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¹ Institute of Cardiovascular Sciences, University of Manitoba, St Boniface General Hospital Research Centre, Winnipeg, Manitoba, Canada, R2H 2A6
² Department of Pharmacology & Physiology, University of Medicine and Dentistry of New Jersey, 185 South Orange Avenue, Newark, NJ 07103, USA

	Abstract

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Chinese hamster ovary cells expressing the bovine cardiac Na⁺–Ca²⁺ exchanger (NCX1.1) accumulated Cd²⁺ after a lag period of several tens of seconds. The lag period reflects the progressive allosteric activation of exchange activity by Cd²⁺ as it accumulates within the cytosol. The lag period was greatly reduced in cells expressing a mutant exchanger, (241-680), that does not require allosteric activation by Ca²⁺ for activity. Non-transfected cells did not show Cd²⁺ uptake under the same conditions. In cells expressing NCX1.1, the lag period was nearly abolished following an elevation of the cytosolic Ca²⁺ concentration. Cytosolic Cd²⁺ concentrations estimated at 0.5–2 pM markedly stimulated the subsequent uptake of Ca²⁺ by Na⁺–Ca²⁺ exchange. Outward exchange currents in membrane patches from Xenopus oocytes expressing the canine NCX1.1 were rapidly and reversibly stimulated by 3 pM Cd²⁺ applied at the cytosolic membrane surface. Exchange currents activated by 3 pM Cd²⁺ were 40% smaller than currents activated by 1 µM cytosolic Ca²⁺. Current amplitudes declined by 30% and the rate of current development fell sharply upon repetitive applications of Na⁺ in the presence of 3 pM Cd²⁺. Cd²⁺ mimicked the anomalous inhibitory effects of Ca²⁺ on outward exchange currents generated by the Drosophila exchanger CALX1.1. We conclude that the regulatory sites responsible for allosteric Ca²⁺ activation bind Cd²⁺ with high affinity and that Cd²⁺ mimics the regulatory effects of Ca²⁺ at concentrations 5 orders of magnitude lower than Ca²⁺.

(Received 19 October 2004; accepted after revision 16 December 2004; first published online 20 December 2004)
Corresponding author J. P. Reeves: Department of Pharmacology & Physiology, University of Medicine and Dentistry of New Jersey, 185 South Orange Avenue, Newark, NJ 07103, USA. Email: reeves{at}umdnj.edu

	Introduction

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Cd²⁺ toxicity is associated with numerous pathogenic conditions, including renal disease, cancer (lungs, prostate and kidney) and cardiovascular disorders, particularly peripheral arterial disease (reviewed in Jarup, 2003; Waalkes, 2003; Waisberg et al. 2003; Navas-Acien et al. 2004). Sources of Cd²⁺ exposure in humans include diet, smoking and industrial pollution (Jarup, 2003). At a cellular level, Cd²⁺ induces the production of reactive oxygen species (ROS), probably through interaction with mitochondria (Wang et al. 2004; Thevenod, 2003; Pourahmad et al. 2003). Cd²⁺-induced ROS production has been implicated in the pathogenesis of each of the conditions listed above (Nath et al. 2000). A major cellular mechanism for protection against Cd²⁺ toxicity is the production of metallothioneins, small (6–10 kDa) proteins containing 30% cysteine residues which bind heavy metal ions such as Zn²⁺ and Cd²⁺ with K_D values of 10^–18 and 10^–22 M, respectively (Kagi & Vallee, 1961; Klaassen et al. 1999). Many cells increase metallothionein production in response to heavy metal ions or oxidative stress (Klaassen et al. 1999). Cellular sensitivity to Cd²⁺ toxicity is inversely correlated with metallothionein expression levels (Klaassen et al. 1999) and metallothionein knockout mice are susceptible to Cd²⁺-induced renal dysfunction at lower doses than wild-type animals (Klaassen & Liu, 1998). The high sensitivity of cardiac myocytes to Cd²⁺ toxicity, e.g. compared with liver, may reflect lower levels, and/or poor induceability, of metallothionein (Nath et al. 2000). Cd²⁺ uptake by cells has been attributed to Ca²⁺ channel activity (Limaye & Shaikh, 1999) but more recent studies suggest that the divalent metal ion transporter DMT1 may be a major contributor to this process (Bressler et al. 2004).

The Na⁺–Ca²⁺ exchanger (NCX) is the major Ca²⁺ efflux mechanism in cardiac myocytes and plays an important role in the regulation of cardiac contractility through its influence on the amount of releasable Ca²⁺ stored within the sarcoplasmic reticulum (reviewed in Blaustein & Lederer, 1999; Shigekawa & Iwamoto, 2001; Philipson et al. 2002). Studies with cardiac sarcolemmal vesicles provided early evidence that the exchanger was capable of transporting Cd²⁺ in exchange for ⁴⁵Ca²⁺ (Trosper & Philipson, 1983). This was later verified by direct measurements of Na⁺-dependent ¹⁰⁹Cd²⁺ fluxes in ferret red blood cells (Frame & Milanick, 1991). Other multivalent metal ions that have been reported to be transported by NCX are La³⁺ (Reeves & Condrescu, 2003b), Ni²⁺ (Egger et al. 1999a,b), Sr²⁺ (Trosper & Philipson, 1983) and Ba²⁺ (Condrescu et al. 1997).

