Abstract
Cd2+ is highly toxic to Staphylococcus aureus since it blocks dithiols in cytoplasmic 2-oxoglutarate dehydrogenase complex (ODHC) participating in energy conservation process. However, S. aureus 17810R is Cd2+-resistant due to possession of cadA-coded Cd2+ efflux system, recognized here as P-type Cd2+-ATPase. This Cd2+ pump utilizing cellular energy—ATP, ∆μ +H (electrochemical proton potential) and respiratory protons, extrudes Cd2+ from cytoplasm to protect dithiols in ODHC, but the mechanism of Cd2+ extrusion remains unknown. Here we propose that two Cd2+ taken up by strain 17810R via Mn2+ uniporter down membrane potential (∆ψ) generated during glutamate oxidation in 100 mM phosphate buffer (high PiB) are trapped probably by high affinity sites in cytoplasmic domain of Cd2+-ATPase, forming SCdS. This stops Cd2+ transport towards dithiols in ODHC, allowing undisturbed NADH production, its oxidation and energy conservation, while ATP could change orientation of SCdS towards facing transmembrane channel. Now, increased number of Pi-dependent protons pumped electrogenically via respiratory chain and countertransported through the channel down ∆ψ, extrude two trapped cytoplasmic Cd2+, which move to low affinity sites, being then extruded into extracellular space via ∆ψ-dependent Cd2+/H+ exchange. In 1 mM phosphate buffer (low PiB), external Cd2+ competing with decreased number of Pi-dependent protons, binds to ψs of Cd2+-ATPase channel, enters cytoplasm through the channel down ∆ψ via Cd2+/Cd2+ exchange and blocks dithiols in ODHC. However, Mg2+ pretreatment preventing external Cd2+ countertransport through the channel down ∆ψ, allowed undisturbed NADH production, its oxidation and extrusion of two cytoplasmic Cd2+ via Cd2+/H+ exchange, despite low PiB.
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Introduction
Cadmium is highly toxic to living organisms, since it blocks sulphhydryl groups in essential proteins (Vallee and Ulmer 1972; Moulis and Thevenod 2010). Some bacteria carry plasmid-linked cadA gene (Novick and Roth 1968; Dyke et al. 1970) conferring Cd2+ resistance expressed as decreased 109Cd uptake (Chopra 1975; Tynecka et al. 1975; Silver et al. 1982). Tynecka et al. (1981a, 1981b) were the first to report that the highly decreased 109Cd uptake by growing cells of Cd2+-resistant Staphylococcus aureus 17810R was due to pH gradient (∆pH)-dependent, nigericin-sensitive cadA-coded Cd2+ efflux system.
Subsequently, Silver and coworkers (Nucifora et al. 1989; Silver et al. 1989) showed that the cadA gene from staphylococcal plasmid pI258 coded the P-type Cd2+-ATPase, belonging to family of membrane-bound, cation-translocating pumps found in eukaryotes and prokaryotes. These pumps located across the membrane maintain homeostasis of essential cations (e.g. Mg2+, Ca2+, K+, Na+) or protons (Apell 2003; Kühlbrandt 2004; Pedersen 2007), and confer resistance to heavy metals (e.g. Cd2+, Zn2+, Cu2+) (Rosen 2002; Nies 2003; Kühlbrandt 2004; Silver and Phung 2005; Argüello et al. 2007, 2011). The best characterized is the P-type Ca2+-ATPase of sarcoplasmic reticulum (SR) for which detailed biochemical and biophysical data (Apell 2003; Toyoshima 2008) and about 50 crystal structures are available (Toyoshima et al. 2013). However, it is still controversial, how ATP energy is transduced to vectorial Ca2+ movement (Scarborough 2003; Toyoshima 2009).
According to sequencing data by Silver and coworkers (Nucifora et al. 1989; Silver et al. 1989), the four cysteine residues present in staphylococcal CadA protein are essential for Cd2+-ATPase activity: the conserved Cys23X2Cys26 in cytoplasmic domain—a possible high affinity Cd2+ binding site, and in conserved Cys371ProCys373 inside transmembrane channel, involved probably in Cd2+ extrusion. The CysX2Cys motif is related to copper-binding region in Cu2+-ATPases (Fan and Rosen 2002) and to mercury-binding region in proteins involved in Hg2+ resistance (Barkay et al. 2003). According to Tsai et al. (1992), staphylococcal P-type Cd2+-ATPase requires only ATP. Here is shown, that the cadA-coded Cd2+ efflux system in Cd2+-resistant S. aureus 17810R (Tynecka et al. Tynecka et al. 1981a, 1981b; Tynecka and Szcześniak 1991) is a P-type Cd2+-ATPase requiring: ATP, electrochemical proton potential (∆μ +H ), high phosphate buffer (PiB) and Pi-dependent protons or Mg2+. The mechanism of Cd2+ extrusion by this staphylococcal Cd2+-ATPase is proposed.
