Ion Channels in T Lymphocytes

  • Michael D. Cahalan
  • K. George Chandy
  • Thomas E. DeCoursey
  • Sudhir Gupta
  • Richard S. Lewis
  • Jeffrey B. Sutro
Part of the Advances in Experimental Medicine and Biology book series (AEMB, volume 213)


Ion channels, proteins that gate the flux of ions across the cell membrane, control an impressive array of physiological processes, including conduction of nerve impulses, synaptic transmission, hormone secretion, generation of the heart beat, initiation of muscle contraction, and transduction of sensory stimuli. To a large extent, this diversity of functions is a reflection of the diversity of ion channel types. Several distinct channel types have been characterized in neurons using a variety of electrophysiological techniques (Table 1; see also ref. 1). In the past several years, some of the same channel types have been observed in cells outside the nervous system, raising the possibility that ion channels may serve functions in electrically non-excitable tissues that are quite distinct from their electrical activities in the nervous system. In fact, a collection of direct and indirect observations suggests that ion channels similar to those found in nerve and muscle participate in the control of biological events associated with cellular proliferation and cytokine production that are crucial to the functioning of the immune system.


Membrane Potential Channel Blocker Sodium Channel Potassium Channel Channel Expression 


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  1. 1.
    B. Hille. Ionic channels of excitable membranes. Sinauer Associates, Sunderland Mass., (1984).Google Scholar
  2. 2.
    M. H. Freedman, M. C. Raff, and B. Gomperts. Induction of increased calcium uptake in mouse T lymphocytes by concanavalin A and its modulation by cyclic nucleotides. Nature 255:378, (1978).ADSCrossRefGoogle Scholar
  3. 3.
    R. Y. Tsien, T. Pozzan, and T. J. Rink. T-cell mitogens cause early changes in cytoplasmic free Ca2+ and membrane potential in lymphocytes. Nature 295:68, (1982).ADSCrossRefGoogle Scholar
  4. 4.
    T. R. Hesketh, G. A. Smith, J. P. Moore, M. V. Taylor, and J. C. Metcalfe. Free cytoplasmic calcium concentration and the mitogenic stimulation of lymphocytes. J. Biol. Chem. 258:4876, (1983).Google Scholar
  5. 5.
    A. Weiss, J. Imboden, D. Shoback, and J. Stobo. Role of T3 surface molecules in human T cell activation: T3-dependent activation results in an increase in cytoplasmic free calcium. Proc. Natl. Acad. Sci. USA 81:4149, (1984).ADSCrossRefGoogle Scholar
  6. 6.
    E. Nisbet-Brown, R. K. Cheung, J. W. W. Lee, and E. W. Gelfand. Antigen-dependent increase cytosolic free calcium in specific human T-lymphocyte clones. Nature 316:545, (1985).ADSCrossRefGoogle Scholar
  7. 7.
    R. B. Whitney, and R. M. Sutherland. Characteristics of calcium accumulation by lymphocytes and alterations in the process induced by phytohemagglutinin. J. Cell. Physiol. 82:9, (1973).CrossRefGoogle Scholar
  8. 8.
    D. L. Birx, M. Berger, and T. A. Fleisher. The interference of T cell activation by calcium channel blocking agents. J. Immunol. 133:2904, (1974).Google Scholar
  9. 9.
    V. C. Maino, N. M. Green, and M. J. Crumpton. The role of calcium ions in initiating transformation of lymphocytes. Nature 251:324, (1974).ADSCrossRefGoogle Scholar
  10. 10.
    A. Mastro, and M. C. Smith. Calcium dependent activation of lymphocytes by ionophore A23187 and a phorbol ester tumor promoter. J. Cell. Physiol. 116:51, (1983).CrossRefGoogle Scholar
