THE INFLUENCE OF MEMBRANE LIPID METABOLITES ON ...

CELLULAR & MOLECULAR BIOLOGY LETTERS

Volume 7, (2002) pp 1095 – 1109 http://www.cmbl.org.pl Received 27 September 2002 Accepted 29 November 2002

THE INFLUENCE OF MEMBRANE LIPID METABOLITES ON LYMPHOCYTE POTASSIUM CHANNEL ACTIVITY ANDRZEJ TEISSEYRE1, KRYSTYNA MICHALAK1 and MAŁGORZATA KULISZKIEWICZ-JANUS2 1 Department of Biophysics, Wrocław Medical University, Chałubińskiego 10, 50-368 Wrocław, Poland, 2Department and Clinic of Haematology and Oncology, Wrocław Medical University, Pasteura 4, 50-367 Wrocław, Poland

Abstract: In the present study, the whole-cell patch-clamp technique was applied to elucidate modulatory effects of high-density lipoproteins (HDL), sphingosine (SPH), sphingosine-1-phosphate (SPP), lysophosphatidic acid (LPA) and sphingosylphosphorylcholine (SPC) on the activity of Kv1.3 channels in human T lymphocytes (TL). Obtained data provide evidence that application of SPC at micromolar concentrations shifts the channel activation midpoint by about 20 mV towards positive membrane potentials. This effect occurs in a concentration-dependent manner and is saturated at SPC concentrations higher than 10 µM. The shift of channel activation midpoint is accompanied by a pronounced slowing of the activation kinetics. The modulatory effect of SPC is clearly voltage-dependent, being most potent at –20 mV and least potent at +60 mV. The steady-state inactivation curve is also shifted by about 20 mV towards positive membrane potentials. The kinetics of channel inactivation and deactivation (closure) remain unaffected upon SPC treatment. In contrast, application of HDL (250 µg/ml), SPH (50 and 100 µM), SPP (10 µM) and LPA (10 and 36 µM) does not exert any modulatory effect on the channel activity. The effect of SPC on Kv1.3 channel gating resembles the effect exerted by extracellular zinc at the concentration of 10 µM. It is concluded that the effect of SPC is specific and may be due to the presence of a choline residue in SPC molecules. The possible mechanism and the physiological significance of this modulatory effect on Kv1.3 channels are discussed. Key Words: Patch-Clamp, Potassium Channel, T Lymphocyte, Lipids

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INTRODUCTION Membrane lipid metabolism is important in regulation of a function of membrane receptors and in extracellular and intracellular signal transduction processes. There is also evidence for a regulatory role of membrane lipid metabolites in channel activity. Available data provide evidence that lysophosphatidic acid (LPA) activates chloride channels in fibroblasts [1]. Moreover, application of sphingosine-1phosphate (SPP) in endothelial cells promotes calcium release from intracellular stores by activating SPP-gated calcium channels in the sarcoplasmic reticulum membrane [2,3]. In many cell types SPP application stimulates calcium release from intracellular stores by acting on its specific membrane receptors [4]. Application of SPP activates acetylcholine-activated potassium channels in guinea-pig atrial cells. Also in this case SPP acts on its specific membrane receptors [4-6]. It is known that some lipid metabolites and lipoproteins significantly influence the T lymphocyte (TL) cell function. Some examples are listed below: I. High density lipoproteins (HDL). It is known that HDL receptors are expressed in TL [7]. Binding of HDL to the receptors provides transportation of some fatty acids and accelerates the cell proliferation rate [8]. II. Lysophospholipids, in particular lysophosphatidic acid (LPA), lysophoshaptidylserine (LPS) and sphingosylphosphorylcholine (SPC). Studies performed on Jurkat T cells provided evidence that LPA, LPS and SPC stimulated calcium release from intracellular stores by acting on specific membrane receptors [9]. It was also shown that LPA treatment stimulates cell proliferation [9]. III. Sphingolipids - sphingosine (SPH) and sphingosine-1-phosphate (SPP). Studies performed on Jurkat T cells demonstrated that SPH inhibited the raise in intracellular calcium concentration upon binding of the anti-CD3 monoclonal antibody [10]. Results obtained on the same cell line provided evidence that SPP inhibited the activity of caspases, and this inhibition blocked the ceramideinduced cell apoptosis. It was shown that the intracellular SPP to ceramide concentration ratio may decide about the cell life or death [11]. It is still unknown whether lipid metabolites modulate the activity of TL ion channels. TL are electrically non-excitable cells. Nevertheless, numerous studies provide evidence that many different ion channels exist in human TL, including several types of potassium channels [12-14]. The prevalent and best characterised potassium channels in TL are voltage-gated channels Kv1.3 [1214]. The channel expression is greatly diversified and is significantly increased upon mitogenic stimulation. The second type are voltage-independent, intermediate-conductance calcium-activated channels IKCa1 [14]. The channel expression is abundant only in activated TL [14]. Activity of potassium channels contributes significantly to processes such as: (i). setting the resting membrane potential, (ii). cell mitogenesis, (iii). volume regulation in TL [12-14]. It is