Materials and methods
Bacterial strains and culture conditions
Cd2+-resistant S. aureus 17810R, carrying cadA gene on penicillinase plasmid pII17810 (Shalita et al. 1980), was described previously (Tynecka et al. 1981a, 1981b). Experiments were performed at 37 °C using early exponential phase cells grown aerobically in 3 % nutrient broth and suspended in 100 mM potassium/sodium phosphate buffer, pH 7 (PiB). Cell suspensions were vigorously aerated for 3 h at 37 °C without exogenous electron donor to deprive cells of endogenous energy reserves (Tynecka and Malm 1995; Tynecka et al. 2001). Next, cells were suspended in PiB of various concentrations, depending on the experiment, at a density of 0.2 mg dry weight/ml and preincubated with 10 mM glutamate for 10 min at 37 °C (glutamate oxidizing cells). In some experiments, cells were suspended in other buffers: 100 mM triethanolamine/phosphate, pH 7, 100 mM Tris/HCl, pH 7.2 or 100 mM MOPS/NaOH, pH 7. Cd2+-sensitive variant strain S. aureus 17810S lacking cadA gene, also described previously (Tynecka et al. 1981a, 1981b), was used in some experiments as a control organism.
Reagents
Inhibitors: 2-heptyl-4-hydroxyquinoline N-oxide (HQNO) and dicyclohexylcarbodiimide (DCCD), and ionophores: valinomycin, nigericin or carbonyl cyanide m-chlorophenyl hydrazone (CCCP) were from Sigma (St. Louis, MO). The following radiolabeled compounds were used: 109Cd (carrier-free) or sodium [U-14C]glutamate (7.4 GBq/mmol)—from Amersham, UK, 86RbCl (1.075 GBq/mmol), sodium [14C]benzoate (407 MBq/mmol), [3H]inulin (3.7 GBq/mmol) or [γ-32P]ATP (111 TBq/mmol)—from NEN™ Life Science Products (Boston, MA), while 32Pi—inorganic orthophosphate (740 MBq/mmol)—from the Institute of Nuclear Research, Świerk, Poland.
Uptake experiments
Uptake of 109Cd at 10 μM (as CdCl2) by glutamate oxidizing cells of strain 17810R and strain 17810S was assayed by filtration procedure, as described previously (Tynecka et al. 1981a, 1981b). These cells suspended in 100 or 1 mM PiB were preincubated at 37 °C for 10 min, with appropriate compounds: MgCl2, MnCl2 or ionophores—nigericin, valinomycin + KCl or CCCP, depending on the experiment, before addition of 10 μM 109CdCl2. In order to determine K m and V max of 109Cd uptake in strain 17810R, the initial influx rate of 109Cd uptake in 1 mM PiB within 1 min at various CdCl2 concentrations was measured.
Uptake of 32Pi (inorganic orthophosphate) by glutamate oxidizing cells of strain 17810R suspended in 100 or 1 mM PiB was assayed by filtration procedure, as described previously (Tynecka and Szcześniak 1991).
Assay of 109 Cd efflux
109Cd efflux was assayed by filtration procedure, as described previously (Tynecka et al. 1981b). 109Cd efflux from washed, glutamate oxidizing cells of strain 17810R was performed after cell preincubation for 20 min with 10 μM 109CdCl2 in 1 mM PiB. After removal of external 109CdCl2 by cell washing at 4 °C, cells were resuspended in 100 mM PiB or in other buffers: 100 mM triethanolamine/phosphate, pH 7, 100 mM Tris/HCl, pH 7.2 or 100 mM MOPS/NaOH, pH 7. DCCD and ionophores: CCCP, valinomycin + KCl or nigericin were added, depending on the experiment. In each experiment 10 mM glutamate was added and all suspensions were prewarmed to 37 °C, before 109Cd efflux was measured. 109Cd efflux from unwashed, glutamate oxidizing cells was also performed after cell preincubation for 20 min with 10 μM 109CdCl2 in 1 mM PiB. Then, PiB concentration was increased at steady-state from 1 mM to 100 mM without cell washing, before 109Cd efflux was measured. Appropriate compounds: MgCl2 or ionophores: CCCP, valinomycin + KCl or nigericin were added at steady-state, depending on the experiment.