  11. 11.
    J. C. Metcalfe, T. Pozzan, G. A. Smith, and T. R. Hesketh. A calcium hypothesis for the control of cell growth. Biochem. Soc. Svmp. 45:1, (1980).Google Scholar
  12. 12.
    G. B. Segel, W. Simon, and M. A. Lichtman. Regulation of sodium and potassium transport in phytohemagglutinin-stimulated human blood lymphocytes. J. Clin-Invest. 64:834, (1979).CrossRefGoogle Scholar
  13. 13.
    P. E. R. Tatham, and P. J. Delves. Flow cytometric detection of membrane potential changes in murine lymphocytes induced by concanavalin A. Biochem. J. 221:137, (1984).Google Scholar
  14. 14.
    V. L. Lew, and H. G. Ferreira. Calcium transport and the properties of a calcium-activated potassium channel in red cell membranes. Curr. Top. Membr. Trans. 10:217, (1978).CrossRefGoogle Scholar
  15. 15.
    T. E. DeCoursey, K. G. Chandy, S. Gupta, and M. D. Cahalan. Voltage-gated K+ channels in human T lymphocytes: a role in mitogenesis? Nature 307:465, (1984).ADSCrossRefGoogle Scholar
  16. 16.
    T. E. DeCoursey, K. G. Chandy, S. Gupta, and M. D. Cahalan. Voltage-dependent ion channels in T-lymphocytes. J. Neuroimmunol. 10:71, (1985).CrossRefGoogle Scholar
  17. 17.
    L. M. Hondeghem, and B. G. Katzung. Time- and voltage-dependent interactions of antiarrhythmic drugs with cardiac sodium channels. Biochim. et Biophvs. Acta 472:373, (1977).CrossRefGoogle Scholar
  18. 18.
    O. P. Hamill, A. Marty, E. Neher, B. Sakmann, and F. J. Sigworth. Improved patch-clamp techniques for high-resolution current recording from cells and cell-free membrane patches. Pflugers Arch. 391:85, (1981).CrossRefGoogle Scholar
  19. 19.
    Y. Fukushima, S. Hagiwara, and R. E. Saxton. Variation of calcium current during the cell growth cycle in mouse hybridoma lines secreting immunoglobulins. J. Physiol. 355:313. (1984).Google Scholar
  20. 20.
    D. L. Ypey, and D. E. Clapman. Development of a delayed outward-rectifying K+ conductance in cultured mouse peritoneal macrophages. Proc. Natl. Acad. Sci. USA 81:3083, (1984).ADSCrossRefGoogle Scholar
  21. 21.
    E. K. Gallin, and P. A. Sheehy. Differential expression of inward and outward potassium currents in the macrophage-like cell line J774.1. J. Physiol. 369:475, (1985).Google Scholar
  22. 22.
    M. D. Cahalan, K. G. Chandy, T. E. DeCoursey, and S. Gupta. A voltage-gated potassium channel in human T lymphocytes. J. Physiol. 358:197, (1985).Google Scholar
  23. 23.
    K. G. Chandy, T. E. DeCoursey, M. D. Cahalan, C. McLaughlin, and S. Gupta. Voltage-gated K channels are required for T lymphocyte activation. J. Exp. Med. 160:369, (1984).CrossRefGoogle Scholar
  24. 24.
    B. Sharma and P. I. Terasaki. In vitro immunization to cultured tumor cells. Cancer Res. 34:115. (1974).Google Scholar
  25. 25.
    D. R. Matteson and C. Deutsch. K channels in T lymphocytes: a patch clamp study using monoclonal antibody adhesion. Nature 307:468, (1984).ADSCrossRefGoogle Scholar
  26. 26.
    Y. Fukushima, S. Hagiwara, and M. Henkart. Potassium current in clonal cytotoxic lymphocytes from the mouse. J. Physiol. 351:645, (1984).Google Scholar
  27. 27.
    K. S. Lee and R. W. Tsien. Mechanism of calcium channel blockade by verapamil, D600, diltiazem, and nitrendipine in single dialysed heart cells. Nature 302:790, (1983).ADSCrossRefGoogle Scholar
  28. 28.
