Am J Physiol Cell Physiol AJP: Lung Cellular and Molecular Physiology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Cell Physiol 295: C624-C631, 2008. First published July 16, 2008; doi:10.1152/ajpcell.00303.2008
0363-6143/08 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
295/3/C624    most recent
00303.2008v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via Web of Science (1)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Jorgacevski, J.
Right arrow Articles by Zorec, R.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Jorgacevski, J.
Right arrow Articles by Zorec, R.

NERVOUS SYSTEM CELL BIOLOGY

Hypotonicity and peptide discharge from a single vesicle

Jernej Jorgacevski,1,* Matjaz Stenovec,1,2,* Marko Kreft,1,2 Aleksandar Bajic,3 Bostjan Rituper,1 Nina Vardjan,1,2 Stanko Stojilkovic,3 and Robert Zorec1,2

1Laboratory of Neuroendocrinology-Molecular Cell Physiology, Faculty of Medicine, University of Ljubljana, Ljubljana, Slovenia; 2Celica Biomedical Center, Ljubljana, Slovenia; and 3Section on Cellular Signaling, Developmental Neuroscience Program, National Institute of Child Health and Human Development, Bethesda, Maryland

Submitted 10 June 2008 ; accepted in final form 11 July 2008


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Neuroendocrine secretory vesicles discharge their cargo in response to a stimulus, but the nature of this event is poorly understood. We studied the release of the pituitary hormone prolactin by hypotonicity, because this hormone also contributes to osmoregulation. In perfused rat lactotrophs, hypotonicity resulted in a transient increase followed by a sustained depression of prolactin release, as monitored by radioimmunoassay. In single cells imaged by confocal microscopy, hypotonicity elicited discharge of the fluorescently labeled atrial natriuretic peptide cargo from ~2% of vesicles/cell. In contrast, KCl-induced depolarization resulted in a response of ~10% of vesicles/cell, with different unloading/loading time course of the two fluorescent probes. In cell-attached studies, discrete changes in membrane capacitance were recorded in both unstimulated and stimulated conditions, reflecting single vesicle fusion/fissions with the plasma membrane. In stimulated cells, the probability of occurrence of full fusion events was low and unchanged, whereas over 95% of fusion events were transient, with the open fusion pore probability, the average pore dwell-time, the frequency of occurrence, and the fusion pore conductance increased. Hypotonicity only rarely elicited new fusion events in silent membrane patches. The results indicate that, in hypotonicity-stimulated lactotrophs, transient vesicle fusion mediates hormone release.

prolactin; hormone secretion; rat lactotrophs; fusion pore; postfusion release regulation

AMONG ITS MANY FUNCTIONS, peptidergic hormone prolactin plays also a role in osmoregulation (14). Previous studies demonstrated prolactin release from isolated tissue (18, 34, 41, 42) or cells in response to hyposmotic stimulation (41, 50), which required extracellular calcium (23, 40, 41). While these findings suggest that prolactin is likely released by exocytosis in response to hyposmotic stress, secretory activity at the single vesicle level in response to hypotonicity was not monitored directly yet. In particular, it is unclear whether the hormone is released rapidly after complete collapse of vesicles into the plasma membrane (24) by full fusion exocytosis (21) or by kiss-and-run exocytosis (1, 13), whereby vesicles transiently fuse with the plasma membrane, forming a channel with the extracellular space, termed the fusion pore.

To address these questions, we studied rat pituitary lactotrophs, which release prolactin stored in a highly aggregated form in large dense-core secretory vesicles (10). To monitor secretory activity, we used perfusion experiments and radioimmunoassays and recorded single exocytotic and vesicle release events in real time, using confocal microscopy, styryl dyes, and fluorescently labeled peptides (2, 45). To directly study fusion pore properties, we used electrophysiological methods to monitor membrane capacitance (Cm), which is linearly related to changes in membrane area (30, 35, 48, 53). Experimental evidence indicates that fusion of secretory vesicles in lactotrophs occurs spontaneously and is facilitated by hypotonicity. Moreover, the results show that hypotonicity-induced hormone discharge from a single vesicle involves transient fusion pore openings.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Measuring prolactin release. For perfusion experiments, 1.2 x 107 lactotroph cells were incubated with preswollen Cytodex-1 beads in 60-mm petri dishes for 24 h. The beads were transferred to 0.5-ml chambers and perfused with Hanks' medium 199 for 90 min at 0.8 ml/min, at room temperature. During perfusion, cells were stimulated with hyposmolar solution or with KCl depolarization. Fractions were collected at 1-min intervals, stored at –20°C, and assayed for prolactin by radioimmunoassay. The primary antibody and standard for the prolactin assay were from the National Pituitary Agency (Dr. A. F. Parlow, Harbor-UCLA Medical Center, Torrance, CA), the secondary antibody was from Sigma, and 125I-labeled prolactin tracer was from PerkinElmer Life Sciences (Boston, MA).

Cell culture and microscopy. Pituitaries were obtained after decapitation from male Wistar rats and enriched for lactotrophs (5). The care of experimental animals was in accordance with the International Guiding Principles for Biomedical Research Involving Animals developed by the Council for International Organizations of Medical Sciences and the Directive on Conditions for Issue of License for Animal Experiments for Scientific Research Purposes (Official Gazette of the Republic of Slovenia 40/85 and 22/87). The animal use procedures were approved by the Veterinary administration of the Republic of Slovenia (approval no. 3440-29/2006). Briefly, lactotrophs were obtained with enzymatic dispersion, enriched with Percoll gradient, plated on glass coverslips coated with poly-L-lysine, and maintained in high-glucose Dulbecco's modified Eagle's medium (Invitrogen) supplemented with 10% newborn calf serum and L-glutamine in an atmosphere of humidified air (92%) and CO2 (8%). The level of lactotroph purification exceeded 80% (data not shown). A plasmid encoding atrial natriuretic peptide-enhanced green fluorescent protein (ANP-EGFP) (a gift from Dr. Ed Levitan) was introduced into the lactotrophs by lipofection (Invitrogen, Carlsbad, CA). Cell-loaded coverslips were placed into a recording chamber on an inverted confocal microscope (LSM 510; Zeiss, Jena, Germany), and extracellular solution (400 µl) containing 4 µM FM 4-64 (Molecular Probes, Leiden, Netherlands) was added. Fluorescence images were acquired with a plan-apochromatic oil-immersion objective (x63, 1.4 numerical aperture) using 488-nm Ar-Ion laser excitation and filtered with LP 580-nm emission filter.

