Zinc ions (Zn2+) are important co-factors in more than 200 enzymes and many non-enzymatic proteins. It is known that Zn is present in the central nervous system, where it raises electrical activity of neurons and may also induce epileptic activity (1,2). Results of electrophysiological studies provide evidence that Zn2+ modulates the activity of many different types of ion channels (2). Among them are: GABAA and NMDA receptors, voltage-gated sodium and calcium channels, voltage-gated and ATP-dependent potassium channels as well as voltage- and ligand-gated chloride channels (2). To the group of voltage-gated potassium channels modulated by Zn2+ belong delayed-rectifier channels in squid giant axons (3,4), Shaker-class channels stably expressed in cell line Sf9 (5), KA channels expressed in rat neurons from suprachiasmatic nucleus (6), granule cells from rat cerebellar slices (7), and in snail neurons (8), as well as in the case of Kv1.1, Kv1.4 and Kv1.5 channels stably expressed in mouse fibroblasts (9). In all the cases it was shown that application of Zn2+ at concentrations up to 1 mM caused a shift of the voltage-dependence of the steady-state activation and inactivation towards positive membrane potentials, a pronounced slowing of the current activation and inactivation rate and a significant reduction of the current amplitudes (3-9). Moreover, it was shown that interactions of Zn2+ with the channels are due to binding to specific binding sites and are not related to the compensation of negative surface charges by divalent cations (5).

Voltage-gated potassium channels Kv1.3 are expressed abundantly in human T lymphocytes (TL), where they play an important role in setting the resting membrane potential, cell mitogenesis, apoptosis and volume regulation (10-13). Kv1.3 channels are also present in rat central nervous system, especially in olfactory bulb neurons, where they play a modulatory role in action potential generation (14-16). The channels are also expressed in human alveolar macrophages (17), rat choroid plexus epithelial cells (18), epithelial cells from rabbit kidney and colon (19), human gliomas (20) and in rat prostate cancer cell lines (21). There is also evidence that the activity of Kv1.3 channels regulates energy homeostasis, body weight and peripheral insulin sensitivity in mice (22-23).

Although the inhibitory effect of Zn2+ on Kv1.3 channels is known since 1985 (24), the effect was studied in detail only recently (25). Obtained data provide evidence that application of 10-100 µM Zn2+ caused a shift of the voltage dependence of both steady-state activation and inactivation towards positive membrane potentials. There was also a significant slowing of the current activation rate and a reduction of the current amplitude to about 70% of the control value. Raising the Zn2+ concentration ftom 100 µM to 2.6 mM caused a concentration-dependent decrease of the current amplitude to about 20% of the control value without any further changes neither of the voltage-dependence of the steady-state activation and inactivation nor of the activation kinetics. The modulatory effect of Zn2+ was not due to the compensation of negative surface charges by Zn (25-26). It was suggested that Zn2+ might bind to two different binding sites. Binding to one of them causes changes of the voltage dependence of the steady-state activation and inactivation and slowing of the activation rate, whereas binding to another one inhibits the current amplitudes (25). The observed modulatory effect of Zn2+ might be of physiological significance (27-28).

One of still unresolved problems concerning the modulatory effect of Zn2+ on Kv1.3 channels is the influence of pH on this effect. It is known that extracellular protons are powerful modulators of the activity of many types of ion channels including Kv1.3 channels (29-30). In case of Kv1.3 channels it was shown that lowering of pHo to less than 6.8 caused an inhibition of the current amplitude, a shift of the voltage-dependence of both steady-state activation and inactivation towards positive membrane potentials and slowing the current activation and inactivation rate (29). The modulatory effect was partially due to the compensation of negative surface charges by protons, and partially to the titration of the histidine residue at position 399 (30). There is indirect evidence that changes in pHo might influence the modulatory effect of Zn2+ on the channels. First, the effect of acidic pHo on the current amplitude, steady-state activation and inactivation and activation kinetics resemble the effect exerted by Zn. Moreover, investigations performed on Shaker-class voltage-gated potassium channels provided evidence that the modulatory effect of Zn2+ on these channels was significantly diminished at pHo =6.5 due to competition between extracellular protons and Zn for the same binding sites containing amino groups (4,5). Results obtained for Kv1.5 channels, which are related to Kv1.3, showed that lowering of pHo mimicked the inhibitory effect of Zn2+ and that Zn2+ and protons bind the same binding sites containing histidine in the channel turret (H463) and arginine near the pore mouth (R487) (31-32).

