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1Department of Neurobiology, Pharmacology, and Physiology, The University of Chicago; and 2Department of Anatomy and Cell Biology, University of Illinois, Chicago, Illinois
Submitted 25 March 2005; accepted in final form 18 May 2005
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ABSTRACT |
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INTRODUCTION |
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; Malinow 2003; Malinow and Malenka 2002) or function (Lee et al. 2003) can change synaptic efficacy. Changes in secretion efficiency of synaptic vesicles has been described (Arancio et al. 1996; Stevens and Wang 1994; Tsien et al. 1996; Zucker and Regehr 2002). Under certain conditions, the presynaptic changes lead to an increase in transmitter secretion by elevating the probability of release at an individual synapse without concomitant changes in the amount of neurotransmitter released per vesicle (Malinow and Tsien 1990; Stevens and Wang 1994). Most studies assume that the transmitter quantal size is invariant; however, there is little direct evidence in support of this assumption at many synapses.
Clear differences in synaptic vesicle diameters have been observed in neurons. At Drosophila's neuromuscular junction, two different glutamatergic motor neurons innervating the same muscle are distinguished by their vesicle sizes (mean diameters of 41 and 48 nm) (Karunanithi et al. 2002). Serotonergic Retzius neurons of leech release 5-HT from three different populations of vesicles, each having a unique vesicle size (Bruns et al. 2000). Two of the populations were dense core vesicles (DCVs) that are thought to store neuropeptides and 5-HT. Furthermore, the Drosophila and leech studies provided evidence that quantal size is a function of vesicle size. Other studies have suggested that vesicle neurotransmitter concentration can be modified (Kullmann and Nicoll 1992; Liao et al. 1992) or that only a fraction of the total vesicle content is released per fusion event (Elhamdani et al. 2001).
In the case of chromaffin cells, a model system for studying exocytosis, it is often assumed that a single population of DCVs supports regulated secretion. However, electron microscopic (EM) studies of bovine (Coupland 1968) and mouse chromaffin cells (Gorgas and Bock 1976) showed that DCV size does not strictly follow a single, normal distribution. In Coupland's 1968 study, if one looks at the size distribution of norepinephrine vesicles (see Fig. 5 of that study), there are too many large-diameter vesicles to fit all the data with a single normal distribution. Physiological evidence for a heterogeneous population of DCVs is emerging. For instance, DCVs respond differentially to secretagogues as a function of the vesicles age (Duncan et al. 2003), and the size of vesicles that undergo regulated secretion are not uniformly distributed (Henkel et al. 2001). In PC12 cells, DCV biogenesis occurs in stages that are marked by changes in vesicle diameter (Tooze et al. 1991).
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; Finnegan et al. 1996; Wightman et al. 1991). Our results showed that quantal sizes were bimodally distributed as small and large events with Q1/3 values of 0.40 and 0.69 pC1/3, respectively (size ratio: 1.73), with amperometry and EM results showing a fivefold difference in the volumes of large and small vesicles. In addition, small amperometric events were kinetically distinct from large events. These results suggest that two separate pools of secretory vesicles give rise to events with different quantal size. The results also imply that under conditions of intense stimulation, as used in this study, vesicles were released in an all-or-nothing fashion. |
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METHODS |
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Animals were housed and handled as required by the Animal Resource Council (University of Chicago, IL). Cells were prepared similar to previously published methods (Obukhov and Nowycky 2002). Briefly, adrenal glands were harvested from 5- to 8-wk-old male C57 BL/6J mice (The Jackson Laboratory, Bar Harbor, ME). The medulla was dissected from the cortex with a surgical scalpel and cut into two pieces. The tissue was digested for 35 min with 1 mg/ml Collagenase P (Roche Diagnostics, Indianapolis, IN) +25 µg/ml DNase (Sigma-Aldrich, St. Louis, MO), followed by a 15-min digestion in 0.03% Trypsin/EDTA (Invitrogen, Grand Island, NY) plus 50 µg/ml DNase. All digestions were carried out in calcium-free buffered saline at 37°C in a shaker water bath. Finally, the digested tissue was transferred to tissue culture media and mechanically dispersed through repeated aspiration (
7 cycles) into a flame-polished Pasteur pipette. Cells were plated on glass cover slips coated with Matrigel (Discovery Labware, Bedford, MA) and maintained in a 37°C, 5% CO2 incubator. The tissue culture media had the following composition: DMEM-F12K, 10% HS, 5% FBS and 1% of a 100x penicillin/streptomycin/glutamine cocktail (all media reagents were purchased from Invitrogen). EM and amperometry experiments were carried out 2 and 3 days post tissue harvest.
