In Paramecium tetraurelia, polyamine-triggered exocytosis is accompanied by the activation of Ca2+-activated currents across the cell membrane (Erxleben, C., and H. Plattner. 1994. J. Cell Biol. 127:935– 945). We now show by voltage clamp and extracellular recordings that the product of current × time (As) closely parallels the number of exocytotic events. We suggest that Ca2+ mobilization from subplasmalemmal storage compartments, covering almost the entire cell surface, is a key event. In fact, after local stimulation, Ca2+ imaging with high time resolution reveals rapid, transient, local signals even when extracellular Ca2+ is quenched to or below resting intracellular Ca2+ concentration ([Ca2+]e ⩽ [Ca2+]i). Under these conditions, quenched-flow/freeze-fracture analysis shows that membrane fusion is only partially inhibited. Increasing [Ca2+]e alone, i.e., without secretagogue, causes rapid, strong cortical increase of [Ca2+]i but no exocytosis. In various cells, the ratio of maximal vs. minimal currents registered during maximal stimulation or single exocytotic events, respectively, correlate nicely with the number of Ca stores available. Since no quantal current steps could be observed, this is again compatible with the combined occurrence of Ca2+ mobilization from stores (providing close to threshold Ca2+ levels) and Ca2+ influx from the medium (which per se does not cause exocytosis). This implies that only the combination of Ca2+ flushes, primarily from internal and secondarily from external sources, can produce a signal triggering rapid, local exocytotic responses, as requested for Paramecium defense.
In most systems analyzed so far, exocytosis is triggered by the increase of intracellular free Ca2+ concentration ([Ca2+]i)1. In fast responding systems such as motor endplates, this increase occurs through an influx of extracellular Ca2+ (Ca2+e), via voltage-dependent Ca2+ channels at active zones where neurotransmitter vesicles are docked. In other systems, Ca2+ is mobilized exclusively or additionally from internal stores (Burgoyne and Morgan, 1993; Cheek and Barry, 1993; Pozzan et al., 1994).
Recent views have emphasized the possible primary importance of subplasmalemmal Ca stores due to their structural and functional coupling with the cell membrane (Berridge, 1995). In most systems, however, such stores are difficult to identify, as is their structural relationship with secretory organelles. The latter relationship is important, considering the rapid decay of local [Ca2+]i increases taking place with space and time (Llinás et al., 1992; Neher and Augustine, 1992; Zucker, 1993). Clearly, therefore, a secretory system operating under defined spatio–temporal conditions offers advantages for analyzing the role of subplasmalemmal Ca stores in regulation of exocytosis. Paramecium tetraurelia cells can be such a system. In fact, each Paramecium cell contains numerous secretory organelles, or trichocysts, attached at the cell membrane, ready for immediate release (Plattner et al., 1991). Single or a few trichocysts are discharged spontaneously, or upon slight irritation of the cell (Haacke-Bell et al., 1990). The other extreme is the synchronous (within <1 s) release of most of the >103 docked trichocysts in response to aminoethyldextran (AED) (Plattner et al., 1984, 1985; Knoll et al., 1991a). Such a massive trichocyst exocytosis can be restricted to discrete sites of the cell surface following either local application of AED (Plattner et al., 1984), or contact with predatory ciliates, whereby it serves a defensive function (Knoll et al., 1991b). With AED, signal transduction is restricted to the somatic (nonciliary) cell membrane (Plattner et al., 1984, 1991; Erxleben and Plattner, 1994).
A Paramecium cell displays an egg case type surface relief with unit fields, kinetids, which are associated with as many alveolar sacs. Alveolar sacs, which are known to be Ca stores (Stelly et al., 1991; Länge et al., 1995), are tightly attached to the cell membrane, with each trichocyst positioned at the edge of four adjacent sacs (Allen, 1988). During AED-triggered exocytosis (Knoll et al., 1993; Stelly et al., 1995) the sacs are rapidly mobilized, with ensuing rapid increase in subplasmalemmal [Ca2+]i (Knoll et al., 1993) and activation of Ca2+-dependent currents whose size roughly correlates with the extent of AED stimulation (Erxleben and Plattner, 1994). Moreover, under these conditions mobilization of Ca2+ from subplasmalemmal stores is accompanied by an influx of Ca2+e (Kerboeuf and Cohen, 1990; Knoll et al., 1992; Erxleben and Plattner, 1994).