Hypotonicity-induced cell swelling was monitored in the equatorial plane of the cells as changes in cell diameter. Initially, cells were kept in normal extracellular solution containing 4 µM FM 4-64. The osmolarity of the solution was then reduced to ~200 mosM by bath application of hyposmolar extracellular solution containing 4 µM FM 4-64. Cell swelling was measured as a change in the cell diameter, determined from fluorescence intensity line profiles drawn perpendicularly over the stained cells with LSM 510 software (Zeiss). The fluorescence peaks marked the cell boundary. The distance between the peaks was determined before and 2 min after hypotonic stimulation. Hypotonicity-induced hormone secretion from a single granule was studied by using FM 4-64 and ANP-EGFP; emission fluorescence was filtered with a 505- to 530-nm band-pass filter. In KCl depolarization experiments, cells were superfused with 100 mM KCl solution. Images were acquired every 493 ms (~2 Hz) and analyzed with an LSM 510 laser-scanning confocal microscope (Zeiss); custom software was written in MATLAB (Math Works, Natick, MA). The average fluorescence intensity changes were monitored within a circular region (diameter 14 pixels) at the locus of a single vesicle by measuring the time required for a 20% to 80% intensity change in the fluorescent signal.

Electrophysiology. Capacitance in attached cells was measured as described previously (26, 53, 45). Briefly, a sine wave voltage (1,591 Hz, 111 mV root mean square; SWAM IIC, Celica, Ljubljana, Slovenia) was applied to the pipette. The pipette potential was held at 0 mV. The resistance of thick-walled pipettes, which were fire polished and coated with Sylgard, was 2–6 M{Omega}. Responses were recorded under basal conditions (bath and pipette contained extracellular solution) in the first set of experiments and under both basal and stimulated conditions (hypotonic stimulation or KCl depolarization) in two other sets, typically for 7 min. During stimulation with hypotonic solution, a bolus of saline was added to reduce osmolarity to ~200 mosM. Stimulation with depolarization was achieved by adding a bolus solution to reach a final KCl concentration of 100 mM. The phase of the lock-in amplifier was adjusted to nullify the changes in the real (Re) portion of the admittance signal by manually generating 10-fF calibration pulses with an in built calibration in patch-clamp instrument.

Transient events in the Im portion of the admittance signal that exhibited measurable projections to the Re portion ("cross talk") were used to calculate fusion pore conductance [Gp = (Re2 + Im2)/Re] (32) and vesicle capacitance {Cv = [(Re2 + Im2)/Im]/{omega}, where {omega} denotes angular frequency}. Fusion pore diameter was determined from the equation Gp = ({pi}r2)/({rho}{lambda}), where r denotes fusion pore radius, {rho} is the resistivity of the saline (100 {Omega}·cm), and {lambda} is the estimated length of a gap junction channel (15 nm) (44). Vesicle diameter was determined from specific membrane capacitance (Cm = 8 fF/µm) and vesicle capacitance (Cv). All recordings were performed at room temperature.

Events were analyzed by using the cursor option in the software subroutine (CellAn, Celica, Slovenia) written for MATLAB. We considered an event to be reversible if an on-step in Im was followed by an off-step within 2 s. The appearance of events was ascertained by progressive filtering. We considered an event to be detectable if the signal-to-noise ratio was at list 3:1 and the event did not exhibit a projection in the current trace. Sigma Plot was used for statistical analysis. Values are presented as means ± SE. Differences between samples were analyzed with t-tests unless stated otherwise.

Solutions. Extracellular solution contained (in mM) 130 NaCl, 5 KCl, 8 CaCl2, 1 MgCl2, 10 D-glucose, 10 HEPES (N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid), and pH 7.2/NaOH. Hypotonic extracellular solution used for cell perfusion contained (in mM) 90 NaCl, 5 KCl, 8 CaCl2, 1 MgCl2, 10 D-glucose, 10 HEPES, and pH 7.2/NaOH. Hypotonic extracellular solution added as bolus to reduced osmolarity of the extracellular solution contained (in mM) 5 KCl, 8 CaCl2, 1 MgCl2, 10 D-glucose, 10 HEPES, and pH 7.2/NaOH. KCl solution added as a bolus contained (in mM) 5 NaCl, 130 KCl, 8 CaCl2, 1 MgCl2, 10 D-glucose, 10 HEPES, and pH 7.2/NaOH. A 100 mM KCl solution was prepared by replacing 95 mM NaCl in the standard saline solution with 95 mM KCl. Osmolarity was measured with a freezing-point osmometer (Osmomat030; Gonotech, Berlin, Germany). Chemicals were from Sigma (Darmstadt, Germany) and were of highest grade of purity available.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Hypotonicity, cell volume increase, and prolactin release. Pituitary lactotrophs of lower vertebrates respond to reduced extracellular osmolarity by increasing prolactin release (18, 34, 41, 42), which reduces ion and water permeability and increases solute retention (7, 22, 33). However, whether hypotonicity affects prolactin release from mammalian lactotrophs is unclear. Here we exposed rat lactotrophs to hypotonicity of ~200 mosM, which is encountered clinically (3, 9).

Under hyposmotic conditions, cells generally swell and undergo a regulatory volume decrease (27). Consistent with previous reports (41), hypotonicity increased the volume of rat lactotrophs, measured by fluorescence intensity line profiles (Fig. 1, A–C) in the presence of FM 4-64, a membrane area marker (39). Hypotonicity increased the interpeak distance, an estimate of cell diameter, by 0.58 ± 0.13 µm (Fig. 1D), significantly different from zero (P < 0.001) (from 10.65 ± 0.16 µm to 11.23 ± 0.22 µm, n = 28), and FM 4-64 peak intensity; regular extracellular solution elicited no such changes (Fig. 1C).