Amperometry
Carbon fiber electrodes were fabricated with 7-µm-diam carbon fibers (Fortafil Fibers, Knoxville, TN). The carbon fiber was threaded through a glass capillary pipette (Drummond Scientific, Broomall, PA) and pulled with a vertical glass pipette puller (Narashige, Tokyo, Japan). An epoxy seal between the carbon fiber and glass was created by dipping the glass-fiber tip into a 14% hardener: resin mixture (w/w), preheated to 100°C (hardener, metaphenylenediamine, and resin, 828 Epon Resin; Miller-Stephenson Chemical, Morton Grove, IL). Electrodes were baked overnight at 66°C. The glass portion of the tip was painted with silicone elastomer (Sylgard, Dow Corning, Midland, MI) and then baked at 66°C for 
1 h. On the day of the experiment, the carbon fiber was cut and visually inspected as described by Bruns et al. (2000). The electrode was pressed gently against the cell during the recording as the collection efficiency is thought to be highest under this recording configuration (Bruns et al. 2000; Schroeder et al. 1992). A newly cut surface or a new electrode was used for each cell to prevent fouling of the electrode. The electrode was backfilled with 3 M KCl. The electrode was held at +800 mV versus silver-chloride using an EPC-7 amplifier (HEKA Electronics, Lambrecht, Germany) to oxidize catecholamine transmitter. The electrode was mounted in a holder and then advanced until it gently touched the cell. The amperometric signal was low-pass filtered at 2 kHz (8-pole Bessel; Warner Instruments, Hamden, CT). A 16-bit A/D converter (National Instruments, Austin, TX) was interfaced with custom data acquisition software. The amperometric signal was acquired at 10 kHz and stored on a personal computer. The rms noise present in the acquired traces was typically under 1.0 pA as determined with the MiniAnalysis program (Synaptosoft, Decatur, GA). Records with rms noise >2 pA were not analyzed. Amperometric spike features, quantal size, and kinetic parameters were analyzed using the amperometric program in Minianalysis. The detection threshold for an event was set four to five times the baseline rms noise, and the spikes were automatically detected. Overlapping events were rejected, and they were considered overlapping if the initial spike had not returned to baseline before the next event occurred. Overlapping events were relatively rare. Rise-time values were measured over 10–90% of the spike's maximal amplitude. The area under individual amperometric spikes is equal to the charge (pC) per release event, referred to as Q. The number of oxidized molecules (N) was calculated using the Faraday equation, N = Q/ne, with n = 2 electrons per oxidized molecule of transmitter; e is the elemental charge (1.603 10-19 coulomb). To make an approximate estimate of the total number of events released over the entire cell, the following assumptions were made. Release was assumed to occur uniformly over the cell (Wick et al. 1997), and the release rate measured at the electrode was multiplied by the ratio of cell surface area/electrode area. The electrode, gently pressed against the cell, was estimated to cover only 6.3% of the cell's surface, and the average rate of 20 events/min detected at the electrode yields a global release rate of 318 events/min. Amperometric data were measured in 32 cells.
Recording solutions and stimulation protocols
Recordings were made from adherent cells that were under constant perfusion (flow rate of 1.0 ml/min, and an approximate chamber volume of 150 µl). All recording solutions had the following standard composition (in mM): 145 NaCl, 2.0 KCl, 10 HEPES, and 1.0 MgCl2. Ca2+-free solutions contained 100 µM EGTA. All solutions used during and after cell permeabilization contained 1.0 mM Na2ATP. The solutions were adjusted to a final pH of 7.20 and an osmolarity of 300 ± 3 mosM/kg. Free-Ca2+ was estimated with WEBMAXCLITE (http://www.stanford.edu/cpatton/maxc.html). All experiments were performed at ambient temperature, ranging from 22° to 26°C. Cells were stimulated repeatedly using the following protocol: 2 min in a Ca2+-free solution, permeabilized with 20 µM digitonin (Ca2+-free) for 10 s, stimulated 2 min with a solution containing free Ca2+ concentration of 100 µM, and washed for 1 min in Ca2+-free media before the cycle was repeated starting at step 2 (permeabilization). Cells were typically stimulated three to four cycles (and at most 6 cycles) or until the cell membrane changed from its initial, bright-field dark appearance to a granular texture (for details, see Jankowski et al. 1992). Quantal size from the first stimulation was compared it to later stimuli; nothing was changed. Digitonin was purchased from Calbiochem (La Jolla, CA). Cells stimulated with nicotine were exposed to 10 µM nicotine for 3 min in 2 mM Ca2+, followed by a 2-min wash without nicotine. Two additional stimulations repetitions were carried out.
Electron microscopy
After the cells were cultured for 48 h (described in Cell culture), they were prepared for EM. Coverslips were rinsed twice with 0.12 M cacodylate buffer (pH 7.4) to remove cell culture media and then fixed with 2% glutaraldehyde and 0.6% paraformaldehyde in 0.06 M cacodylate buffer pH 7.4 for 1 h. The coverslips were again rinsed three times and then postfixed for 1 h in a mixture of 1% osmium tetroxide in 0.06 M cacodylate buffer pH 7.4 containing 1.5% potassium ferricyanide, which was added immediately before use. The samples were washed three times, dehydrated in a graded series of EtOH (50–100%), with the 70% step containing 1% uranyl acetate (30 min), and finally dehydrated in propylene oxide. The samples were infiltrated with Durcupan (Fluka), and the coverslips were inverted over 15-mm round-well molds with the tissue side down and polymerized for 48 h at 60° C. After polymerization, the resin plug with cover slip was suspended in hydroflouric acid for 10 min to dissolve the glass. Areas of interest were cut out with a jewelers saw, attached with Superglue to a blank Beem capsule stub, and sectioned parallel to the plane of the cover slip with a diamond knife. Sections with a silver interference color were collected on 1 x 2 mm Formvar-coated slot grids and counterstained with uranyl acetate and lead citrate. The sections were viewed in a Jeol 1220 electron microscope at 80 kV, and images were taken on EM film (Eastman Kodak, Rochester, NY) at x12,000 magnification.
Vesicle size measurements
Digital images of EM micrographs were made and analyzed at a final magnification of x100,000 or x200,000. Random single sections from cells (11 cells total) that possessed multiple DCVs and well-fixed cellular constituents (mitochondria, nuclear material and plasma membrane) were selected for analysis. The criteria for DCVs that were included in the size distribution are as follows: the vesicle had to have an electron dense core and an intact, continuous vesicle membrane surrounding the dense core, and the membrane had to be approximately circular in geometry. All vesicles meeting these criteria were included in the analysis. The vesicle diameters were measured from 11 cells. The area of each vesicle was measured using Image-J software (National Institutes of Health, Bethesda, MD) or SigmaScan software (SPSS, Chicago, IL) (as described previously, Colliver et al. 2000). The two programs were compared and found to yield results that were essentially identical. Diameter was calculated from vesicle area [diameter = 2*(Area/
)0.5]. Two investigators made independent measurements from the same single sections, and resulting vesicle size distributions were essentially identical. Corrected vesicle diameter was calculated using a previously published algorithm (Parsons et al. 1995). This algorithm is dependent on section thickness and the empirically measured vesicle diameter (referred to as apparent diameter). Section thickness was estimated to be 60 nm based on the silver interference color of the sections (Sakai 1980). Apparent vesicle diameters were transformed to corrected diameters prior to statistical or graphical analysis of the data. Widest vesicle diameter was measured from serial sections using Image-J (National Institutes of Health). Four to five sections through individual cells (3 cells in total) were aligned using cellular landmarks, such as mitochondria and large DCVs. Individual DCVs (meeting the morphological criteria described in the preceding text) were followed through multiple sections to determine which section captured the vesicle at its widest diameter. Small vesicles that were only obvious in a single section were considered to represent the vesicle captured at its greatest diameter.
