leukocytes were loaded with FluoZin-3 and the resulting zinc-dependent fluorescence monitored by flow cytometry. Granulocyte, monocyte, and lymphocyte populations were distinguished by forward and sideward scatter. (A) Time dependent depiction of fluorescence. After 300 s, the indicated concentrations of HgCl2 were added (arrows). Representative data from one out of n = 3 donors are shown. (B, C) Fluorescence at 600 s after addition of HgCl2 was used to calculate the concentration of free zinc (B) and the increase of the free zinc concentration relative to the concentration in untreated control cells (C). Data are shown as means ± SD of n = 3 donors. Values in (B) that differ statistically significant from untreated controls are indicated (**p b 0.01, one-way ANOVA with Dunnett's Post Hoc test).

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and Barford, 2005). This involves reversible oxidation to disulfides and sulfenic acid, and further, irreversible oxidation to sulfinic and sulfonic acid (Samet and Tal, 2010). The objective of this study is to investigate the impact of Hg2+ on phosphatase activity in the human T-cell line Jurkat. It compares direct inhibition of PTP by Hg2+, the role of zinc, and oxidation by Hg2+induced ROS formation. We found that Hg2+ inhibits MKPs by an oxidative mechanism, involving reversible and irreversible oxidation and leading to increased MAPK phosphorylation.

Isolation and culture of primary human leukocytes. Primary human leukocytes were isolated from heparinized whole blood of three healthy, consenting donors from the institute of immunology (2 females, 1 male) by addition of 1 part of 6% hydroxyethyl starch solution to 2 parts of blood. After erythrocyte sedimentation for 30 min at room temperature, the leukocyte fraction was washed twice with PBS. The pellet was resuspended in distilled water to lyse remaining erythrocytes. Lysis was stopped by addition of 10× PBS, followed by transfer into culture medium (RPMI 1640 containing the same supplements as above).

Materials and methods

Detection of free zinc and reactive oxygen species with fluorescent probes. Cells were loaded with FluoZin-3AM (1 μM) or dihydrorhodamine 123 (DHR123, 1 μg/ml) in loading buffer (25 mM HEPES, pH 7.35, 120 mM NaCl, 5.4 mM KCl, 5 mM glucose, 1.3 mM CaCl2, 1 mM MgCl2, 1 mM NaH2PO4, 0.3% bovine serum albumin) for 30 min at 37 °C. Subsequently, cells were washed twice with measurement buffer (loading buffer w/o albumin) and transferred into a 96 well plate. Fluorescence was recorded on a Tecan Ultra 384 fluorescence well plate reader (Tecan, Crailsheim, Germany) using an excitation wavelength of 485 nm and measuring the emission at 535 nm. Cells were maintained at 37 °C during the entire experiment and measurements were made in 2-min intervals. Flow cytometric zinc measurements were done on a BD FACSCalibur. Forward and sideward scatter were used to distinguish monocyte, granulocyte, and lymphocyte populations. Free zinc concentrations were

Materials. RPMI 1640 cell culture medium, penicillin, streptomycin, L-glutamine, and phosphate buffered saline (PBS, 1× and 10×) were purchased from Lonza (Verviers, Belgium). Low endotoxin fetal calf serum (FCS) was obtained from PAA (Cölbe, Germany) and was heat inactivated for 30 min at 56 °C prior to use. Western Blot antibodies were purchased from New England Biolabs, Frankfurt, Germany. All other chemicals were of analytical quality and obtained from standard sources. Cell culture. Jurkat cells (a human acute T-cell leukemia cell line) were seeded in RPMI 1640 supplemented with 10% FCS, 2 mM Lglutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin. Cells were cultured at 37 °C, 100% humidity in a mixture of 95% air and 5% CO2.

Fig. 2. Formation of reactive oxygen species after incubation with Hg2+. (A, B, C) Jurkat T-cells were loaded with DHR 123 and the resulting fluorescence measured in 2-min intervals. After 10-min recording of the baseline, 5 or 25 μM HgCl2 were added (arrows) either to cells without antioxidants (A), or pre-incubation with N-acetylcysteine (B) or ascorbic acid (C) for 30 min. Measurements are shown as means of triplicates ± SD from one representative experiment out of n = 3. (D) Cellular viability and induction of apoptosis were measured by flow cytometry after staining with propidium iodide and Annexin V. Data are shown as means from n = 4 independent experiments ± SD. Values that differ statistically significant from untreated controls are indicated (**p b 0.01, one-way ANOVA with Dunnett's Post Hoc test).

