[Ca2+]. and at -22 Mv in 5 Mm [Ba2+]0. 5

Journal of Physiology (1991), 443, pp. 175-192 175 With 8 figures Printed in Great Britain

VOLTAGE-ACTIVATED CALCIUM AND POTASSIUM CURRENTS IN HUMAN PANCREATIC fl-CELLS BY R. P. KELLY, R. SUTTON* AND F. M. ASHCROFT From the University Laboratory of Physiology, Parks Road, Oxford OXi 3PT and the *Nuffield Department of Surgery, John Radcliffe Hospital, Headington OX3 9DU (Received 26 February 1991)

SUMMARY 1. The whole-cell configuration of the patch clamp technique was used to study inward and delayed outward currents in fl-cells isolated from human pancreatic islets. 2. The delayed outward current activated at about -20 mV and increased linearly with further depolarization. The instantaneous current-voltage (I-V) relation, measured by tail current analysis, reversed at -70 mV. This is close to the K+ equilibrium potential and suggests the outward current is carried primarily by potassium ions. In support of this idea, outward currents were abolished when internal K+ was replaced by the impermeant cation N-methyl-D-glucamine (NMG). 3. The voltage dependence of K+ current activation could be fitted by a sigmoidal function with a mid-point at +1 mV. K+ currents showed voltage-dependent inactivation which was half-maximal at -25 mV. 4. Inward currents were studied after outward currents were suppressed by replacing internal potassium with NMG. In 5 mm [Ca2+]O, the inward current activated between -50 and -40 mV, had a peak amplitude at -10 mV and reversed at potentials positive to +60 mV. The voltage dependence of inward current activation was sigmoidal with half-maximal activation at -10 mV in 5 mM [Ca2+]. and at -22 mV in 5 mm [Ba2+]0. 5. Inward currents were unaffected by tetrodotoxin (TTX), but could be blocked by cadmium ions. Barium was also capable of carrying inward current. This pharmacology is consistent with inward currents flowing through Ca2+ channels. 6. The inactivation of the inward current was dependent on calcium entry. In two- pulse experiments, the voltage dependence of inactivation was U-shaped, and resembled that of the calcium current. Barium currents showed little inactivation. 7. In two-pulse experiments the degree of inward current inactivation during the test pulse was related to the amount of calcium entry during the first pulse. Calcium entering at more positive potentials was less effective at producing inactivation. 8. Calcium and barium currents also showed a slow, voltage-dependent inactivation when the holding potential was changed between - 100 and -40 mV. This inactivation developed with a time course of seconds. 9. The Ca2+ and K+ currents described here are similar to those reported for rodent fl-cells and indicate the rodent f8-cell provides a good model for that of man. MS 9183 176 R. P. KELLY, R. SUTTON AND F. M. ASHCROFT

INTRODUCTION The regulation of calcium entry is central to the control of insulin release from the pancreatic fl-cell. Calcium influx is determined by fl-cell electrical activity which consists of slow oscillations in membrane potential (slow waves) between a hyperpolarized potential and a relatively depolarized plateau potential upon which Ca2+-dependent action potentials are superimposed (see review by Henquin & Meissner, 1984). The duration of the slow waves increases with the glucose concentration until above about 15 mM-glucose electrical activity becomes continuous. There is good evidence that the action potentials are due to Ca2+ entry through voltage-dependent Ca21 channels; action potential repolarization results from activation of a delayed outward potassium current, with some possible contribution from Ca2+-activated K+ channels (see review by Ashcroft & Rorsman, 1989). The precise mechanisms which generate, sustain and repolarize the plateau potential are unclear but it is believed that the plateau is due to a Ca21 current (Cook, Porte & Crill, 1981; Meissner & Schmeer, 1981) and that the inactivation of this current may be involved in plateau repolarization (Satin & Cook, 1989). Only a few studies of fl-cell electrophysiology have been carried out using human tissue. These studies show that, as in rodent ,-cells, glucose closes an ATP-sensitive K+ channel (Ashcroft, Kakei, Gibson, Gray & Sutton, 1989b: Misler, Gee, Gillis, Scharp & Falke, 1989), thus depolarizing the fl-cell membrane and eliciting Ca2+- dependent electrical activity (Falke, Gillis, Pressel & Misler, 1989). To date, however, all data on the voltage-activated currents that underlie this electrical activity have been obtained from rodent fl-cells or fl-cell lines. We have therefore undertaken an analysis of voltage-activated currents in human f-cells to provide information on the currents responsible for the generation of slow waves, action potentials and Ca2+ entry in man. Some of these results have previously appeared in abstract form (Kelly, Sutton & Ashcroft, 1989; Ashcroft, Gray, Kelly & Sutton, 1989a).