    P. Bregestovski, A. Redkozubov, and A. Alexeev. Elevation of intracellular calcium reduces voltage-dependent potassium conductance in human T cells. Nature 319:776, (1986).ADSCrossRefGoogle Scholar
  29. 29.
    T. E. DeCoursey, K. G. Chandy, M. Fischbach, N. Talal, S. Gupta, M. D. Cahalan. Two types of K channels in T lymphocytes from MRL mice. Biophvs. J. 47:387a, (1985).Google Scholar
  30. 30.
    T. E. DeCoursey, K. G. Chandy, S. Gupta, and M. D. Cahalan. Two types of potassium channels in murine T lymphocytes. J. Gen. Physiol. (in press).Google Scholar
  31. 31.
    E. K. Gallin. Electrophysiological properties of macrophages. Fed. Proc. 43:2385, (1984).ADSGoogle Scholar
  32. 32.
    S. Ikeda and F. Weight. Inward rectifying K+ currents recorded from rat basophilic leukemic cells by whole cell patch clamp. Neurosci. Abstr. 10:870, (1984).Google Scholar
  33. 33.
    L. Schlichter, N. Sidell, and S. Hagiwara. Potassium channels mediate killing by human natural killer cells. Proc. Natl. Acad. Sci. USA 83:451, (1986).ADSCrossRefGoogle Scholar
  34. 33a.
    J. H. Russell and C. B. Dobos. The role of monovalent cations in the interaction between the cytotoxic T lymphocyte and its target. Eur. J. Immunol. 11:840, (1981).CrossRefGoogle Scholar
  35. 34.
    W. Schwartz and H. A. Kolb. Voltage-dependent kinetics of an anionic channel of large unit conductance in macrophages and myotube membranes. Pflugers Arch. 402:281, (1984).CrossRefGoogle Scholar
  36. 35.
    M. E. Krouse, G. T. Schneider, and P. W. Gage. A large anion-selective channel has seven conductance levels. Nature 319:58, (1986).ADSCrossRefGoogle Scholar
  37. 36.
    P. T. A. Gray and J. M. Ritchie. Ion channels in Schwann and glial cells. TINS 9:411, (1986).Google Scholar
  38. 37.
    M. M. Bosma. Chloride channels in neoplastic B lymphocytes. Biophvs. J. 49:413a, (9186).Google Scholar
  39. 38.
    C. Deutsch, D. Krause, and S. C. Lee. Voltage-gated potassium conductance in human T lymphocytes stimulated with phorbol ester. J. Physiol. 372:405, (1986).Google Scholar
  40. 39.
    S. Lee, D. Krause, and C. Deutsch. Increased voltage-gated K+ conductance in T-lymphocytes stimulated with phorbol ester. Biophvs. J. 47:147a, (1985).Google Scholar
  41. 40.
    T. E. DeCoursey, K. G. Chandy, M. Fischbach, N. Talal, S. Gupta, and M. D. Cahalan. Potassium channel expression in proliferating murine T lymphocytes. Fed. Proc. 44:1310, (1985).Google Scholar
  42. 41.
    T. E. DeCoursey, K. D. Chandy, S. Gupta, and M. D. Cahalan. Mitogen induction of ion channels in murine T lymphocytes. J. Gen. Physiol. (in press).Google Scholar
  43. 42.
    S. C. Lee, D. E. Sabath, C. Deutsch, and M. B. Prystowsky. Increased voltage-gated potassium conductance during interleukin 2-stimulated proliferation of a mouse helper T lymphocyte clone. J. Cell. Biol. 102:1200, (1986).CrossRefGoogle Scholar
  44. 43.
    R. S. Lewis, K. G. Chandy, S. Gupta, and M. D. Cahalan. Changes in K channel expression during the life cycle of murine T lymphocytes. Neurosci Abstr. in press, (1986).Google Scholar
  45. 44.
    K. G. Chandy, T. E. DeCoursey, M. Fischbach, N. Talal, M. D. Cahalan, and S. Gupta. Altered K+ channel expression in abnormal T lymphocytes from mice with the lpr gene mutation. Science 233:1197, (1986).ADSCrossRefGoogle Scholar
  46. 45.
    G. B. Mills, R. K. Cheung, S. Grinstein, and E. W. Gelfand. Increase in cytosolic free calcium concentration is an intracellular messenger for the production of interleukin 2 but not for the expression of the interleukin 2 receptor. J. Immunol. 134:1640, (1985).Google Scholar