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calculated as previously described (Haase et al., 2006), using a dissociation constant of the FluoZin-3/Zn2+ complex of 8.9 nM (Krezel and Maret, 2006). Analysis of cellular viability. Cells were treated with mercuric chloride for 24 h under normal culture conditions and analyzed with the human Annexin V-FITC kit (Bender Med Systems, Vienna, Austria) according to the manufacturer's instructions, using a BD FACSCalibur flow cytometer. Viable cells were identified by being negative for both Annexin V and propidium iodide staining, apoptotic cells were positive for Annexin V but negative for counterstaining with propidium iodide to discriminate them from necrotic cells.

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tion at 412 nm. Concentrations were calculated using a calibration curve prepared from 0 to 100 μM cysteine. Western blotting. Sample preparation and analysis by Western Blot were performed as previously described (Haase et al., 2008). Statistical analysis. Statistical significance was calculated by oneway ANOVA followed by Dunnetts Post Hoc Test using GraphPad Prism. A p value ≤ 0.05 was considered statistically significant. All experiments presented have been performed at least three times independently with comparable results. Results

Measurement of total phosphatase activity and free thiols in cellular lysate. Cell lysate was prepared by sonication in reaction buffer (20 mM HEPES/NaOH, 20 mM MgCl2, pH 7.5) and incubated in 96 well plates as indicated in the figure legends. To measure phosphatase activity, para-nitrophenyl phosphate (pNPP) was added to a final concentration of 1 mM. After 1 h, the reaction was stopped by addition of an equal volume NaOH (1 M). Formation of p-nitrophenolate was quantified by its absorption at 405 nm. To measure reduced thiols, lysates were incubated with 5,5′-Dithiobis(2-nitrobenzoic acid) (Ellman's reagent) for 15 min, measuring the absorp-

Zn2+-release and induction of ROS by Hg2+ Zinc-release in response to addition of HgCl2 was compared in the monocyte, granulocyte, and lymphocyte subpopulations of primary human leukocytes (Fig. 1). Zinc was quantified by flow cytometry with the fluorescent probe FluoZin-3. This probe is insensitive toward mercury and several biologically relevant ions such as calcium and magnesium (Haase et al., 2009; Zhao et al., 2008; Zhao et al., 2009). Hence, the fluorescent signal results from intracellular free zinc, but

Fig. 3. Role of reactive oxygen species in Hg2+-mediated zinc release. (A, B, C) Cells were loaded with FluoZin-3 and the resulting fluorescence measured in 2-min intervals. After 10-min recording of the baseline, 5 or 25 μM HgCl2 were added (arrows) either to cells without antioxidants (A), or after pre-incubation with N-acetylcysteine (B) or ascorbic acid (C) for 30 min. Data are shown as means of triplicates ± SD from one out of n = 3 independent experiments. (D) Potential interference by N-acetylcysteine or ascorbic acid with the detection of zinc (500 nM) was measured with FluoZin-3 free acid in a cell free environment, using the same concentrations as above. Data are shown as means from n = 3 independent experiments ± SD.

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not mercury. Hg2+-concentrations of up to 5 μM led to a linear increase of fluorescence, whereas at 25 μM a biphasic effect with rapid initial release of zinc was observed (Fig. 1A). This effect was detected in all cell types, but was most pronounced in lymphocytes. In these cells, Hg2+-treatment led to the highest absolute intracellular free zinc concentration and the strongest increase compared to control levels (Fig. 1B, C). The largest fraction of lymphocytes are T-cells. Hence, the following experiments were performed in Jurkat cells, a human T-cell line. The redox-sensitive fluorescent dye DHR123 indicated a concentrationdependent production of ROS after addition of 5 or 25 μM HgCl2 (Fig. 2A), which was inhibited by the reducing agents N-acetylcysteine (N-AC) and ascorbic acid (AA) (Fig. 2B, C). Notably, the mercury concentrations of up to 25 μM that were used in this experiment were not cytotoxic and, as indicated by Annexin V staining, cell death did not involve apoptosis (Fig. 2D). Zinc itself is redox-inactive in biological systems, but many zincbinding sites in proteins contain cysteine thiols. Upon their oxidation, protein-bound zinc is released (Maret, 2006). Hence, elevated free zinc after Hg2+-treatment could not only result from direct replacement of Zn2+ by Hg2+, but also from thiol oxidation. To elucidate the role of ROS in Hg2+-mediated zinc release, Jurkat cells were loaded with FluoZin-3 and the effect of N-AC and AA was investigated (Fig. 3A–C). The elevation of free intracellular zinc was reduced by AA and completely abrogated by N-AC, pointing toward an oxidative mechanism rather than direct competition with Hg2+ for common binding sites. Although both antioxidants may bind divalent metal ions, it was