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known that blockage of Kv1.3 channels inhibits both the cell mitogenesis and volume regulation [12-14]. Therefore, some non-peptide Kv1.3 channel blockers have been tested as prospective immunosuppresive agents [12-14]. Taking into account the importance of potassium channel activity on the TL cell function, it was of interest to investigate the regulatory role of membrane lipid metabolites n the channel activity. In our studies, the influence of HDL, SPC, LPA, SPH and SPP on the activity of Kv1.3 channels was investigated. The results demonstrate that a modulatory effect was observed upon SPC application. The effect occurs at micromolar concentrations and is probably due to specific interactions of SPC molecules with binding sites on the channels. Other compounds did not exert any modulatory effect on the channels. Results of the present study were presented as an abstract [15] and as a part of a review article [16]. MATERIALS AND METHODS Cell separation, solutions and pipettes Human TL were separated from peripheral blood samples from 16 healthy donors using a standard method [17]. After separation, cells were cultured for at least 24 hours in the standard RPMI-1640 Medium (SIGMA) supplemented with 5% vol/vol Horse Serum (SIGMA). Upon experiment, the cells were placed in the external solution containing (in mM): 150 NaCl, 4.5 KCl, 1 CaCl2, 1 MgCl2, 10 HEPES, pH=7.35, 300 mOsm. The pipette solution contained (in mM): 70 KF, 80 KCl, 1 CaCl2, 1 MgCl2, 10 HEPES, 10 EGTA, pH=7.2, 280 mOsm. The calculated concentration of free calcium in the internal solution was 100 nM. The chemicals were provided by the Polish Chemical Company (POCH, Gliwice, Poland), except for HDL, SPH, LPA, SPHP, SPP, HEPES and EGTA, which were purchased from SIGMA. Dishes with cells were placed under an inverted Olympus IMT-2 microscope. Lipid solutions 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 MΩ. Electrophysiological recordings Whole-cell potassium currents in TL were recorded applying the patch-clamp technique [18]. The currents were recorded using an EPC-7 Amplifier (List Electronics, Darmstadt, Germany), low-pass filtered at 3 kHz, digitised using the LAB MASTER TL-1 (Axon Instruments, USA) 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 40 ms, holding potential – 90 mV. The linear (ohmic) component of the current was subtracted on-line from the final record. Inactivation kinetics was studied using another protocol containing