In this study we examined the influence of changes of pHo on the modulatory effect of Zn2+ on Kv1.3 channels. Since Kv1.3 channels predominate in human TL, these cells were used as the model system in the experiments. The results provide evidence that, in contrast to what was obtained for Shaker-class channels, the modulatory effects exerted by lowering of pHo to 6.4 and application of Zn2+ on Kv1.3 channels were additive and that the modulatory effect of Zn2+ did not depend on the changes of pHo in the range from 6.4 to 8.4. The inhibitory effects of extracellular protons and Zn2+ were significantly diminished when the holding potential was changed from -90 mV to -60 mV, whereas the proton- and Zn2+-induced shift of the voltage-dependence the activation midpoint remained unaffected when changing the holding potential.

The preliminary results were published in the form of an abstract (33).


MATERIALS AND METHODS
Cell separation, solutions and pipettes. Human TL were separated from peripheral blood samples from 10 healthy donors using a standard method described elsewhere (34).

Upon experiment, the cells were placed in the external solution containing (in mM): 150 NaCl, 4.5 KCl, 1 CaCl2, 1 MgCl2, 10 HEPES, 10 Glucose, pH=7.35 adjusted with NaOH, 300 mOsm. In case of the solutions with pH of 6.4 and 8.4, 10 mM MES and 10 mM TRIS was applied, respectively, insteat of HEPES. The pipette solution contained (in mM): 150 KCl, 1 CaCl2, 1 MgCl2, 10 HEPES, 10 EGTA; pH=7.2 adjusted with KOH and osmomolarity - 300 mOsm. The concentration of free calcium in the internal solution was below 100 nM, assuming the dissociation constant for EGTA at pH=7.2 of 10-7 M (35). Such a low calcium concentration was applied in order to prevent the activation of calcium-activated IKCa1 channels (35). The reagents were provided by the Polish Chemical Company (POCH, Gliwice, Poland), except for HEPES, MES, TRIS, EGTA and ZnCl2, which were purchased from SIGMA. Dishes with cells were placed under an inverted Olympus IMT-2 microscope. External solutions containing Zn were applied using a fast perfusion system RSC 200 (Bio-Logic, Grenoble, France). Pipettes were pulled from a borosilicate glass (Hilgenberg, Germany) and fire-polished before the experiment. The pipette resistance was in the range of 3-5 M.

Electrophysiological recordings. Whole-cell potassium currents in TL were recorded applying the patch-clamp technique (36). The currents were recorded using an EPC-7 Amplifier (List Electronics, Darmstadt, Germany), low-pass filtered at 3 kHz, digitised using the CED Micro 1401 (Cambridge, UK) analogue-to-digital converter with the sampling rate of 10 kHz. A standard protocol of depolarising voltage stimuli contained 7 pulses in the range from -60 mV to +60 mV (20 mV increment) applied every 10 s; pulse duration was 40 ms and holding potential - 90 mV. The linear (ohmic) component of the current was subtracted off-line from the final record. In other experiments, a sequence containing 10 voltage ramps applied every 20 s depolarising the cell membrane from -100 mV to +40 mV, ramp duration - 340 ms, holding potential -90 mV, was applied. The linear (ohmic) component of the current was subtracted off-line from the final record. The data were analysed using the WCP J. Dempster Program.

Since the number of active channels varied significantly among the cell population, the magnitude of the inhibitory effect caused by Zn2+ and protons at pHo = 6.4 is presented in terms of the relative peak current recorded applying the voltage ramp protocol at +40 mV. The activation midpoint of the currents is presented in terms of the voltage of half-to-peak current (V1/2) defined as the membrane potential at which the ramp current reaches the half of its maximal value.

The activation kinetics was fitted by applying a power function, given by an equation: I(t) = Ip(1-exp(-t/n))2, where: n - activation time constant.