Assay of 109Cd content in subcellular fractions
109Cd distribution among subcellular fractions obtained from glutamate oxidizing cells of strain 17810R and strain 17810S preloaded with 10 μM 109CdCl2 in 100 or 1 mM PiB and preincubated at 37 °C with appropriate compounds: MgCl2 or ionophores—CCCP, valinomycin + KCl or nigericin, depending on the experiment, was determined according to Tynecka et al. (2001).
Assay of enzyme activity
Activity of 2-oxoglutarate dehydrogenase complex (ODHC) was measured in the cytoplasmic fraction obtained from glutamate oxidizing cells of strain 17810R suspended in 100 or 1 mM PiB and preincubated at 37 °C with CdCl2, according to the method described previously (Tynecka and Malm 1996).
Assay of membrane potential (∆ψ) and pH gradient (∆pH)
The values of ∆ψ and ∆pH in glutamate oxidizing cells of strain 17810R suspended in 100 or 1 mM PiB were determined by a filtration procedure from the steady-state distribution of 100 μM 86Rb in the presence of 10 μM valinomycin or of 20 μM sodium [14C]benzoate, respectively, as described previously (Tynecka et al. 1999). [3H]inulin served as a marker for extracellular water.
Phosphorylation assay
Membrane fraction of strain 17810R was obtained according to the procedure described previously (Tynecka and Malm 1996; Tynecka et al. 2001). Phosphorylation assay was performed as described elsewhere (Tsai and Lynn Linet 1993) with some modifications. To 200 μl of membrane fraction (2.4 mg protein/ml), 2 μl of 1.2 mM EDTA were added, followed by incubation for 10 min, and then 2 μl of 50 μM CdCl2 or equivalent volume of deionized water, followed by incubation for 5 min. The reaction was started by addition of 10 μCi of [γ-32P]ATP and 2 μl of 0.8 M MgCl2. The reaction mixture was incubated at room temperature, then the reaction was stopped after 60 s by addition of equivalent volume of ice-cold 20 % TCA. After 10 min incubation on ice, the membranes were collected by centrifugation (14,000 rpm, 5 min). In order to assay the effect of alkali or hydroxylamine, the pellets were incubated with 100 μl of 0.5 M KOH for 5 min on ice or with 200 μl of 0.1 M sodium acetate containing 260 mM hydroxylamine for 10 min at room temperature. After incubation, equivalent volume of ice-cold 10 % TCA was added. In each case, the collected pellets were washed with water and then twice with 50 mM H3PO4/NaOH, pH 2.4. Then, the pellets were dissolved in 10 % SDS at 100 °C and suspended in a standard sample buffer used for acidic SDS-PAGE, as described elsewhere (Fairbanks and Avruch 1972). Gels were run at 40 mA for 4–5 h at room temperature. After electrophoresis, autoradiography of the dried gels was performed at 4 °C for 48 h.
Reproducibility of results
The experimental data shown in each figure are the mean ± SD from at least three independent experiments.
Results
Highly decreased 109Cd accumulation in Cd2+-resistant S. aureus 17810R oxidizing glutamate in 100 mM phosphate buffer, pH 7 (high PiB).
First, membrane proteins of S. aureus 17810R harbouring cadA gene were phosphorylated by [γ-32P]ATP (Fig. 1). The protein band of about 100 kDa was strongly phosphorylated, when Cd2+ was present. Intensity of this band was decreased by alkali or hydroxylamine, which is typical for phosphoenzyme intermediate of P-type ATPases (Tsai and Lynn Linet 1993). This suggests that the band strongly phosphorylated in strain 17810R in the presence of Cd2+ (Fig. 1) may correspond to CadA protein, having also molecular weight of about 80 kDa (Nucifora et al. 1989; Tsai and Lynn Linet 1993).