Statistical analyses and plots were performed in Origin (Origin, Northampton, MA). To select the model that gave the best-fit to the size distributions, we applied the corrected Akaike Information Criterion (AICc) for small sample sizes (n) (Anderson 2002). This method selects the model that best represents the data while minimizing the number of parameters used in the model. The AICc equation is as follows
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), and the degree that one model better describes the data compared with a second model can be expressed numerically as a probability as follows: difference in AICc values for model 1 and 2: 
AIC = AICc1 – AICc2; probability of model 1 giving the best fit: Prob1 = 100 [exp-0.5(AIC)/(1 + exp–0.5(AIC))]. Probability of model 2 yielding the best fit: Prob2 = 100 –Prob1. |
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RESULTS |
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Previous studies using whole mouse adrenal glands identified two major chromaffin cell types, namely epinephrine- and norepinephrine-storing cells (Gorgas and Bock 1976; Kobayashi and Coupland 1977). These catecholamine transmitters react differently with aldehyde fixatives, which allows the two cell types to be distinguished by the appearance of their DCVs, the organelles that concentrate the transmitter (Coupland 1968; Kobayashi and Coupland 1993). Norepinephrine reacts with fixative aldehydes and osmium tetroxide (Coupland and Hopwood 1966a,b), which impart a core that is very electron dense and often surrounded by a halo of membrane (Coupland 1968). In contrast, epinephrine reacts poorly with fixatives, and as a result epinephrine-storing cells exhibit DCVs that are relatively less electron dense (Coupland 1968). The cells in our study generally fit the description of norepinephrine-storing cells that possess DCVs with electron dense cores, as described by Thureson-Klein et al. (1984), even though we did not select for this type of cell.
A quantitative analysis of DCV size revealed a striking heterogeneity within individual cells. Random sections from cells that possessed multiple DCVs and well-fixed cellular constituents (mitochondria, nuclear material, and plasma membrane) were selected for analysis (see Fig. 1A). The criteria for DCVs that were included in the size distribution can be found in METHODS. All vesicles meeting these criteria were included in the analysis. Typically the larger DCVs possessed an excess membrane giving a halo appearance around the vesicle's core, and the cores of smaller vesicles often appeared to have a tightly associated membrane (see Fig. 1B) (Gorgas and Bock 1976). Under physiological conditions, catecholamines fill the intra-vesicular space (either in a freely diffusible state or bound to a protein matrix); therefore the entire area within the vesicle membrane was measured and then converted to a diameter (Colliver et al. 2000) and then transformed using an algorithm that corrects for sectioning errors (Parsons et al. 1995) (described in more detail in the following text). Typically, individual cells appeared to have a collection of smaller vesicles with diameters ranging from 80 to approximately 200 nm, and a second group of larger vesicles with diameters >200 nm (see Fig. 1C). Cells with a single Gaussian distribution of vesicle sizes were not observed.
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Vesicle size measured from single EM sections is known to result in an apparent diameter that is less than the correct diameter. In the past, a variety of corrections have been applied to adjust the data (Parsons et al. 1995). To verify the accuracy of the correction that we used in this study, serial sections were analyzed to determine the vesicle's widest diameter. Figure 2A shows a region of a cell enriched in DCVs. The vesicles with large diameters span three to four sections (Fig. 2B), and smaller vesicles typically span one to three sections (Fig. 2C; sections are 60 nm in diameter). The results from three cells (four to five sections analyzed per cell) were pooled (both small and large vesicles were combined) and averaged, which yielded a widest diameter of 194 nm. Single-section results measured from the same sections produced an average apparent diameter of 164 nm (18% less than the widest diameter). Transforming the data from a single section using an algorithm that corrects for sectioning errors (Parsons et al. 1995) yielded a mean diameter of 202 nm ("corrected" diameter). The corrected and widest diameter results were in good agreement (4.2% difference).
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Quantal size measured from individual chromaffin cells suggests two populations of vesicles
Amperometry was used to assay catecholamine release from mouse adrenal chromaffin cells. Release was stimulated by the application of 100 µM Ca2+ to digitonin permeabilized cells (Jankowski et al. 1992). Amperometric events were rare in the absence of Ca2+, even after digitonin permeabilization, which suggests that the secretory machinery functions properly in the presence of digitonin (Graham et al. 2002; Holz et al. 1989; Jankowski et al. 1992, 1993). Figure 3A shows a representative amperometric trace with multiple individual events occurring over a 2-min stimulation period. On average the electrode detected 20 events/min, which provides an estimated rate of release of 318 events/min over the entire surface of the cell (the electrode samples 6% of the cell surface, see METHODS for calculation). The amperometric events from Fig. 3A are shown on an expanded time scale in B. Each amperometric event, corresponding to an individual release event, has been aligned such that they all begin at the same time. The events differ considerably in amplitude and overall shape. The majority of events appear as a single spike that rises rapidly and falls at a slower rate; however, in some cells, a fraction of the events exhibit the small elevation from baseline prior to the spike known as a "foot," which is thought to arise from neurotransmitter release from a partially open fusion pore (Chow et al. 1992).