confirmed by in vitro measurements that they do not interfere with the detection of zinc by FluoZin-3 (Fig. 3D). Inhibition of phosphatases by Hg2+ and Zn2+ In order to investigate phosphatase activity, cleavage of the chromogenic phosphatase substrate pNPP was analyzed in Jurkat cell lysate. Hg2+ and Zn2+ inhibited phosphatase activity in a concentration dependent manner. Inhibition by zinc was reversed by the chelator N,N,N′,N′-tetrakis-2(pyridyl-methyl)ethylenediamine (TPEN) (Fig. 4A), but not by the reducing agent Tris(2-carboxyethyl) phosphine (TCEP), which does not act as a metal ion chelator (Fig. 4B). In contrast, inhibition by Hg2+ was not reversible by addition of TPEN (Fig. 4C). This observation excluded an involvement of released Zn2+, which would have been chelated by TPEN. Because TPEN binds Hg2+ with an even higher affinity than Zn2+ (Anderegg et al., 1977), it was also unlikely that Hg2+ inhibited phosphatases directly. Furthermore, no inhibition by Hg2+ was observed in the presence of TCEP, substantiating an oxidative mechanism of phosphatase inhibition (Fig. 4D). In the next experiment, it was tested if phosphatase inhibition was still reversible when TPEN and TCEP were added 30 min after the metal ions. The inhibition by Zn2+ was reversible by TPEN administration at any time point investigated (Fig. 5A, B). In contrast, inhibition by Hg2+ was only completely blocked when TCEP was present from the beginning (Fig. 5A), but was not reversible if TCEP was added after 30 min (Fig. 5B). This indicates the existence

Fig. 4. Roles of zinc and oxidation in phosphatase-inhibition by Hg2+. Total phosphatase activity in Jurkat lysate was measured by pNPP hydrolysis in the presence of ZnSO4 (A, B) or HgCl2 (C, D). Experiments were performed either in the presence of the chelator TPEN (50 μM; A, C) or the reducing agent TCEP (250 μM; B, D). All data are shown as means from n = 3 independent experiments ± SD.

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of at least two different oxidation states after treatment with mercuric ions; a first one that is reversible by TCEP, and is converted into a second, non-reversible state. Complete phosphatase inhibition was observed at relatively moderate Hg2+ concentrations, which were not cytotoxic. This was somewhat surprising, because extensive thiol oxidation should have severely affected cellular viability. Hence, we compared phosphatase activity in the lysate to the total amount of reduced thiols, measured with DTNB (Fig. 6). After Hg2+ treatment, a concentration-dependent decrease in reduced thiols and phosphatase activity was observed. Yet, phosphatases were affected to a significantly higher extent than thiol oxidation (Fig. 6A). Notably, a slight preference for phosphatase inactivation was also observed when H2O2 was used as an oxidizing agent, but the difference was not nearly as prominent as with Hg2+ (Fig. 6B).

Impact of Hg2+ on MAPK phosphorylation One function of phosphatases is the maintenance of signaling quiescence (Samet and Tal, 2010). Hence, phosphatase inhibition should lead to phosphorylation signals, even in the absence of physiological stimulation. Accordingly, there was a concentrationdependent phosphorylation of the MAPKs ERK1/2 and p38 upon treatment of cells with HgCl2 (Fig. 7A). The effect of Hg2+ was dependent on ROS, because it was absent in cells that had been pretreated with the antioxidant N-AC. In contrast, MAPK phosphor-

Fig. 5. Effect of time on the reversibility of phosphatase inhibition by Hg2+. Jurkat lysate was incubated with TPEN (250 μM) and TCEP (250 μM) either before (A) or 30 min after (B) addition of ZnSO4 (25 μM) or the indicated concentrations of HgCl2. After further 5 min, total phosphatase activity in the lysate was measured by addition of pNPP. Data are shown as means of n = 3 independent experiments ± SD.

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ylation in response to stimulation with the lectin phytohemagglutinin (PHA) remained unaffected by N-AC (Fig. 7B). Another difference was that MEK1/2, the upstream MAPKK of ERK1/2, was phosphorylated only after treatment with PHA, whereas activation of ERK1/2 by Hg2+ occurred without phosphorylation of MEK1/2 (Fig. 7B). Next, we compared the impact of Hg2+ and Zn2+ on the dephosphorylation of ERK1/2. Again, mercuric chloride had a concentration-dependent effect, inhibiting dephosphorylation of p38 and ERK1/2 (Fig. 7C). Although dephosphorylation was inhibited by Hg2+ and Zn2+, only the inhibition by Hg2+ was reversed in the presence of the reducing agents AA and N-AC (Fig. 7D).