METHODS Cells Human pancreata were removed (with permission) from normoglyeaemic, heart-beating, cadaver organ donors and islets of Langerhans isolated by collagenase digestion as described in detail elsewhere (Gray et al. 1984; Ashcroft et al. 1989b). The donors were aged between 17 and 64 and died from head injuries, cerebral or subdural haemorrhage. After overnight culture, islets were dispersed into single cells, plated onto glass cover-slips and maintained in short-term tissue culture for up to 4 days (Ashcroft, Ashcroft & Harrison, 1988; Ashcroft et al. 1989b). The secretory responses of human islets prepared in this way are similar to those of rodent islets and have been reported elsewhere (Grant, Christie & Ashcroft, 1980; Harrison, Christie & Gray, 1985). Recording conditions Standard patch clamp methods were used to record whole-cell currents (Hamill, Marty, Neher, Sakmann & Sigworth, 1981). Pipettes were pulled from 1 5 mm capillaries (Boralex, Rochester Scientific, USA), coated close to their tips with Sylgard to reduce their capacitance and fire-polished immediately before use. They had resistances between 3 and 5 MQ when filled with 140 mM-KCl. Current signals were recorded with an EPC-5 patch clamp amplifier (List Elektronic, Darmstadt, Germany) without series resistance compensation. The reference potential for all measurements Ca2+ AND K+ CURRENTS IN HUMAN /-CELLS 177 was the zero current potential of the pipette before establishment of the seal. Except where indicated, the holding potential was -70 mV. Pulse protocols were generated using a programmable stimulator (Stiihmer Elektronik, Gottingen, Germany) and pulses were applied at a frequency of 0 5 Hz or less. The data were digitized using a modified digital audio processor (Sony PCM 701ES) and stored on videotape for later analysis. Small pieces of cover-slip were placed in the recording chamber, which had a volume of 1 ml. All experiments were done at room temperature (18-22 °C). Data analysis For measurement of current amplitudes, the recorded currents were amplified, low-pass filtered (-3 dB down at 1 kHz) using an 8-pole Bessel filter (Frequency Devices, Haverhill, MA, USA), digitized at 5 kHz using a 12-bit ADC (CED 1401, Cambridge Electronic Design, UK) and analysed using the program VCAN (written by John Dempster, University of Strathclyde). Current-voltage relations were corrected for leak by linear extrapolation. of currents elicited by 30 mV hyperpolarizing steps from the holding potential: no voltage-dependent currents were active in this potential range. Data are expressed as the means +S.E.M. For the illustrations in this paper the recorded currents were amplified ( x 5 or x 10) and low-pass filtered and then digitized at 2 kHz, using a 12-bit (ADC (INDEC) controlled by a PDP 11/73 computer (Digital Equipment Corporation). Solutions For recording K+ currents, the intracellular solution contained (K+-int, mM): 107 KCI, 1 CaCl2, 11 EGTA (free Ca2+ about 0-06 /M), 11 HEPES (pH 7-2 with KOH, total K+ about 130 mM), 2 MgSO4, 3 K-ATP. For recording inward currents, the intracellular solution contained (NMG-int, mM): 145 N-methyl-D-glucamine (NMG), 120 HCI, 2 CaCl2. 10 EGTA, 5 HEPES (pH 7-2 with HCI), 4 MgCl2, 3 Na-ATP. The Ca2+ concentration was buffered to prevent Ca2+-dependent inactivation of Ca2+ currents and ATP was included to block ATP-sensitive K+ currents. The extracellular solution contained either 5 CaCl2 (Ca2+-ext) or 5 BaCl2 (Ba2+-ext) in addition to (mM): 5 KCl, 135 NaCl, 2 MgSO4, 5 HEPES (pH 7-4 with NaOH). The extracellular solution contained 10 sM- tetrodotoxin (TTX), except when testing the effect of Cd2+ as in Fig. 4. Cd2+ was applied from a puffer pipette containing 1 mM-CdCl2 positioned at a distance of about 20,um from the cell using a Picospritzer (General Valve Corporation).

RESULTS Outward currents Membrane currents elicited by depolarizing voltage steps from a holding potential of -70 mV to potentials between -60 and + 30 mV in a human pancreatic f-cell are shown in Fig. IA. At -40 mV a small inward current is elicited which is followed at more positive potentials by an increasingly large delayed outward current. The current-voltage relation for the maximal outward current elicited during a 200 ms depolarization is plotted as the open circles in Fig. 1B. Outward currents are activated at potentials positive to about -30 mV, and their amplitude and rate of activation increase with depolarization. The reversal potential of the outward current was determined from the instantaneous current-voltage relation (Fig. 1B, filled circles). This was obtained by activating the current with a constant 400 ms depolarization to 0 mV and then returning the membrane to test potentials between -30 and -90 mV. The instantaneous amplitudes of the tail current flowing on repolarization (Fig. 1 C) was plotted against the test potential. The instantaneous current was determined by fitting a single exponential to the tail currents at time points greater than 2 ms (to 178 R. P. KELLY, R. SUTTTON AND F. M. ASHCROFT avoid uncompensated capacitative transients) and extrapolating back to zero time. Figure 1C shows that the tail currents reverse at a potential between -80 and -70 mV. The mean instantaneous I-V relation crosses the zero current axis close to -70 mV (Fig. IB). This is close to the calculated reversal potential for potassium

A +30 mV +20 +0 B I(pA/pF) ~-20-10 60] 80 pA 20s40- <20 ms C 0 mV/ -50mVmV V 20-

-60 -70 mVV -70 ~~~~~~~~~-100-80 -70 -60 -40 -20 J 20 -801 Vm (MV) 10lpA / ~~6 ms -90