The data are given as the mean ± standard error. All experiments were carried out at room temperature (22-24 °C).


RESULTS
Figure 1A shows an example of whole-cell currents recorded in a TL applying a standard protocol of depolarising stimuli described in Materials and Methods. The currents were evoked upon a membrane depolarisation to potentials more positive than -40 mV. The currents were completely blocked upon application of 5 mM 4-aminopiridine (4-AP, Fig. 1B), which selectively blocks Kv1.3 channels in TL (37). This indicates that the recorded currents were predominantly due to the activation of Kv1.3 channels.

Fig. 1. A) Example of the whole-cell currents in a TL recorded applying a standard protocol of depolarizing voltage pulses (see Materials and Methods). For clarity, only the first 30 ms of the records are shown. B) Relative peak current to voltage relationship obtained under control conditions (open squares, n=6) and upon application of 5 mM 4-AP (filled squares, n=6). Data points were connected with a point-to-point line.

Figure 2 depicts the whole-cell potassium currents recorded in a TL applying a standard protocol of depolarising stimuli at pHo = 7.35 (A) and when the pHo was lowered to 6.4 (B). Apparently, lowering of the pHo slowed remarkably the current activation rate (Fig. 2B, D). Moreover, at pHo =6.4 the currents were evoked only at the potential of 0 mV and more positive, suggesting that the channel activation threshold was shifted towards positive membrane potentials. Finally, there was a reduction of the current amplitude to about a half of the control value (Fig. 2B). Effect of application of 300 µM Zn2+ at pHo =6.4 is shown in Figure 2C. Apparently, application of Zn2+ slowed additionally the current activation rate (Fig. 2C, D). Moreover, when Zn2+ was applied at pHo =6.4, the currents were activated only at the potential of +20 mV and more positive. This may suggest that application of Zn2+ at pHo =6.4 produced a shift of the channel activation threshold towards positive membrane potentials, such as it occurred at pHo =7.35 (25). Finally, Zn2+ application caused additionally a decrease of the current amplitudes to about a half of the value obtained in the absence of Zn2+.

Fig. 2. Whole-cell potassium currents recorded in a TL applying a standard sequence of depolarising voltage pulses (see Materials and Methods): A) pHo=7.35, B) pHo=6.4, C) upon application of 300 µM Zn at pHo=6.4, D) normalised currents recorded at +60 mV at pHo=7.35 (upper trace), 6.4 (middle trace) and upon application of 300 µM Zn at pHo=6.4 (lower trace).

The activation time constants (n) calculated for the currents recorded at pHo =7.35, 6.4 and upon appication of 300 µM Zn2+ at pHo =6.4 are depicted in Figure 3. The n values calculated at pHo =6.4 were higher than the values obtained at pHo =7.35 for any voltage between 0 and +60 mV (at -20 mV calculation of the tn was not possible at pHo =6.4). The n values calculated at +40 and +60 mV in the presence of Zn2+ at pHo =6.4 were significantly higher than the values calculated in the absence of Zn2+ at this value of pHo . Thus, the data confirm the hypothesis that the current activation rate was slowed at pHo =6.4 and that application of Zn2+ at this pHo slowed additionally the activation rate more significantly than extracellular protons.

Fig. 3. Activation time constants (defined in Materials and Methods) as a function of membrane potential: empty squares-currents recorded at pHo=7.35 (n=5); filled squares - currents recorded at pHo=6.4 (n=5); empty triangles - currents recorded upon application of 300 µM Zn at pHo=6.4. Data points were connected by a point-to-point line.

In order to study in a further detail the influence of Zn2+ application at pHo =6.4 on the current amplitude and activation midpoint, another series of experiments applying the voltage ramp protocol (see Materials and Methods) was performed. Figure 4 depicts an example of ramp currents recorded in a TL at pHo =7.35 (upper trace), 6.4 (middle trace) and upon application of 100 µM Zn2+ at pHo =6.4 (lower trace).