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diameter3) (Bruns et al. 2000; Finnegan et al. 1996; Wightman 1991). The Q1/3 distribution for a single cell does not appear as a single Gaussian distribution (Fig. 3C). Rather the data are best described as the sum of two populations each one with its own Gaussian distribution. The first group of events is clustered at 0.4 pC1/3. The second type of event has a broader distribution centered at 0.7 pC1/3. Hence, neither vesicle diameter nor Q1/3 distributions measured from individual cells appeared to be composed of a single uniform population of vesicles. Vesicle size and quantal size measured from cell populations are best described as bimodally distributed
Combining vesicle size measurements from 11 cells resulted in a distribution that is best described as the sum of two Gaussians. Vesicle diameter was measured from single sections (11 sections, 1 per cell) and transformed to the corrected diameter, as described in the preceding text. The resulting distribution was fit best as a bimodal distribution with small and large vesicles centered at 179 and 310 nm, respectively (see Fig. 4A). Size distributions made from individual cells (see the Figs. 1C and 2E) or from multiple cells (Fig. 4A) share a similar bimodal distribution; this argues that the two populations of vesicles exist in the same cell.
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The apparent bimodal nature of both vesicle size and Q1/3 are in agreement (see Table 1). The size ratio of large and small vesicles is 1.73 (corrected diameter in nm: 310/179), which is identical to the ratio of the two quantal size populations, 1.73 (Q1/3: 0.69/0.40). The average ratio of quantal size and vesicle size translates to a fivefold difference in volume (volume diameter3, 1.73). In summary, both measurements are consistent with the hypothesis that mouse chromaffin cells have two populations of vesicles with different sizes (summarized in Table 1).
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The concentration of transmitter in each vesicle population was estimated from the vesicle size and quantal size data sets. The corrected vesicle diameter was used to calculate volume, and the mean charge was used to estimate the number of transmitter molecules (see Table 1). Assuming that the small quantal events arise from small vesicles, we calculate an intra-vesicular transmitter concentration of 0.11 M. Similarly, the large vesicles are estimated to have a transmitter concentration of 0.11 M. These concentrations are similar to previous estimates where both vesicle and quantal sizes were determined (Bruns et al. 2000). The results suggest that transmitter concentration remains constant while vesicle size changes, a conclusion similar to that found in other studies (Albillos et al. 1997; Bruns et al. 2000; Colliver et al. 2000).
Distinct kinetic properties are consistent with two populations of vesicles
Groups of amperometric events were averaged to better observe kinetic properties of small events. Amperometric events were binned according to size, in Fig. 5, and presented as ensemble averaged spikes (each ensemble was composed of 50 individual amperometric events). The top inset (Fig. 5I1) highlights the rapid rising phase of the small events. Figure 5I2 superimposes a small event group with a large event group, after scaling them to the same amplitude. The scaled small event group does not overlay exactly with the large event group and in particular the small event group has a slower falling phase.
Figure 6A plots the relationship between amperometric event amplitude and Q. The data can be described as the composite of two separate linear processes. The fits for small and large events were made to regions where overlap was minimal. Small events are fit to a line with a modest slope compared with the line used to fit large events. The intermediate region appears as a transition between the two types of vesicles. This data show that small events grow more slowly in amplitude as quantal size increases when compared with large events.
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One possible concern with the data outlined in this study is that the two populations of vesicles were an artifact resulting from the use of digitonin to permeabilize the cells. Figure 7 compares quantal sizes from digitonin permeabilized cells with those recorded after stimulating intact cells with nicotine (10 µM). Figure 7A shows a Q1/3 distribution obtained from a single nicotine-stimulated cell, whereas C shows averaged data from multiple nicotine-stimulated cells (n = 6). Both histograms are best fit as the sum of two Gaussian distributions (Fig. 7, A and C). The data from intact cells stimulated with nicotine are very similar to cells permeabilized with digitonin (n = 9) and then exposed to Ca2+ (compare Fig. 7, A and B, or C and D). Our data suggest that two populations of vesicles are an intrinsic property of the cells.
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DISCUSSION |
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One possible concern is that the digitonin used to permeabilize cells was somehow responsible for producing the two populations of amperometric events observed. Digitonin stimulation of chromaffin cells has been used for over 20 years (Dunn and Holz 1983; Wilson and Kirshner 1983). Jankowski et al. (1992) first described the method of permeabilizing cells with digitonin for use in amperometry experiments. This same group later compared secretion stimulated by nicotine and digitonin and found very similar results for each stimulation method (Jankowski et al. 1993). Our own data are consistent with Jankowski et al. (1993). Figure 7 compares release stimulated by digitonin with that of nicotine shows that the Q1/3 histograms from both data sets were quite similar and that both were best fit as the sum of two Gaussians.
Another possible concern is that the assignment of small events as an independent population is flawed due to the possibility that they arise from release sites far from the electrode. Under these conditions, the electrode would capture only a fraction of the total secreted transmitter. Previous theoretical and experimental descriptions of distant release events predict attenuation in quantal size with concurrent lengthening of rise-time and half-width values (Bruns et al. 2000; Haller et al. 1998; Schroeder et al. 1992). The rapid rise times of small events, shown in Fig. 5, are not consistent with release at a distant site. Furthermore, Fig. 6B shows that the smallest events, had the fastest rise times and half-widths of all the small events (also see Fig. 5I1). Kinetic values grew with increasing quantal size over the range of small events (0.2–0.4 pC1/3); this is the opposite relationship predicted for distant release events. Our data are consistent with the Bruns et al. (2000)'s model, which suggests that diffusional losses do not significantly alter size distributions. Therefore we believe that the small quantal events reported here were faithfully sampled with our recording configuration.