In the present study, a more precise correlation between electrophysiological and morphometric data has been carried out by using [Ca2+]-fluorochrome analysis and rapid confocal laser scanning microscopy (CLSM), with quenched- flow/freeze-fracture to reveal membrane fusion events. Our goal was to analyze the origins of subplasmalemmal Ca2+ signals and their importance for exocytotic events.
Materials And Methods
Paramecium tetraurelia wild-type (7S) cells were cultivated monoxenically to stationary phase, with Enterobacter aerogenes added, as previously described (Plattner et al., 1984, 1985), using a medium supplemented with stigmasterol (Sigma Chemical Co., St. Louis, MO), 5 mg/liter, and [Ca2+] of 0.1 mM.
The method applied for voltage clamp and for extracellular current recordings as well as the solutions used were as described previously (Erxleben and Plattner, 1994). Briefly, the anterior pole of a cell was sucked into a holding pipette (covering ∼1/2 of the cell surface). Spontaneous currents were registered after cells were impaled by the microelectrode or, for extracellular recordings, after immobilization in the holding pipette. Spontaneous current fluctuations were registered in parallel to the occurrence of single or several secretory events. Since the number of truly spontaneous current events was very variable and usually quite low (in the order of 1/1–5 min), cells were also triggered to different extents by local pressure application of 0.01% AED (Plattner et al., 1984, 1985) from a pipette positioned about two cell lengths away. Pressure and duration of the pulse were adjusted such that only a small number of trichocysts were released. Pressure application of solution without AED evoked neither currents nor exocytosis.
Secretagogue-induced currents were recorded under voltage clamp conditions or as extracellular currents, as described previously (Erxleben and Plattner, 1994). Discharge of trichocysts was observed under a microscope with 40× phase contrast water-immersion optics and recorded with a CCD camera. To establish the temporal relationship between the currents elicited by AED and the discharge of trichocysts as observed under the microscope, AED-induced currents were displayed on an oscilloscope, from which they were recorded by a second CCD camera. The video signals of both cameras, one attached to the microscope and the other recording the oscilloscope traces, were combined by a digital video mixer (WJ-AVE5; Panasonic, Osaka, Japan). The time resolution for the observation camera was 20 ms. For analysis, half frames of the combined video signal were analyzed on a monitor. For processing of figures, the video signal was digitized with a frame grabber (miroVideo D1; MicroComputer products AG, Braunschweig, Germany).
Estimation of Cell Surface Area and of the Number of Subplasmalemmal Ca Stores
Cell size was estimated from pictures taken with a phase contrast microscope, 40× objective, including a grating scale in samples. On prints of 1,125× magnification, median views of cells were subdivided into 5 mm discs. Cell surface area and cell volume were computed assuming rotational symmetry. Cilia and crenulation of the somatic surface area could be neglected for the following reasons: (a) Cilia do not contribute to Ca2+ fluxes during AED-triggered exocytosis (Plattner et al., 1984; Erxleben and Plattner, 1994). (b) Crenulation may cause only <20% surface increase. (c) Our main goal was to estimate the number of alveolar sacs per cell reflected by the number of surface fields, or kinetids, per cell.
The average area of a kinetid was determined at the light and electron microscope level. On LM micrographs (100× oil immersion lens and grating scale included in samples), kinetid sizes were evaluated at 2,130× magnification. The same was done on EM micrographs obtained as follows. Cells sandwiched between thin copper sheets were injected into melting propane, freeze-fractured, and Pt/C replicated in a BAF300 freeze-fracture device from Balzers S.P.A. (Balzers, Liechtenstein). Replicas were evaluated at EM magnifications controlled by latex particles of defined size (Serva, Heidelberg, Germany) and reproduced at 11.400× magnification. Longitudinal and perpendicular dimensions of kinetids were computed from LM and EM micrographs by evaluating as many periods in strictly vertical view from as many cells as possible.