The activity of NCX is allosterically regulated by cytosolic Ca²⁺ ions (DiPolo, 1979), which interact with regulatory Ca²⁺ binding sites within the exchanger's central hydrophilic domain (Levitsky et al. 1994). Various studies have reported a broad range of K_h values (the value of cytosolic Ca²⁺ ([Ca²⁺]_i) at which the activation of NCX activity was half-maximal) for allosteric Ca²⁺ activation (22–600 nM; see Reeves & Condrescu, 2003a) and references cited therein). Recently, we demonstrated that exchange activity was also activated by La³⁺ at estimated concentrations of 5–15 pM (Reeves & Condrescu, 2003b). Here we present evidence that Cd²⁺ is an even more potent activator of NCX. Thus, Cd²⁺ greatly stimulated exchange activity in Chinese hamster ovary (CHO) cells expressing the wild-type exchanger, but had little effect on CHO cells expressing a constitutively active mutant in which the Ca²⁺ regulatory sites had been deleted. Cd²⁺ activated exchange activity in the transfected cells at concentrations of 0.5–2 pM. Experiments with excised patches from oocytes expressing NCX1.1 showed that 3 pM Cd²⁺ rapidly and reversibly activated outward exchange currents. The results suggest that toxic effects of low cytosolic concentrations of Cd²⁺ might include the inappropriate activation of the Na⁺–Ca²⁺ exchanger.

	Methods

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Cells

Chinese hamster ovary (CHO) T cells (CHO K1 cells expressing the human insulin receptor (Langille et al. 1999), kindly provided by Dr Michael Czech, University of Massachusetts Medical Center, Worcester, MA, USA), were transfected with the mammalian expression vector pcDNA3 containing the coding sequence for the bovine cardiac Na⁺–Ca²⁺ exchanger (Aceto et al. 1992). Cells expressing NCX activity were selected using the ionomycin-treatment procedure of Iwamoto et al. (1998). Cells expressing the constitutive (241-680) deletion mutant were prepared similarly. In this mutant, a large part of the exchanger's central ‘regulatory’ domain has been deleted, including the regulatory Ca²⁺ binding sites. The (241-680) mutant does not require allosteric activation by cytosolic calcium for activity (Matsuoka et al. 1993). The cells were grown in F-12 medium supplemented with 10% fetal calf serum, 2 mM L-glutamine, 100 U ml^–1 penicillin, 100 µg ml^–1 streptomycin, and 20 µg ml^–1 gentamicin.

Solutions

Na-PSS contained 140 mM NaCl plus 5 mM KCl, K-PSS contained 140 mM KCl, and 20/120 Na/K-PSS contained 20 mM NaCl plus 120 mM KCl. All solutions also contained 1 mM MgCl₂, 10 mM glucose and 20 mM 3-(N-morpholino)propanesulphonic acid (Mops), buffered to pH 7.4 with Tris. Biochemicals were purchased from Sigma, unless indicated otherwise, and cell culture media, including fetal bovine serum, was from Life Technologies.

Fura-2 imaging

Cells were grown on 25 mm circular cover slips and loaded with fura-2 by incubating the cover slips for 30 min at room temperature in Na-PSS containing 1 mM CaCl₂, 1% bovine serum albumin, 0.25 mM sulfinpyrazone (to retard fura-2 transport from the cell) and 3 µM fura-2 AM (Molecular Probes). The cover slips were then washed in Na-PSS + 1 mM CaCl₂, placed in a stainless steel holder (bath volume 0.8 ml; Molecular Probes), and viewed in a Zeiss Axiovert 100 microscope coupled to an Attofluor digital imaging system. Alternating excitation at 334 and 380 nm was obtained through the use of appropriate filters, and emission was observed at > 510 nm. Forty to sixty individual cells were selected and monitored simultaneously from each cover slip. Results are presented as the ratio (R) of fluorescence intensities obtained at excitation wavelengths of 334 and 380 nm. Maximal ratios (R_max) for Cd²⁺ of 4.5–5 were obtained by treating cells with 10 µM ionomycin and applying 25 mM CdCl₂ in K-PSS. Approximate Cd²⁺ concentrations can be computed from the relation provided by Grynkiewicz et al. (1985), i.e. [Cd²⁺] = K_D(S_f/S_b)(R – R_min)/(R_max – R). In this expression, R_min is the minimal value of R obtained in the absence of Cd²⁺, and in calibrated experiments with Ca²⁺ in our laboratory, R_min is approximately equal to 0.4. S_f/S_b is the ratio of intensities at 380 nM excitation in the absence of Cd²⁺ and when fura-2 is saturated with Cd²⁺; again, by analogy to experiments with Ca²⁺, we assume the S_f/S_b is slightly higher than R_max or 5. Full calibrations with Cd²⁺ were not conducted because both R_min and background fluorescence would be difficult to determine after experiments conducted with Cd²⁺ due to its very high affinity for fura-2.

Experimental protocol: CHO cells

Cells were treated with ATP, a purinergic agonist and thapsigargin (Tg), an irreversible inhibitor of the sarco(endo)reticulum Ca²⁺-ATPase (Lytton et al. 1991), to release Ca²⁺ from the endoplasmic reticulum (ER). This was done to ensure that any change in the fura-2 ratio was due to the influx of Ca²⁺ or Cd²⁺ rather than the release of Ca²⁺ from internal stores. To assay Cd²⁺ transport by Na⁺–Ca²⁺ exchange, cells were exposed to 0.1 mM CdCl₂ under conditions where Cd²⁺ uptake would occur in exchange for internal Na⁺. In one set of experiments (Fig. 2), cells were treated with 1 µg ml^–1 gramicidin in 20/120 Na/K-PSS + 0.3 mM EGTA to permeabilize the plasma membrane to monovalent cations. This treatment ‘clamped’ the cytosolic Na⁺ concentration at 20 mM (Condrescu et al. 1997). Cd²⁺ uptake was then initiated by manually applying 4 ml of 20/120 Na/K-PSS containing 0.1 mM CdCl₂ over a period of 15 s. In other experiments (Figs 3, 4 and 5), Cd²⁺ uptake was initiated by applying 4 ml of K-PSS + 0.1 mM CdCl₂ to the cells. All experiments were carried out at room temperature. Statistical analyses utilized Student's two-tailed t test for unpaired samples.