Previous studies found that DCVs in both norepinephrine- and epinephrine-storing cells exhibited broad size distributions (Gorgas and Bock 1976; Kobayashi and Coupland 1977). Gorgas and Bock (1976) reported that a cell type, which resembled norepinephrine-storing chromaffin cells (also see Thureson-Klein et al. 1984), possessed a vesicle size distribution that was skewed toward large vesicles. In addition, our DCV size distribution histogram is similar to that shown in Coupland (1968) when one looks at norepinephrine storing cells alone. We found vesicles where the membrane fit tightly while in others the membrane fit more loosely, as previously described (Gorgas and Bock 1976). Surprisingly, because we did not select for either epinephrine or norepinephrine storing cells, all the cells in our study had a majority of vesicles with intensely osmiophilic dense cores, a characteristic of norepinephrine storing DCVs; thus it appears that our EM preparations lacked epinephrine cells which are thought to represent 55% of the cells in the gland (Gorgas and Bock 1976).
A recent study provided evidence for two types of vesicles in rat adrenal chromaffin cells; in one type, the vesicular membrane fit tightly around the dense cores, whereas in the other, the vesicular membrane was loosely fitting, leaving an electron-lucent halo (Pothos et al. 2002). Although, we did not attempt to categorize the vesicles into two groups based on their appearance as done by Pothos et al. (2002), large vesicles tended to have halos. Their study also reported that there was an increased number of vesicles with a halo after a 40-min stimulation (Pothos et al. 2002). The length of the stimulation used in that study makes it difficult to interpret whether the different types of vesicles are lost at different rates, or if some of the vesicles were newly formed. It is important to note that the rate of recycling for some DCVs has been reported to occur in <10 min (Bauer et al. 2004a,b), and de novo, TGN-derived, DCV synthesis has been suggested to occur in 30 min in PC-12 cells (Tooze et al. 1991).
Previous amperometric studies in either murine or bovine chromaffin cells posited the existence of a single, broad distribution of Q1/3 values. Murine chromaffin cells were reported to have mean charge and mean amplitude values similar to our own (Colliver et al. 2000; Sorensen et al. 2003); however, these studies did not present the data as a distribution. Bovine chromaffin cells also have broad Q1/3 distributions (Albillos et al. 1997; Glavinovic et al. 1998; Wightman et al. 1995). Therefore the data presented in this study are not qualitatively different from these earlier studies. We fit Q1/3 with a bimodal distribution, even though these earlier studies did not, because our statistical analysis indicated that the sum of two Gaussians yielded a significantly better fit than did a single Gaussian, and the addition of a third Gaussian did not improve the fit (Table 2). Because bimodal Q1/3 distributions were observed in individual cells as well as ensembles of cells, the results cannot be explained as separate distributions originating in different types of chromaffin cells. Our data are consistent with a recent study by Tang et al. (2005), who also concluded that rat adrenal chromaffin cells possess multiple populations of vesicles.
The data in this manuscript show a tight correlation between vesicle size (as measured by EM with appropriate corrections) and Q1/3. The simplest explanation for the tight correlation between vesicle sizes as measured with EM compared with amperometry is that the stimuli used in this study results in complete empting of vesicles. Other studies have also found a close relationship between quantal size and vesicle size in cells that possess small and large secretory vesicles (Bruns et al. 2000; Chen et al. 1995). Bruns et al. (2000), using amperometric and EM methods, distinguished three different vesicle populations in leech neurons. Due to a segregated cellular distribution of the three kinds of vesicles, they were able to pair morphological and secretion data to convincingly show a linear relationship between vesicle size and quantal size. Karunanithi et al. (2002) showed that the two types of nerve terminals that form the Drosophila neuromuscular junction were distinguishable by their vesicle sizes, which had mean diameters of 41 and 48 nm. This difference in vesicle diameter leads to a 70% increase in vesicle volume and a corresponding increase in quantal size (Karunanithi et al. 2002). These same authors also showed that some individual synaptic boutons exhibited more than one class of quantal event (Wong et al. 1999). Proteins that regulate vesicle formation may change vesicle size (for review, see Murthy and De Camilli 2003). Alterations in the expression levels of adaptor proteins, which are known to be involved in vesicle formation, can modify vesicle size (Fergestad et al. 1999; Karunanithi et al. 2002; Morgan et al. 1999; Nonet et al. 1999; Zhang et al. 1998), which gives rise to a change in quantal size (Fergestad et al. 1999; Karunanithi et al. 2002; Zhang et al. 1998). Please note that incomplete emptying of the vesicles would introduce errors into our calculations of vesicle size and neurotransmitter concentration, using amperometric data. It would not change the observation that chromaffin cells possess two populations of vesicles.
Amperometric events from the two vesicle populations exhibited distinct kinetic properties. The amperometric spike's amplitude for small quantal events increased slowly with increasing quantal size, whereas large events exhibited a much larger increase in amplitude as a function of quantal size. These unique kinetics may reflect differences in rates of fusion pore dilation or possibly different rates of transmitter escape from the DCV's protein matrix (for review of each possibility, Travis and Wightman 1998).
One could ask why have two populations of vesicles if they both store the same neurotransmitter? If the large vesicles are less fusigenic, then weak stimulation would preferentially release small vesicles, whereas strong stimulation would release both large and small vesicles. In this way cells would have tight control of low levels of release, as each vesicle that fused with the plasma membrane only releases a small amount of transmitter, but strong stimulation is able to release large amounts of catecholamines for rapid responses to stress or danger etc. A similar mechanism has been observed in parotid salivary cells where distinct vesicles are believed to mediate basal and stimulated secretion (Huang et al. 2001). Similarly, Aplysia bag cells release different peptide products from distinct vesicle classes (Sossin et al. 1990).