Quenched-Flow/Freeze-Fracture Analysis with Normal or Reduced Extracellular Ca2+
Cells were processed in a quenched-flow device according to Knoll et al. (1991a) and Plattner et al. (1994). The medium contained 0.1 mM Ca2+. Cells were mixed with either equal parts of culture medium (negative control) or with 0.02% AED (positive control). Aliquots were mixed with equal parts of 9 mM EGTA for 200 ms and then with two parts of 0.02% AED for an additional 80 ms. After mixing in the apparatus, samples were frozen in melting propane for subsequent freeze-fracture. Evaluation of trichocyst docking sites, with or without exocytotic membrane fusion, was as described previously (Knoll et al., 1991a; Plattner et al., 1994).
Ca2+ Imaging by Conventional Microscopy or CLSM
Experiments were conducted with single cells bathed in a defined microdroplet of medium containing 0.1 mM Ca2+. Cells were microinjected with the following fluorochromes: 100 μM calcium green-2 (CaGr-2), 100 μM Fluo-3, or 50 μM Fura red (final concentrations in cells after injection of 10% of the cell volume size; Table I). Fluorochromes, all obtained from Molecular Probes (Eugene, OR), were allowed to spread for 2 min to reach the outermost cell periphery.
In some experiments, cells preloaded with Fura red were flushed with high [Ca2+]e and quantitatively analyzed for cortical [Ca2+] transients and exocytotic response. In more expansive experiments, [Ca2+]e was reduced by superfusion (for up to 1 s, from a distance of ∼10 μm, in a direction tangential to the cell surface) with a local flush, using a pipette (2 μm inner diameter) filled with 10 mM EGTA and 50 μM fluorescein (Sigma Chemical Co.) using a home made device operated at 2.5 kPa. By the calibration of the fluorescein signal, the EGTA concentration at the discrete site of the cell surface was estimated to be ∼5 mM, corresponding to [Ca2+] resting levels inside the cell (50–100 nM). Overall [Ca2+]e was determined separately by addition of a calibrated Fura red solution to the microdroplet. At the end of an EGTA flush, AED was applied to the same site using a second pipette close to the EGTA pipette but vertical to the cell surface. To control the propagation of AED and the concentration sensed by the cell surface with this set up, fluorescein was sometimes also added to the secretagogue.
Fluorescence evaluation was either in a conventional mode, with 1–2 s filter changes, using an inverted microscope (ICM 405; Zeiss Inc., Oberkochen, Germany), or a microscope (Axiovert; Zeiss Inc.) equipped with a confocal imaging system (Noran Instruments, Bruchsal, Germany). A minimum of 33 ms frame sequences allowed documentation of [Ca2+]i fluorescence transients with CLSM, in some cases alternating with phase contrast imaging. Excitation wavelength was 488 nm (Ar-laser emission) for CaGr-2, Fluo-3, or fluorescein, or 490 nm (±5 nm, band pass filter) for Fura red. Emitted wavelength registered was ⩾515 nm for CaGr-2, Fluo-3, or fluorescein, or ⩾560 nm for Fura red (detected by a moonlight camera; Panasonic, combined with a 560 nm dichroic mirror and a 560-nm-long pass filter).
To minimize false or irrelevant fluorescence signals, evaluation included (a) calibration of fluorochrome signals with standard Ca2+ solutions in microdroplets, (b) intermittent registration of signals with exocytosis recording in phase contrast, (c) overlapping use of fluorochromes with different properties, (d) mimicking shape changes via mechanical deformation without exocytosis triggering, (e) background reduction by digital image subtraction in rapid CLSM series, (f) transformation of signals into false colors, and (g) evaluation of important cell regions by line scans. We could thus either follow semi-quantitatively rapid [Ca2+] changes in the CLSM mode, or we could follow quantitatively longer lasting [Ca2+] changes in the conventional microscope.
Electrophysiological Recordings Show Correlation of Subplasmalemmal Ca2+ Transients with Exocytosis
Flushing a Paramecium cell with AED causes massive exocytosis which, under voltage clamp conditions, is paralleled by an outward current of 3 × 10−9 As lasting ∼2 s (Fig. 1,A, top). Some smaller peaks may precede the main current peak. As shown in more detail below, the magnitude of the current is proportional to the number of trichocysts released. The currents display a fast rising phase and a slower, approximately exponential decay (Fig. 1,B). A second pulse of AED after 20 s provokes little if any additional trichocyst release and causes a series of only small currents (Fig. 1 A, bottom).