Figure 1
View larger version (14K):
[in this window]
[in a new window]

  		Fig. 1. Hypotonic stimulation induced cell swelling. A: confocal images of isolated lactotroph stained with 4 µM FM 4-64 before (0 s) and 2 min after bath application of regular extracellular solution. Scale bar = 5 µm. B: confocal images of a lactotroph before and after hypotonic stimulation. Note an increase in the cell diameter 120 s after hypotonic stimulation. Thin lines perpendicular to the plasma membrane indicate where fluorescence intensity profiles were obtained. C: superimposed fluorescence intensity profiles in the cell before (black line) and 120 s after (gray line) stimulation with normal extracellular solution (left) and hypotonic extracellular solution (right). Note a clear increase in the cell diameter indicated by the shift of the fluorescence intensity peaks, which mark the cell perimeter. AU, arbitrary units. D: cell diameter change (means ± SE) in controls and in cells stimulated by hypotonic extracellular solution (Hypo). Numbers adjacent to the bars indicate the number of cells analyzed. *P < 0.001 vs. control.

 
Perfused cells spontaneously released prolactin at room temperature, and hypotonicity evoked a transient increase in prolactin release, followed by a sustained depression (Fig. 2), likely reflecting reduced basal spontaneous secretion, which is controlled by spontaneous voltage-gated calcium influx (47). The transient increase did not reflect depletion of the pool of readily releasable vesicles, since KCl depolarization of cells yielded a sustained increase in prolactin release that continued after the stimulus was discontinued (Fig. 2).


Figure 2
View larger version (28K):
[in this window]
[in a new window]

  		Fig. 2. Prolactin release (PRL) induced by hyposmolarity and depolarization in perfused pituitary cells. Both profiles are representative of four independent experiments. Shaded area indicates the duration of stimulation.

 
Hypotonicity and hormone discharge from a single vesicle. To determine whether hypotonicity induces hormone discharge from a single vesicle, we transfected cells with a construct that encodes ANP-EGFP and is targeted to secretory vesicles (19). In transfected lactotrophs, ANP-EGFP fluorescence (Fig. 3A) had a punctate appearance and colocalized with prolactin-containing vesicles, as reported previously (45).


Figure 3
View larger version (34K):
[in this window]
[in a new window]

  		Fig. 3. Induced hormone release from single vesicles. A: confocal image of hypotonicity triggered peptide release from a lactotroph granule at time of fusion (0 s; left) and 40 s later (right). Scale bar: 5 µm. Bottom: atrial natriuretic peptide-enhanced green fluorescent protein (ANP-EGFP; green) and FM 4-64 (red) fluorescence from a selected vesicle (squares at top). Scale bar: 0.5 µm. B: increase in FM 4-64 fluorescence (top) normalized to minimal and maximal fluorescence (thin solid lines) and ANP-EGFP fluorescence decrease (black trace, bottom) in response to hypotonic stimulation. Gray trace indicates baseline fluorescence before opening of the vesicle fusion pore and increase in fluorescence due to pH neutralization after fusion pore opening. Dotted lines represent 20% and 80% fluorescence intensity levels. C: the rise times ({tau}20-80%, means ± SE) of FM 4-64 and ANP-EGFP fluorescence. Numbers adjacent to bars indicate the number of vesicles analyzed. D: high potassium triggered fluorescent peptide release from lactotroph vesicles displayed at stimulation onset (0 s; left) and 40 s after (right). Scale bar: 5 µm. Note substantial increase in number of labeled vesicles that discharged fluorescent peptides and were loaded with FM 4-64 (circles at right) in comparison to vesicle cargo discharge in A. White arrowheads indicate selected initially nonfluorescing vesicles that fused with the plasma membrane upon stimulation and were loaded with FM 4-64. Bottom: ANP-EGFP (green) and FM 4-64 (red) fluorescence in a selected vesicle (squares at top). Scale bar: 0.5 µm. E: FM 4-64 fluorescence increase (top) normalized to minimal and maximal fluorescence (thin solid lines) and ANP-EGFP fluorescence decrease (black trace, bottom) in response to 100 mM K+. F: the rise times ({tau}20-80%, means ± SE) of FM 4-64 and ANP-EGFP signals. Numbers adjacent to bars indicate the number of vesicles analyzed. *P < 0.01.

 
To analyze the properties of stationary vesicles at the plasma membrane, we monitored changes in the fluorescence intensity of labeled vesicles (Fig. 3A, insets). In lactotrophs exposed to hypotonic solution, ANP-EGFP fluorescence displayed complex time-dependent changes. Initially, fluorescence intensity increased. Since EGFP is pH sensitive (38), the increase likely indicates alkalinization of the vesicle lumen after the exposure to the extracellular space through the fusion pore. Thereafter, fluorescence intensity decreased, likely reflecting ANP-EGFP discharge into the extracellular space (Fig. 3B, bottom), since FM 4-64 fluorescence increased simultaneously (45; Fig. 3B, top). FM-dyes are widely used to study exo-/endocytosis, given that these amphiphilic styryl fluorescent dyes are virtually nonfluorescent in water but show a dramatic increase in the quantum yield upon incorporation into the cell membranes or any other hydrophobic environment (6).

In 4 of 11 image planes at cell base, 7 of 448 vesicles (2 ± 1 vesicles/cell image) that were initially labeled by ANP-EGFP and exhibited green fluorescence dimming were loaded with FM 4-64. This corresponds to hypotonicity-induced release from 2.4 ± 1.6% (n = 4) of ANP-EGFP-labeled vesicles determined as a fraction (%) of responding vesicles per cell image. Measurements of the time required for a 20–80% change of the fluorescent signal ({tau}20-80%) revealed that ANP-EGFP fluorescence decayed in 5.5 ± 1.4 s, which appeared slower than vesicle loading with FM 4-64 (2.8 ± 0.6 s; Fig. 3C), however, not significantly different (P = 0.23, n = 7). Upon hypotonic stimulation in four cells, eight vesicles loaded FM 4-64 with a {tau}20-80% of 2.6 ± 0.5 s (not shown) but did not fluoresce green before stimulation, consistent with the view that not all exocytotic vesicles are preloaded with ANP-EGFP (45).