Fig. 1. A, whole-cell currents recorded from a single human fl-cell in response to depolarizations to between -60 and + 30 mV from a holding potential of -70 mV. B, 0, mean current-voltage relation of the maximal outward current for six cells. *, mean instantaneous current-voltage relation for the same six cells. Following a 400 ms depolarization to 0 mV the membrane was returned to test potentials between -30 and -90 mV (see inset, C). The instantaneous amplitude of the tail current is plotted against the test potential. The current amplitude was divided by the cell capacitance to allow for the differences in cell size. Mean capacitance = 6 2 + 0-8 pF. C, tail currents recorded on repolarization from 0 mV to potentials between -50 (uppermost trace) and -90 mV (lowest trace). Stimulation frequency, 0 5 Hz. Bath solution, Ca2+-ext. Pipette solution, K+-int.

(about -84 mV), suggesting that the outward current is primarily carried by K+ ions. In confirmation of this idea, outward currents were abolished when internal K+ was replaced by the impermeant cation N-methyl-D-glucamine (as in Figs 3-8). The deviation of the reversal potential from EK could be due to the presence of a junction potential (about 3 mV, Fenwick, Marty & Neher, 1982; Ashcroft, Kakei & Kelly, 1989c), or result from an incomplete selectivity of the channel for potassium (estimated permeability ratio PNa/PK= 0016, assuming no junction potential). The time constants of the K+ tail currents increased with depolarization, roughly e-fold every 40 mV, from a mean value of 8-2+0-5 ms (n = 5) at -90 mV to 37 4+0 8 ms (n = 5) at -20 mV. At -40 mV, the time constant for deactivation was 16 7 + 14 ms. Ca2+ AND K+ CURRENTS IN HUMAN ,/-CELLS 179

Inactivation of outward currents Although inactivation of the outward current was not generally observed during a 400 ms depolarization, longer depolarizations did produce inactivation. Figure 2A (inset) shows outward currents elicited at 0 mV in the absence of a pre-pulse and A O mV B l5LThmV-OmV -50 mV -70 mV -70 mV-

0 1 1

05j -70 mV 0.5- -40 mV -20 mV A 10 pAL50 ms o

0 0 -0 -60 -40 -20 -40 -20 0 20 40 Vm (mV) Vm (mV) Fig. 2. Potassium current inactivation and activation. A, normalized inactivation curve (below). A 15 s inactivating pre-pulse to different potentials was followed by a 200 ms test depolarization to 0 mV and the potential was subsequently stepped to -50 mV as shown in the protocol above. The instantaneous current amplitude at -50 mV was determined from the initial value of an exponential fitted to the tail current. Inactivation is expressed as the ratio of the instantaneous tail current at -50 mV in the presence of a pre-pulse (I) to that in its absence (Ic). The data points are drawn to eqn (1) of the text with V0.5 =- 24-7 mV and k = 7-3 mV. Inset, current elicited at 0 mV in the absence of a pre-pulse (-70 mV) and following a pre-pulse to -40 or -20 mV. B, normalized activation curve (below). The membrane was depolarized for 400 ms to potentials between -20 and +30 mV and then returned to -50 mV, as shown in the protocol above. Activation is given as the amplitude of the instantaneous tail current at -50 mV, expressed as a fraction of its maximum value. Symbols indicate different cells. The curve is drawn to eqn (1) of the text with V0.5 = 1-3 mV and k =-8-5 mV. Bath solution, Ca2+-ext. Pipette solution, K+-int. following a 15 s inactivating pre-pulse to -40 or -20 mV; a pre-pulse to -40 mV decreases the current amplitude while a pre-pulse to -20 mV inactivates the current by more than 50 %. The voltage dependence of inactivation is illustrated in Fig. 2A. The continuous curve is drawn to the function, I/IC = 1/{1+exp [(VO.6-V)/k]}, (1) where I is the amplitude of the instantaneous tail current at -50 mV in the presence of an inactivating pre-pulse and I is its amplitude in the absence of a 180 R. P. KELLY, R. SUTTON AND F. M. ASHCROFT pre-pulse. V is the voltage during the inactivating pulse, V0.5 is the potential at which inactivation is half-maximal and k is the slope factor. For each of three individual experiments the values of VO.5 and k were determined by fitting this function to the data using a non-linear least-squares routine. The mean values obtained were V.5 = -24-7+341 mV and k = 7-3+226 mV (n = 3) and were used to calculate the curve shown in Fig. 2A. Activation of outward currents Figure 2B shows the voltage-dependence of activation of the delayed outward current. For each experiment, the values of the potential at which activation is half- maximal (VO.5) and the slope factor (k) were determined using eqn (1). The mean values obtained for the activation parameters were VO.5 = 1-3+IP0 mV and k = -8-5+009 mV (n = 4) and were used to calculate the curve shown in Fig. 2B. Inward currents Inward currents were isolated by replacing internal K+ with 145 mM-N-methyl-D- glucamine (NMG). NMG has been shown to be effective in abolishing outward currents in rodent f-cells and in GH3 cells (Fernandez, Fox & Krasen, 1984; Rorsman & Trube, 1986). With outward currents blocked, depolarizing voltage steps from a holding potential of -70 mV to potentials less negative than -50 mV elicited slow inward currents (Fig. 3A). The presence of TTX in the external solution indicates these currents do not flow through Na+ channels. The inward currents increased in amplitude with depolarization, reaching a maximum at -10 mV, and then decreased in size as the potential was made more positive (Fig. 3A and B). With 5 mM [Ca2+]., the leak-corrected current-voltage relation did not cross the voltage axis at potentials negative to + 60 mV (Fig. 3 C). The inward current activated during a voltage step peaks rapidly (within 5 ms at -10 mV), and then relaxes during a sustained depolarization. The maximal current amplitude and the degree of inactivation during a 200 ms depolarization were quite variable from cell to cell, as has been reported for calcium currents in f-cells from other species (Rorsman & Trube, 1986; Plant, 1988; Satin & Cook, 1988). The voltage dependence of the peak Ca2+ permeability is given for five cells in Fig. 3D. Because of the Ca2+ gradient, which causes rectification of the single-channel current-voltage relation (Rorsman et al. 1988), it is more correct to plot Ca2+ permeability than Ca2+ conductance. The Ca2+ permeability (Pca) was determined from: PCa I /4( VF2) f ([Ca2+l -[Ca2+] exp (- 2VF/RT) (2) / (RT) ( 1-exp (-2VF/RT)) p ( where ICa is the peak Ca2+ current, V is the membrane potential, [Ca2+]i and [Ca2+]. are the intracellular and extracellular Ca2+ concentrations respectively, and R, T and F have their usual meanings. [Ca2+]i was taken as 0-06 JM. The curve in Fig. 3D is the best fit of all the data points to eqn (1); V0.5 = - 10-8 mV and k = 10 mV. The mean value of V0.5 was - 12-6 + 2-6 mV and of k was 11 5 +0 4 mV (n = 5). Figure 4 shows that the inward current is blocked by cadmium; 1 mM-Cd2+ in Ca2 -ext solution was puffed onto the cell from a distance of about 20 ,tm for 1 s. The inward current was rapidly, completely and reversibly blocked: Fig. 4C shows the Ca2+ AND K+ CURRENTS IN HUMAN /9-CELLS 181