Whether neurotransmitter is released via full-fusion or a kiss-and-run mechanism and whether vesicles empty completely, or not, are still controversial topics. Cell-attached capacitance studies have functionally defined full-fusion as a patch of vesicle membrane stably incorporated into the plasma membrane for seconds; in a fraction of these events, it was possible to measure a conductance for the fusion pore prior to final dilation (Albillos et al. 1997; Klyachko and Jackson 2002; Tabares et al. 2003). On the other hand, kiss-and-run events were defined as reversible changes in membrane capacitance; such events occurred much less frequently (Albillos et al. 1997; Klyachko and Jackson 2002). Other techniques, though, provide strong evidence for kiss-and-run (Taraska et al. 2003). A cell-attached capacitance-amperometry study was able to shift release between full-fusion and kiss-and-run by varying the Ca2+ concentration. In this study, Ales et al. (1999) were able to demonstrate that under recording conditions that favored kiss-and-run, the quantal size and vesicle size maintained a linear relationship just as it had under conditions that favored full-fusion (Ales et al. 1999). These results argued that vesicles emptied completely.
Although examples of complete release of transmitter have been proposed based on EM and amperometry results (Bruns et al. 2000; Colliver et al. 2000), other amperometry studies have yielded results that can be interpreted as partial emptying of vesicles. In dopamine neurons, flickers in the fusion pore result in the partial release of vesicular dopamine (Staal et al. 2004). The amount of secretion is variable in the porocytosis model as well (Kriebel et al. 2005). Elhamdani et al. (2001) recorded amperometric events while applying different patterns of electrical stimulation to chromaffin cells; they found that quantal size increased as a function of increased intracellular Ca2+. The strongest stimulation conditions were thought to reflect complete emptying of vesicles (Elhamdani et al. 2001). In our study, we used a relatively intense stimulus to elicit catecholamine release, which would be consistent with the complete emptying of the DCVs found in the chromaffin cells. Our data suggest that under these release conditions, the amperometric events are truly quantal and represent the neurotransmitter content of individual vesicles.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: C. P. Grabner, Dept. of Neurobiology, Pharmacology, and Physiology, The University of Chicago, 947 E. 58th St., Chicago, IL 60637 (E-mail: cpg22{at}email.med.yale.edu)
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REFERENCES |
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Ales E, Tabares L, Poyato JM, Valero V, Lindau M, and Alvarez de Toledo G. High calcium concentrations shift the mode of exocytosis to the kiss-and-run mechanism. Nat Cell Biol 1: 40–44, 1999.[CrossRef][Medline]
Anderson Ba. Model Selection and Multimodel Inference. A Practical Information-Theoretic Approach. New York: Springer-Verlag, 2002.
Arancio O, Kiebler M, Lee CJ, Lev-Ram V, Tsien RY, Kandel ER, and Hawkins RD. Nitric oxide acts directly in the presynaptic neuron to produce long- term potentiation in cultured hippocampal neurons. Cell 87: 1025–1035, 1996.[CrossRef][Web of Science][Medline]
Bauer RA, Khera RS, Lieber JL, and Angleson JK. Recycling of intact dense core vesicles in neurites of NGF-treated PC12 cells. FEBS Lett 571: 107–111, 2004a.[CrossRef][Web of Science][Medline]
Bauer RA, Overlease RL, Lieber JL, and Angleson JK. Retention and stimulus-dependent recycling of dense core vesicle content in neuroendocrine cells. J Cell Sci 117: 2193–2202, 2004b.
Bruns D, Riedel D, Klingauf J, and Jahn R. Quantal release of serotonin. Neuron 28: 205–220, 2000.[CrossRef][Web of Science][Medline]
Chen G, Gavin PF, Luo G, and Ewing AG. Observation and quantitation of exocytosis from the cell body of a fully developed neuron in Planorbis corneus. J Neurosci 15: 7747–7755, 1995.
Chow RH, von Ruden L, and Neher E. Delay in vesicle fusion revealed by electrochemical monitoring of single secretory events in adrenal chromaffin cells. Nature 356: 60–63, 1992.[CrossRef][Medline]
Colliver TL, Pyott SJ, Achalabun M, and Ewing AG. VMAT-mediated changes in quantal size and vesicular volume. J Neurosci 20: 5276–5282, 2000.
Coupland RE. Determining sizes and distribution of sizes of spherical bodies such as chromaffin granules in tissue sections. Nature 217: 384–388, 1968.[CrossRef][Medline]
Coupland RE and Hopwood D. Mechanism of a histochemical reaction differentiating between adrenaline- and noradrenaline-storing cells in the electron microscope. Nature 209: 590–591, 1966a.[CrossRef][Medline]
Coupland RE and Hopwood D. The mechanism of the differential staining reaction for adrenaline-and noreadrenaline-storing granules in tissues fixed in glutaraldehyde. J Anat 100: 227–243, 1966b.[Web of Science][Medline]
Duncan RR, Greaves J, Wiegand UK, Matskevich I, Bodammer G, Apps DK, Shipston MJ, and Chow RH. Functional and spatial segregation of secretory vesicle pools according to vesicle age. Nature 422: 176–180, 2003.[CrossRef][Medline]
Dunn LA and Holz RW. Catecholamine secretion from digitonin-treated adrenal medullary chromaffin cells. J Biol Chem 258: 4989–4993, 1983.
Elhamdani A, Palfrey HC, and Artalejo CR. Quantal size is dependent on stimulation frequency and calcium entry in calf chromaffin cells. Neuron 31: 819–830, 2001.[CrossRef][Web of Science][Medline]
Fergestad T, Davis WS, and Broadie K. The stoned proteins regulate synaptic vesicle recycling in the presynaptic terminal. J Neurosci 19: 5847–5860, 1999.