Cd2+-resistant S. aureus 17810R took up only 0.5 ± 0.15 nmol 109Cd/mg dry wt (Fig. 2a) and accumulated in cytoplasm merely 0.37 ± 0.1 nmol 109Cd/mg protein (Fig. 2b). Under similar conditions, the Cd2+-sensitive variant strain S. aureus 17810S lacking cadA gene, took up 20 ± 1.2 nmol 109Cd/mg dry wt (Fig. 2a) and accumulated in cytoplasm 21 ± 1.5 nmol 109Cd/mg protein (Fig. 2b) down ∆ψ (membrane potential) via high affinity Mn2+ uniporter sensitive to Mn2+ or valinomycin + K+ (Fig. 2a). As was already reported (Tynecka et al. 1981a, 1981b; Tynecka and Malm 1995, 1996: Tynecka et al. 1989), two Cd2+ accumulated by strain 17810S in a transport cycle, blocked vicinal dithiols in dihydrolipoate and dihydrolipoate dehydrogenase in the cytoplasmic 2-oxoglutarate dehydrogenase complex (ODHC) in Krebs cycle located in the first energy coupling site of respiratory chain (Tynecka et al. 1999). These dithiols are the only Cd2+-sensitive targets in glutamate-linked energy conservation system in strain 17810S; their blocking stopped endogenous NADH production, its oxidation via respiratory chain, generation of electrochemical proton potential (∆μ +H ) and consequently ∆μ +H -dependent processes without direct blocking of solute transporters and ATP synthase (Tynecka and Malm 1995, 1996; Tynecka et al. 1989, 2001).
The Cd2+-resistant strain 17810R did not accumulate Cd2+ (Fig. 2a, b), although cells of strain 17810R and 17810S oxidizing glutamate generated ∆μ +H of similar value expressed as protonmotive force (∆p) of about −191 ± 5 mV. Data in Fig. 2a, b suggest that two Cd2+ transported by strain 17810R via Mn2+ uniporter down ∆ψ of −161 ± 5 mV were extruded by Cd2+ efflux system described by Tynecka et al. 1981a, 1981b, which was recognized here as a P-type Cd2+-ATPase. Cd2+ extrusion by this Cd2+ pump via Cd2+/H+ exchange before reaching Cd2+-sensitive targets—dithiols in ODHC, allowed undisturbed NADH production (5.4 ± 0.6 nmol NADH/min/mg protein), and consequently its oxidation via respiratory chain, ∆μ +H generation and energy conservation (data not shown), rendering host cells Cd2+-resistant.
Since nigericin, collapsing ∆pH but stimulating ∆ψ, did not increase Cd2+ uptake by strain 17810R (Fig. 2a), this suggested that Cd2+ extrusion by Cd2+-ATPase from glutamate oxidizing cells was not energized by ΔpH. According to chemiosmotic principles (Mitchell 1966), the enhanced transport of inorganic phosphate (Pi) by strain 17810R via H+/32Pi symport consuming ΔpH (Tynecka and Szcześniak 1991), could stimulate generation of membrane potential (∆ψ). It is probable that according to Rosenberg and Friedberg (1984) the H+/Pi cotransport by strain 17810R in high PiB could result in phosphate polymerization and accumulation of additional protons in cytoplasm. We suggest that these Pi-dependent protons pumped electrogenically via respiratory chain could return through the transmembrane channel of Cd2+-ATPase down ∆ψ and extruded two cytoplasmic Cd2+ into extracellular space via ∆ψ-dependent Cd2+/H+ exchange. This was confirmed by dependence of Cd2+ extrusion on high PiB, since below 25 mM PiB, linear Cd2+ uptake by strain 17810R insensitive to Mn2+ was observed (data not shown). To explain the mechanism of Cd2+ extrusion by P-type Cd2+-ATPase in high Pi, 109Cd uptake by strain 17810R was first characterized in 1 mM PiB and then requirements for its net extrusion were studied.
Uptake of 109Cd by Cd2+-resistant S. aureus 17810R oxidizing glutamate in 1 mM phosphate buffer, pH 7 (low PiB).
The markedly decreased 32Pi uptake by strain 17810R from 350 nmol 32Pi/mg dry wt/20 min in high PiB to 150 nmol 32Pi/mg dry wt/20 min in low PiB could result in decreased number of Pi-dependent protons pumped electrogenically via respiratory chain. Under these conditions, strain 17810R took up 10 ± 1.3 nmol 109Cd/mg dry wt, insensitive to Mn2+ (Fig. 2c); about 8 ± 0.9 nmol Cd2+/mg protein were found in cell wall and membrane and similar amount of 109Cd—in cytoplasm (Fig. 2d), which was only about half of that accumulated by strain 17810S in high PiB (Fig. 2b). We suggest that in low PiB the external Cd2+ could compete with decreased number of Pi-dependent protons for entry into cytoplasm down ∆ψ through transmembrane channel of Cd2+-ATPase. Therefore, the first cytoplasmic Cd2+ could be extruded from strain 17810R via exchange with external Cd2+ via Cd2+/Cd2+ exchange, while the second cytoplasmic Cd2+ was absent, suggesting its net extrusion. External 109Cd uptake in low PiB showed linear dependence on Cd2+ concentration (data not shown) and high K m = 112 ± 2.3 μM and V max = 9.1 ± 1.2 nmol Cd2+/mg dry wt/min, suggesting that Cd2+-ATPase channel may function now as low affinity second pathway transporting external Cd2+ down ∆ψ instead of protons towards Cd2+–sensitive targets—dithiols in ODHC.