View larger version (13K): [in this window] [in a new window]   Figure 2.  Cd2+ uptake by CHO cells expressing NCX1.1 and the constitutive mutant (241-680) A, cells were treated with gramicidin in 20/120 Na/K-PSS + 0.3 mM EGTA as described in Methods. Cd2+ uptake was initiated by the application of 0.1 mM CdCl2 in 20/120 Na/K-PSS. Traces are the average of 4–5 experiments with parental CHO cells (trace 3), cells expressing the wild-type NCX1.1 (trace 2) and cells expressing the (241-680) mutant (trace 1). B, data in A on an expanded time scale. Small vertical displacements were introduced to equalize the starting ratios for each trace. Standard error bars are shown for each data point.

View larger version (11K): [in this window] [in a new window]   Figure 3.  Persistent Ca2+ activation of Cd2+ uptake A, cells expressing NCX1.1 were incubated in Na-PSS + 0.3 mM EGTA and, for trace 1, ATP + Tg was applied to release Ca2+ from the ER. Cd2+ uptake was then initiated at the arrow by applying 0.1 mM CdCl2 in K-PSS. For trace 2, the addition of ATP + Tg was omitted. B, an identical experiment was carried out with cells expressing the (241-680) mutant. Traces for single cover slips are shown; similar results were obtained in 3 additional experiments.

View larger version (23K): [in this window] [in a new window]   Figure 5.  Cd2+ and Ca2+ uptakes by the (241-680) mutant The experiment was essentially identical to that described in Fig. 4, except that cells expressing the (241-680) mutant were used. The slope of the regression line in D is 1.66 ± 0.22; this value is significantly different from the slope of 1.0 for the dashed (additive) line (P < 0.005) and also from the slope of the regression line in Fig. 4 (P < 0.005).

Fluorescence was measured at 510 nm using a cuvette based Photon Technology International RF-M 2001 fluorometer. A quartz cuvette contained 3.0 ml of 0.15 M KCl, 10 mM Mops/Tris, 0.1 mM EGTA and 0.1 µM fura-2, pH 7.1 (room temperature). Excitation spectra (300–450 nm) were measured after successive additions of aliquots (0.5–2 µl) of 20 mM CdCl₂. Free Cd²⁺ concentrations were computed using the MAXC program (Bers et al. 1994) to correct for effects of pH and ionic strength, assuming pK_D values of 16.1 and 10.1 for the unprotonated and monoprotonated forms of EGTA (Sillin & Martell, 1971). All fluorescence values were corrected for background fluorescence (no fura-2).

Experimental protocol: giant excised patches

The canine cardiac exchanger, NCX1.1, was expressed in Xenopus laevis oocytes and Na⁺–Ca²⁺ exchange activity was measured using the giant excised patch technique, as previously described (Trac et al. 1997). Xenopus laevis were anaesthetized in 250 mg l^–1 ethyl p-aminobenzoate (Sigma) in deionized ice-water for 30 min. Oocytes were surgically removed and the frog was subsequently killed by excision of the heart. All procedures were conducted in accordance with the Canadian Council on Animal Care and were approved by the University of Manitoba Protocol Management and Review Committee. Oocytes were washed in Solution A containing (mM): 88 NaCl, 15 Hepes, 2.4 NaHCO₃, 1.0 KCl, 0.82 MgSO₄; pH 7.6 at room temperature (RT). Follicles were teased apart and the oocytes transferred to 5 ml of Solution A containing 3500 U ml^–1 collagenase (Type II; Worthington) and incubated at RT for 45–60 min with gentle agitation. The oocytes were washed several times in Solution B containing (mM): 88 NaCl, 15 Hepes, 2.4 NaHCO₃, 1.0 KCl, 0.82 MgSO₄, 0.41 mM CaCl₂, 0.3 mM Ca(NO₃)₂, 1 mg ml^–1 BSA (Fraction V; Sigma); pH 7.6 at RT, and transferred to 5 ml of 100 mM K₂HPO₄; pH 6.5 at RT, containing 1 mg ml^–1 BSA. Following incubation at RT for 12 min with gentle agitation, the oocytes were washed in Solution B at RT. Defolliculated stage V–VI oocytes were selected and incubated at 18°C in Solution B (minus BSA) until injection the following day with 23 ng of NCX1.1 cRNA. Outward Na⁺–Ca²⁺ exchange currents were activated by switching from Li⁺- to Na⁺-based bath solutions containing (mM): 100 Li⁺- or Na⁺-aspartate, 20 CsOH, 20 Mops, 20 TEA-OH, 10 EGTA, 0 or 7.23 Ca(OH)₂ to generate free Ca²⁺ concentrations of 0 and 1 µM, respectively, or 7.41 Cd(OH)₂ to generate a free Cd²⁺ concentration of 3 pM, 1.09–1.36 Mg(OH)₂; pH 7.0 at 30°C. Free Ca²⁺, Cd²⁺ and Mg²⁺ concentrations were calculated using WEBMAX STANDARD software (C. W. Patton, 2004; http://www.stanford.edu/~cpatton/webmaxc/webmaxcS.htm)

	Results

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Titration of fura-2 wth Cd²⁺

Figure 1A displays excitation spectra for fura-2 in 150 mM KCl + 0.1 mM EGTA after successive additions of Cd²⁺. The fluorescence at 340 nm excitation in the absence of Cd²⁺ (EGTA) was subtracted from the corresponding fluorescence values at each Cd²⁺ concentration and the differences (F₃₄₀ – F₃₄₀(EGTA)) are plotted versus the calculated free Cd²⁺ concentrations in Fig. 1B. The free Cd²⁺ concentrations were computed as described in Methods. The K_D of fura-2 for Cd²⁺ was 0.7 pM. The exceedingly high affinity of fura-2 for Cd²⁺ means that even large increases in the 334/380 ratio in fura-2-loaded cells correspond to very low free cytosolic [Cd²⁺] concentrations; thus, a ratio of 3.0 corresponds to a free [Cd²⁺] concentration of 5 pM (see Methods).