Fig. 4. Whole-cell potassium ramp currents recorded applying a sequence of voltage ramps (defined in Materials and Methods). Depicted currents were recorded at pHo=7.35 (upper trace), at pHo=6.4 (middle trace) and upon application of 100 µM Zn at pHo=6.4 (lower trace).

Apparently, the currents at pHo =6.4 were activated with a time delay in relation to the currents recorded at pHo =7.35. Considering the voltage ramp protocol, the time delay means that the currents at pHo =6.4 were activated at higer (more positive) membrane potentials than the currents recorded at pHo =7.35. Moreover, the currents recorded at pHo =6.4 exhibited a significantly smaller amplitude than the currents recorded at pHo =7.35. Application of 100 µM Zn2+ at pHo =6.4 produced additionally both a pronounced decrease of the current amplitude and a shift of the current activation threshold towards positive membrane potentials. The magnitude of this shift was calculated in terms of the shift of V1/2 parameter defined in Materials and Methods.

The two bars: "pH=6.4" and "Zn at pH=6.4" in Figure 5 depict the average values of the V1/2 shift at pHo =6.4 and upon Zn2+ application at pHo =6.4, respectively. At pHo =6.4 the V1/2 was shifted by 16.2±1.43 mV (n=15) towards positive membrane potentials. Application of Zn2+ at pHo =6.4 caused an additional shift of the V1/2 by 19.3±1.48 mV (n=15) towards positive membrane potentials. Since the V1/2 values obtained for Zn2+ concentrations of 100 µM, 300 µM and 1 mM were comparable one to another, they were averaged. The total shift of the V1/2 caused by Zn2+ and extracellular protons at pHo =6.4 was 35.5 mV (n=15, see last bar in Fig. 5) and was much bigger than the shift induced by Zn2+ alone (p< 0.05, Student t-test).

Fig. 5. The shift of the V1/2 value when lowering the pHo from 7.35 to 6.4 (left, n=15), upon application of 100 µM Zn at pHo=6.4 (center, n=15), and the summaric effect (right).

It is shown that the relative peak current at pHo =6.4 was decreased to 0.55±0.055 (n=15) of the curent recorded at pHo =7.35 (see Fig. 6). Application of 100 µM Zn2+ at pHo =6.4 reduced additionally the current amplitude to 0.56±0.05 (n=10) of the value in the absence of Zn2+ (see Fig. 6). Taken together, the current amplitude was reduced by Zn2+ and protons to 0.26±0.05 (n=10) of the control value, and this reduction was significantly (p<0.05, Student t-test) more potent than when Zn2+ was applied alone.

Fig. 6. The relative peak ramp current recorded when lowering the pHo from 7.35 to 6.4 (left, n=15), upon application of 100 µM Zn at pHo=6.4 (center, n=10), and the summaric effect (right).

The magnitude of the V1/2 shift and current amplitude reduction upon Zn application was also measured at pHo =7.35 and 8.4. First, the effect of raising pHo from 7.35 to 8.4 on the whole-cell potassium ramp currents was examined. In contrast, to what was observed at pHo =6.4, raising the pHo to 8.4 did not change significantly the V1/2 value (not shown). There was only a small increase of the current amplitude to 1.06±0.05 (n=10) of the control value (not shown), and this increase was statistically insignficant (p>0.05, Student t-test).

The magnitude of the Zn2+-induced shift of the V1/2 value at pHo =6.4, 7.35 and 8.4 is summarised in Figure 7. The V1/2 values were shifted by 19.3±1.48 mV (n=15); 22.8±0.49 mV (n=10) and 22.7±1.2 mV (n=10), respectively. These shifts were not significantly different one from another (p>0.05, one-way ANOVA). This may suggest that the Zn2+-induced shift of the V1/2 value was not affected by pHo in the range from 6.4 to 8.4.

Fig. 7. Effect of pHo on the Zn-induced shift of the V1/2 value. Further description in the text.