External Cd2+ accumulated by strain 17810R in low PiB blocked dithiols in ODHC, which stopped NADH production (from 5.4 ± 0.6 to 0.2 ± 0.1 nmol NADH/min/mg protein) and consequently its oxidation via respiratory chain, but ∆μ +H generation was unaffected (Δp = −210 ± 4 mV). This suggests that according to Mitchell (1966), Cd2+ toxicity to cell respiration could result in conversion of the reversible FoF1-ATP synthase into hydrolytic direction, which working now as Cd2+-insensitive, anaerobic proton pump—FoF1-ATPase (Tynecka et al. 1990), could continue ∆μ +H generation. We suggest that Δψ of −195 ± 4 mV could energize transport of the second cytoplasmic Cd2+ via Mn2+ uniporter, while ΔpH of 15 ± 2 mV could support its extrusion via Cd2+/H+ exchange, as confirmed by absence of the second Cd2+ in cytoplasm (Fig. 2c, d). Thus, Cd2+-ATPase extruded in low PiB also two cytoplasmic Cd2+, but only external Cd2+ reached dithiols in ODHC through the channel via Cd2+/Cd2+ exchange, disturbing energy conservation and Cd2+ resistance of strain 17810R.
We also considered in strain 17810R a controversial problem– existence of low affinity sites on external surface of P-type ATPases (McIntosh 2000; Apell 2003; Scarborough 2003; Toyoshima 2009). First, Silver and coworkers (Nucifora et al. 1989, Silver 1996) recognized during sequencing studies some negatively charged amino acid residues (Glu, Asp) on extracellular surface of CadA protein. It is known (Williams 1978; Barber 1980) that such residues create at physiological pH the surface potentials (ψs) on biological membranes, protected by cations of various protective abilities (Mg2+ > Ca2+ > K+ > Na+), depending on their concentration and/or affinity. We suggest that Cd2+-ATPase channel in strain 17810R may also possess two negatively charged residues forming surface potential (ψs) functioning as low affinity sites to which protons or external Cd2+ may bind before entering the channel, but this depends on PiB concentration.
According to Fig. 2c, 1 mM Mg2+ prevented external Cd2+ uptake by strain 17810R in 50 %, while 5 mM Mg2+ (further called Mg2+) stopped it. These data confirm existence of low affinity ψs sites on extracellular surface of Cd2+-ATPase channel in strain 17810R. Protection of ψs by Mg2+ against external Cd2+ binding and its countertransport through the channel towards dithiols in ODHC allowed undisturbed energy conservation and Cd2+ resistance. In contrast, Cd2+ uptake by strain 17810S was Mg2+-insensitive (data not shown), suggesting ψs absence in Mn2+ uniporter.
The ionophore studies in low PiB showed that nigericin, collapsing ΔpH, doubled Cd2+ uptake by strain 17810R (Fig. 3a). Probably, by stopping ΔpH-dependent efflux of the second cytoplasmic Cd2+ energized by the reversed Cd2+-insensitive FoF1-ATPase, nigericin could unmask Δψ-dependent Cd2+ transport via Mn2+ uniporter, sensitive to Mn2+ (Fig. 3a). Now, strain 17810R accumulating in cytoplasm two Cd2+ down Δψ—via transmembrane channel (Cd2+/Cd2+ exchange) and via Mn2+ uniporter (Fig. 3a, b), became Cd2+-sensitive, like strain 17810S (Fig. 2a, b). However, Mg2+ pretreatment of strain 17810R before nigericin addition, prevented external Cd2+ binding to ψs of Cd2+-ATPase and also stopped Cd2+ countertransport through the channel, rendering host cells Cd2+-resistant, despite low PiB (Fig. 3a).