View larger version (13K): [in this window] [in a new window]   Figure 1.  Titration of fura-2 by Cd2+ A, excitation spectra were obtained after successive additions of Cd2+ to a solution containing 0.1 µM fura-2 and 0.1 mM EGTA as described in Methods. Emission was monitored at 510 nm. B, [Cd2+] dependence of fluorescence values at 340 nm excitation (F340) in A after subtracting the fluorescence in the absence of Cd2+ (F340(EGTA)). The data were fitted to a hyperbolic curve (continuous line) with a KD of 0.7 pM (Sigma Plot).

Cd²⁺ uptake by CHO cells expressing NCX1.1 and the constitutively active (241-680) mutant

The cells were loaded with fura-2 and then treated with gramicidin, a channel-forming monovalent cation ionophore, in 20/120 Na/K-PSS to clamp cytosolic [Na] at 20 mM (see Methods). In these experiments, Ca²⁺ had been released from the ER by applying ATP + thapsigargin (Tg) 10 min before beginning the recordings. Upon the addition of 0.1 mM Cd²⁺, the fura-2 ratios increased in cells expressing the wild-type NCX1.1 (Fig. 2A, trace 2), indicating Cd²⁺ uptake by the cells. Parental CHO cells, which do not exhibit Na⁺–Ca²⁺ exchange activity, showed no Cd²⁺ uptake under the same conditions (Fig. 2A, trace 3). Cd²⁺ uptake was terminated by the application of 20/120 Na/K-PSS containing 0.3 mM EGTA. The fura signal was nearly constant following EGTA addition, indicating that once Cd²⁺ entered the cell cytosol, it was removed slowly if at all. This probably reflects the inability of organelle or plasma membrane transporters to transport Cd²⁺ under these conditions. The time course of Cd²⁺ uptake was sigmoidal, with a prolonged lag phase before Cd²⁺ uptake attained a linear rate. For the five cover slips comprising the data in trace 2, the mean lag period (± S.E.M.), defined as the time elapsing between the addition of Cd²⁺ and the maximal velocity of Cd²⁺ uptake (maximal rate of increase in the fura-2 ratio for each cover slip), was 126 ± 8 s. Similar sigmoidal progress curves have been observed for Ca²⁺ uptake (Reeves & Condrescu, 2003a) and for La³⁺ uptake (Reeves & Condrescu, 2003b) and reflect positive feedback due to the interaction of these ions with the regulatory Ca²⁺ binding sites. By analogy to the results with Ca²⁺ and La³⁺, the duration of the lag phase probably reflects the time required for cytosolic [Cd²⁺] to increase to levels that activate NCX.

To test this interpretation, Cd²⁺ uptake was monitored in cells expressing the (241-680) mutant of the exchanger. The activity of this mutant is constitutive, i.e. it is not allosterically regulated by divalent cations, and one would therefore expect the lag phase to be absent in the mutant, as shown previously for Ca²⁺ and for La³⁺ uptakes. The results in Fig. 2A (trace 1) show that the lag period for Cd²⁺ uptake was reduced in the mutant cells, with a mean value, defined above, of 84 ± 7 s (P < 0.01 versus trace 2). The uptake curves in Fig. 2A are displayed on an expanded time scale in Fig. 2B, with small displacements of the curves along the vertical axis so that each cell type started from the same ratio.

Persistent Ca²⁺ activation of Cd²⁺ uptake

If the lag in Cd²⁺ uptake observed in cells expressing the wild-type exchanger is due to the time required for allosteric activation to occur, the lag should be reduced or eliminated if the exchanger is activated by Ca²⁺. The data in Fig. 3A confirm this prediction. Fura-2-loaded cells were incubated in Na-PSS + 0.3 mM EGTA and then ATP + Tg was added to release Ca²⁺ from the ER. A transient increase in [Ca²⁺]_i was observed that rapidly decayed back to baseline levels as Ca²⁺ was cleared from the cytosol (Fig. 3A, trace 1). Immediately after the [Ca²⁺]_i transient, K-PSS containing 0.1 mM Cd²⁺ was applied to the cells. As shown, Cd²⁺ uptake under these conditions began with only a barely discernable lag phase. In contrast, when the addition of ATP + Tg was omitted (Fig. 3A, trace 2), practically no Cd²⁺ uptake was observed initially, although a gradual increase in Cd²⁺ uptake was found after several minutes. The greatly prolonged lag phase in Fig. 3A (trace 2) compared with Fig. 2A (trace 2) reflects the absence of gramicidin in the experiments in Fig. 3A; gramicidin enhances NCX activity through cytosolic Na loading and membrane depolarization, thereby increasing the rate of Cd²⁺ uptake and shortening the lag phase. Note that the fura-2 ratio at the time of Cd²⁺ addition was nearly the same with or without the addition of ATP + Tg. This illustrates the phenomenon we have termed ‘persistent Ca²⁺ activation’, which refers to the observation that after allosteric activation of the exchanger at high [Ca²⁺]_i, the activated state is maintained for tens of seconds following the return of [Ca²⁺]_i to low values (for full details, see Reeves & Condrescu, 2003a and Chernysh et al. 2004). The mechanism underlying persistent Ca²⁺ activation is unclear; it might reflect a long-lasting elevation of subplasmalemmal [Ca²⁺]_i following the decline in bulk cytosolic [Ca²⁺]_i or an intrinsic hysteresis in allosteric Ca²⁺ activation.

Figure 3B displays the results of a similar experiment carried out with cells expressing the constitutively active (241-680) mutant. In this case, the release of Ca²⁺ with ATP + Tg had essentially no effect on Cd²⁺ uptake. Note that the abbreviated lag phase seen with this mutant was still evident after ATP + Tg addition.