It is known that application of Zn reduced also the current amplitude in a concentration-dependent manner. Experiments with the voltage ramps were carried out at pHo =6.4, 7.35 and 8.4 and at Zn2+ concentrations of 100, 300 µM and 1 mM. Results are summarised in Figure 8 (data obtained at 300 µM Zn2+ concentration is ommitted for clarity). Apparently, the relative peak currents recorded at both Zn2+ concentrations at pHo =6.4, 7.35 and 8.4 did not differ significantly one from another (p>0.05, one-way ANOVA). Thus, similarly to what was observed in case of the shift of the V1/2 value, the inhibitory effect of Zn2+ on the currents is pHo -independent in the range from 6.4 to 8.4.

Fig. 8. The inhibitory effect of Zn applied at concentrations of 100 µM (black bars, n=10) and 1 mM (grey bars, n=10) on the relative peak ramp currents at +40 mV at pHo=6.4, 7.35 and 8.4.

Available data provide evidence that the inhibitory effect of Zn2+ on the whole-cell potassium currents recorded from the holding potential of -90 mV may be abolished or even reversed when the holding potential is changed to -60 or -40 mV (6,38). This is presumably due to removal of the steady-state inactivation, which compensates the inhibitory effect of Zn2+ on the currents (6,38). Our results published previously showed that application of Zn2+ shifted the inactivation midpoint (Vi) of the Kv1.3 currents from an average value of -53.06±0.44 mV (n=10) under control conditions to -36.05±0.48 mV (n=10) upon application of 100 µM Zn2+ (25). This may suggest, that the steady-state inactivation of Kv1.3 channels, significant at the holding potential of -60 mV in the absence of Zn2+, is abolished upon Zn2+ application. It is therefore possible that the inhibitory effects of Zn2+ and protons might be diminished or even abolished when changing the holding potential from -90 to -60 mV.

In order to proove this hypothesis, experiments using a modified sequence of valtage ramps starting from the holding potential of -60 mV were performed. Effect of the holding potential change on the inhibitory effect of Zn2+ on the currents recorded at pHo =7.35 and 6.4 is depicted in Fig. 9A and 9B, respectively (since the currents recorded at pHo =7.35 and 8.4 were not significantly different from each other, experiments at pHo =8.4 were not performed).

Fig. 9. The inhibitory effect of Zn applied at pHo=7.35 (A) and 6.4 (B) on the ramp current amplitudes reocrded at +40 mV from the holding potential of -90 mV (black bars, n=10) and -60 mV (grey bars, n=10).

Apparently, for the currents recorded from the holding potential of -60 mV at pHo =7.35 the magnitude of the inhibitory effect of Zn2+ was greatly reduced at all the concentrations used. At 50 and 100 mM the inhibitory effect of Zn2+ was fully abolished, even a small amplification of the currents was sometimes observed in the presence of 100 µM Zn2+. A small inhibitory effect was observed upon application of 300 µM Zn2+, but a significant block occurred only at the concentration of 1 mM. A similar situation occurred when Zn2+ was applied at pHo =6.4 (Fig. 9B). However, since the inhibitory effects exerted by protons and Zn2+ were additive, the degree of inhibition of the currents recorded from the holding potential of -60 mV was more pronounced than the inhibition at 7.35. It was shown that application of 100 µM Zn2+ at pHo =6.4 inhibited the current amplitude to 0.79±0.05 of the control value. The inhibition was statistically significant (p<0.05, Student t-test). This was in contrast to what was observed at 100 µM Zn2+ concentration at pHo =7.35 (compare Fig. 9A and 9B).

Finally, we examined whether the change of the holding potential affected the Zn2+- and proton - induced shift of the V1/2 value of the currents.

Fig. 10. The shift of V1/2 values caused by extracellular protons at pHo=6.4 (black bars, n=10) and Zn (grey bars, n=10) calculated for the currents recorded from the holding potentials of -90 and -60 mV.

Obtained results provide evidence that, in contrast to what was observed for the inhibitory effects, both shifts remained unaffected when the holding potential was changed from -90 to -60 mV.


