Cell Calcium 38 (2005) 87–99
Calcium gradients and exocytosis in bovine adrenal chromaffin cells
Fernando D. Marengo a,b,∗
b
a Department of Physiology, UCLA, School of Medicine, Los Angeles, CA 90095, USA
Laboratorio de Fisiologı́a y Biologı́a Molecular, Instituto de Fisiologı́a, Biologı́a Molecular y Neurociencias, Facultad de Ciencias Exactas y Naturales,
Universidad de Buenos Aires, Ciudad Universitaria – Pabellón II – 2◦ piso, IFIByNE-CONICET, Buenos Aires CP 1428, Argentina
Received 21 December 2004; received in revised form 15 May 2005; accepted 1 June 2005
Available online 1 August 2005
Abstract
The relationship between the localized Ca2+ concentration and depolarization-induced exocytosis was studied in patch-clamped adrenal
chromaffin cells using pulsed-laser Ca2+ imaging and membrane capacitance measurements. Short depolarizing voltage steps induced Ca2+
gradients and small “synchronous” increases in capacitance during the pulses. Longer pulses increased the capacitance changes, which
saturated at 16 fF, suggesting the presence of a small immediately releasable pool of fusion-ready vesicles. A Hill plot of the capacitance
changes versus the estimated Ca2+ concentration in a thin (100 nm) shell beneath the membrane gave n = 2.3 and Kd = 1.4 ␮M. Repetitive
stimulation elicited a more complex pattern of exocytosis: early pulses induced synchronous capacitance increases, but after five or more pulses
there was facilitation of the synchronous responses and gradual increases in capacitance continued between pulses (asynchronous exocytosis)
as the steep submembrane Ca2+ gradients collapsed. Raising the pipette Ca2+ concentration led to early facilitation of the synchronous response
and early appearance of asynchronous exocytosis. We used this data to develop a kinetic model of depolarization-induced exocytosis, where
Ca2+ -dependent fusion of vesicles occurs from a small immediately releasable pool with an affinity of 1–2 ␮M and vesicles are mobilized to
this pool in a Ca2+ -dependent manner.
© 2005 Elsevier Ltd. All rights reserved.
Keywords: Pulsed-laser imaging; Ca2+ signalling; Secretion; Immediately releasable pool
1. Introduction
Several lines of evidence led to the conclusion that in neuroendocrine cells, such as adrenal chromaffin cells, the trigger for exocytosis—or more specifically vesicle fusion—in
response to membrane depolarization is a Ca2+ elevation near
the cell surface. First, exocytosis is more potent when elicited
by Ca2+ entry through Ca2+ channels than by Ca2+ released
from intracellular stores [1–3]. Second, higher Ca2+ concentrations (1–10 ␮M) appeared to be required to trigger exocytosis when Ca2+ is elevated uniformly via cell dialysis with
Ca2+ buffers (0.17 ␮M–9 mM) than when measured following membrane depolarization [3]. Third, photolytic release
of Ca2+ from “caged Ca2+ compounds”, which induces large
uniform Ca2+ elevations (10–50 ␮M), elicits a rapid exo∗
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0143-4160/$ – see front matter © 2005 Elsevier Ltd. All rights reserved.
doi:10.1016/j.ceca.2005.06.006
cytotic response, suggesting the requirement for relatively
high-Ca2+ concentrations [4]. And finally, Ca2+ gradients
that might provide such locally elevated Ca2+ concentrations
have been measured in adrenal chromaffin cells in response
to depolarization [2,5–7].
A Ca2+ signal may be involved in other steps of the exocytotic process besides vesicle fusion. For instance, Ca2+
has been implicated in vesicle movement between different pools and facilitation during repetitive stimuli [4,8,9].
In addition, Ca2+ may play a role in endocytosis [10]. However, these processes may have different sensitivity to Ca2+
and different temporal characteristics compared to membrane
fusion. Furthermore, separate pathways of regulated exocytosis with distinct kinetics and Ca2+ sensitivity may operate
in chromaffin cells [8,11]. Thus, to understand the mechanisms involved in Ca2+ -induced exocytosis it is necessary
to characterize the temporal and spatial properties of Ca2+
gradients evoked by membrane depolarization. In practice,
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F.D. Marengo / Cell Calcium 38 (2005) 87–99
such measurements are difficult for rapid events, and a tradeoff must be made between temporal and spatial resolution.