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7 long-lasting (1s) depolarising pulses applied every 30 s in the range from 0 to +60 mV (10 mV increment, holding potential –90 mV). Tail currents were recorded with the following protocol: cells were briefly (20 ms) depolarised to +40 mV from the holding potential of –90 mV and then repolarised to –60 mV. The linear (ohmic) component of the current was subtracted on-line by the P/4 procedure. The data are given as the mean ± standard error. All experiments were carried out at room temperature (22-24 oC). Data analysis Since the number of active channels was varying significantly among the cell population, the steady-state activation of the channels was presented in terms of a relative chord conductance (gKrel) defined by an equation: gKrel = gK/gKcontr60, where: gK – chord conductance, gKcontr60 – chord conductance recorded under control conditions at the membrane potential of +60 mV. Steady-state inactivation of the channels was presented in terms of a relative peak current (Irel) defined by an equation: Irel =I/I-120, where: I – peak current recorded at +40 mV from a given holding potential, I-120 – peak current recorded at the same potential from the holding potential of -120 mV. The chord conductance was calculated according to the definition: gK = Ip/(V-Vrev), where: Ip – amplitude of the current, V – membrane potential, Vrev – reversal potential of the current. The voltage dependence of steady-state activation was fitted by a Boltzmann function given by an equation: gKrel(V) = 1/[1+exp-(V-Vn)/kn], where - Vn activation midpoint, kn - steepness of the voltage dependence. The activation kinetics was fitted by applying a power function given by an equation: I(t) = Ip[1-exp(-t/τn)]2, where: τn - activation time constant. The voltage dependence of steady-state inactivation was fitted by a Boltzmann function as in the case of steady-state activation with the inactivation midpoint (Vi) applied instead of Vn and steepness (ki) applied instead of kn. Inactivation and deactivation kinetics were fitted by applying the single exponential function. RESULTS Fig. 1A shows an example of the whole-cell potassium currents recorded in a TL under control conditions applying a standard protocol of depolarising voltage stimuli described in Materials and Methods. The currents are evoked upon membrane depolarisation from the holding potential of –90 mV to voltages higher than –40 mV and fully activated at all potentials more positive than –20 mV. Experiments performed in our laboratory [19] provide evidence that the currents reverse at the potential near –80 mV, that is close to the estimated equilibrium potential for potassium ions under physiological conditions (-87 mV). The currents are blocked to less than 5% of the control value upon an application of 5 mM 4-AP, which selectively blocks Kv1.3 channels in human TL [19]. Moreover, the currents are not affected by 500 nM clotrimazole – a specific blocker of TL IKCa1 channels [19]. These results are in agreement with


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data obtained by other authors [12] and indicate that the recorded currents are predominantly due to an activation of Kv1.3 channels (see also Discussion).

Fig. 1. Whole cell potassium currents recorded in a TL under control conditions (A), upon an application of 10 µM SPC (B) and during wash-out (C). Currents recorded at +60 mV were off-scale.

The effect of an application of 10 µM SPC on the currents is depicted in Fig. 1B. Upon SPC application, the current at –20 mV is completely abolished and the current recorded at 0 mV is much smaller than in the control record. Also the amplitude of the current recorded at +20 mV in the presence of SPC is reduced compared to the control. This indicates that the channel activation threshold is shifted towards positive membrane potentials upon SPC application. All the currents recorded in the presence of SPC exhibit a significantly slower activation kinetics. The effect of SPC on the channel activity is reversible (see Fig. 1C). In order to present the effect of application of SPC on the channel activation in a further detail, the steady-state activation as a function of membrane potential is depicted in Fig. 2A-C for three chosen SPC concentrations. The effect of SPC application on the channel activation is clearly concentration-dependent (compare Figs 2A,B,C). The shift of the steady-state activation can be described more precisely in terms of the difference value (δVn) between the Vn calculated upon SPC application and under control conditions.

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Fig. 2. Steady-state activation of the currents in terms of a relative chord conductance (defined in Materials and Methods) vs. membrane potential upon application of: A) 1 µM SPC (n=10), B) 5 µM SPC (n=5), C) 10 µM SPC (n=10). Open squares – control conditions, filled squares – SPC application. The data were fitted by a Boltzman function (defined in Materials and Methods).