Finnegan JM, Pihel K, Cahill PS, Huang L, Zerby SE, Ewing AG, Kennedy RT, and Wightman RM. Vesicular quantal size measured by amperometry at chromaffin, mast, pheochromocytoma, and pancreatic beta-cells. J Neurochem 66: 1914–1923, 1996.[Web of Science][Medline]
Glavinovic MI, Vitale ML, and Trifaro JM. Comparison of vesicular volume and quantal size in bovine chromaffin cells. Neuroscience 85: 957–968, 1998.[CrossRef][Web of Science][Medline]
Gorgas K and Bock P. Morphology and histochemistry of the adrenal medulla. I. Various types of primary catecholamine-storing cells in the mouse adrenal medulla. Histochemistry 50: 17–31, 1976.[CrossRef][Web of Science][Medline]
Graham ME, O'Callaghan DW, McMahon HT, and Burgoyne RD. Dynamin-dependent and dynamin-independent processes contribute to the regulation of single vesicle release kinetics and quantal size. Proc Natl Acad Sci USA 99: 7124–7129, 2002.
Haller M, Heinemann C, Chow RH, Heidelberger R, and Neher E. Comparison of secretory responses as measured by membrane capacitance and by amperometry. Biophys J 74: 2100–2113, 1998.[Web of Science][Medline]
Henkel AW, Horstmann H, and Henkel MK. Direct observation of membrane retrieval in chromaffin cells by capacitance measurements. FEBS Lett 505: 414–418, 2001.[CrossRef][Web of Science][Medline]
Heynen AJ, Quinlan EM, Bae DC, and Bear MF. Bidirectional, activity-dependent regulation of glutamate receptors in the adult hippocampus in vivo. Neuron 28: 527–536, 2000.[CrossRef][Web of Science][Medline]
Holz RW, Bittner MA, Peppers SC, Senter RA, and Eberhard DA. MgATP-independent and MgATP-dependent exocytosis. Evidence that MgATP primes adrenal chromaffin cells to undergo exocytosis. J Biol Chem 264: 5412–5419, 1989.
Huang AY, Castle AM, Hinton BT, and Castle JD. Resting (basal) secretion of proteins is provided by the minor regulated and constitutive-like pathways and not granule exocytosis in parotid acinar cells. J Biol Chem 276: 22296–306, 2001.
Jankowski JA, Schroeder TJ, Ciolkowski EL, and Wightman RM. Temporal characteristics of quantal secretion of catecholamines from adrenal medullary cells. J Biol Chem 268: 14694–14700, 1993.
Jankowski JA, Schroeder TJ, Holz RW, and Wightman RM. Quantal secretion of catecholamines measured from individual bovine adrenal medullary cells permeabilized with digitonin. J Biol Chem 267: 18329–18335, 1992.
Karunanithi S, Marin L, Wong K, and Atwood HL. Quantal size and variation determined by vesicle size in normal and mutant Drosophila glutamatergic synapses. J Neurosci 22: 10267–10276, 2002.
Kirkup L and Mulholland M. Comparison of linear and non-linear equations for univariate calibration. J Chromatogr A 1029: 1–11, 2004.[CrossRef][Web of Science][Medline]
Klyachko VA and Jackson MB. Capacitance steps and fusion pores of small and large-dense-core vesicles in nerve terminals. Nature 418: 89–92, 2002.[CrossRef][Medline]
Kobayashi S and Coupland RE. Two populations of microvesicles in the SGC (small granule chromaffin) cells of the mouse adrenal medulla. Arch Histol Jpn 40: 251–259, 1977.[Medline]
Kobayashi S and Coupland RE. Morphological aspects of chromaffin tissue: the differential fixation of adrenaline and noradrenaline. J Anat 183: 223–235, 1993.[Web of Science][Medline]
Kriebel ME, Keller B, Silver RB, and Pappas GD. Porocytosis: a transient pore array secretes the neurotransmitter packet. Anat Rec B New Anat 282: 38–41, 2005.[Medline]
Kullmann DM and Nicoll RA. Long-term potentiation is associated with increases in quantal content and quantal amplitude. Nature 357: 240–244, 1992.[CrossRef][Medline]
Lee HK, Takamiya K, Han JS, Man H, Kim CH, Rumbaugh G, Yu S, Ding L, He C, Petralia RS, Wenthold RJ, Gallagher M, and Huganir RL. Phosphorylation of the AMPA receptor GluR1 subunit is required for synaptic plasticity and retention of spatial memory. Cell 112: 631–643, 2003.[CrossRef][Web of Science][Medline]
Liao D, Jones A, and Malinow R. Direct measurement of quantal changes underlying long-term potentiation in CA1 hippocampus. Neuron 9: 1089–1097, 1992.[CrossRef][Web of Science][Medline]
Malinow R. AMPA receptor trafficking and long-term potentiation. Philos Trans R Soc Lond B Biol Sci 358: 707–714, 2003.
Malinow R and Malenka RC. AMPA receptor trafficking and synaptic plasticity. Annu Rev Neurosci 25: 103–126, 2002.[CrossRef][Web of Science][Medline]
Malinow R and Tsiem RW. Presynaptic enhancement shown by whole-cell recording of long-term potentiation in hippocampal slices. Nature 346: 177–180, 1990.[CrossRef][Medline]
Morgan JR, Zhao X, Womack M, Prasad K, Augustine GJ, and Lafer EM. A role for the clathrin assembly domain of AP180 in synaptic vesicle endocytosis. J Neurosci 19: 10201–10212, 1999.
Murthy VN and De Camilli P. Cell biology of the presynaptic terminal. Annu Rev Neurosci 26: 701–728, 2003.[CrossRef][Web of Science][Medline]
Nonet ML, Holgado AM, Brewer F, Serpe CJ, Norbeck BA, Holleran J, Wei L, Hartwieg E, Jorgensen EM, and Alfonso A. UNC-11, a Caenorhabditis elegans AP180 homologue, regulates the size and protein composition of synaptic vesicles. Mol Biol Cell 10: 2343–2360, 1999.