KCl depolarization generated a much more robust response (Fig. 3D). In 9 of 10 image planes at cell base, 66 of 585 ANP-EGFP-labeled vesicles (13 ± 7 vesicles/cell image) discharged their cargo and were loaded with FM 4-64 upon KCl depolarization (Fig. 3E); this corresponds to fluorescent cargo release from 9.8 ± 3.2% of ANP-EGFP-labeled vesicles. Thus, four- to fivefold more vesicles discharged ANP-EGFP and were loaded with FM 4-64 in response to KCl depolarization than in response to the hypotonic solution. The ANP-EGFP fluorescence decay (5.0 ± 0.5 s) differed significantly from the increase in FM 4-64 fluorescence (3.1 ± 0.3 s) in KCl-depolarized vesicles (P < 0.01, n = 66; Fig. 3F), consistent with our previous report (45). These results show that hypotonicity induces cargo release from vesicles associated with the plasma membrane. However, a significantly smaller fraction of vesicles responded to hypotonicity than to KCl depolarization.

Transient fusion pore events and hypotonicity. Next we studied fusion pore properties. For this, high-resolution changes in Cm, which are related to surface area changes, were recorded. Discrete steps in Cm correspond to elementary membrane fusion and fission events (35, 48). In 11 of 34 cells exposed to hypotonicity (total recording time, 5.06 h; average, 536 s per patch), we observed spontaneous discrete steps in Cm of around 1 fF (range, 0.4 to 6 fF). In 10 of the cells, discrete steps were seen under resting conditions and after stimulation. In one cell, hypotonicity induced discrete Cm steps in a previously silent patch. In addition, the spontaneous discrete Cm steps were silenced after stimulation in one cell. Thus, rather than generating new discrete steps, hypotonicity modulates the activity of discrete Cm steps present under nonstimulated conditions.

Irreversible Cm steps, representing full fusion/fission (Fig. 4A, left), and reversible Cm steps (Fig. 4A, right), likely representing transient kiss-and-run events, were observed in nonstimulated and hypotonic-stimulated cells (Fig. 4B). Reversible Cm steps in the imaginary part (Im) of the admittance signal were often accompanied by projections to the real part (Re) of the admittance signal (Fig. 4B). The amplitude distribution of the upward steps and of the downward steps that followed within 2 s was best fitted with a regression line (n = 619; correlation coefficient r = 0.93; Fig. 4C), with a slope near 1. This finding supports the idea that reversible Cm steps reflect the fusion and subsequent fission of a single vesicle. The amplitude of upward Cm steps, corresponding to Cv (Fig. 4D), were similar before and after hypotonic stimulation (Kolomogorov-Smirnov test, P > 0.05; n = 179 and n = 440, respectively). Thus, hypotonicity had little effect on the amplitude of these events. The calculated diameter of vesicles was 150–450 nm (Fig. 4D), consistent with the size of prolactin-containing vesicles in lactotrophs (2, 43, 53).


Figure 4
View larger version (34K):
[in this window]
[in a new window]

  		Fig. 4. Capacitance steps under control (isosmotic) and hyposmotic conditions in lactotrophs. A: irreversible (left) and reversible (right) capacitance steps were observed under both conditions. B: capacitance steps were more frequent under hyposmotic conditions. Shown is a 10-s epoch at spontaneous conditions (left) and a 10-s epoch recorded 4 min after the stimulation (right). Note that transient Im steps often exhibit cross talk with the Re trace even though the phase angle was set correctly (asterisks indicate calibration pulse in Im trace of 10 fF without projection to the Re signal). Transient events with detectable projections were used to calculate fusion pore conductance (Gp) and vesicle capacitance (Cv) (shown in insets, after hyposmotic stimulation). C: correlation between the amplitudes of downward and upward steps; the line represents the best fit of the form: [y = (0.11 ± 0.05) + (0.92 ± 0.01) x x; correlation coefficient r = 0.93; n = 619]. D: histogram showing the frequency distribution of Cv of reversible events before (n = 179) and after (n = 440) the hyposmotic shock (by Kolomogorov-Smirnov test; P > 0.05). Top abscissa shows the corresponding vesicle diameters, estimated by assuming that vesicles are spheres and the specific membrane capacitance is 8 fF/µm2.

 
To study whether transient fusion proceeds into full fusion (12, 15), we analyzed fusion events and the mode of secretory activity after hypotonic stimulation and after KCl depolarization. The frequency of transient events, measured in 10-s epochs, increased from 0.07 ± 0.01 (n = 761 events in 895 epochs) to 0.10 ± 0.01 events/s (P < 0.05; n = 440 events in 428 epochs) after hypotonic stimulation but was sixfold higher after KCl depolarization (0.66 ± 0.06 events/s; P < 0.01; n = 2,372 events in 366 epochs) (Fig. 5A). The frequency of irreversible Cm steps increased only slightly, however, not significantly different (P > 0.05), and was considerably lower than the frequency of transient events occurring spontaneously (0.006 ± 0.001 events/s; n = 35 events) or in response to hypotonicity (0.007 ± 0.001; n = 29 events) or KCl depolarization (0.009 ± 0.001; n = 32 events) (Fig. 5A). Therefore, transient, reversible fusion events were the predominant mode of exocytosis before and after stimulation, consistent with a previous report (48).


Figure 5
View larger version (19K):
[in this window]
[in a new window]

  		Fig. 5. Hyposmotic stress and KCl depolarization increases the appearance of reversible but not irreversible steps. A: mean frequencies of reversible and irreversible events at spontaneous conditions (0.074 ± 0.007 events/s for reversible events and 0.006 ± 0.001 events/s for irreversible events), after hypotonic stimulation (0.101 ± 0.012 events/s for reversible events/s and 0.007 ± 0.001 events/s for irreversible events), and after KCl depolarization (0.655 ± 0.063 events/s for reversible and 0.009 ± 0.001 events/s for irreversible events). Numbers above bars indicate the number of events. *P < 0.05 and **P < 0.01. B: relationship between the average Cv before and after stimulations. Plots were fitted with regression lines [y = (–0.08 ± 0.38) + (1.14 ± 0.25) x x; r = 0.85; n = 10 patches for hypotonic stimulation, and y = (–0.06 ± 0.31) + (1.05 ± 0.17) x x; r = 0.93; n = 8 patches for KCl depolarization].