Thus, confocal spot detection gets excellent time resolution
in one location, whereas pulsed-laser imaging provides high
spatial resolution at a single time. Then, successive measurements can be made at different locations or times to get spatial
and temporal information, respectively [6,7,12,13]. Alternatively, confocal line scan provides good time resolution,
but the spatial information is limited to one direction [14].
These “state-of-the-art” techniques have further limitations.
The amount of data collected may be limited by the highintensity laser illumination, the spatial resolution is limited
by the microscope optics and the dynamics of Ca2+ distribution is affected by the presence of Ca2+ indicators, which
bind significant amounts of Ca2+ . An alternative approach
to experimental measurements is the use of mathematical
models to simulate Ca2+ gradients [15,16]. For example, a
radial diffusion model has been used to simulate Ca2+ gradients generated upon opening of Ca2+ channels in small
spherical cells [17,18]. These studies have provided considerable information on possible types of Ca2+ gradients, but
they unfortunately rely upon parameter selection for endogenous Ca2+ buffer concentrations, affinities and kinetic rate
constants.
Recently, others [19] and we [7] have used a combination of high-resolution Ca2+ measurement and mathematical modeling to address these issues. In our case, we used
pulsed-laser Ca2+ imaging together with computer modeling
to obtain information on local Ca2+ signals at high time resolutions in adrenal chromaffin cells [7] and recently extended
the model to consider the pattern of Ca2+ gradient development, dissipation and clearance in response to repetitive
depolarizing stimuli [20].
Patch-clamp experiments in adrenal chromaffin cells have
shown that only a small pool of vesicles is available for release
during single short depolarizing stimulus [3,21]. For chromaffin cells in culture this immediately releasable pool (IRP)
consists of 10–20 vesicles [21]. It was suggested that IRP is a
sub-pool of the ready releasable pool of vesicles and closely
spatially associated with Ca2+ channels [22].
Repetitive depolarizing stimuli resulted in exocytosis of
more vesicles, with exocytosis continuing between depolarizing pulses [3,8,23–25]. Since short depolarizing pulses only
elicit the release of a small number of vesicles present in IRP,
it follows that repetitive stimuli can mobilize secretory granules into the pool of vesicles that fuses [21]. In relation to this
hypothesis, recently, it was proposed that vesicles could be
mobilized to closer positions with respect to Ca2+ channels
in a Ca2+ -dependent fashion [26]. In addition, experiments
using flash photolysis of caged Ca2+ molecules reveal several different kinetic phases of exocytosis, suggesting that
the ready releasable pool (first phase) is replenished from
a rapidly mobilized pool for the second phase and from a
large slowly mobilizable depot pool for the slowest phase
[4,27]. A number of questions remain unresolved regarding
these pools and their Ca2+ regulation during depolarization-
induced exocytosis. For instance, what is the Ca2+ sensitivity
of exocytosis in a context of more physiological stimulation
(in comparison with photolysis experiments); is the mobilization of granules Ca2+ dependent, and what is the Ca2+
sensitivity of this mobilization?
Previous work analyzed the relationship between exocytosis and Ca2+ released globally by flash photolysis or tried
to estimate the relevant localized Ca2+ concentration theoretically. Here, we make simultaneous measurements of
Ca2+ gradients and exocytosis by determining changes in
membrane capacitance, for single pulse and repetitive depolarizing simuli. We used the fluorescence measurements of
Ca2+ changes and computer modeling to estimate the Ca2+
signal near the membrane and used this data to investigate the
Ca2+ dependence of depolarization-induced exocytosis and
evaluate various kinetic models of secretion.
2. Materials and methods
2.1. Whole-cell patch-clamp and membrane capacitance
measurements
Chromaffin cells were prepared from bovine adrenal
medullae by enzymatic digestion [28] and kept 1–4 days in
culture prior to use [7].
Conventional whole-cell recordings were performed as
detailed in Monck et al. [6]. The patch-clamp set up comprised a patch-clamp amplifier (Model Axopatch 200A, Axon
Instruments Inc., Foster City, CA, USA), a data acquisition
interface (Model IDA15125, Indec Systems Inc., Mountain
View, CA, USA) and a personal computer.
Chromaffin cells were washed in an extracellular medium
comprising 120 mM NaCl, 20 mM Hepes, 4 mM MgCl2 ,
5 mM CaCl2 , 5 mg/ml glucose and 1 ␮M tetrodotoxin (pH
7.25). The standard internal solution used in the patch-clamp
pipettes contained 125 mM Cs d-glutamate, 30 mM Hepes,
8 mM NaCl, 1 mM MgCl2 , 2 mM Mg-ATP, 0.3 mM GTP,
0.3 mM Cs-EGTA and 0.2 mM rhod-2 triammonium salt [29]
(pH 7.2). This solution allows measurement of Ca2+ currents
because Na+ and K+ currents are prevented. The holding
potentials have not been corrected for junction potentials [30].