Valinomycin + K+ collapsing Δψ, did not affect external Cd2+ uptake by strain 17810R (Fig. 3a), although Δψ-dependent Cd2+ transport via Mn2+ uniporter into cytoplasm of strain 17810S was stopped by this ionophore (Fig. 2a). Therefore, valinomycin-insensitive Cd2+ uptake by strain 17810R may represent only Δψ-independent external Cd2+ binding to cell wall and only to one ψs site of Cd2+-ATPase, prevented by Mg2+ (Fig. 3a, c), while external Cd2+ binding to the second ψs site was probably prevented by protons countertransported down unaffected ΔpH.
CCCP also doubled 109Cd uptake by strain 17810R (Fig. 3a), although Cd2+ accumulation in cytoplasm was stopped (Fig. 3d), since Δψ for Cd2+ transport through the channel was blocked by CCCP. This means that two external Cd2+ could bind without energy to two ψs sites in Cd2+-ATPase channel (Fig. 3d), prevented by Mg2+ (Fig. 3a). These CCCP data strongly confirm existence of two low affinity ψs sites in Cd2+-ATPase channel of strain 17810R.
Restoration of Cd2+ resistance in S. aureus 17810R by extrusion of Cd2+ preaccumulated in 1 mM phosphate buffer, pH 7 (low PiB).
Cd2+-preloaded cells of strain 17810R in low Pi were washed and resuspended in high PiB. Since in these Cd2+-poisoned cells of strain 17810R the NADH production was blocked and consequently its oxidation, therefore ∆μ +H generation and Cd2+ extrusion via Cd2+/H+ exchange were also stopped. Therefore, under such conditions only the Cd2+-insensitive proton pump—the reversed FoF1-ATPase could provide protons for Cd2+-ATPase to start Cd2+ extrusion from dithiols in ODHC. We suggest that these protons could bind easily to ψs of Cd2+-ATPase channel in washed cells, since there was no extracellular Cd2+ to compete. Finally, protons countertransported through the channel down Δψ displaced Cd2+ from dithiols in ODHC, which was evidenced by undisturbed ODHC activity (5.6 ± 0.8 nmol NADH/min/mg protein).
Cd2+ extrusion was inhibited in 50 % by DCCD, blocking H+ channel of FoF1-ATPase and also by CCCP or valinomycin + K+ collapsing Δψ (Fig. 4b). This suggests that Cd2+ could be removed from cell wall and ψs of strain 17810R without energy, but Cd2+ extrusion from dithiols in ODHC requiring H+ and Δψ, was stopped as evidenced by inhibited ODHC activity with all three compounds (from 1.4 ± 0.2 to 1.5 ± 0.3 nmol NADH/min/mg protein). Only nigericin collapsing ΔpH, allowed Cd2+ extrusion down undisturbed Δψ (Fig. 4b). Since CCCP or valinomycin + K+ prevented Δψ-dependent proton countertransport through the channel and also stopped Cd2+ extrusion, this suggests that dithiols in ODHC (S–S−) may function in glutamate-linked energy conservation process probably as Δψ generation site.
According to Fig. 4c, other 100 mM buffers containing glutamate-Tris/HCl, pH 7.2 or MOPS/NaOH, pH 7.0, did not initiate Cd2+ extrusion from washed cells of strain 17810R. Only 100 mM triethanolamine/phosphate buffer pH 7, triggered total Cd2+ efflux (Fig. 4c) sensitive to CCCP or valinomycin + K+ in 50 %, but insensitive to nigericin (Fig. 4d). These data strongly confirm requirement of high Pi and of Pi-dependent protons for net Cd2+ extrusion by Cd2+-ATPase.
Cd2+ efflux triggered by 100 mM PiB from unwashed cells of strain 17810R was incomplete (Fig. 5a), since only Cd2+ from cell wall and ψs could be released without energy, but not Cd2+ from cytoplasm requiring protons and Δψ, as evidenced by blocked ODHC activity (1.4 ± 0.3 nmol NADH/min/mg protein). Only high PiB plus Mg2+—preventing external Cd2+ countertransport down Δψ through the channel, allowed Cd2+ extrusion from dithiols in ODHC via Cd2+/H+ exchange (Fig. 5a), as evidenced by unblocked ODHC activity (5.5 ± 0.8 nmol NADH/min/mg protein). This Cd2+ efflux was equally affected by ionophores (Fig. 5b), as that from washed cells (Fig. 4b).