Activation of Ca²⁺ uptake by cytosolic Cd²⁺

We modified the protocol illustrated in Fig. 3 to determine whether cytosolic Cd²⁺ could substitute for Ca²⁺ as an allosteric activator of Na⁺–Ca²⁺ exchange activity. In Fig. 4A (trace 1), 0.1 mM Cd²⁺ was applied to the cells in K-PSS following Ca²⁺ release with ATP and Tg, and after 2 min, Cd²⁺ uptake was terminated by the addition of 0.3 mM EGTA in Na-PSS. Cytosolic Cd²⁺ was retained within the cells for a subsequent 1.5 min interval. K-PSS containing 0.1 mM Ca²⁺ was then applied to the cells and a nearly immediate uptake of Ca²⁺ was observed. Although the rate of Ca²⁺ uptake appears to be linear in this trace, the behaviour of the individual cells being monitored was decidedly non-linear, as discussed below. As shown by trace 2 in Fig. 4A, the rate of Ca²⁺ uptake was much lower if the prior application of Cd²⁺ was omitted.

View larger version (23K): [in this window] [in a new window]   Figure 4.  Cd2+ activates NCX activity A, for trace 1, an experiment similar to that shown for trace 1 in Fig. 3 was carried out, except that Na-PSS + 0.3 mM EGTA was applied after 1.5 min of Cd2+ uptake. After an additional 2 min, Ca2+ uptake by NCX activity was initiated by applying 0.1 mM CaCl2 in K-PSS. For trace 2, K-PSS + 0.3 mM EGTA was applied between 60 and 150 s instead of K-PSS + 0.1 mM CdCl2. B, traces from 7 cells out of the 59 cells monitored for trace 2 in A. C, traces from 7 cells out of the 69 cells monitored for trace 1 in A. D, values of the initial fura-2 ratio just prior to Ca2+ addition (trace 1, A; averaged over 230–260 s) are correlated with the final fura-2 ratio (averaged over 330–350 s). The regression line (slope = 2.82 ± 0.30) is shown by the continuous line. The dashed line (slope = 1) is the relation expected if the final ratio were merely additive with respect to the initial ratio. Traces from a single cover slip are shown; similar results were obtained in 2 additional experiments.

The Cd²⁺-treated cells in Fig. 4C each show different values for the initial fura ratio just prior to the addition of Ca²⁺; this reflects the variability among the different cells in the cytosolic Cd²⁺ concentration achieved during the 1.5 min exposure to 0.1 mM Cd²⁺. There was a strong correlation between the level of cytosolic Cd²⁺ and the subsequent rate of Ca²⁺ uptake. This is illustrated in Fig. 4D, in which the ‘final’ fura ratio, averaged over the time period between 330 and 350 s in this experiment, is plotted versus the ‘initial’ fura ratio prior to Ca²⁺ addition, averaged over 230–260 s. The correlation is highly significant (R² = 0.58; P << 10^–3), indicating that there was a positive correlation between the cytosolic Cd²⁺ concentration and the degree of NCX activation. The dashed line in Fig. 4D displays the regression line that would be expected if the presence of Cd²⁺ merely produced an additive increment with respect to the final fura ratio. Clearly the response of the cells was much greater than additive.

Figure 5 depicts the results of an experiment identical to that shown in Fig. 4, except that cells expressing the constitutively active (241-680) mutant were used. As shown in Fig. 5A, these cells displayed high rates of Ca²⁺ uptake irrespective of whether the cells were (trace 1) or were not (trace 2) pre-exposed to Cd²⁺. Traces for representative individual cells are shown in Fig. 5B and C for traces 2 and 1, respectively. Note that Ca²⁺ uptake began without a lag phase in cells expressing the (241-680) mutant (see also Reeves & Condrescu, 2003a). Following Cd²⁺ uptake (trace 1, Fig. 5A), the fura-2 ratio increased at a greater rate (2.6-fold) than without Cd²⁺ (trace 2). Figure 5D shows that Cd²⁺-treated cells expressing the (241-680) mutant displayed a positive correlation between the initial and final fura-2 ratios after Ca²⁺ addition (R² = 0.5; P << 10^–3). However, in contrast to the cells expressing the wild-type exchanger (Fig. 4D), the slope of the regression line is only slightly higher than expected for an additive effect (dashed line).

The data presented above indicate that Cd²⁺ allosterically activates exchange activity in cells expressing the wild-type NCX1.1. For the experiment in Fig. 4, Cd²⁺ concentrations yielding an average fura-2 ratio of 1.2 produced a large increase in the rate of Ca²⁺ uptake. This ratio corresponds to an estimated free Cd²⁺ concentration of 0.74 pM (see Methods). NCX is not fully activated at this concentration, however, since many of the individual cells continue to show lag periods before Ca²⁺ uptake is observed (Fig. 4C). Full activation of NCX would be indicated by an immediate increase in [Ca²⁺]_i upon application of Ca²⁺ in K-PSS, similar to the results shown in Fig. 5 for the (241-680) mutant. The lag periods are nevertheless much shorter than seen in the absence of Cd²⁺ addition (Fig. 4B), consistent with partial activation of the exchanger at these Cd²⁺ concentrations.