We considered that the cells are “leaky” when the leak current measured at the normal holding potential of −70 mV
was bigger than −5 pA.
The cell membrane capacitance was measured with a software phase-sensitive detector as described before [31,32].
Recordings were performed in the whole-cell mode of the
patch-clamp technique. The command voltage applied to the
cell was composed of the sum of a sinusoidal voltage (833 Hz,
80 mV peak to peak) and a holding potential of −70 mV. The
membrane current was measured at two-phase angles, Iϕ + 0
and Iϕ + 90 , relative to the applied sinusoidal potential. The
phase-tracking technique [32] was used to select the phase
angle ϕ. The output at Iϕ + 0 represents changes in the real part
of the cell admittance, and the output at Iϕ + 90 reflect changes
F.D. Marengo / Cell Calcium 38 (2005) 87–99
in the imaginary part of the cell admittance, from which we
can determine the changes in membrane capacitance.
2.2. Measurement of Ca2+ gradients with pulsed-laser
imaging
Ca2+ gradients were measured using pulsed-laser Ca2+
imaging of whole-cell patch-clamped cells [6,20]. The imaging system consists of an inverted epi-fluorescence microscope (Model IX-70, Olympus), a peltier-cooled chargecoupled device (CCD) camera (Model PXL, Photometrics
Ltd., Tucson, AZ, USA) with a 1317 × 1035 pixel CCD
chip (Model KAF 1400, Kodak), and a host Pentium microcomputer. Illumination is achieved with a high-intensity
pulsed coaxial flash lamp dye laser (LumenX Model LS1400, Phase-R Corporation, New Durham, NH, USA), which
provides short (350 ns) high-intensity pulses of illumination
[33]. Lasing dye Coumarin 525 (0.02 mM in methanol) gives
an appropriate emission spectrum (500–540 nm) for excitation of rhod-2. The patch-clamp set-up was used for voltageclamp recording and precise synchronization of the laser
pulses with the Ca2+ currents. The laser light is focused into
a multimode fiber optic (General Fiber Optics Corporation,
Fairfield, NJ, USA), which is coupled to the epi-fluorescence
port of the microscope through an adaptor containing a fused
silica plano-convex lens (Oriel Corporation, Stratford, CT,
USA). This arrangement results in global illumination of the
microscope field of view. The epi-fluorescence filter block
contains a 570 nm DRLP dichroic mirror and 585 nm EFLP
emission filter (Chroma Optical, Brattleboro, VT, USA). A
89
high-numerical aperture objective (N.A. 1.4, plan apo 60×;
Olympus America Inc., Melville, NY, USA) was used to
image the cells.
After allowing 10 min for the Ca2+ indicator to diffuse
into the cell, fluorescence measurements were taken as image
pairs, a control image with no depolarization (i.e., constant holding potential) and a stimulus image with a depolarizing step pulse. The results are shown as ratio images
(Figs. 1A and 4A) of the stimulus image divided by the corresponding control image. Taking the ratio of the images in
this way corrects for spatial differences in cell thickness (light
pathlength), indicator concentration and accessible cytosolic
volume [34]. Pairing of the control and stimulus images minimizes the effects of cell movements. The ratio images (Ft /F0 )
are used to analyze the gradients and to estimate the free Ca2+
concentrations as described below.
2.3. Estimation of Ca2+ concentration
The ratio of the stimulus image divided by the control
image is displayed as a pseudo color image representing the
fractional change in fluorescence. Since the ratio corrects
for differences in indicator concentration, indicator excluded
volume and light pathlength, the images showing the fractional change in fluorescence represent spatial maps of the
Ca2+ changes, provided that the cell does not move between
the control and stimulus images. Rhod-2 does not undergo
a shift in either excitation or emission spectra on binding
Ca2+ [29], so we cannot use a ratiometric calibration scheme
[35]. Instead, the change in Ca2+ concentration was estimated
Fig. 1. Ca2+ gradients and exocytosis induced by single-pulse depolarizing stimuli. Top: sequence of dynamic ratio (Ft /F0 ) images taken at the end of
depolarizing pulses (+20 mV from a −70 mV resting potential) of different durations in rhod-2-loaded chromaffin cells. The profiles (below the images) show
the ratio values along the lines through the images. Middle: cellular capacitance (Cm ) increases induced on stimulation with 10, 50 and 80 ms pulses. Bottom:
The Ca2+ currents recorded at the same time. Note: the Ca2+ currents appear over-filtered because the data was stored at the same rate as the capacitance data
(9.6 ms/point).