Discussion Mercury affects multiple functions of the immune system, resulting either in autoimmunity, immunosuppression or inflammation (Vas and Monestier, 2008). T-cells are involved in all of these events, but so far, molecular explanations how mercury affects T-cell function remain to be found. The aim of this study was to investigate the mechanism by which mercuric ions interact with MAPK signal transduction in T-cells, and especially the impact on PTPs that dephosphorylate MAPKs. Measuring zinc-release in HgCl2-treated leukocytes loaded with a zinc-selective fluorescent probe demonstrated that incubation with

Fig. 6. Comparison of the effect of Hg2+ and H2O2 on phosphatase inhibition and thiol oxidation. Jurkat lysate was incubated for 30 min with the indicated concentrations of HgCl2 (A) or H2O2 (B), followed by measurement of free thiols with DTNB or phosphatase activity with pNPP. Measurements are shown as means of triplicates ± SD from one out of n = 3 independent experiments.

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mercuric chloride significantly elevates the intracellular free zinc concentration. It can be excluded that the FluoZin-3 signal was caused by Hg2+ and not Zn2+, because the probe is insensitive to mercury (Haase et al., 2006; Haase et al., 2009). Calcium release is frequently observed in cells treated with mercury (Rossi et al., 1991). Nevertheless, FluoZin-3 is also insensitive toward Ca2+ (Zhao et al., 2008). Actually, some signals of fluorescent probes for calcium in mercury-treated cells are in fact caused by zinc (Haase et al., 2009). The raise in free intracellular zinc was most pronounced in the lymphocyte fraction, which consists mainly of T-cells. Accordingly, if there is a role for zinc release in the impact of mercury on signal transduction, it should be strongest in lymphocytes. Zinc itself is not redox-active in biological systems, but its coordination sites in proteins frequently contain cysteine thiols. Upon oxidation of these thiols, zinc is displaced from its binding sites (Maret, 2006). Mercury-induced zinc release was blocked by N-acetylcysteine and ascorbic acid, indicating oxidative release of zinc. However, phosphatase inhibition by mercuric chloride was unaffected by TPEN, which binds zinc with high affinity (1015.58 M−1) (Anderegg et al., 1977) and abrogates inhibition of phosphatases by zinc. This excludes an involvement of zinc in phosphatase inhibition in response to treatment with mercuric chloride. Notably, TPEN binds mercuric ions with even higher affinity (1025.05 M−1) (Anderegg et al., 1977). Complete prevention of enzyme inactivation by the non metalbinding reducing agent TCEP, in combination with no reversal by high affinity chelator TPEN, clearly indicates that PTP are inhibited by a ROS-dependent mechanism, but not mercuric ion binding. Redox regulation is an important mechanism in signal transduction, because it allows transient inactivation of phosphatases (Lee et al., 1998). The active site of the PTP family, which includes dual specificity Tyr/Thr MAPK phosphatases, contains a characteristic (H) CX5R motif. The cysteine has an unusually low pKA, in order to form a thiolate that can perform a nucleophilic attack during catalysis. This makes the cysteine also highly susceptible to oxidation (Samet and Tal, 2010). Accordingly, treatment with H2O2 inhibited phosphatase

activity to a higher degree than it caused thiol oxidation. This effect was far more pronounced with mercuric chloride, indicating that phosphatases' active site cysteines are an exceedingly sensitive target for Hg2+. One explanation for the high preference for oxidative inhibition of active site cysteines compared to the total cellular thiol content could be that mercury might not only induce ROS-production, but also interfere with the mechanisms that recycle oxidized phosphatases. Thiols can be oxidized to several different species. These include disulfides and sulfenic, sulfinic, and sulfonic acid. Formation of disulfides or sulfenamide protects the oxidized catalytic cysteine from oxidation to the other oxides, as demonstrated for PTP 1B (Salmeen et al., 2003). In general, disulfides and sulfenic acid can be converted back into thiols by reducing agents or cellular redox mechanisms, whereas the other two are considered irreversible (Samet and Tal, 2010). Oxidation by low concentrations of H2O2 is reversible, whereas high concentrations lead to irreversible oxidation. In this case, the irreversibly oxidized state consisted mainly of sulfinic acid (Kamata et al., 2005). As seen in Fig. 5A, phosphatases remained active if TCEP was present from the beginning of mercuric chloride incubation. However, TCEP was not able to reverse inhibition if it was added later (Fig. 5B). This indicates a first, reversible oxidation, presumably to sulfenic acid. If this is not immediately reversed by TCEP, it is followed by a second, irreversible step to sulfinic and/or sulfonic acid. There are 107 members of the different PTP families in the human genome. Among them there exist more than a dozen dual specificity phosphatases that dephosphorylate MAPKs, and many of these phosphatases are expressed in cells of the immune system (Liu et al., 2007; Mustelin et al., 2005). As shown in Figs. 4 and 6, Hg2+ inhibits the entire pNPP-hydrolyzing activity in the lysate of Jurkat T-cells. This indicates that not only single phosphatases are inhibited, but at least a large proportion of the cellular capacity to cleave phosphate-ester bonds. After treatment of Jurkat cells with Hg2+, an increase of general tyrosine phosphorylation has been reported (Colombo et al., 2004), which could result from inhibition of PTPs. Also, it was shown that