 
To further test whether the repetitive transient fusion events can be attributed to the same vesicles in particular patches, we analyzed the amplitude of Cm steps in each recording separately. Plots of average Cv amplitude before and after stimulation show a high correlation coefficient (r = 0.85 for hypotonicity and r = 0.93 for depolarization; n = 10 and 8 patches, respectively) and regression line slopes near 1 (1.14 ± 0.25 and 1.05 ± 0.17, respectively; Fig. 5B). Therefore, the average Cv amplitudes before and after stimulation were similar.

Hypotonicity increases fusion pore dwell-time and conductance. The secretory output from a cell can be increased in several ways. It can increase by an increase in the frequency of fusion events (Fig. 5A), but also by a prolonged fusion pore dwell-time and a wider fusion pore diameter. Hypotonicity prolonged the fusion pore dwell-time (0.16 ± 0.01 vs. 0.20 ± 0.01 s; P < 0.05; n = 716 events and n = 440 events, respectively; Fig. 6A) to approximately the same extent as depolarization (0.20 ± 0.01 s; n = 2,372 events), consistent with a previous report (48). The probability of observing an open fusion pore (calculated as the sum of all pore dwell-times divided by the duration of each recording) increased from 0.012 ± 0.002 under resting conditions (n = 37 recordings) to 0.019 ± 0.005 (P < 0.05; n = 11 recordings) after hypotonic stimulation. An even greater increase was observed after KCl depolarization (0.110 ± 0.04; P < 0.01; n = 8 recordings) (Fig. 6B).


Figure 6
View larger version (19K):
[in this window]
[in a new window]

  		Fig. 6. Hyposmotic stress increases the average fusion pore conductance, pore dwell-time, and the probability of open fusion pore state. A: the average pore dwell-time increased significantly (0.16 ± 0.01 s at spontaneous conditions; n = 716 events) after hyposmotic stimulation (0.20 ± 0.01 s; n = 440 events) and KCl depolarization (0.20 ± 0.01 s; n = 2,372 events). Numbers above bars indicate the number of fusion events. B: hyposmotic stress increased the probability of observing an open fusion pore from 0.012 ± 0.002 (n = 37 patches) to 0.019 ± 0.005 (n = 11 patches), and depolarization increased the probability to 0.110 ± 0.04 (n = 8 patches). Numbers above bars indicate the number of patches. C: the mean Gp increased from 47 ± 4 pS (n = 261) to 73 ± 5 pS (n = 142) after hyposmotic stimulation and to 68 ± 4 (n = 200) pS after KCl depolarization. Numbers above bars indicate the number of fusion events with measurable Gp. *P < 0.05 and **P < 0.01.

 
Transient exocytosis could constrain peptide discharge from large dense-core vesicles owing to a narrow fusion pore (4, 8, 48, 49). Narrow fusion pore can act as a resistor in series and thus can reduce the signal in Im part of admittance by producing a correlated increase in Re part (cross talk) (32). Therefore, we analyzed transient fusion events with detectable cross talk to calculate the fusion pore conductance [Gp = (Re2 + Im2)/Re] and vesicle capacitance {Cv = [(Re2 + Im2)/Im]/{omega}} (32; Fig. 4C). The percentage of reversible events that exhibited cross talk was similar under isotonic, spontaneous (34%), and hypotonic conditions (32%). However, after KCl depolarization, only 8% of events exhibited cross talk. The mean fusion pore conductance (Gp) of spontaneous events with measurable cross talk was 47 ± 4 pS (range, 8–251 pS; n = 261 events). Gp increased to 73 ± 5 pS (range, 13–288 pS; P < 0.01; n = 142 events) after hypotonic simulation and to 68 ± 4 pS (range 9–537 pS; P < 0.01; n = 200 events) after KCl depolarization (Fig. 6C). Therefore, the mean pore diameter for events with measurable Gp increased from 0.94 ± 0.04 nm under spontaneous conditions to 1.18 ± 0.04 nm after hypotonic stimulation and to 1.14 ± 0.04 nm after KCl depolarization.


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The present study shows that mammalian lactotrophs release prolactin in a hypotonic environment and that this involves transient vesicle fusion pore openings.

Cell swelling and surface area changes in hypotonic conditions. Cells generally swell in response to hyposmotic conditions (27). Hypotonic stimulation of lactotrophs induced sustained cell swelling for more than 120 s. (Fig. 1). Using data on the cell diameter increase and assuming spherical geometry of lactotrophs, we estimate that the cell surface area increased by ~11% under our experimental conditions. Simultaneously, the peak fluorescence intensity profile also increased, indicating enhanced labeling of plasma membrane with FM 4-64. This may reflect exocytotic addition of vesicles into the membrane. After full fusion with the plasma membrane, a single spherical dense-core vesicle 200 nm in diameter (43) may increase the membrane surface by 0.126 µm2. Thus, an 11% increase in the surface area would correspond to ~78 vesicles fusing with the plasma membrane throughout the cell. Alternatively, increased staining of plasma membrane by FM 4-64 may be attributed to membrane expansion that changes local membrane curvature in response to hypotonic cell stimulation. Measurements with preloaded fluorescent peptides showed that one to five vesicles fused with the plasma membrane per 2-µm-thick optical slice (Fig. 3). In a cell 10–12 µm in diameter, this corresponds to a maximum of 25–30 fusing vesicles. These considerations support the notion that hypotonicity-induced cell swelling is associated with an increased surface membrane area. However, it is not clear why the actual number of vesicles that fused with the plasma membrane differed from the estimate. One possibility is that there is no simple relationship between cell swelling, surface membrane increase, and FM 4-64 staining, which is thought to reflect the membrane surface area (25). Another likely possibility is that hyposmotic challenge of lactotrophs resulted in expansion of the existing plasma membrane reservoirs-membrane folds, as described previously in astrocytes (36, 37) and in hepatocytes (17). Thus, both the exocytotic vesicle membrane additions and unfolding of the lactotrophs plasma membrane likely contributed to the increased plasma membrane staining by FM 4-64.