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F.D. Marengo / Cell Calcium 38 (2005) 87–99
from the fractional fluorescence change (stimulus/control
ratio, Ft /F0 ), as described previously [6]. We used a value
of 1880 nM for the Kd [36] and 0.013 for α (the ratio of
the fluorescence of free and Ca2+ -bound rhod-2, determined
in vitro using internal solution with “zero” Ca2+ (10 mM
EGTA) and saturating Ca2+ , respectively). Assuming a resting value of 100 nM for the resting Ca2+ concentration, these
values give an estimated peak Ca2+ concentrations of around
200–250 nM at the end of a 40 ms depolarizing stimulus. The
conversion to Ca2+ has some caveats, as we do not know the
precise value of the resting Ca2+ or the in situ calibration
for the indicator dye. However, attempts to saturate rhod2 “in situ” using ionomycin gave Ft /F0 values >8, which
indicates that the resting Ca2+ concentration was less than
200 nM. Estimates of the Ca2+ change using different resting
free Ca2+ concentrations in the range 50–200 nM give peak
Ca2+ increases of about 2–2.5 times the resting concentration.
The Ca2+ concentration estimates are relatively insensitive to
the value of α used, since the amounts of Ca2+ -indicator complex are small (∼5–10%). Thus, provided the in situ value of
α is less than 0.05 the estimates of Ca2+ remain accurate. In
addition, the Ca2+ calibration procedure assumes equilibrium
conditions and during the Ca2+ current, the Ca2+ indicators
will not be at equilibrium with the free Ca2+ ions. Therefore,
we chose to express most of the data as the fractional fluorescence change (Ft /F0 ), as this best represents the measured
parameter.
2.4. Analysis of Ca2+ gradients
Ratio images representing the fractional fluorescence
change (Ft /F0 ) are used to display the spatial distribution
of Ca2+ concentration. The ratios are made after averaging a 3 × 3 kernel for each pixel in the original fluorescence images. Thus, each averaged region corresponds to a
square with sides three times the measured pixel size of 113,
or 339 nm. To analyze the Ca2+ gradients, we used crosssectional profiles through the center of the cell. The value
chosen for each position in the profile was taken with no
further averaging. The focus of the cell was always chosen
as the image with the maximum visible diameter, which in
a perfectly spherical cell would represent a section through
the center of the cell. To maximize the signal available for
analysis, the direction of the profile was usually chosen from
the brightest point around the periphery through the center
of the cell. However, we tried to avoid the regions beneath
the pipette (at 2–4 o’ clock in the displayed images), as these
were often distorted by the presence of either out-of-focus
light or reflected light from the pipette.
pool, four-state model:
k1
k1
γ(Ca)
A B C −→E
k−1 k−2
In this model, the chromaffin granules are placed in three
pools: a small immediately releasable pool (IRP, pool C)
where vesicles are available for fusion (exocytosed state, E);
a larger releasable pool (pool B), where vesicles are available for rapid mobilization as occurs during facilitation and
repetitive stimulation; and a depot pool (pool A), which is a
larger pool that can slowly replenish the other pools. In this
scheme, pool B would include the ready releasable pool plus
the slowly releasable pool determined in photolysis experiments [27]. The fusion rate constant, γ (Ca), was made
Ca2+ -dependent by introducing a Ca2+ sensor with four Ca2+ binding sites, which was modeled as an independent kinetic
scheme, as follows for four binding sites:
The rate constants represent the independent binding to
sites with an affinity of β/α. It is assumed that the fully
occupied state modulates the fusion rate constant, so the rate
constant for fusion was taken as [S4 ]/[Stotal ] × γ. This method
of incorporating the Ca2+ sensitivity is similar to that previously used by Bertram et al. [39].
To simulate this model, we set up a series of first-order
differential equations for these reactions, for example:
dB
= k1 A + k−2 C − (k−1 + k2 )B
(1)
dt
Similar equations were determined for A, C and E and another
set of equations set up for S0 , S1 , S2 , S3 and S4 . The equations
were numerically integrated using an Euler finite difference
algorithm implemented in Visual Basic 6.0 (Microsoft, Redmond, WA, USA).