C /Ca A -80 -40 80 Vm (mV)

-60 mV

-10 mV -6

TV-# 10 pA 50 ms D Pca/Pca,max 0fa 11 +40

0 mV

10 pA 50 ms 0 4 Vm (mV) Fig. 3. Ca2+ inward currents. A and B, currents recorded when intracellular K+ was replaced by NMG, in response to depolarizations to between -60 and -10 mV (A) and between 0 and 40 mV (B) in 10 mV steps. Holding potential -70 mV. The traces have not been corrected for leak. C, mean current-voltage relation for five cells (leak corrected); the current amplitudes from different cells were divided by the cell capacitance to compensate for differences in cell size. Mean capacitance = 4-5 + 0X6 pF. D, voltage dependence of the Ca2+ permeability obtained as described in the text. Ca2+ permeability (Pca) is expressed as a fraction of the maximum Ca2+ permeability (PC.,max) Symbols refer to five different cells. The line is drawn to eqn (1) of the text with V0.6 = - 10-8 mV and k = 10 mV. Bath solution, Ca2+-ext, 10 /SM-TTX. Pipette solution, NMG-int. A B C

10 pAl 50 ms Fig. 4. Inhibition of the inward current by Cd2 . Currents elicited by voltage steps toO mV from a holding potential of -70 mV before (A), during (B), and 8 s after (C) application of 1 mM-Cd2+ from a puffer pipette. Bath solution, Ca2+-ext. Pipette solution, NMG-int. 182 R. P. KELLY, R. SUTTON AND F. M. ASHCROFT recovery of the inward current about 8 s after ceasing cadmium application. The concentration of Cd2+ at the cell cannot accurately be estimated, but it must be considerably lower than 1 mm. From the results above, the inward currents in the human f8-cells can be identified as calcium currents by (i) the very positive reversal potential, (ii) the similarities of

A 0 mV

=70 mV

-60 mV B 1.0 h

10 pA 50 ms 0.8X

10 mV 0.6|

0.4

0.2-

1+40 mV -80 -40 0 40 80 Vm (mV)

Fig. 5. Inactivation of calcium currents. A, voltage protocol (above) and current traces (below) elicited by a 200 ms test pulse to 0 mV following 150 ms pre-pulses to -60, -10 and +40 mV. A 10 ms gap separates the two pulses. B, mean voltage dependence of inactivation (three cells). Inactivation (h) is expressed as the current amplitude (I) during a test pulse to 0 mV following a pre-pulse of variable amplitude is divided by that obtained following a pre-pulse to -60 mV (Ic) Bath solution, Ca2+-ext, 10 1tM-TTX. Pipette solution, NMG-int.