Cell volume increased by ~17% after hypotonic stimulation, as estimated from the change in cell diameter. When the osmolarity of the extracellular solution was reduced by ~30%, regulatory volume decrease was absent over the 2 min after the stimulation. Consistent with this finding, regulatory decreases in cell volume occur over 1–2 h (41), and cell volume and prolactin release are closely associated as extracellular osmolarity decreased (51).

Prolactin release from cell populations and from single vesicles in hypotonicity. To study hypotonicity-induced hormone discharge, we measured prolactin release from perfused rat lactotrophs (47) and monitored the dynamics of single large dense-core vesicles by fluorescence and electrophysiology techniques (4, 45, 53).

The amount of stimulus-induced prolactin release, estimated by integrating the signal over the basal level (Fig. 2), was ~30 ng during hypotonic stimulation and ~360 ng during KCl depolarization. Since the density of cells was similar under both conditions, the reduced amount of hormone release after hypotonic stimulation may reflect reduced release of prolactin per cell. However, it cannot be entirely excluded that lactotrophs are functionally heterogeneous cells (29) and may thus exhibit distinct sensitivity to the two stimuli.

To examine this question further, we transfected cells with the ANP-EGFP construct that targets secretory vesicles (19). Prolactin (23 kDa; 14) and ANP.emd (32 kDa) appear to colocalize in the same vesicles (45) and have similar molecular weights; hence measurements of ANP-EGFP discharge likely reflect the rate of prolactin release from a single vesicle. We also monitored single vesicle exocytosis with FM 4-64, which stains the membrane and the matrix of individual vesicles (2) when a fusion pore forms between the docked vesicle and the plasma membrane. Since fewer vesicles discharged their cargoes after hypotonic stimulation than after KCl depolarization (Fig. 3), the reduced prolactin release elicited by hypotonic stimulation vs. KCl depolarization (Fig. 2) is likely due to the reduced discharge of hormone per cell rather than due to response of a smaller, distinct fraction of cells in a heterogeneous population of lactotrophs (29).

Sustained prolactin release from KCl-stimulated cell population of lactotrophs (Fig. 2) likely reflects the discharge from readily docked/primed (45) vesicles as well as the newly recruited vesicles. In contrast, hypotonicity-evoked transient increase followed by the subsequent depression appears to preferentially involve hormone discharge from vesicles in the postfusion state only. As discussed below, in the majority of patches, hypotonicity failed to evoke new fusion events in previously silent patches and thus affected vesicle discharge properties at the postfusion state.

Transient fusion events are the predominant form of exocytosis in hypotonicity. In neurons, kiss-and-run exocytosis is unexpectedly common (16), although the appearance of this mode of exocytosis is debated (20). Transient fusion pore openings, although uncommon, were identified in neuroendocrine cells by electrophysiological measurements (28). Using cell-attached patch-clamp recordings, we found that reversible and irreversible steps occur spontaneously in lactotrophs. Irreversible steps (Fig. 3A, left) most likely represent full fusion exocytosis (35), whereas reversible steps (Fig. 4A, right, and Fig. 4B) probably represent kiss-and-run exocytosis (1, 13). Hypotonic stimulation increased the occurrence of transient fusion events, whereas it did not affect the occurrence of full fusion events (Fig. 5A).

The main obstacle for hormone release from a single vesicle is the diffusion through fusion pore, which opens transiently in our experiments (Fig. 4B). Increased hormone release from a single vesicle may be attained if stimulation affected the fusion pore dwell-time, prolonging the connection between vesicle lumen and the extracellular space (46, 48). Indeed, in our experiments, stimulation prolonged dwell-time and increased fusion pore open probability (Fig. 6, A and B). On the other hand, the pore diameter should be sufficiently wide to permeate the relatively large peptidergic hormones. The results revealed that stimulation affected fusion pore properties by also increasing the average Gp, which reflects widening of fusion pore diameter (Fig. 6C). Moreover, the incidence of events with measureable Gp was fourfold smaller in KCl-stimulated cells, which shows that KCl is a stronger modulator of fusion pore diameter in comparison to hypotonicity. This is consistent with the difference in prolactin release responses of cell populations to the two respective stimuli (Fig. 2).

Hypotonicity modulates postfusion vesicle hormone discharge. Reversible exocytosis has been measured in various cell types (11, 16, 28, 31, 45, 48, 52) and appears to be the dominant mode of exocytosis in both nonstimulated and stimulated lactotrophs (48). Of 11 patches in which we observed discrete steps in Cm, in only one did hypotonic stimulation induce steps in a previously silent patch of membrane. This indicates that hypotonicity mainly affects the postfusion events, of which the molecular nature and Ca2+ dependence awaits further studies.

In summary, the exposure of perfused rat lactotrophs to hypotonic stimulation caused a transient increase in prolactin release, followed by a sustained decrease below the basal level, which contrasts the sustained KCl-evoked release. This is likely due to the reduced fraction of release competent vesicles per cell in hypotonic vs. KCl depolarization. Vesicle release competence appears to be affected by postfusion pore properties, whereby transient vesicle fusion mediates hormone release.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by Grants P3 521 0381, P3 0310 0381, and Z3-3510-1683 from the Ministry of Education, Sciences and Sports of the Republic of Slovenia and by R01-NS36665-05 from the National Institutes of Health.


   DISCLOSURE
 
The authors declare that they have no competing financial interests.


   ACKNOWLEDGMENTS
 
We thank Dr. Ed Levitan for the ANP-EGFP construct and Mrs. Sonja Grilc and Dr. Maja Potokar for valuable help and discussions.