  47. 46.
    E. W. Gelfand, R. W. Cheung, and S. Grinstein. Role of membrane potential in the regulation of lectin-induced calcium uptake. J. Cell. Physiol. 121:533, (1984).CrossRefGoogle Scholar
  48. 47.
    E. D. Murphy. Lymphoproliferation (lpr) and other single locus models for murine lupus. In Immunologic Defects in Laboratory Animals. M. E. Gershwin, B. Merchant eds. Plenum Press, New York, (1981) Vol 2 pp. 143.CrossRefGoogle Scholar
  49. 48.
    A. N. Theofilopoulos, R. A. Eisenberg, M. Bourdon, J. S. Crowell, and F. J. Dixon. Distribution of lymphocytes identified by surface markers in murine strains with systemic lupus erythematosus-like syndromes. J. Exp. Med. 149:516, (1979).CrossRefGoogle Scholar
  50. 49.
    D. E. Lewis, J. V. Giorgi, and N. L. Warner. Flow cytometry analysis of T cells and continuous lines from autoimmune MRL/l mice. Nature 289:298, (1981).ADSCrossRefGoogle Scholar
  51. 50.
    H. C. Morse, W. F. Davidson, R. A. Yetter, E. D. Murphy, J. B. Roths, and R. L. Coffman. Abnormalities induced by the mutant gene lpr: expansion of a unique lymphocyte subset. J. Immunol. 129:2612, (1982).Google Scholar
  52. 51.
    F. J. Dumont, R. C. Habbersett, E. A. Nichols, J. A. Treffinger, and A. S. Tung. A monoclonal antibody (100C5) to the Lyt-2- T cell population expanding in MRL/MpJ-lpr/lpr mice detects a surface antigen normally expressed on Lyt-2+ cells and B cells. Eur. J. Immunol. 13:455, (1983).CrossRefGoogle Scholar
  53. 52.
    F. J. Dumont, R. C. Habbersett, and E. A. Nichols. A new lymphocyte surface antigen defined by a monoclonal antibody (9F3) to the T cell population expanding in MRL/MpJ-lpr/lpr mice. J. Immunol. 133:809, (1984).Google Scholar
  54. 53.
    F. Takei. Unique surface phenotype of T cells in lymphoproliferative autoimmune MRL/Mp-lpr/lpr mice. J. Immunol. 133:1951, (1984).Google Scholar
  55. 54.
    H. S. Oettgen, C. Terhorst, L. C. Cantley, and P. M. Rosoff. Stimulation of the T3-T cell receptor complex induces a membrane-potential-sensitive calcium influx. Cell 40:583, (1985).CrossRefGoogle Scholar
  56. 55.
    Y. Fukushima and S. Hagiwara. Voltage-gated Ca2+ channel in mouse myeloma cells. Proc. Natl. Sci. USA 80:2240, (1983).ADSCrossRefGoogle Scholar
  57. 56.
    Y Fukushima and S. Hagiwara. Currents carried by monovalent cations through calcium channels in mouse neoplastic B lymphocytes. J. Physiol. 358:255, (1985).Google Scholar
  58. 57.
    E. K. Gallin. Calcium- and voltage-activated potassium channels in human macrophages. Biophys. J. 46:821, (1984).ADSCrossRefGoogle Scholar
  59. 58.
    D. Nelson, E. R. Jacobs, J. M. Tang, J. M. Zeller, and R. C. Bone. Immunoglobulin G induces single channels in human alveolar macrophage membranes. J. Clin. Invest. 76:500, (1985).CrossRefGoogle Scholar

Copyright information

© Plenum Press, New York 1987

Authors and Affiliations

  • Michael D. Cahalan
    • 1
  • K. George Chandy
    • 2
  • Thomas E. DeCoursey
    • 3
  • Sudhir Gupta
    • 2
  • Richard S. Lewis
    • 1
  • Jeffrey B. Sutro
    • 1
  1. 1.Department of Physiology & BiophysicsUniversity of CaliforniaIrvineUSA
  2. 2.Division of Basic and Clinical Immunology, Department of MedicineUniversity of CaliforniaIrvineUSA
  3. 3.Department of PhysiologyRush Medical CollegeChicagoUSA

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