the activation properties, amplitude and time course to those of calcium currents in rodent fl-cells, (iii) the fact that they were not blocked by 10 4tM-TTX, and (iv) their sensitivity to Cd2+. Inactivation of calcium currents The amplitude of the inward current during a 200 ms pulse was always seen to decline when Ca2+ was the current carrier, although the extent of this inactivation varied significantly from cell to cell. That the current relaxation is due to inward Ca2+ AND K+ CURRENTS IN HUMAN /-CELLS 183 current inactivation rather than to the development of an outward current, is indicated by the fact that it occurs even when internal potassium is replaced with N- methyl-D-glucamine, and that, for currents of similar amplitude, inactivation is less at more positive potentials where K+ currents would be expected to be greater. Calcium-activated K+ currents might be expected to show a similar voltage- dependence to the Ca2` current inactivation shown here, but these currents should be abolished when K+ is replaced by NMG. Activation of calcium-dependent chloride channels cannot contribute to the decline of the inward current significantly since inactivation is maximal at - 10 mV, which is close to the chloride reversal potential. Inactivation was most pronounced at potentials close to - 10 mV and was less for weaker or for stronger depolarizations (Fig. 3A and B). This resembles the voltage- dependence of the Ca2+ current, which peaks at -10 mV and is smaller at potentials approaching Eca (Fig. 3C). The correlation between the amplitude of the Ca2+ current and the degree of inactivation suggests that Ca2+ current inactivation in human f8-cells, like that in rodent f8-cells (Plant, 1988), may be dependent on Ca2+ entry. The contributions of potential and calcium entry to inactivation of the calcium current were investigated using a two-pulse voltage protocol (Eckert & Tillotson, 1981), as shown in Fig. 5A. A constant test pulse to 0 mV from a holding potential of -70 mV was preceded by a 150 ms test pulse to potentials between -60 and + 40 mV. A gap of 10 ms between the inactivating and test pulses allowed activation to return to the steady state (Gillespie & Meves, 1980; Plant, 1988). Figure 5A shows how the current during the test pulse is affected by pre-pulses to -60, -10 and + 40 mV. With a pre-pulse to -60 mV virtually no inward current is activated, and the current during the test pulse is large. When the current during the pre-pulse is large (as during the pre-pulse to - 10 mV), the amplitude of the test current is greatly reduced. However, the test pulse current increases again in amplitude following a pre-pulse to +40 mV, which elicits little inward current. The relationship between the amplitude of the test current and the pre-pulse potential was investigated in three cells. The degree of inactivation is plotted as a function of pre-pulse potential in Fig. 5B. Inactivation is initiated at potentials close to those at which ICa is activated, peaks at - 10 mV (where the Ca2+ current is largest), and decreases again as the potential of the pre-pulse approaches ECa. The restoration of the current amplitude at positive potentials indicates that a 150 ms depolarization is not effective per se in inactivating the calcium current. Rather, the correlation between the voltage dependence of the Ca2+ current and its inactivation suggest that calcium current inactivation is primarily mediated by depolarization- induced calcium entry. Figure 6A shows calcium currents elicited by pulses to -30 mV and to + 10 mV. Although the amplitudes of the two currents are very similar, the current at -30 mV inactivates much more rapidly than that at +10 mV: the relaxing portions of the current traces can be well fitted by single exponentials with time constants of 32 ms (-30 mV) and 113 ms (+ 10 mV). This suggests that Ca2+ entering at the more negative potential is more effective in producing inactivation. To examine this point more fully, we measured the relation between the extent of inactivation and the amount of Ca2+ entry during the pre-pulse (Fig. 6B). Charge 184 R. P. KELLY, R. SUTTON AND F. M. ASHCROFT entry was calculated by integrating the current during the pre-pulse (after correcting for a linear leak), and the amount of Ca2+ entry was obtained by dividing the charge by zF, where z is the charge on the ion (+ 2), and F is the Faraday constant. Surprisingly, the extent of inactivation of the test current depends not only on the

B 1.0- -60 A N/Ic -50 0.8- - = ~~~~~~-30mV\ +30 0.6- -40

0.4- 1 50 pA -0 100 0.2- L 50 ms

0 0 10 20 Ca2+ entry (mol x 10-18) Fig. 6. The effect of potential on Ca2+-dependent inactivation. A, different time courses of Ca2+ current inactivation elicited by potential steps to -30 and + 10 mV. The traces are from the same cell and are not leak corrected: after leak correction the two currents are of almost identical amplitude. B, Ca2+ current inactivation as a function of Ca2+ entry for one cell. The degree of inactivation (estimated as in Fig. 5) is plotted against the total Ca2+ entry during the pre-pulse. The latter was calculated as described in the text. The membrane potential during the pre-pulse is indicated next to each point. Bath solution, Ca2+-ext, 10 ,uM-TTX. Pipette solution, NMG-int. amount of calcium entry, but also on the potential at which that calcium entry occurred. Specifically, calcium entry during a more depolarized pre-pulse is less effective at producing inactivation than (a similar amount of) Ca2+ influx at a more negative potential. The effects of potential on Ca2+ current inactivation shown in Figs 5 and 6 (i.e. depolarization appearing to decrease inactivation of the Ca2+ current) are inconsistent with a Hodgkin-Huxley mechanism of voltage-dependent inactivation but provide support for a mechanism of inactivation based on Ca2+ entry. Effects of barium as the charge carrier We next investigated inward current inactivation using barium as a charge carrier, as this ion has been shown to produce no inactivation of calcium currents (Eckert & Chad, 1984; Plant, 1988). Figure 7 shows results of a two-pulse inactivation experiment in 5 mM-Ba2+ solution (Ba2+-ext). Identical results were obtained in two other cells. Barium currents show very little inactivation during the course of a 200 ms depolarization Ca2+ AND K+ CURRENTS IN HUMAN /-CELLS 185 and the size of the current during the pre-pulse has little effect on the amplitude of the test current (Fig. 7A). The current-voltage relation for the current during the pre-pulse is plotted as the open circles in Fig. 7B; the filled circles indicate the extent ofinactivation of the current during the test pulse. Because there is a significant Ba2+

A B

Vm (mV) c

PBa/PBa, ma . +410 mV 01.5-

1. rwIpvvo*mA-.