Small numbers of trichocysts can be quantified in the LM (Plattner et al., 1984, 1985). Exocytosis of one or a few trichocysts occurs upon slight irritation of a cell (HaackeBell et al., 1990), i.e., during immobilization and impalement by a microelectrode. This, however, does not appear to be the immediate trigger for spontaneous exocytosis or spontaneous currents we observed, since we allowed cells to stabilize before recordings were started. Stochastic current peaks are observed during both intracellular and extracellular recordings (Fig. 2). Extracellularly recorded currents are of a different size but of similar shape, as are currents recorded under voltage clamp conditions (Fig. 1,A, top). Spontaneous signals representing 36 events were averaged in Fig. 2,B. Currents with a mean amplitude of 34 pA show a fast rising phase (rise time, tr = 7 ms) and a half width of t1/2 = 21 ms. These properties characterize single events, corresponding to the peak in Fig. 3,D (arrow) which is associated with the release of a single trichocyst. The histogram of Fig. 2,C clearly shows the absence of discrete steps in the current size, i.e., absence of quantal current events. Fig. 3 reveals that a minimal current peak accompanies release of an individual trichocyst. A further example of simultaneous current and exocytosis registration involving a larger number of trichocysts is documented in Fig. 4.
Fig. 5 correlates the charge of the electrical events with the number of trichocysts released. Data scatter is large but not unexpected for several reasons. (a) Trichocyst countings are not absolutely certain. (b) The Ca2+-activated currents recorded are not caused simply by mobilization from discrete stores but may be amplified to a variable extent by Ca2+ influx from the medium. (c) While exocytosis is always accompanied by a current, spontaneous currents can be observed that are not accompanied by spontaneous exocytosis. This may be due to the fact that not all potential exocytosis sites are occupied by a trichocyst (Plattner et al., 1991; Knoll et al., 1991a, 1993). Within these limitations, we calculated the unit current event as 1.21 ± 0.97 pC per exocytotic event (Fig. 5). By EGTA injection we have ascertained that the currents described depend on a subplasmalemmal [Ca2+]i increase (Fig. 6).
In conclusion, our recordings show that the number of trichocysts released by exocytosis in response to AED is paralleled by a nonstepwise increase in subplasmalemmal Ca2+-dependent currents.
Quenched-Flow/Freeze-Fracture Analysis Shows only Partial Reduction of Exocytotic Membrane Fusion with Low [Ca2+]e, while under Normal Conditions Internal and External Ca2+ Sources Contribute to Exocytosis
Exocytosis of trichocyst contents is monitored in the LM by decondensation (stretching) to the typical needles seen outside a cell (see Fig. 9, D and F and Fig. 10, M and O). Since this requires Ca2+e (Bilinski et al., 1981), the effects of low [Ca2+]e on membrane fusion had to be analyzed separately in quenched-flow/freeze-fracture experiments (Fig. 7). This also allows rigorous mixing of cells with EGTA and, thus, restriction of the time of EGTA application since this could potentially affect cell function (see below). Trichocyst docking sites were rated as resting stages with rosettes (aggregates of integral membrane proteins; Plattner et al., 1991), or as activated stages with exocytotic openings of variable diameters. Fig. 7 includes data from controls (left) which were mockstimulated with culture medium (0.1 mM [Ca2+]e) and processed by the same method. In this case, no activated docking sites were observed, and resting stages in these controls are used as reference (100%). In the presence of Ca2+e, AED activates all exocytotic sites (Fig. 7, middle). EGTA application for 200 ms, followed by 80 ms AED triggering, allows activation of only ∼40% of the potential trichocyst docking sites (Fig. 7, right). 80 ms was selected because synchronous exocytosis is normally completed within this time (Knoll et al., 1991a). Columns in these data pairs do not add up to identical values because they represent medians derived from different individual cells, with variable numbers of observations.