DISCUSSION
Results of our study provide evidence that lowering of pHo from 7.35 to 6.4 shifts the activation midpoint of the currents by about 16 mV towards positive membrane potentials, reduces the current amplitude to about a half of the control value and slows markedly the activation kinetics. This resembles the modulatory effects exerted on the currents upon Zn application. In contrast, raising of pHo from 7.35 to 8.4 does not affect significantly the currents. These results are generally in agreement with those obtained by Deutsch and Lee, who observed similar modulatory effects on the currents at pHo below 7.2, but not above 7.2 (29). Our results shaw also that, both, the proton- and Zn-induced inhibition of the current amplitude and the shift of the activation midpoint are additive. Moreover, obtained data provide evidence that the effects exerted by Zn do not depend on the pHo in the range from 6.4 to 8.4.

These results may suggest that the modulatory effects caused by extracellular protons and Zn2+ occur independently on each other and probably involve different mechanisms. In case of protons, available data provide evidence that the effects on Kv1.3 channels are due both to the compensation of negative surface charges and to titration of the histidine at position 399 (29,30). Recently obtained data showed that mutations of His399 abolished the proton-induced slowing of the inactivation rate but did not affect significantly the proton-induced shift of the activation midpoint (30). Thus, it can be suggested that the shift of the activation midpoint and inhibition of the current amplitude is due to the compensation of negative surface charges by extracellular protons. On the other hand, our results published previously demonstrated that the modulatory effect of Zn2+ on Kv1.3 channels was not due to the surface charge compensation, but to another mechanism including binding of Zn2+ to two different sites (25). Thus, the additivity of the shift of the activation midpoint and the degree of the current inhibition by extracellular protons and Zn2+ might be due to the fact that these ions act on Kv1.3 channels by different mechanisms on different binding sites. On the other hand, it might be possible that Zn2+ and protons compete for binding on the histidine 399 and that application of Zn2+ might diminish the proton-induced slowing of the inactivation kinetics. Recently obtained data (Teisseyre - unpublished observations) showed that lowering of pHo to about 5.5 slowed the inactivation rate of the currents similarly to what was observed by other investigators (29,30). However, it remained questionable whether application of Zn2+ at this pHo affected the pH-induced slowing of the inactivation rate (Teisseyre - unpublished data). This might correspond to our results published previously (25), which showed that application of Zn2+ at 100 µM concentration did not change significantly the inactivation rate of the currents. Thus, one can suggest that Zn2+ and protons probably do not compete for the binding site at histidine 399.

Our results are in contrast to the data obtained for other Shaker-type voltage-gated potassium channels expressed in neurons and in cell line Sf9, in which the modulatory effect of Zn was significantly diminished at pHo =6.5 due to a competition between protons and Zn2+ for the same binding sites containing amino groups (5). Moreover it was shown that raising of pHo from 7.3 to 8.0 augmented the Zn2+-induced raising of the activation time constant (5), this result is also opposite to our data. In case of the Shaker-type channels extracellular protons inhibited the modulatory effect exerted by Zn2+ with the apparent pK=7.3 (5). Also the results obtained recently for Kv1.3-related Kv1.5 channels showed that Zn2+ and protons bind the same binding sites containing histidine in the channel turret (H463) and arginine near the pore mouth (R487) (31-32). These differences might indicate that the mechanism of pHo -influence on the modulatory effects exerted by Zn2+ on Kv1.3 and Kv1.5 channels is different.