The δVn values obtained for SPC concentrations of 1,5,10 and 50 µM are: +0.53 mV (n=10), +8.51 mV (n=5), +21.41 mV (n=10), and +19.8 mV (n=4), respectively (the “+” sign stands for a shift towards positive potentials). These results provide evidence that the effect of SPC on the channel activation is concentration-dependent in the range from 1 to 10 µM and is saturated at concentrations higher than 10 µM. It is worth noting that the gKrel at very positive potentials (+40 and +60 mV) for control conditions and upon an application of 10 µM SPC are not significantly different (see Fig. 2C). This indicates that the SPC-induced shift of the steady-state inactivation is voltagedependent. As it was mentioned, application of SPC slows the activation kinetics markedly. Fig. 3A-C depicts the activation time constants (τn) as a function of membrane potential under control conditions and upon application of 1, 5 and 10 µM SPC, respectively. Fig. 3D shows the difference value (δτn) between the records obtained upon SPC application and under control conditions as a function of the membrane potential. The effect of SPC on the activation kinetics is both concentration- and voltage-dependent. It is most pronounced at 0 mV (results obtained at –20 mV were off-scale) and least pronounced at +60 mV.


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Fig. 3. Activation time constants vs. membrane potential for control conditions and upon application of: A) 1 (n=10), B) 5 (n=5), C) 10 µM SPC (n=10). Open squares – control conditions, filled squares – SPC application. D) Difference value of the activation time constants (defined in Results) upon application of: 1 (open squares, n=10), 5 (filled squares, n=5), 10 µM SPC (filled triangles, n=10). The data in A-D were fitted by a point-to-point line.

The effect of SPC application on the inactivation process was also investigated. Fig. 4A depicts the relative peak current recorded at +40 mV as a function of holding potential under control conditions and upon an application of 10 µM SPC. The steady-state inactivation curve is apparently shifted towards positive membrane potentials. The difference value of the inactivation midpoint (δVi) defined similarly to the activation midpoint, is +19.79 mV (n=4). This value is comparable to the shift of the activation. Fig. 4B depicts the effect of an application of 10 µM SPC on the inactivation kinetics. In contrast to the activation, inactivation kinetics is not changed significantly upon SPC application. The effect of SPC application on the channel deactivation (closure) was also investigated applying the tail current protocol described in Materials and M. Fig. 5 depicts the currents recorded applying the tail current protocol under control conditions and and upon an application of 10 µM SPC. It can be seen that application of SPC slows markedly the current activation kinetics upon membrane depolarisation, whereas the current amplitude is not reduced significantly. Tail currents recorded upon membrane repolarisation under control conditions and upon SPC application are not significantly different.

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Fig. 4. The influence of SPC application on the inactivation of the currents. A) Steadystate inactivation of the currents in terms of a relative peak current (defined in Materials and Methods) vs. holding potential upon an application of 10 µM SPC (n=4). Open squares – control conditions, filled squares – SPC application. The data were fitted by a Boltzman function (defined in Materials and Methods). B) Inactivation time constants vs. membrane potential upon an application of 10 µM SPC (n=4). Open squares – control conditions, filled squares – SPC application. The data were fitted by a point-topoint line.

Deactivation time constants at –60 mV (τd) are 4.69±0.89 ms (n=5), and 5.21±0.69 ms (n=5), under control conditions and upon SPC application, respectively. These values are not significantly different from each other. Thus, in contrast to the activation, the channel deactivation is not altered significantly upon SPC application.

Fig. 5. Tail currents recorded after a brief membrane depolarisation (the protocol described in detail in Materials and Methods) under control conditions (upper trace) and upon an application of 10 µM SPC (lower trace).

Results of experiments with other lipid metabolites are presented in Figs 6-9. Fig. 6A-B depicts the influence of 50 and 100 µM of sphingosine (SPH) on the


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steady-state activation and activation kinetics, respectively. The Vn values are – 15.08±0.9 mV (n=5), -7.65±2.05 mV (n=5) and –15.79±1.18 mV (n=5) for control conditions and upon an application of 50 and 100 µM of SPH, respectively. Both the activation midpoint values and the activation kinetics were not changed significantly upon SPH application (see also Fig. 6A-B).