Cd²⁺ activates NCX activity in excised patches

Excised patches from frog oocytes expressing the canine wild-type NCX1.1 were utilized to determine directly whether Cd²⁺ activates NCX activity. The data in Fig. 6 show that, in the absence of cytosolic Ca²⁺, the application of 100 mM Na⁺ to the cytosolic surface of the excised patch produces only a small outward exchange current. Much larger currents were observed when Na⁺ was applied in the presence of 3 pM Cd²⁺, also in the absence of cytosolic Ca²⁺. Outward currents quickly attained a peak and then declined to a lower steady-state value; the decline reflects an inactivation process that is initiated by the binding of Na⁺ to cytosolic translocation sites (Na⁺-dependent or I₁ inactivation; reviewed in Hilgemann, 1996). The increase in current in the presence of Cd²⁺ reflects allosteric activation of the exchanger since no effects of Cd²⁺ were observed after destroying the regulatory properties of the exchanger with -chymotrypsin (Fig. 6, right traces). Chymotrypsin proteolyses the exchanger's hydrophilic regulatory domain so that exchange currents no longer display normal regulatory properties such as Na⁺-dependent inactivation or allosteric Ca²⁺ activation (Hilgemann, 1990). The peak and steady-state currents elicited by Cd²⁺ were inhibited 69 ± 3% (n = 7) and 89 ± 3% (n = 4), respectively, by 3 µM KB-R7943 (data not shown), an inhibitor of Na⁺–Ca²⁺ exchange activity (Iwamoto et al. 1996); this provides a further indication that they originated from NCX activity as opposed to a non-specific effect of Cd²⁺ on membrane conductance.

View larger version (10K): [in this window] [in a new window]   Figure 6.  Cd2+ activates outward exchange currents in excised giant patches Representative traces of outward Na+–Ca2+ exchange currents are shown, before and after deregulation of NCX1.1 with -chymotrypsin (1 mg ml–1 for 1 min). Outward current was activated by rapid application of 100 mM Na+ to the cytoplasmic side of the patch in the absence and presence of 3 pM cytosolic Cd2+. Cytosolic Ca2+ was absent throughout these experiments. Note that 3 pM Cd2+ caused a significant activation of the Na+–Ca2+ exchanger. There were no discernible effects of Cd2+ once the exchanger was de-regulated with -chymotrypsin.

View larger version (13K): [in this window] [in a new window]   Figure 7.  Activation of NCX activity by Cd2+ is rapid and reversible Outward NCX currents were generated by the rapid application of 100 mM Na+ in alternating 32 s active (100 mM Na+) and inactive (100 mM Li+) episodes. A, application of 3 pM Cd2+ in the absence of cytosolic Na+ (episode a) or simultaneously with cytosolic Na+ (episode c). B, Cd2+-activated exchange current generated after a 32 s pre-incubation with Li+-based 3 pM Cd2+ solution. C, comparison of currents generated with or without Cd2+ pre-incubation. Current integrals were normalized to those obtained in the absence of Cd2+.

The traces in Fig. 8 depict alternating 32 s active and inactive episodes in which NCX was activated by the continuous presence of either 1 µM Ca²⁺ (panel A) or 3 pM Cd²⁺ (panel B). Here, each trace constituted one episode with no activating divalent cations and four consecutive episodes in the presence of either 1 µM Ca²⁺ or 3 pM Cd²⁺ designated 1–4 (the fourth episode is not shown for brevity). Figure 8C and D shows the pooled data on currents integrated over the 32 s active episodes in experiments using Ca²⁺ or with Cd²⁺, respectively. The integrals were normalized to the largest integral in each series (always episode 1 for Cd²⁺).

View larger version (22K): [in this window] [in a new window]   Figure 8.  Attenuation of serial Cd2+-activated currents A, a series of Na+–Ca2+ exchange currents from the same patch was generated by serial applications of 100 mM Na+ in the presence of 1 µM Ca2+. B, the experimental protocol is same as in A, except that 3 pM Cd2+i was used instead of 1 µM Ca2+i. C and D show the pooled data on currents integrated over the 32 s active episodes in experiments using Ca2+ or Cd2+, respectively. The integrals were normalized to the largest integral in each series (always episode 1 for Cd2+). The values of normalized integral in the absence of regulators were 0.17 ± 0.02 (n = 6) and 0.29 ± 0.03 (n = 10) for regulation by Ca2+ and by Cd2+, respectively. Thus, in comparison to currents in the absence of activator, Cd2+-activated currents were 40% smaller than Ca2+-activated currents. The integrals in episodes 1–4 in the presence of Ca2+ (C) were not significantly different (ANOVA). For Cd2+, however (D), the integrals for episodes 2, 3 and 4 were each significantly different from the first episode (by ANOVA).

Subsequent applications of Na⁺ in the presence of Cd²⁺ yielded substantially reduced amplitudes of outward exchange currents (panel D), as well as further reductions in the rate of current development (panel B). In comparison to the currents seen in the absence of activator, currents activated by 3 pM Cd²⁺ were 40% lower than currents activated by 1 µM Ca²⁺ (see figure legend). We did not attempt to measure a full concentration profile for Cd²⁺ because of the alterations Cd²⁺ induced in the kinetic features of exchange currents after the initial pulse (Fig. 8B) and because full recovery from Na⁺-dependent inactivation was incomplete in the presence of Cd²⁺ (data not shown).