Fig. 7. Effect of Hg2+ on MAPK phosphorylation. Phosphorylation of MAPKs was analyzed by Western blotting with phosphorylation state specific antibodies. (A) Jurkat cells were incubated with the indicated concentrations of HgCl2, and samples analyzed for phosphorylation of p38 (on Thr180/Tyr182) and ERK1/2 (on Thr202/Tyr204). (B) After preincubation with N-acetylcysteine for 30 min, cells were incubated with HgCl (25 μM) or PHA (10 μg/ml) for further 30 min and samples analyzed for phosphorylation of MEK (on Ser217/221), p38, and ERK1/2. (C, D) Jurkat cells were incubated with 10 μg/ml PHA for 25 min to induce ERK1/2 phosphorylation and then lysed by sonication. To allow dephosphorylation by intrinsic phosphatase activity, lysates were incubated for 30 min, supplemented either with the indicated concentrations of HgCl2 alone (C), or with HgCl2 or ZnSO4 (25 μM each) alone or in the presence of N-acetylcysteine (1 mM) or TCEP (250 μM) (D). All data shown are representative for n = 3 independent experiments.

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mercury activates p38 via ROS production (Kim et al., 2002, 2005; Kim and Sharma, 2004). These observations correspond well with a mechanism involving general inhibition of phosphatases by ROS after treatment with mercuric chloride. MAPKs are important regulators of the immune system and control activation and survival of T-cells (Dong et al., 2002). Mercury activates all major families of MAPK, including p38, JNK, and ERK1/2 (Barnes and Kircher, 2005; Kim et al., 2002, 2005; Kim and Sharma, 2004; Turney et al., 1999). For ERK1/2, the effect of mercury is twoedged; in addition to a slight activation of unstimulated T-cells, ERK1/2 phosphorylation triggered by the T-cell receptor is inhibited by Hg2+, because it blocks upstream signaling pathways (Mattingly et al., 2001; Ziemba et al., 2006). Our experiments indicate that activation of ERK1/2 by mercuric chloride is not mediated by the same mechanisms as conventional T-cell stimulation, e.g. activation by the T-cell mitogen PHA. In contrast to PHA, activation of ERK1/2 with HgCl2 occurred while the upstream kinase MEK1/2 remained unphosphorylated. Also, ERK1/2 phosphorylation was blocked by NAC, whereas PHA-mediated phosphorylation was insensitive to this reducing agent. This indicates that mercury preserves spontaneous phosphorylation by a redox-dependent mechanism, independent of upstream phosphorylation. The in vivo relevance of the mercury concentrations used in this study can be estimated by comparing them to human autopsy material (Bjorkman et al., 2007; Falnoga et al., 2000; Nylander et al., 1987; Nylander et al., 1989). Here, even the maximum total mercury concentration observed in blood (27.4 μg/L = 0.14 μM) was nearly one order of magnitude below the minimum concentrations required to see any effect in our phosphatase activity assays (Bjorkman et al., 2007). Furthermore, these results include organic and inorganic mercury, and a large proportion of inorganic mercury is probably bound to proteins, and not available to bind phosphatases. However, interaction between mercury and T-cells does not necessarily have to occur in peripheral blood. In some parts of the body, such as brain, thyroid gland, pituitary gland or renal cortex, accumulation occurs up to 0.5 μg/g wet tissue, which corresponds to micromolar concentrations (Bjorkman et al., 2007; Nylander et al., 1987, 1989). Furthermore, extreme occupational exposure can even lead to significantly higher concentrations of mercury in those organs (Falnoga et al., 2000). Hence, in mercury accumulating tissues, or following high level exposure, concentrations have been reported that could be sufficient to induce oxidative inhibition of MAPK dephosphorylation.

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