Fig. 6. The influence of SPH application on the currents. A) Steady-state activation of the currents in terms of a relative chord conductance (defined in Materials and Methods) vs. membrane potential for control conditions (open squares, n=10) and upon an application of 50 µM SPH (filled squares, n=10), and 100 µM SPH (open triangles, n=4). The data were fitted by a Boltzman function (defined in Materials and Methods). B) Activation time constants vs. membrane potential for control conditions (open squares, n=10) and upon an application of 50 µM SPH (filled squares, n=10), and 100 µM SPH (open triangles, n=4). The data were fitted by a point-to-point line.

Fig. 7A-B shows the effect of an application of 10 µM of sphingosine-1phosphate (SPP) on the steady-state activation and activation kinetics, respectively. The steady-state activation is not shifted upon SPP application: the activation midpoint values are –23.93±3.86 mV (n=5) and –22.38±1.21 mV (n=5) for control conditions and upon an application of 10 µM SPP, respectively. Activation kinetics is not altered upon SPP application (see Fig. 7B). Fig. 8A-B depicts the effect of an application of 10 and 36 µM of lysophosphatidic acid (LPA) on the steady-state activation and activation kinetics, respectively. The steady-state activation is not affected significantly upon LPA application. The activation midpoint values are -15.23±0.88 mV (n=5), -16.58±0.49 mV (n=5) and -20.53±1.83 mV (n=5) for control conditions and upon an application of 10 and 36 µM LPA, respectively. Activation kinetics is not changed upon LPA application (see Fig. 8B). Fig. 9A-B shows the influence of high-density lipoproteins (HDL) on the steadystate activation and activation kinetics, respectively.

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Fig. 7. The influence of SPP application on the currents. A) Steady-state activation of the currents in terms of a relative chord conductance (defined in Materials and Methods) vs. membrane potential for control conditions (open squares, n=4) and upon an application of 10 µM SPP (filled squares, n=4). The data were fitted by a Boltzman function (defined in Materials and Methods). B) Activation time constants vs. membrane potential for control conditions (open squares, n=4) and upon an application of 10 µM SPP (filled squares, n=4). The data were fitted by a point-to-point line.

Fig. 8. The influence of LPA application on the currents. A) Steady-state activation of the currents in terms of a relative chord conductance (defined in Materials and Methods) vs. membrane potential for control conditions (open squares, n=10) and upon an application of 10 µM LPA (filled squares, n=10), and 36 µM LPA (open triangles, n=4). The data were fitted by a Boltzman function (defined in Materials and Methods). B) Activation time constants vs. membrane potential for control conditions (open squares, n=10) and upon an application of 10 µM LPA (filled squares, n=10), and 36 µM LPA (open triangles, n=4). The data were fitted by a point-to-point line.

The examined cells were incubated in the presence of HDL at a concentration of 250 µg/ml for at least 24 hours. The activation midpoint values are -19.7±0.72 mV (n=10) and -19.45±0.25 mV (n=10) for controls and HDL-treated cells,


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respectively. Neither the steady-state activation nor the activation kinetics were changed in the HDL-treated cells.

Fig. 9. The influence of cell incubation with HDL on the currents. A) Steady-state activation of the currents in terms of a relative chord conductance (defined in Materials and Methods) vs. membrane potential for control conditions (open squares, n=15) and in cells incubated with 250 µg/ml of HDL (filled squares, n=15). The data were fitted by a Boltzman function (defined in Materials and Methods). B) Activation time constants vs. membrane potential for control conditions (open squares, n=15) and in HDL-treated cells (filled squares, n=15). The data were fitted by a point-to-point line.