Obukhov AG and Nowycky MC. TRPC4 can be activated by G-protein-coupled receptors and provides sufficient Ca(2+) to trigger exocytosis in neuroendocrine cells. J Biol Chem 277: 16172–16178, 2002.
Parsons TD, Coorssen JR, Horstmann H, and Almers W. Docked granules, the exocytic burst, and the need for ATP hydrolysis in endocrine cells. Neuron 15: 1085–1096, 1995.[CrossRef][Web of Science][Medline]
Pothos EN, Mosharov E, Liu KP, Setlik W, Haburcak M, Baldini G, Gershon MD, Tamir H, and Sulzer D. Stimulation-dependent regulation of the pH, volume and quantal size of bovine and rodent secretory vesicles. J Physiol 542: 453–476, 2002.
Sakai T. Relation between thickness and interference colors of biological ultrathin section. J Electron Microsc 29: 369–375, 1980.
Schroeder TJ, Jankowski JA, Kawagoe KT, Wightman RM, Lefrou C, and Amatore C. Analysis of diffusional broadening of vesicular packets of catecholamines released from biological cells during exocytosis. Anal Chem 64: 3077–3083, 1992.[Medline]
Sorensen JB, Nagy G, Varoqueaux F, Nehring RB, Brose N, Wilson MC, and Neher E. Differential control of the releasable vesicle pools by SNAP-25 splice variants and SNAP-23. Cell 114: 75–86, 2003.[CrossRef][Web of Science][Medline]
Sossin WS, Sweet-Cordero A, and Scheller RH. Dale's hypothesis revisited: different neuropeptides derived from a common prohormone are targeted to different processes. Proc Natl Acad Sci USA 87: 4845–4848, 1990.
Staal RG, Mosharov EV, and Sulzer D. Dopamine neurons release transmitter via a flickering fusion pore. Nat Neurosci 7: 341–346, 2004.[CrossRef][Web of Science][Medline]
Stevens CF and Wang Y. Changes in reliability of synaptic function as a mechanism for plasticity. Nature 371: 704–707, 1994.[CrossRef][Medline]
Tabares L, Lindau M, and Alvarez de Toledo G. Relationship between fusion pore opening and release during mast cell exocytosis studied with patch amperometry. Biochem Soc Trans 31: 837–841, 2003.[CrossRef][Web of Science][Medline]
Tang KS, Tse A, and Tse FW. Differential regulation of multiple populations of granules in rat adrenal chromaffin cells by culture duration and cyclic AMP. J Neurochem 92: 1126–1139, 2005.[CrossRef][Web of Science][Medline]
Taraska JW, Perrais D, Ohara-Imaizumi M, Nagamatsu S, and Almers W. Secretory granules are recaptured largely intact after stimulated exocytosis in cultured endocrine cells. Proc Natl Acad Sci USA 100: 2070–2075, 2003.
Thureson-Klein A, Harless S, and Klein R. Ultrastructural changes in adrenaline- and SGC-cells after morphine coincide with alterations of adrenaline and dopamine levels. Cell Tissue Res 236: 53–65, 1984.[CrossRef][Web of Science][Medline]
Tooze SA, Flatmark T, Tooze J, and Huttner WB. Characterization of the immature secretory granule, an intermediate in granule biogenesis. J Cell Biol 115: 1491–1503, 1991.
Travis ER and Wightman RM. Spatio-temporal resolution of exocytosis from individual cells. Annu Rev Biophys Biomol Struct 27: 77–103, 1998.[CrossRef][Web of Science][Medline]
Tsien JZ, Huerta PT, and Tonegawa S. The essential role of hippocampal CA1 NMDA receptor-dependent synaptic plasticity in spatial memory. Cell 87: 1327–1338, 1996.[CrossRef][Web of Science][Medline]
Wick PF, Trenkle JM, and Holz RW. Punctate appearance of dopamine-beta-hydroxylase on the chromaffin cell surface reflects the fusion of individual chromaffin granules upon exocytosis. Neuroscience 80: 847–860, 1997.[CrossRef][Web of Science][Medline]
Wightman RM, Jankowski JA, Kennedy RT, Kawagoe KT, Schroeder TJ, Leszczyszyn DJ, Near JA, Dilberto Jr, EJ, and Viveros OH. Temporally resolved catecholamine spikes correspond to single vesicle release from individual chromaffin cells. Proc Natl Acad Sci USA 88: 10754–10758, 1991.
Wightman RM, Schroeder TJ, Finnegan JM, Ciolkowski EL, and Pihel K. Time course of release of catecholamines from individual vesicles during exocytosis at adrenal medullary cells. Biophys J 68: 383–390, 1995.[Web of Science][Medline]
Wilson SP and Kirshner N. Calcium-evoked secretion from digitonin-permeabilized adrenal medullary chromaffin cells. J Biol Chem 258: 4994–5000, 1983.
Wong K, Karunanithi S, and Atwood HL. Quantal unit populations at the Drosophila larval neuromuscular junction. J Neurophysiol 82: 1497–1511, 1999.
Zhang B, Koh YH, Beckstead RB, Budnik V, Ganetzky B, and Bellen HJ. Synaptic vesicle size and number are regulated by a clathrin adaptor protein required for endocytosis. Neuron 21: 1465–1475, 1998.[CrossRef][Web of Science][Medline]
Zucker RS and Regehr WG. Short-term synaptic plasticity. Annu Rev Physiol 64: 355–405, 2002.[CrossRef][Web of Science][Medline]
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ABSTRACT
INTRODUCTION
) and half-width (
) are separable into 2 components. Both rise time and half-width increase in value and follow one another closely over the range where small events predominate, from 0.2 to 0.4 pC1/3. Rise time and half-width decline slightly in the 0.6- to 0.85-pC1/3 range and then become more dispersed at larger values. Smaller events tend to have a slower rate of transmitter release relative to large events as seen in the rapid rise in amplitude for the large events. Data are presented as means ± SE.