Cd²⁺ inhibits exchange activity in CALX1.1

The Na⁺–Ca²⁺ exchanger from Drosophila (CALX1.1) is anomalous in that Ca²⁺ acts as an allosteric inhibitor of Na⁺–Ca²⁺ exchange activity rather than an activator. This is evident in Fig. 9, when the effects of 1 µM Ca²⁺ are compared on exchange currents generated by NCX1.1 (trace 1) and CALX1.1 (trace 3). Note that large outward currents developed in the absence of Ca²⁺ in patches containing CALX1.1 (trace 3). As shown previously (Hryshko et al. 1996), application of Ca²⁺ after a steady state was attained led to a substantial decrease in current which gradually recovered when the Ca²⁺ was removed. Trace 4 shows that a similar inhibition of the CALX1.1 steady-state current is elicited by 3 pM Cd²⁺; as discussed above, Cd²⁺ activated currents generated by NCX1.1 (trace 2). These results are entirely consistent with the notion that Cd²⁺ acts at the high affinity regulatory Ca²⁺ binding site for both exchangers. Pooled results are shown in Fig. 9B, illustrating the inhibitory effects of 3 pM Cd²⁺ on CALX1.1-mediated currents. Like Ca²⁺ (Omelchenko et al. 1998), Cd²⁺ inhibited steady-state currents to a greater extent than peak currents. However, the inhibitory effects of 1 µM Ca²⁺ on steady-state (75 ± 3%, n = 36) and peak currents (58 ± 3%, n = 39; data not shown) were substantially more robust than for 3 pM Cd²⁺; this is consistent with the reduced effectiveness of 3 pM Cd²⁺ in activating exchange currents in NCX1.1 compared with 1 µM Ca²⁺. We conclude that the anomalous inhibitory effects of Ca²⁺ on CALX1.1 currents are mimicked by Cd²⁺.

View larger version (15K): [in this window] [in a new window]   Figure 9.  Inhibition of CALX-1-mediated exchange currents by Ca2+ and Cd2+ A, representative traces demonstrating the activation of NCX1.1-mediated Na+–Ca2+ exchange currents and the inhibition of CALX1.1-mediated Na+–Ca2+ exchange currents by Ca2+i and Cd2+i. Both Ca2+ and Cd2+ were added to the cytoplasmic side of the patch in the continuous presence of 100 mM Na+i. B, pooled data on the inhibitory effects of Cd2+i on CALX1.1-mediated Na+–Ca2+ exchange currents. The effects of Cd2+ on peak and steady-state currents, and current integrals are shown (n = 6). Note that the inhibition of steady-state current and the current integral is significantly greater than the inhibition of peak current.

	Discussion

Top Abstract Introduction Methods Results Discussion References

The time course of Cd²⁺ uptake displayed a pronounced lag phase before uptake became linear. Similar lags were seen with Ca²⁺ and La³⁺ as transport substrates (Reeves & Condrescu, 2003a,b); see also Fig. 4B for Ca²⁺), and in these instances we demonstrated that the sigmoidal time course reflected a positive feedback between cation entry and allosteric activation of NCX. This conclusion was strongly supported by the finding that the constitutively active mutant (241-680) did not display a lag phase for Ca²⁺ or La³⁺ uptake. In the case of Cd²⁺, however, a definite lag phase was observed in cells expressing (241-680), although it was significantly shorter in duration than the lag for the wild-type NCX1.1 (84 ± 7 versus 126 ± 8 s; P < 0.01) (Fig. 2B). The origin of the lag phase in cells expressing the mutant exchanger is uncertain. One possibility is that it reflects the titration by Cd²⁺ of binding sites with a higher affinity for Cd²⁺ than fura-2. Metallothioneins would provide binding sites with the appropriate affinity, since their K_D value for Cd²⁺ has been reported to be 10^–22 M (Kagi & Vallee, 1961; Klaassen et al. 1999).

The data in Figs 3–5 provide compelling support for the idea that the sigmoidal time course of Cd²⁺ uptake reflects the activation of NCX activity by positive feedback as Cd²⁺ accumulates within the cell. Thus, activation of NCX by the release of Ca²⁺ from the ER greatly reduced the lag period in cells expressing NCX1.1 (Fig. 3A); in contrast, Ca²⁺ release had no effect on Cd²⁺ uptake by (241-680) (Fig. 3B). As noted in Results, the enhanced rate of Cd²⁺ uptake following Ca²⁺ release in Fig. 3A (trace 1) provides an illustration of persistent Ca²⁺ activation, where the allosterically activated state of NCX is retained for a period of several tens of seconds after cytosolic [Ca²⁺]_i declines to low values. This phenomenon is fully described in Reeves & Condrescu (2003a) and Chernysh et al. (2004). We initially suggested that the transient persistence of allosteric Ca²⁺ activation at low [Ca²⁺]_i might be due to local elevations in [Ca²⁺]_i beneath the plasma membrane that decay slowly following the return of bulk [Ca²⁺]_i to low values (see Reeves & Condrescu, 2003a). Another possibility, however, is that allosteric Ca²⁺ activation displays an intrinsic hysteresis, i.e. activation of NCX requires more Ca²⁺ than is needed to maintain the activated state once it is formed. Both possibilities are currently under investigation.

Moreover, cytosolic Cd²⁺ greatly reduced the lag time for Ca²⁺ uptake in cells expressing NCX1.1. In Fig. 4A (trace 1), Ca²⁺ uptake by reverse NCX activity began nearly immediately in NCX1.1 cells that had taken up a modest amount of Cd²⁺ prior to initiating Ca²⁺ uptake. In the absence of cytosolic Cd²⁺, however, Ca²⁺ uptake was very slow. Comparison of traces for individual cells in Fig. 4B and C showed that Cd²⁺ reduced the duration of the lag period during Ca²⁺ uptake. Moreover, the extents of Ca²⁺ uptake among the cells in the population showed a greater-than-additive correlation with the initial level of cytosolic Cd²⁺ (Fig. 4D), suggesting that the degree of activation of NCX increased with the cytosolic Cd²⁺ concentration. The range of initial ratios due to Cd²⁺ accumulation than resulted in greater-than-additive Ca²⁺ uptake in this experiment was 1–2, which corresponds to approximate free Cd²⁺ concentrations of 0.5–2 pM (see Methods). We suggest that cytosolic Cd²⁺ partially activates Na⁺–Ca²⁺ exchange activity within this range of concentrations, resulting in a reduced lag period for further activation of NCX activity by Ca²⁺ as it enters the cell.