These data show that exocytotic membrane fusion of about half of the docking sites, can be induced by mobilization of Ca2+ from internal pools. We conclude that Ca2+ mobilization from subplasmalemmal pools operates at its limits and is normally intensified by a Ca2+ influx.
Ca2+ Imaging Shows a Rapid Subplasmalemmal [Ca2+] Transient by AED Stimulation Even with Low [Ca2+]e, while Such a Transient Achieved by Rapid [Ca2+]e Increase Alone Does Not Induce Exocytosis
Different fluorochromes were used to overcome problems specific of different experiments. Fura red allows quantitation, although with low time resolution. CaGr-2 and Fluo-3 combined with CLSM allow only semi-quantitative analyses, though in the sub-second range, whereby Fluo-3 has the disadvantage of being partially sequestered into phagosomes, while providing significantly higher quantum yield than CaGr-2. The occurrence of similar signals over the same time periods made us confident, however, about the relevance of our findings.
To establish whether a Ca2+ influx would also suffice to trigger exocytosis or, alternatively, whether alveolar sacs may be the primary Ca2+ source during AED stimulation, as suggested above, we first worked with media in which Ca2+e was chelated by EGTA. Long term experiments, however, were impossible, because at low [Ca2+]e (⩽50 μM) the membrane of many cells becomes leaky to a broad spectrum of small molecules (Hille, 1992). As Fig. 8 shows, increasing [Ca2+]e to 10 mM suffices to produce an immediate strong cortical [Ca2+] transient, yet without inducing any exocytotic responses. This largely excludes any CICR mechanism.
CLSM using CaGr-2 f/fo imaging shows that a cortical [Ca2+] transient occurs within ∼100 ms of AED application (Fig. 9, A–F), i.e., within the time required for exocytosis (Knoll et al., 1991a), as documented by the intermittent phase contrast pictures in Fig. 9, A–F. Controls carried out to exclude a role of cell deformation included analyses with more strict time scales, revealing a continuous build up of the cortical fluorescence signal irrespective of cell deformation (Fig. 9, G–N), and local mechanical deformation of cells which failed to produce any comparable signal (Fig. 9, O–Q). Therefore, we are confident that the cortical Ca2+ signal parallels exocytosis, as documented above independently by electrophysiology and by quenched-flow/freeze-fracture analysis.
Next we analyzed whether Ca2+ mobilization from stores might suffice to trigger exocytosis. Results obtained by fluorescein-tagged EGTA solution with subsequent AED application are shown in Fig. 10, A–H. Under these conditions no trichocyst release occurs. Reliability of the superfusion approach was checked independently with fluorescein-tagged AED in the presence of 0.1 mM [Ca2+]e (Fig. 10, I–P). Since trichocyst decondensation (expansion during expulsion of contents) requires Ca2+e (Bilinski et al., 1981), comparison of Figs. 10, A–H (+ EGTA) and 10, I–P (+ Ca2+e) shows successful chelation of Ca2+e. Next we have loaded cells with Fluo-3. After 2 min, each of these cells was superfused with EGTA and exposed to a local AED flush (without fluorescein). This final set of experiments, documented in Fig. 10, Q–T, clearly shows a cortical [Ca2+]i increase already recognizable within ∼100 ms with little diffusion throughout the cell. Under such conditions quenched-flow/freeze-fracture has revealed occurrence of membrane fusions.
Alveolar Sacs Are the Most Likely Structural Equivalents of Subplasmalemmal Ca2+ Pools
As shown above, the extent of trichocyst exocytosis is paralleled by the size of currents observed. Assuming alveolar sacs to be the primary source of Ca2+ for Ca2+-activated currents a hypothesis remained to be tested: the ratio of maximal versus minimal currents should reflect the ratio of total cell surface versus unit area of cell surface. These units, the kinetids, also correspond with the number of alveolar sacs which occur all over a cell. Current ratios would roughly have to reflect area ratios, even when currents were superimposed by Ca2+ influx.