   FOOTNOTES
 

Address for reprint requests and other correspondence: R. Zorec, Institute of Pathophysiology, Faculty of Medicine, Univ. of Ljubljana, LN-MCP, Zaloska 4, 1000 Ljubljana, Slovenia (e-mail: robert.zorec{at}mf.uni-lj.si)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

* Jernej Jorgacevski and Matjaz Stenovec contributed equally to this work. Back


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
1. Alvarez de Toledo G, Fernández-Chacón R, Fernández J. Release of secretory products during transient vesicle fusion. Nature 363: 554–558, 1993.[CrossRef][Web of Science][Medline]

2. Angleson J, Cochilla A, Kilic G, Nussinovitch I, Betz W. Regulation of dense core release from neuroendocrine cells revealed by imaging single exocytic events. Nat Neurosci 2: 440–446, 1999.[CrossRef][Web of Science][Medline]

3. Arieff AI, Llach F, Massry SG. Neurological manifestations and morbidity of hyponatremia: correlation with brain water and electrolytes. Medicine (Baltimore) 55: 121–129, 1976.[Medline]

4. Barg S, Olofsson C, Schriever-Abeln J, Wendt A, Gebre-Medhin S, Renström E, Rorsman P. Delay between fusion pore opening and peptide release from large dense-core vesicles in neuroendocrine cells. Neuron 33: 287–299, 2002.[CrossRef][Web of Science][Medline]

5. Ben-Tabou S, Keller E, Nussinovitch I. Mechanosensitivity of voltage-gated calcium currents in rat anterior pituitary cells. J Physiol 476: 29–39, 1994.[Abstract/Free Full Text]

6. Betz WJ, Bewick GS. Optical analysis of synaptic vesicle recycling at the frog neuromuscular junction. Science 255: 200–203, 1992.[Abstract/Free Full Text]

7. Brown PS, Brown SC. Vertebrate endocrinology: fundamentals and biomedical implications: regulation of water and electrolytes. In: Osmoregulatory Actions of Prolactin and Other Adenohypophysial Hormones. New York: Academic, vol. 2, 1987, p. 211–238.

8. Burgoyne R, Fisher R, Graham M, Haynes L, Morgan A. Control of membrane fusion dynamics during regulated exocytosis. Biochem Soc Trans 29: 467–472, 2001.[CrossRef][Web of Science][Medline]

9. Çelik JB, Gezginç K, Çelik Ç, Çapar M. Toxicity symptoms following hysteroscopic surgery: a case report. J Turkish German Gynecol Assoc 5: 321–323, 2004.

10. Dannies P. Manipulating the reversible aggregation of protein hormones in secretory granules: potential impact on biopharmaceutical development. BioDrugs 17: 315–324, 2003.[CrossRef][Web of Science][Medline]

11. Dietl P, Haller T. Persistent fusion pores but transient fusion in alveolar type II cells. Cell Biol Int 24: 803–807, 2000.[CrossRef][Web of Science][Medline]

12. Elhamdani A, Palfrey H, Artalejo C. Quantal size is dependent on stimulation frequency and calcium entry in calf chromaffin cells. Neuron 31: 819–830, 2001.[CrossRef][Web of Science][Medline]

13. Fesce R, Grohovaz F, Valtorta F, Meldolesi J. Neurotransmitter release: fusion or 'kiss-and-run'? Trends Cell Biol 4: 1–4, 1994.[CrossRef][Medline]

14. Freeman M, Kanyicska B, Lerant A, Nagy G. Prolactin: structure, function, and regulation of secretion. Physiol Rev 80: 1523–1631, 2000.[Abstract/Free Full Text]

15. Fulop T, Radabaugh S, Smith C. Activity-dependent differential transmitter release in mouse adrenal chromaffin cells. J Neurosci 25: 7324–7332, 2005.[Abstract/Free Full Text]

16. Gandhi S, Stevens C. Three modes of synaptic vesicular recycling revealed by single-vesicle imaging. Nature 423: 607–613, 2003.[CrossRef][Web of Science][Medline]

17. Graf J, Rupnik M, Zupancic G, Zorec R. Osmotic swelling of hepatocytes increases membrane conductance but not membrane capacitance. Biophys J 68: 1359–1363, 1995.[Web of Science][Medline]

18. Grau E, Nishioka R, Bern H. Effects of osmotic pressure and calcium ion on prolactin release in vitro from the rostral pars distalis of the tilapia Sarotherodon mossambicus. Gen Comp Endocrinol 45: 406–408, 1981.[CrossRef][Web of Science][Medline]

19. Han W, Ng Y, Axelrod D, Levitan E. Neuropeptide release by efficient recruitment of diffusing cytoplasmic secretory vesicles. Proc Natl Acad Sci USA 96: 14577–14582, 1999.[Abstract/Free Full Text]

20. He L, Wu L. The debate on the kiss-and-run fusion at synapses. Trends Neurosci 30: 447–455, 2007.[CrossRef][Web of Science][Medline]

21. Heuser J, Reese T. Evidence for recycling of synaptic vesicle membrane during transmitter release at the frog neuromuscular junction. J Cell Biol 57: 315–344, 1973.[Abstract/Free Full Text]

22. Hirano T. The spectrum of prolactin action in teleosts. In: Comparative Endocrinology: Developments and Directions. New York: Liss, 1986. p. 53–74.

23. Huang C, Kuo W, Tsai T, Wu D, Hsieh Y, Wang P, Cheng C, Liu J. Prolactin secretion and intracellular Ca2+ change in rat lactotroph subpopulations stimulated by thyrotropin-releasing hormone. J Cell Biochem 87: 126–132, 2002.[CrossRef][Web of Science][Medline]

24. Katz B. Nerve, Muscle and Synapse. New York: McGraw Hill, 1966.

25. Kilic G. Exocytosis in bovine chromaffin cells: studies with patch-clamp capacitance and FM1-43 fluorescence. Biophys J 83: 849–857, 2002.[Web of Science][Medline]

26. Kreft M, Zorec R. Cell-attached measurements of attofarad capacitance steps in rat melanotrophs. Pflügers Arch 434: 212–214, 1997.[CrossRef][Web of Science][Medline]

27. Lang F, Busch G, Ritter M, Völkl H, Waldegger S, Gulbins E, Häussinger D. Functional significance of cell volume regulatory mechanisms. Physiol Rev 78: 247–306, 1998.[Abstract/Free Full Text]

28. Lindau M, Almers W. Structure and function of fusion pores in exocytosis and ectoplasmic membrane fusion. Curr Opin Cell Biol 7: 509–517, 1995.[CrossRef][Web of Science][Medline]

29. Lledo P, Guerineau N, Mollard P, Vincent J, Israel J. Physiological characterization of two functional states in subpopulations of prolactin cells from lactating rats. J Physiol 437: 477–494, 1991.[Abstract/Free Full Text]