These conclusions are strongly supported by the behaviour in identical experiments of the cells expressing the constitutive mutant (Fig. 5). Here, Ca²⁺ uptake began immediately (i.e. without a lag period) whether or not cytosolic Cd²⁺ was present (Fig. 5A–C). A positive correlation was also observed between the extent of Ca²⁺ uptake and the initial ratio due to Cd²⁺, but in the case of the mutant, the slope of the regression line was only slightly higher that for an additive relationship. The average rate of Ca²⁺ uptake was 2.6-fold higher in cells that had pre-accumulated Cd²⁺ than in cells that had not, suggesting the Cd²⁺ might stimulate the activity of the mutant as well as the wild-type. However, much of this effect was probably due to the reduced amount of free fura-2 available for interacting with Ca²⁺ when Cd²⁺ is present. For the Cd²⁺-treated (241-680) cells, the average initial ratio prior to Ca²⁺ addition was 1.2, which, under our experimental conditions, corresponds to titration of approximately 50% of the fura-2. Thus, for a given amount of Ca²⁺ uptake, the increase in the fura-2 ratio would be greater in the Cd²⁺-treated cells simply because only half the amount of free fura-2 is available to be titrated by the incoming Ca²⁺. This might also account for the slight departure from additivity between the initial and final ratios discussed above for Fig. 5D; indeed, the slope of the regression line is 66 ± 22% higher than the slope of the line indicating additivity, consistent with our estimate that the presence of Cd²⁺ titrated approximately half of the fura-2.

Experiments with excised patches from oocytes expressing the canine NCX1.1 provide further evidence that Cd²⁺ activates exchange activity at picomolar concentrations. Cd²⁺ (3 pM) markedly increased outward exchange currents but had no effect following treatment of the cytosolic membrane surface with chymotrypsin (Fig. 6). The chymotrypsin treatment abrogates all ionic regulatory features of the exchanger, including allosteric Ca²⁺ activation (Hilgemann, 1990). Thus, chymotrypsin-treated membranes are functionally equivalent to the (241-680) mutant in terms of their regulatory properties (see also Matsuoka et al. 1993). Cd²⁺ had no effect on membrane currents in the absence of cytosolic Na⁺ (Fig. 7) and Cd²⁺-activated outward exchange currents were blocked by KB-R7943, an inhibitor of NCX activity; both observations indicate that the effects of Cd²⁺ were solely due to their activation of outward NCX currents. The activating effects of Cd²⁺, like those of Ca²⁺, developed within the mixing time of the solution changes and were rapidly reversible (Fig. 7). In patches from oocytes expressing the Drosophila exchanger CALX1.1, the effects of Cd²⁺ were similar to the anomalous effects of Ca²⁺ in inducing an inhibition of outward currents, rather than an activation. Cd²⁺ (3 pM) was somewhat less effective than Ca²⁺ (1 µM) both in activating exchange currents in NCX1.1 (Fig. 8C and D) and in inhibiting exchange currents in CALX-1 (Fig. 9B).

The findings with excised patches indicate that Cd²⁺ was a faithful mimic of Ca²⁺ in terms of its regulatory effects on the exchanger. Two observations, however, suggest that there are subtle differences between Cd²⁺- and Ca²⁺-activated currents. First, outward currents developed more slowly upon application of cytosolic Na⁺ when Cd²⁺ was the activating divalent cation (Figs 7 and 8). This occurred whether Cd²⁺ was applied simultaneously with Na⁺ or tens of seconds prior to Na⁺ (Fig. 7). Second, the rate of current development and current amplitude are both reduced upon multiple applications of Na⁺ in the presence of Cd²⁺ (Fig. 8) whereas Ca²⁺-activated currents showed much slower rates of decay under similar conditions. The significance of this finding is uncertain but one possible explanation is that Cd²⁺ retards the recovery of the exchanger from Na⁺-dependent inactivation. However, we could not obtain complete recovery curves. Even after very prolonged interpulse intervals, recovery was never complete and the kinetic features of currents changed considerably between the first and subsequent pulses (Fig. 8). We conclude that, although Cd²⁺ mimics most of the regulatory properties of Ca²⁺ with respect to exchange activity, additional effects of Cd²⁺ are also evident.

With the exceptions noted above, Cd²⁺ and Ca²⁺ appear to behave similarly with respect to allosteric activation of NCX. Importantly, however, Cd²⁺ acts at concentrations nearly five orders of magnitude less than Ca²⁺. We previously reported that La³⁺ also activates NCX activity within an estimated concentration range of 5–15 pM (Reeves & Condrescu, 2003b). Our findings with Cd²⁺ and La³⁺ raise the issue of whether endogenous heavy metal ions might activate NCX under either physiological or pathophysiological conditions. It seems likely that metallothioneins, with their extraordinarily high affinity for heavy metal ions (see introduction), would keep the free concentrations of these ions well below the levels needed to activate NCX under physiological conditions. However, cardiac tissue appears to be particularly susceptible to Cd²⁺ toxicity, perhaps because metallothioneins are less well expressed in cardiac myocytes, and are also less inducible, than in other tissues (Nath et al. 2000). The toxic effects of Cd²⁺ in cardiac myocytes and many other tissues are widely attributed to ROS generation, possibly through interactions with components of the mitochondrial electron transfer chain (Thevenod, 2003; Pourahmad et al. 2003; Wang et al. 2004). The results presented here raise the additional possibility that the toxic profile for Cd²⁺ in cardiac myocytes might include an interference in the allosteric Ca²⁺ regulation of NCX activity at extremely low cytosolic concentrations of Cd²⁺.

	Footnotes

 
The laboratories of Drs Hryshko and Reeves contributed equally to this work.

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	Acknowledgements

 
We thank Larissa Bonilla (J.P.R. lab) for excellent technical assistance. This work was supported by National Heart, Lung and Blood Institute Grant HL-49932 and American Heart Associate Grant 0151201T (JPR) and by an operating grant from the Canadian Institutes of Health Research (LVH).

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