Another finding, which is important in our study, is the fact that the Zn2+-induced inhibition of the current amplitude is significantly diminished when the holding potential was changed from -90 mV to -60 mV. At Zn2+ concentrations below 300 µM the inhibitory effect is completely abolished or the currents are even enhanced in the presence 100 µM Zn2+. This is likely to be due to removal of the steady-state inactivation of the channels at the potential of -60 mV, which compensates the inhibitory effects of Zn2+. The additive inhibitory effects exerted by Zn2+ and protons at pHo =6.4 are also diminished when changing the holding potential, however, less potently. A significant decrease of the current amplitude at pHo =7.35 occurs at Zn2+ concentration of 1 mM, whereas application of 100 µM Zn2+ is sufficient to significantly reduce the amplitude at pHo =6.4. In contrast, the Zn2+- and pHo -induced shift of the V1/2 parameter towards positive membrane potentials remains unchanged when changing the holding potential from -90 to -60 mV. These effects have possible physiological significance. First, because the holding potential of -60 mV is close to the resting membrane potentials of lymphocytes, which oscillates around -60 mV in case of non-stimulated cells (39,40). On the other hand, it is known that the concentration of free Zn2+ in the blood plasma is in submicromolar range, which is probably too low to exert any effects on Kv1.3 channels (41). Nevertheless, experiments performed in vitro provided evidence that application of Zn2+ at concentrations up to 200 µM stimulated the lymphocyte mitogenesis, whereas in the presence of 800 µM Zn2+ the mitogenesis was inhibited (28). Since it is known that inhibition of Kv1.3 channels inhibits the lymphocyte mitogenesis (10-13), it can be suggested that inhibition of TL mitogenesis by Zn2+ may be related to the inhibitory effect on Kv1.3 channels. Our results provided evidence that inhibition of Kv1.3 currents recorded from the holding potential of -60 mV by Zn2+ is significant at 1 mM concentration. This concentration is close to 800 µM, which is sufficient for inhibition of the mitogenesis. The stimulatory effect on lymphocyte mitogenesis exerted by Zn2+ at concentrations up to 200 µM may be related to the Zn2+ -induced shift of the activation midpoint of the channels towards positive membrane potentials. Our results demonstrate that the magnitude of this shift remained unchanged when the holding potential was changed from -90 to -60 mV. Since it is known that the resting membrane potential in TL is set primarily by the activity of Kv1.3 channels (9-13), it can be suggested that the shift of the activation midpoint towards positive membrane potentials causes the membrane depolarisation. It was shown that the lymphocyte cell membrane is depolarised from the resting value of ca. -60 mV to about -25 mV during the first 2-4 hours following mitogenic stimulation (42). This is probably one of the initial key events of lymphocyte mitogenesis (43). It is therefore possible that the stimulatory effect on lymphocyte mitogenesis exerted by Zn2+ at concentrations up to 200 µM was related to the membrane depolarisation. It should be pointed out that Kv1.3 currents recorded from the holding potential of -60 mV were not blocked at Zn2+ concentrations as high as 200 µM. More studies need to be done to elucidate the influence of Zn2+ application on the resting membrane potential in TL and to correlate the changes in the membrane potential with the cell mitogenesis.

It is also known that Kv1.3 channels are present in the central nervous system, especially in olfactory bulb neurons (14,16), where they stabilise tonic firing of action potentials (15). This part of brain is particularly rich in Zn2+ - the physiological concentrations of this ion may reach 100-300 µM (1,2). The resting potential in case of neurons is closer to -60 than to -90 mV. Thus, taking into account our results presented here and published previously (25), physiological application of Zn2+ causes primarily the shift of the activation and inactivation midpoints of Kv1.3 channels towards positive membrane potentials. Simultaneous lowering of pHo causes additionally a shift of the activation midpoint towards positive membrane potentials. Probably the inactivation midpoint is also shifted towards positive membrane potentials when pHo is lowered - more experiments will be performed in order to proove this hypothesis.

Results presented in this study were obtained at pHo = 6.4. This value is out of physiologically relevant range. However, the proton-induced shift of the activation midpoint towards positive membrane potentials was also observed at pHo = 6.8 (data not shown). The value of 6.8 was considered as physiologically relevant for studying the influence of acidification on activation of the Kv1.5 channels, which are closely related to Kv1.3 (32). Application of Zn2+ at pHo = 6.8 caused additionally a shift of the activation midpoint towards positive membrane potentials, such as it occurred at pHo = 6.4 (data not shown). Therefore, it can be suggested that the proton and Zn2+-induced shift of the Kv1.3 channel activation midpoint occurs under physiological conditions in neurons. It remains to be elucidated how such a shift modulates the electrical activity of neurons.

Acknowledgements: The Authors would like to express best thanks to our colleague from the Biophysics Department - dr Andrzej Po³a for his kind help in providing blood samples for lymphocyte isolation. This work was supported by the Polish State Committee for Scientific Research (KBN): Grant No 2 PO5A 010 27 and the Medical University Grant No 452. The experimental procedure is in accordance with the Good Medical Practice and was approved by the Committee of Bioethics at the Wroc³aw Medical University.


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