To answer the problem, we established morphometric parameters for the cells used in electrophysiological recordings. Viable or cryofixed cells were analyzed by both LM and EM methods (Table I) revealing ∼3,100 surface units. For comparison, current ratios are compiled in Table II. The minimal current, i.e., the current per trichocyst discharge, as derived from extracellular recordings (Fig. 5), is 1.2 × 10−12 As. Correction by a factor of 2 is required, since the current is recorded from ∼1/2 of the cell surface, i.e., the part in the holding pipette (see Materials and Methods). The corrected value of 2.4 × 10−12 As is in agreement with the value obtained by intracellular voltageclamp recording, i.e., 2.5 × 10−12 As. Maximal currents recorded extracellularly during maximal AED stimulation, after correction for recording area, amount to 6 × 10−9 As (Table II). The ratio of maximal to minimal currents is ∼2.5 × 103, a value closely approaching the area ratio of 3.1 × 103, i.e., the estimated number of cortical Ca stores per cell (Table I). This is another aspect supporting our suggestion that alveolar sacs may be the internal source of Ca2+ relevant for exocytotic membrane fusion.
Current Flow Reflects Extent of Cell Surface Area Activated, Rather than Fusion Pore Conductance–Superposition of Ca2+ Activation from Stores by a Ca2+ Influx
Could spontaneous currents reveal fusions of single trichocysts with the cell membrane during exocytosis? From work in other secretory cells, it is known that during fusions an electric discharge occurs as soon as the fusion pore opens (Almers, 1990; Lindau, 1991). The magnitude of the current, or its charge, which depends on the potential difference between the two fusing compartments, can be recorded as a current transient preceding the stepwise increase in membrane capacitance, reflecting the increase of surface membrane, as measured by the patch clamp technique. From the current transient and the capacitance change, one can calculate the potential of the fusing vesicle.
If one applies the same considerations to Paramecium, one can compare the measured spontaneous electrical signal with that expected for a fusion event. Considering a trichocyst surface of ∼10 μm2 (estimated from electron micrographs) and assuming the usual 1 μF × cm−2 for biomembranes (Cole, 1972; Kado, 1993), a capacitance of 10−13 farad would result. From the recorded transients (e.g., in Fig. 1 B) the charge can be calculated to be ∼2.5 × 10−12 As. The potential difference between the cell membrane and the trichocyst membrane thus calculated would amount to 2.5 × 10−12 As/10−13 farad (25 V). However, this value is orders of magnitude above physiological values, and this excludes that the spontaneous current events we have recorded is due to the transient capacitative discharges of the fusing trichocyst membrane. This interpretation is supported by the occurrence of the same currents in the trichocyst-free mutant, tl, where spontaneous and AED-elicited currents were quite similar to those in wildtype cells (Erxleben and Plattner, 1994). Therefore, we favor the interpretation which relates current events, no matter whether spontaneous or triggered, to the activation of a variable number of alveolar sacs with a superimposed Ca2+ influx (see below).
The concomitant abolition of both Ca2+ currents and exocytosis by EGTA injection supports a causal relationship between [Ca2+]i increase and exocytotic membrane fusion (Fig. 6). Discharge of subplasmalemmal Ca stores is probably a primary response to AED triggering, mainly for two reasons. First, some Ca2+ signal and some membrane fusion occur also with [Ca2+]e ⩽ [Ca2+]i at rest. Second, rapid Ca2+ influx caused by 10 mM [Ca2+]e causes a cortical [Ca2+] transient of ⩾900 nM, but no exocytosis (see Results and below). Both largely exclude a CICR mechanism, as also suggested by 45Ca2+ release studies with isolated alveolar sacs (Länge et al., 1995) and by patch clamp analysis of reconstituted Ca2+ release channels from Paramecium (Zhou et al., 1995).
From all our observations we come to the following conclusion. With “normal” [Ca2+]e (0.1 mM), Ca2+ release from alveolar sacs is clearly accompanied by a Ca2+ influx through the cell membrane, resulting in a diffusion to the entire cell (data not shown) and increased exocytotic activity, while the signal remains locally restricted with low [Ca2+]e (Fig. 10, Q–T). Both phenomena, normally acting in concert and possibly involving site-directed Ca2+ release and/or influx, support rapid trichocyst exocytosis responses.