Lack of HDL effect on the channel activity was also observed upon an application of 250 µg/ml of HDL to the extracellular solution during a wholecell recording (not shown). DISCUSSION Results of our study demonstrate that the activity of potassium channels in TL is modulated by SPC at micromolar concentrations. Both the steady-state activation and inactivation curves are shifted by about +20 mV towards positive membrane potentials. This shift is accompanied by a significant slowing of the current activation rate, whereas the inactivation and deactivation rates remain unchanged upon SPC application. The modulatory effect of SPC is concentration- and voltage-dependent. Several lines of evidence support the idea that SPC affects Kv1.3 channels, but not IKCa1 channels. First, the current activation is clearly voltage-dependent. Moreover, the currents are blocked by 5 mM 4-AP and are insensitive to 500 nM clotrimazole [19]. Finally, the channel activity does not depend on intracellular free calcium concentration in the range from 100 nM to 1 µM [20]. All this properties are characteristic of Kv1.3 channels [12-14]. If the activated channels were IKCa1 channels, the currents would be voltage-independent, insensitive to 5 mM 4-AP, but sensitive to clotrimazole at nanomolar concentrations, and would be strictly dependent on intracellular calcium concentration in the

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submicromolar range [12-14]. None of such phenomena were observed in our experiments. Other lipid metabolites applied in our experiments did not exert any modulatory effect on the channel activity. HDL do not modulate the activity of Kv1.3 channels neither in the cells incubated with HDL nor in the case of an HDL application upon the whole cell recording. Application of LPA, SPH and SPP also produced no modulatory effect on the channels, although the chemical structures of SPH, SPP and SPC are similar. Thus, it can be concluded that the modulatory effect on Kv1.3 channels is not a common feature of lipid metabolites, but may be due to specific interactions of SPC molecules with binding sites on the channels. The fact that the modulatory effect of SPC is more potent at negative than at positive potentials may suggest that a positive charge might be involved in the binding. SPC molecules differ from SPH and SPP by the presence of a choline residue that contains positively charged nitrogen. This nitrogen group may be involved in the binding of SPC to a binding site. Whether this hypothesis is true, remains to be elucidated. The modulatory effect of SPC on Kv1.3 channels resembles the effect exerted by extracellular zinc [19]. Available data provide evidence that an application of zinc in a concentration range of 10-100 µM shifts both the steady-state activation and inactivation curves towards positive membrane potentials and slows markedly the activation rate of the currents [19]. In contrast, inactivation and deactivation rates remain unaffected upon zinc treatment. The shift of the activation midpoint upon an application of 10 µM Zn2+ is ca. +17.5 mV [19], which is comparable to the value obtained upon an application of 10 µM SPC (this study). However, the modulatory effect on the channel gating exerted by zinc is saturated at a higher concentration (20 µM [19]) and the shift of the steady-state activation curve in the presence of saturating zinc concentrations is more pronounced (ca. +30 mV [19]). Moreover, zinc application reduces the current amplitude at +60 mV to ca. 75% of the control value [19]. Such a reduction was not observed in the experiments with SPC. It is also worth noting that raising the zinc concentration in the range from 100 µM to 2.6 mM produces an additional inhibitory effect on the currents, probably due to an interaction with a distinct binding site [19]. The highest SPC concentration applied in our experiments was 50 µM. Thus, it is not known whether such an effect would occur upon SPC application. The mechanism of zinc interaction with potassium channels has been investigated thoroughly [21-23]. According to the model, zinc binds to a binding site when the channel is closed and stabilizes the closed and non-inactivated state by a direct interaction with the gating charge [21-23]. This may explain the shift of the steady-state activation and inactivation in the presence of zinc. Binding of zinc also prolongs the time to the first opening during the channel activation. This causes the slowing of the activation rate of the whole-cell currents. Once the channel opens, zinc ions are removed from the binding site and cannot re-bind until the channel closes [21]. Thus, the deactivation kinetics