Discussion
Bacterial Cd2+-ATPases belong to the superfamily of P-type ATPases and to the P1 subfamily of soft metal ions pumps (Kühlbrandt 2004). Studies on Cd2+-ATPase in S. aureus (Nucifora et al. 1989; Silver et al. 1989; Tsai et al. 2002) and in Listeria monocytogenes (Bal et al. 2003; Wu et al. 2006a) established amino acid sequence of CadA protein, its membrane topology and suggested involvement in Cd2+ extrusion of four cysteine residues present in this protein, but the mechanism of how Cd2+ is extruded from Cd2+-resistant S. aureus remains so far unknown. Also studies on Cd2+-ATPase in other microorganisms did not explain this mechanism (Schwager et al. 2012; Schurig-Briccio and Gennis 2012; Chien et al. 2013; Maynaud et al. 2014). As found here, the cadA-coded Cd2+ efflux system in S. aureus 17810R described by Tynecka et al. (1981a, 1981b), appeared to be the P-type Cd2+-ATPase. Figure 6 presents a proposed scheme for Cd2+ extrusion via Cd2+/H+ exchange mechanism by the native Cd2+-ATPase in S. aureus 17810R oxidizing glutamate in high PiB.
We propose that two Cd2+transported by strain 17810R down Δψ via Mn2+ uniporter in high PiB are trapped by high affinity sites—dithiols in cytoplasmic domain of staphylococcal Cd2+-ATPase, which were recognized by Silver and coworkers (Nucifora et al. 1989; Silver et al. 1989). We suggest that Cd2+ trapping (SCdS) stops its transport towards dithiols in cytoplasmic ODHC, allowing undisturbed NADH production, its oxidation via respiratory chain and ∆μ +H generation, while ATP could change SCdS orientation from facing cytoplasm towards facing transmembrane channel, probably by a tilting mechanism suggested for Ca2+-ATPase (Albers 1967; Post et al. 1972; Higgins and Linton 2001). Finally, increased number of Pi-dependent protons pumped electrogenically via respiratory chain, could compete with external Cd2+ for binding to ψs of Cd2+-ATPase. Then, entering the channel, protons displaced from high affinity sites the two trapped cytoplasmic Cd2+, which were transferred through the channel towards low affinity sites ψs, being then extruded into extracellular space via Cd2+/H+ exchange against electrochemical and concentration gradients (Tynecka et al. 1981a, 1981b), rendering host cells Cd2+-resistant. In Listeria monocytogenes (Wu et al. 2006b) the cadA-coded Cd2+-ATPase also extruded two Cd2+.
According to the proposed concept (Fig. 6), the Pi-dependent protons and Cd2+ binding ligands in Cd2+-ATPase channel of strain 17810R seem to play vital role in Cd2+ extrusion. We suggest that the negative charges exposed in the channel by successive proton movement to ψs, CysProCys and CysCys (recognized by Nucifora et al. 1989; Silver et al. 1989; Tsai et al. 2002) can be the driving force for uphill Cd2+ pumping through the channel into opposite direction–the extracellular environment. Also Ca2+ efflux by Ca2+-ATPase from mammalian mitochondria was Pi-dependent (Roos et al. 1980; Nicholls and Akerman 1982; Ligeti and Lukács 1984). Proton requirement for Cd2+ extrusion in strain 17810R, is in accord with Scarbourough`s considerations (Scarborough 2003)—that after phosphorylation reaction, something must weaken ion binding site, allowing Ca2+ release into extracellular space. However, the role of protons in SR Ca2+-ATPase has been controversial for many years (Ueno and Sekine 1981; Levy et al. 1990; Andersen and Vilsen 1995; Karjalainen et al. 2007; Toyoshima 2009; Fibich and Apell 2011; Bublitz et al. 2013).
Our earlier observations (Tynecka et al. 1981a, 1981b) suggested that protons and external Cd2+ could compete for entry into cytoplasm through the channel of Cd2+-ATPase down Δψ. Our present data confirm that due to decreased number of Pi-dependent protons in low PiB the external Cd2+ could bind to ψs of Cd2+-ATPase channel and then driven down Δψ through the channel via Cd2+/Cd2+ exchange, blocks dithiols in ODHC, rendering host cells Cd2+-sensitive, like strain 17810S.