30. Lledo P, Vernier P, Vincent J, Mason W, Zorec R. Inhibition of Rab3B expression attenuates Ca(2+)-dependent exocytosis in rat anterior pituitary cells. Nature 364: 540–544, 1993.[CrossRef][Web of Science][Medline]

31. Lollike K, Borregaard N, Lindau M. Capacitance flickers and pseudoflickers of small granules, measured in the cell-attached configuration. Biophys J 75: 53–59, 1998.[Web of Science][Medline]

32. Lollike K, Lindau M. Membrane capacitance techniques to monitor granule exocytosis in neutrophils. J Immunol Methods 232: 111–120, 1999.[CrossRef][Web of Science][Medline]

33. McCormick SD. Endocrine control of osmoregulation in teleost fish. Am Zool 41: 781–794, 2002.[CrossRef][Web of Science]

34. Nagahama Y, Nishioka R, Bern H, Gunther R. Control of prolactin secretion in teleosts, with special reference to Gillichthys mirabilis and Tilapia mossambica. Gen Comp Endocrinol 25: 166–188, 1975.[CrossRef][Web of Science][Medline]

35. Neher E, Marty A. Discrete changes of cell membrane capacitance observed under conditions of enhanced secretion in bovine adrenal chromaffin cells. Proc Natl Acad Sci USA 79: 6712–6716, 1982.[Abstract/Free Full Text]

36. Olson JE, Li GZ. Increased potassium, chloride, and taurine conductances in astrocytes during hypoosmotic swelling. Glia 20: 254–61, 1997.[CrossRef][Web of Science][Medline]

37. Pangrsice T, Potokar M, Haydon PG, Zorec R, Kreft M. Astrocyte swelling leads to membrane unfolding, not membrane insertion. J Neurochem 99: 514–523, 2006.[CrossRef][Web of Science][Medline]

38. Patterson G, Knobel S, Sharif W, Kain S, Piston D. Use of the green fluorescent protein and its mutants in quantitative fluorescence microscopy. Biophys J 73: 2782–2790, 1997.[Web of Science][Medline]

39. Rizzoli S, Richards D, Betz W. Monitoring synaptic vesicle recycling in frog motor nerve terminals with FM dyes. J Neurocytol 32: 539–549, 2003.[CrossRef][Web of Science][Medline]

40. Seale A, Fiess J, Hirano T, Cooke I, Grau E. Disparate release of prolactin and growth hormone from the tilapia pituitary in response to osmotic stimulation. Gen Comp Endocrinol 145: 222–231, 2006.[CrossRef][Web of Science][Medline]

41. Seale A, Richman Nr Hirano T, Cooke I, Grau E. Cell volume increase and extracellular Ca2+ are needed for hyposmotically induced prolactin release in tilapia. Am J Physiol Cell Physiol 284: C1280–C1289, 2003.[Abstract/Free Full Text]

42. Seale A, Riley L, Leedom T, Kajimura S, Dores R, Hirano T, Grau E. Effects of environmental osmolality on release of prolactin, growth hormone and ACTH from the tilapia pituitary. Gen Comp Endocrinol 128: 91–101, 2002.[CrossRef][Web of Science][Medline]

43. Smets G, Velkeniers B, Finne E, Baldys A, Gepts W, Vanhaelst L. Postnatal development of growth hormone and prolactin cells in male and female rat pituitary. An immunocytochemical light and electron microscopic study. J Histochem Cytochem 35: 335–341, 1987.[Abstract]

44. Spruce A, Breckenridge L, Lee A, Almers W. Properties of the fusion pore that forms during exocytosis of a mast cell secretory vesicle. Neuron 4: 643–654, 1990.[CrossRef][Web of Science][Medline]

45. Stenovec M, Kreft M, Poberaj I, Betz W, Zorec R. Slow spontaneous secretion from single large dense-core vesicles monitored in neuroendocrine cells. FASEB J 18: 1270–1272, 2004.[Abstract/Free Full Text]

46. Sun J, Wu X, Wu L. Single and multiple vesicle fusion induce different rates of endocytosis at a central synapse. Nature 417: 555–559, 2002.[CrossRef][Web of Science][Medline]

47. Van Goor F, Zivadinovic D, Martinez-Fuentes A, Stojilkovic S. Dependence of pituitary hormone secretion on the pattern of spontaneous voltage-gated calcium influx. Cell type-specific action potential secretion coupling. J Biol Chem 276: 33840–33846, 2001.[Abstract/Free Full Text]

48. Vardjan N, Stenovec M, Jorgacevski J, Kreft M, Zorec R. Subnanometer fusion pores in spontaneous exocytosis of peptidergic vesicles. J Neurosci 27: 4737–4746, 2007.[Abstract/Free Full Text]

49. Vardjan N, Stenovec M, Jorgacevski J, Kreft M, Zorec R. Elementary properties of spontaneous fusion of peptidergic vesicles: fusion pore gating. J Physiol 585: 655–651, 2007.[Abstract/Free Full Text]

50. Wang X, Sato N, Greer M, Greer S, McAdams S. Significant qualitative differences exist between thyrotropin and prolactin secretory dynamics induced by pituitary cell swelling. Proc Soc Exp Biol Med 198: 612–619, 1991.[CrossRef][Medline]

51. Weber GM, Seale AP, Richman III NH, Stetson MH, Hirano T, Grau EG. Hormone release is tied to changes in cell size in the osmoreceptive prolactin cell of a euryhaline teleost fish, the tilapia, Oreochromis mossambicus. Gen Comp Endocrinol 138: 8–13, 2004.[CrossRef][Web of Science][Medline]

52. Zhou Z, Misler S, Chow R. Rapid fluctuations in transmitter release from single vesicles in bovine adrenal chromaffin cells. Biophys J 70: 1543–1552, 1996.[Web of Science][Medline]

53. Zorec R, Sikdar S, Mason W. Increased cytosolic calcium stimulates exocytosis in bovine lactotrophs. Direct evidence from changes in membrane capacitance. J Gen Physiol 97: 473–497, 1991.[Abstract/Free Full Text]





This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
295/3/C624    most recent
00303.2008v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via Web of Science (1)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Jorgacevski, J.
Right arrow Articles by Zorec, R.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Jorgacevski, J.
Right arrow Articles by Zorec, R.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2008 by the American Physiological Society.