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of the channels is not affected upon zinc application [21]. Whether this model is relevant for the modulatory effect of SPC, remains to be elucidated. The modulatory effect of SPC on Kv1.3 channels may be of physiological significance. It is known that lysophospholipids, in particular LPA, are present at micromolar concentrations in the ascitic fluid and plasma of human ovarian cancer patients and may play a role in the growth of ovarian cancer cells [9]. Ascitic fluid contains large amounts of TL [9]. Available data provide evidence that both SPC and LPA stimulate calcium release from intracellular stores in Jurkat TL when applied at a 10 µM concentration [9]. However, the effects of application of LPA and SPC on Jurkat cell proliferation are different - LPA stimulates the cell proliferation, whereas SPC inhibits it [9]. Kv1.3 channels are also present in Jurkat T cells [12-14]. Results of our study demonstrate that LPA does not affect the activity of Kv1.3 channels. Application of SPC shifts the Kv1.3 channel activation threshold towards positive membrane potentials. It is known that the resting membrane potential in quiescent TL is set primarily by the activity of Kv1.3 channels [12-14]. Thus, application of SPC causes the depolarisation of the TL cell membrane. It is known that depolarisation of the TL membrane inhibits cell proliferation [12-14]. It can therefore be suggested that the inhibition of Jurkat T cell proliferation upon SPC treatment may be due to the modulatory effect on Kv1.3 channels. It would be worth investigating whether SPC application inhibits the proliferation of freshly isolated TL. Acknowledgements. The authors would like to express their best thanks to Dr. Andrzej Poła from the Department of Biophysics, Wrocław Medical University, for his kind help in providing blood samples for lymphocyte isolation. This work was supported by Wrocław Medical University Grant No. 562. REFERENCES 1. Moolenaar, W. LPA: a novel lipid mediator with diverse biological actions. Trends Cell Biol. 4 (1994) 213-218. 2. Mattie, M., Brooker, G. and Spiegel, S. Sphingosine-1-phosphate, a putative second messenger mobilizes calcium from internal stores via an inositol triphosphate-independent pathway. J. Biol. Chem. 269 (1994) 3181-3188. 3. Kim, S., Lakhani, V., Costa, D., Sharara, A., Fitz, G., Huang, L., Peters, K. and Kindman, L. Sphingolipid-gated Ca2+ release from intracellular stores of endothelial cells is mediated by a novel Ca2+ permeable channel. J. Biol. Chem. 270 (1995) 5266-5269. 4. Koppen van, Ch., Meyer zu Heringdorf, D., Laser, K., Zhang, Ch., Jakobs, K., Bünemann, M. and Pott L. Activation of high affinity G protein-coupled plasma membrane receptor by sphingosine-1-phosphate. J. Biol. Chem. 271 (1996) 2082-2087. 5. Bünemann M., Brandts, B., Meyer zu Heringdorf, D., Koppen van, Ch., Jakobs K. and ,Pott L. Activation of muscarinic K+ current in guinea-pig

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19. Teisseyre, A. and Mozrzymas, J.W. Inhibition of the Activity of T Lymphocyte Kv1.3 Channels by Extracellular Zinc. Biochem. Pharmacol. 64 (2002) 595-607. 20. Teisseyre, A. Regulacja funkcji kanałów potasowych w limfocytach T przez wewnątrzkomórkowe wtórne przekaźniki na przykładzie wapnia, cyklicznego AMP i związków lipidowych. Post. Hig. Med. Dośw. 54 (2000) 381-390. 21. Gilly, F. and Armstrong, C.M. Divalent cations and the activation kinetics of potassium channels in squid giant axons. J. Gen. Physiol. 79 (1982) 965996. 22. Spires, S. and Begenisich, T. Chemical properties of the cation binding site on potassium channels. J. Gen. Physiol. 100 (1992) 181-193. 23. Spires, S and Begenisich, T. Modulation of potassium channel gating by external divalent cations. J. Gen. Physiol. 104 (1994) 675-692.