400 -40. 40o 0 Vm (mV)

Fig. 7. Voltage dependence of activation and inactivation in 5 mM-Ba2+ solution, A, Ba2+ currents elicited at 0 mV following pre-pulses to -60, 0 and + 40 mV (pulse protocol as in Fig. 5). B, 0, the relationship between the pre-pulse potential and the amplitude of the Ba2+ current during the pre-pulse, i.e. the voltage dependence of the Ba2+ current. 0, the relationship between the pre-pulse potential and the Ba2+ current amplitude during the test pulse, a measure of the voltage dependence of Ba2+ current inactivation. C, voltage dependence of the Ba2+ permeability. Ba2+ permeability (PB.) is expressed as a fraction of the maximum Ba2+ permeability (PB,.max) The line is drawn to eqn (1) of the text with V0.5 = -22 mV and k = 10 mV. Bath solution, Ba2+-ext, 10 /LM-TTX. Pipette solution, N'MG-int. current at -60 mV, we have not attempted to correct for run-down of inward current as described for Ca2+ currents (see legend, Fig. 5B); however, this is unlikely to cause an error as there was almost no run-down of Ba2+ currents during the course of these experiments. 186 R. P. KELLY, R. SUTTON AND F. M. ASHCROFT In 5 mM-Ba2+ solution, the current-voltage relation was shifted by between 5 and 10 mV to more negative potentials; barium currents were elicited at -60 mV and were maximal at -20 mV. This shift in the I-V relation is probably due to an effect on surface potential (Frankenhaeuser & Hodgkin, 1957), with Ba2' being less

A -40- Vm(mV) 6] 0

/Ca (pA) 1[0pA -100- C

-20

-30- 10 pAL20 ms 0 60 120 180 Time (s) Fig. 8. Voltage-dependent inactivation of calcium and barium currents. A, effect of changing the holding potential from -100 to -70 and -40 mV (above) on the maximum amplitude ofthe calcium current elicited by steps toO mV (below). B, Ca2+ currents elicited by depolarization to 0 mV from holding potentials of -100, -70 and -40 mV. Bath solution, Ca2+-ext, 10 ,uM-TTX. Pipette solution, NMG-int. C, Baa2 currents elicited by depolarization to 0 mV from holding potentials of -100, -70 and -40 mV. The Ba2+ current baseline shows an inward shift at a holding potential of -40 mV, due to a sustained inward current which has not inactivated after 30 s. Bath solution, Ba2+-ext, 10 /LM-TTX. Pipette solution, NMG-int. effective than Ca2+ at screening negative membrane surface charge. A similar shift is found for the activation curve (Fig. 7 C), which has a mid-point of -22 mV in 5 mM- Ba2+ compared with one of - 10-8 mV in 5 mM-calcium. These values are similar to those reported for rodent fl-cells (about -27 mV in 5 mM-Ba2+; P. A. Smith, C. M. S. Fewtrell & F. M. Ashcroft, unpublished observations). The lack of Ba2+ current inactivation further supports the view that the inactivation of Ca2+ currents seen in Fig. 5 is dependent on Ca2+ entry. It also indicates that there is no potential-dependent inactivation on a millisecond time- scale in human fl-cells. Voltage-dependent inactivation Although no evidence for voltage-dependent inactivation of calcium currents was apparent during a 200 ms pulse, a slow inactivating effect of depolarization was found by changing the holding potential. Figure 8A illustrates the effect of holding potential on the Ca21 current amplitude at 0 mV. Representative calcium current Ca2+ AND K+ CURRENTS IN HUMAN /1-CELLS 187 traces are shown in 8B. The currents elicited at 0 mV are largest when the holding potential is -100 mV, are partially inactivated at -70 mV and greatly (> 80 %) inactivated at a holding potential of -40 mV. These effects of holding potential develop on a time scale of seconds. The decrease in the Ca2+ current amplitude is also accompanied by a reduction in the rate and extent of inactivation. The decrease in the peak calcium current amplitude at more depolarized holding potentials could either be due to increased steady-state calcium-dependent inactivation, caused by Ca2+ entry through Ca2+ channels open at more depolarized potentials, or to a genuine inactivating effect of voltage. To distinguish between these possibilities, the experiment was repeated in 5 mM-Ba2+, where the effects of potential may be seen in isolation. Figure 8C shows barium currents elicited at 0 mV with holding potentials of -100, -70 and -40 mV. The current evoked from a holding potential of -100 mV shows a transient component which is not observed with more depolarized holding potentials. This component may be a T-type calcium current similar to that reported in rat pancreatic f-cells (Ashcroft, Kelly & Smith, 1990) but was not further investigated. The sustained component of the barium current is similar to L-type currents in mouse f-cells described elsewhere (Rorsman et al. 1988). It is unaltered in amplitude between -100 and -70 mV, but is almost totally inactivated at a holding potential of -40 mV. These results suggest that voltage-dependent inactivation contributes to the slow inactivation of inward currents at -40 mV. Figure 8A also indicates that calcium currents in the f8-cell are subject to run- down, as is common for L-type calcium currents in other cell types (Fenwick et al. 1982; Belles, Malecot, Hescheler & Trautwein, 1988). Run-down is present even though the intracellular solution contains ATP and EGTA and it appears to be accelerated by more depolarized holding potentials (Fig. 8B).