However, we found that Mg2+ can protect strain 17810R against Cd2+ poisoning in low PiB. According to William’s model (Williams 1978) and our CCCP data, Mg2+ can prevent external Cd2+ binding to ψs and stops its countertransport through the channel towards dithiols in ODHC. In energized cells, Mg2+ can be displaced transiently by respiratory protons, but still prevents external Cd2+ countertransport. Therefore, even a decreased number of Pi-dependent protons pumped electrogenically during glutamate oxidation in low PiB, but protected by Mg2+, could enter the channel to extrude two trapped cytoplasmic Cd2+ via energy-dependent Cd2+/H+ exchange. Discharge of ∆μ +H by protons allowed Mg2+ return to ψs. Such Mg2+ oscillation can maintain undisturbed NADH production, its oxidation, energy conservation and Cd2+ resistance, despite low PiB. Also Ca2+ influx and Δψ disruption in mitochondria were prevented by Mg2+ (Sharikabad et al. 2001; Racay 2008).
Net Cd2+ extrusion requires also steady-state thermodynamic equilibrium between activities of two energy-dependent membrane systems—Mn2+ uniporter and Cd2+-ATPase. Some changes, e.g. increased Cd2+ concentration (Tynecka et al. 1981b), alkaline pH (Tynecka et al. 1981a) or decreased PiB concentration shown here, disturb equilibrium and consequently Cd2+ resistance. Therefore, the Cd2+-ATPase cooperating with Pi-dependent protons and utilizing cellular energy (ATP and ∆μ +H ) can protect against Cd2+ poisoning the vital dithiols in ODHC.
However, we found that Cd2+, which blocked dithiols in ODHC in low PiB could be also extruded. As was already mentioned, in these Cd2+-poisoned cells of strain 17810R, only the Cd2+-insensitive, reversed FoF1-ATPase could pump protons to start the Cd2+ efflux process. Besides, we increased the PiB concentration to 100 mM (high PiB) and also inhibited external Cd2+ countertransport down Δψ through the channel either by cell washing or by Mg2+ pretreatment. In both situations, the countertransport of Pi-dependent protons through the channel was restored, leading to Cd2+ displacement from dithiols in ODHC. We suggest that the displaced Cd2+ could be trapped by high affinity sites of Cd2+-ATPase, forming SCdS, while ATP could change SCdS orientation towards facing transmembrane channel. Now, Cd2+ displaced by protons from high affinity sites via Cd2+/H+ exchange moves towards low affinity sites (ψs) of Cd2+-ATPase. From here, Cd2+ is displaced into extracellular space also via Cd2+/H+ exchange. Gradual Cd2+ extrusion by Cd2+-ATPase restored gradually: NADH production, its oxidation, ∆μ +H generation via respiratory chain, reversal of FoF1-ATPase into biosynthetic direction and energy conservation, rendering host cells again Cd2+-resistant. DCCD—blocking H+ channel of FoF1-ATPase or CCCP and valinomycin + K+ collapsing Δψ, prevented Cd2+ extrusion, confirming the requirement of Pi-dependent protons and of Δψ for the Cd2+ efflux process.
To summarize, these studies provide for the first time the novel data on the so far unknown mechanism of Cd2+ extrusion by cadA-coded P-type Cd2+-ATPase in S. aureus 17810R, oxidizing glutamate in high PiB. Energy-dependent Cd2+ extrusion by this pump via Cd2+/H+ exchange mechanism renders host cells Cd2+-resistant, since the toxic Cd2+ could not reach the primary Cd2+-sensitive targets—dithiols in ODHC via two routes—Mn2+ uniporter or transmembrane channel, allowing undisturbed glutamate-linked energy conservation process. Moreover, the vital role of Pi-dependent protons or Mg2+ and of cellular energy (ATP and ∆μ +H ) in Cd2+ extrusion by Cd2+-ATPase is underlined.
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Acknowledgments
Zofia Tynecka would like to thank Prof. Keith Dyke, Wadham College, University of Oxford, for the generous gift of Cd2+-resistant S. aureus 17810R and its Cd2+-sensitive variant strain S. aureus 17810S. These studies were supported by a grant 6 P04C 020 13 from the State Committee for Scientific Research, Warsaw, Poland. We dedicate this work to the memory of our friend and collegue Zofia Goś-Szcześniak.
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Tynecka, Z., Malm, A. & Goś-Szcześniak, Z. Cd2+ extrusion by P-type Cd2+-ATPase of Staphylococcus aureus 17810R via energy-dependent Cd2+/H+ exchange mechanism. Biometals 29, 651–663 (2016). https://doi.org/10.1007/s10534-016-9941-5
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DOI: https://doi.org/10.1007/s10534-016-9941-5