Relationship to Models of Ca2+ Dynamics during Triggered Exocytosis
In Paramecium, alveolar sacs have been identified as Ca pools by their Ca2+ sequestration (Stelly et al., 1991) and Ca2+ release (Länge et al., 1995) properties, both after isolation and by in situ Ca imaging methods (ESI, SIMS) (Knoll et al., 1993; Stelly et al., 1995). These organelles are connected to the cell membrane by electron-dense connections ∼15 nm long (Plattner et al., 1991). Thus, they clearly fulfill the criteria of subplasmalemmal pools.
The capacitative model of Ca2+ entry (Putney, 1986, 1990, 1993) assumes functional coupling between emptying intracellular Ca stores and Ca2+ influx. Structural coupling of Ca stores with the cell membrane has been discussed for a large number of secretory systems (Berridge, 1995), although the structural identification of such pools is still questioned. In these secretory systems however, Ca2+ release is sustained by inositol 1,4,5-trisphosphate, a second messenger for which we have no evidence in Paramecium (Länge et al., 1995; Zhou et al., 1995). Thus a capacitative model sensu strictu cannot be envisaged in our cells.
In some secretory cells, Ca2+ release has been defined as quantal (Muallem et al., 1989; Meyer and Stryer, 1990; Tregear et al., 1991; Cheek et al., 1993; Parys et al., 1993) in the sense that [Ca2+]i and, in parallel, Ca2+-dependent exocytosis is increased by increasing concentrations of secretagogues. So far, however, no strictly quantal pool mobilization could be demonstrated (Taylor and Potter, 1990; Bootman 1994a,b). Our analyses with Paramecium have revealed in contrast a correlation between the size of Ca2+-activated currents, the extent of exocytosis achieved, and the amount of subplasmalemmal Ca stores available, with no sign of discrete current steps. This discrepancy may be caused by either the different processes of Ca2+ release from the stores, the different Ca2+ sensitivity of Ca2+-activated channels along the cell surface (Erxleben and Plattner, 1994), or by the superposition to the release of a slightly variable Ca2+ influx.
One argument for Ca2+ release as the primary event induced by AED is the relatively small current elicited by a second application (20 s later). By comparison, the ability of the electric system (i.e., the Ca2+-activated channels) to generate a second current pulse in response to increased [Ca2+]i, can be regarded as instantaneous. Since we have no evidence of an AED receptor, desensitization is a less likely explanation. Potential Ca2+ release channels recently analyzed by electrophysiology after reconstitution show pharmacological characteristics (Zhou et al., 1995) which are practically identical to those for Ca2+ release from isolated sacs (Länge et al., 1995). Although the molecular identity of the channels actually involved in Ca2+ influx during AED stimulation still has to be established, it certainly differs from those in the Paramecium ciliary membrane (Erxleben and Plattner, 1994; Plattner et al., 1994; Zhou et al., 1995). Another subject of further analysis is the significance in Paramecium of Ca2+ influx, whether it serves directly to amplify the subplasmalemmal signal (Putney, 1990, 1993; Berridge, 1995) or to refill subplasmalemmal stores during or immediately after their release.
In the Paramecium system, local Ca2+ transients are sustained by a local interplay between endogenous and exogenous sources. This combination is needed for the rapid local trichocyst exocytosis, with ensuing efficient defensive function activation within tens of milliseconds. Prerequisite to this activity is the proper assembly of surface cell components, i.e., trichocysts and subplasmalemmal Ca2+ stores, with the concomitant activation of both Ca2+ release and influx. The special design of a Paramecium cell, investigated by appropriate methods, has allowed us to reveal some features in our system which might turn out to be applicable also to higher eukaryotic cells.
We thank Jochen Deitmer for communicating some early observations to the problem, Jochen Hentschel for his help with the CLSM set-up, and Mary Anne Cahill for reading an early version of the manuscript.
Abbreviations used in this paper
This paper is supported by SFB156 as well as DFG grants Pl78/11 and Pl78/12, all from the Deutsche Forschungsgemeinschaft (DFG).
Please address all correspondence to H. Plattner, Faculty of Biology, University of Konstanz, P.O. Box 5560, D-78434 Konstanz, Germany. Tel.: (49) 7531-88-2228. Fax: (49) 7531-88-2245.
C. Erxleben and N. Klauke contributed equally to this publication.