DISCUSSION Although most recordings of fl-cell electrical activity have been obtained from mouse fl-cells (Ashcroft & Rorsman, 1990), similar electrical activity has recently been found in man (Falke et al. 1989). In the following discussion we therefore assume that the electrical activity of mouse f8-cells is typical of human fl-cells.

Potassium currents The properties of the delayed outward K+ current we describe are similar to those described in mouse fl-cells (Rorsman & Trube, 1986). The greatest differences are in the potential dependence of activation and inactivation which are shifted to more positive potentials in our experiments; it is likely that this results from the higher extracellular Ca2+ concentration. Activation of the delayed outward K+ current is probably responsible for the repolarization of the action potential. The peak of the action potential is normally around -10 mV. In our solutions only about 20 % of the maximum outward conductance is activated at this potential. A considerably greater fraction, however, is likely to be activated under physiological conditions because of the reduced Ca2+ 188 R. P. KELLY, R. SUTTON AND F. M. ASHCROFT concentration. For example, 50 % of the maximum conductance would be activated at -10 mV if there were a surface potential shift of only 10 mV (which is not unreasonable) between, our solution and one containing 2-6 mM-Ca2±. It seems probable that the outward K+ current is also involved in the plateau potential of the f-cell (see also Ashcroft & Rorsman, 1989). At this potential ( -40 mV), outward currents are not significantly activated, even allowing for surface potential shifts; the time constant of deactivation of the outward current is also slow (- 20 ms). This suggests that the slow decay of the outward current will produce a gradual depolarization and so ultimately trigger a new action potential. In other words, the 'pacemaker potential' between successive spikes results from deactivation of the outward current. The plateau potential is thus determined by the balance between the declining K+ current and the maintained Ca2+ current. The voltage-dependence of the delayed outward K+ current argues against a role for this current in burst repolarization. Voltage-dependent inactivation of outward currents may be of physiological significance during sustained depolarizations, such as those found during the plateau of the slow wave and the continuous depolarization produced by high glucose concentrations (> 16 mM). This inactivation may partly explain the observation that the action potential duration increases throughout the burst. Calcium currents The amplitude, activation properties and time course of the inward current in human fl-cells are similar to those of calcium currents measured in rodent f-cells (Rorsman & Trube, 1986; Rorsman & Hellman, 1988; Plant, 1988; Hiriart & Matteson, 1988). Our results suggest that Ca21 current inactivation comprises both fast and slow components, as described for HIT T15,8-cells (Satin & Cook, 1989). In this section we first discuss the properties of fast and slow inactivation and then consider the role of these processes in shaping fl-cell electrical activity. Fast inactivation We have defined fast inactivation as that which occurs during a short (200 ms) depolarization. Evidence supporting the view that fast inactivation of calcium currents in human fl-cells is primarily calcium-dependent can be summarized as follows: (1) the rate and extent of inactivation vary with the amplitude of the Ca2+ current; (2) in two-pulse experiments, the voltage-dependence of inactivation is U- shaped and parallels that ofthe Ca2+ current itself; (3) inactivation is greatly reduced in barium solutions. The conclusion that fast inactivation is primarily calcium- dependent is in agreement with Plant (1988) who demonstrated calcium-dependent inactivation of calcium currents in mouse fl-cells. It has been argued that if inactivation is purely calcium-dependent, the inactivation of the test current in a two-pulse protocol should dependent only on the quantity of calcium entering during the first pulse and not on the potential at which it entered (Plant, 1988). However, in our experiments Ca2+ current inactivation during the test pulse was determined by both the amount of Ca2+ entry during the pre-pulse and by the pre-pulse potential itself. That is, the same amount of Ca2+ entry at a more negative pre-pulse potential produced greater inactivation of the test Ca2+ AND K+ CURRENTS IN HUMAN /-CELLS 189 pulse Ca2+ current. Neither a potential-dependent component of inactivation nor an underestimation of Ca2+ entry at more positive potentials (due to a non-linear leak), can account for this effect; indeed, both would act to produce an apparent increase in inactivation. One possible interpretation of our results is that hyperpolarization enhances the sensitivity of the Ca21 channel to calcium. An alternative explanation is provided by consideration of the domain theory of calcium entry. If the elevation of cytosolic free Ca21 is localized to small 'domains' centered on active Ca21 channels (as proposed in the models of Eckert & Chad, 1984; Simon & Llina's, 1985; Sherman, Keizer & Rinzel, 1990), a small depolarization will open a few channels, producing localized regions of high [Ca2+]i. A large depolarization, producing the same macroscopic Ca2+ current, will activate more Ca2+ channels but the single-channel currents will be smaller because of the reduced driving force and the rectification of the I-V relation (Rorsman et al. 1988). Thus the stronger depolarization will produce a more diffuse distribution of calcium and a lower concentration at the intracellular mouth of each Ca2+ channel. This will result in a reduced rate and extent of inactivation at the more positive potential as is seen in our experiments. It can also be argued that the 'domains' are not completely isolated from each other. This is suggested by examination of Ca2+ current inactivation during a voltage step to 0 mV from different holding potentials. As the holding potential is made more positive, the number of channels available for activation decreases, producing a reduction in the whole-cell Ca2+ current. Since the single-channel current, and thus the amount of calcium entering through each channel, remains constant there should be no change in the rate of inactivation if the 'domains' are completely isolated. However, Ca2+ current inactivation is reduced with positive holding potentials. This suggests that during depolarizations from more negative holding potentials the calcium 'domains' may overlap and produce a higher local Ca2+ concentration. Our results do not allow us to draw any conclusion about the way in which Ca2+ interacts with the Ca2+ channel to produce inactivation. One possibility is that Ca2+ interacts directly with the channel protein. It has also been proposed that Ca2+ activates a phosphatase which dephosphorylates the Ca2+ channel and thereby causes it to enter the inactivated state (Eckert & Chad, 1984), since calcineurin, a calcium and calmodulin-dependent phosphatase, accelerates inactivation of Ca2+ currents in molluscan neurons in a Ca2+-dependent manner (Chad & Eckert, 1986). If this were also the case in f-cells, an attractive explanation of our results would be that neighbouring Ca2+ domains function independently on a short time scale (as in a 150 ms pulse) but interact on a slow time scale due to the diffusion of the activated phosphatase (as when the holding potential is altered). Finally, it must not be forgotten that in the intact cell the degree of calcium- dependent inactivation will largely be determined by the rate and capacity of intracellular Ca2+ buffering. Slow inactivation Although inactivation of calcium currents during a 200 ms pulse seems to be almost entirely calcium dependent, a slow voltage-dependent component of inactivation may also be present. Evidence in favour of this idea is the dramatic 190 R. P. KELLY, R. SUTTTON, AND F. M. ASHCROFT decrease in the amplitude of Ba2" currents produced by a holding potential of -40 mV (Fig. 8; see also Satin & Cook, 1989). This decrease occurs slowly, taking many seconds to reach a steady state. A similar slow inactivation is seen when Ca2+ is the charge carrier and presumably consists of both Ca2+- and voltage-dependent components. Interestingly, unlike Ba2+ currents, a reduction in Ca2+ current amplitude was also found when the holding potential was changed from -100 to -70 mV. We attribute this to a small amount of Ca2+ entry at potentials between - 100 and -70 mV; single-channel recordings have demonstrated Ca2" channel activity at these potentials in mouse 8-cells (P. A. Smith, C. M. S. Fewtrell & F. M. Ashcroft, unpublished observations). Our results contrast with those of Plant (1988) who found that changing the holding potential between - 100 and -50 mV had no effect on calcium current amplitude in the mouse f-cell. We have no explanation for this difference. Physiological roles The calcium current seems sufficient to account for the upstroke of the action potential, as it does in rodent fl-cells (Rorsman & Trube, 1986; Ashcroft & Rorsman, 1990). Inward currents activate between -50 and -60 mV in our solutions and therefore presumably about 10 mV more negative at physiological Ca2+ levels. This voltage dependence argues that the Ca2+ current also contributes to the plateau potential, which has a threshold of around -50 mV. This is in agreement with the observations that the slow waves are abolished in Ca2+-free solution (Meissner & Schmeer, 1981) or in the presence of Ca2+ channel blockers (Meissner & Preissler, 1979). Thus we hypothesize that activation of the inward current both initiates and maintains the burst. At the plateau potential, the Ca2+ current inactivates very slowly on a time scale of seconds; indeed, significant current remains even after 20 s. This suggests that a maintained Ca2+ current underlies the plateau and that the duration of the plateau will be influenced by the rate of Ca2+ current inactivation (see also Satin & Cook, 1989; Ashcroft & Rorsman, 1989). Likewise, the decrease in the amplitude and frequency of the Ca2+ spikes during the plateau (Henquin & Meissner, 1984) can be ascribed to the combined effects of calcium-mediated and voltage- dependent inactivation. The currents reported here resemble those present in rodent fl-cells, confirming that the rodent fl-cell is an acceptable model for that of man. A similar correspondence has also been found for the K-ATP channel (Ashcroft et al. 1989b; Misler et al. 1989). These findings are particularly important given the limited availability of human tissue.

We thank the Wellcome Trust, the Novo-Nordisk UK Fund, the Medical Research Council, the Juvenile Diabetes Foundation International and the British Diabetic Association for financial support (to F. M. A. and Dr D. Gray, Nuffield Department of Surgery). F. M. A. was a Royal Society 1983 University Research Fellow. Ca2+ AND K+ CUTRRENTS IN HUMlAN /1-CELLS 191

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