The Ca2+ input function is provided from a Ca2+ diffusion
model that we have developed to simulated measured Ca2+
gradients, as described in detail elsewhere [7,20]. As this
model considers a spherical cell divided in concentric shells
of 100 nm width, the input of the secretion model is the Ca2+
concentration in the 100 nm width external shell simulated
by the Ca2+ diffusion model. This condition assumes that
the vesicles sensitive to be exocytosed in response to Ca2+
elevation are in close apposition to plasma membrane. This is
an important difference from previous studies in chromaffin
cells, where the Ca2+ input function was an arbitrary function,
as this model has been developed to simulate the measured
Ca2+ distribution in patch-clamped adrenal chromaffin cells.
2.5. Simulation of exocytosis
3. Results and discussion
The kinetics of exocytosis was modeled using sequential
multi-pool models, similar to those previously developed by
Neher and co-workers [27,37,38]. We used a simple three-
We investigated the relationship between Ca2+ and
depolarization-induced exocytosis in patch-clamped adrenal
F.D. Marengo / Cell Calcium 38 (2005) 87–99
chromaffin cells using pulsed-laser Ca2+ imaging and membrane capacitance measurements.
3.1. Ca2+ gradients and exocytosis induced by a single
depolarizing stimuli
Fig. 1 shows an experiment where a patch-clamped cell,
held at −70 mV, was stimulated with single voltage pulses of
different durations (10–80 ms) to +20 mV. These short depolarizing pulses led to the development of submembrane Ca2+
gradients, where the Ca2+ concentration decreased progressively from the border towards the center of the cell (Fig. 1A)
[7]. The size of the Ca2+ signal at the cell border increased
with pulse length, as did the spatially averaged signal. There
was also an increase of the signal in the center of the cell
when pulses are longer than 20 ms.
Brief depolarizing pulses also induced exocytosis (Fig. 1),
as shown by the increase in the cell membrane capacitance
(Cm ), which occurred as a consequence of dense core granule
fusion with the plasma membrane. During single-pulse stimulation, exocytosis always occurred during the pulse, without
additional increases in membrane capacitance after the end
of the pulse. We will refer to such increases in Cm as synchronous exocytosis. The synchronous exocytotic response
increased with pulse duration (Fig. 1B) and showed saturation (Fig. 2A). This saturation is presumably due to depletion
of a small pool of fusion ready secretory granules, previously
described as the immediately releasable pool (IRP) [21]. Fitting the data in Fig. 2A with a single exponential gives an
asymptote value of about 16 fF. The vesicle size in adrenal
91
chromaffin cells has been estimated to be 1.3 fF [40], suggesting that the IRP is comprised of about 12 vesicles under
our experimental conditions. This is smaller than the pool
size (34 fF) previously reported for rat and bovine chromaffin cells [21,41], and markedly smaller than the 39–52 fF
determined in slices from mouse adrenal glands [22]. These
differences can be partially explained by the fact that we
used an external Ca2+ concentration of 5 mM while previous
studies [21,41] were performed in 10 mM Ca2+ . In addition, the differences between our results and mouse slices
can be due to the poor coupling between Ca2+ channels and
secretion of primary cultured cells in comparison with the
chromaffin in situ [42]. It should be mentioned here that
when very long (i.e., 500 or 1000 ms) pulses were applied
an additional exocytotic component became detectable (data
not shown), which is presumably due to vesicle replenishment [22]. However, this component was too slow, and was
not noticeable in the time frame studied in this paper for single
pulses.
As mentioned above, brief depolarizing pulses caused an
increase in Cm restricted to the period of the stimuli, with
no additional changes after the end of the pulse. This observation suggests that synchronous exocytosis is triggered by
the high-Ca2+ gradient generated near the plasma membrane
during the depolarization, and dissipated rapidly thereafter.
The increase in cell capacitance induced by single depolarizing pulses correlated with Ca2+ entry, measured as the time
integral of Ca2+ currents (Fig. 2B) and with the intracellular
Ca2+ signal near the cell membrane (Fig. 2C), represented by
the averaged Ft /F0 value at the cell border.
Fig. 2. Analysis of the dependence of exocytosis on voltage pulse duration and Ca2+ concentration. (A) Summary of capacitance increases after stimulation
with depolarizing pulses of different lengths. The graph was fitted (using the individual experimental points) with a single exponential function (y = ae−λt ,
a = 15.83 ± 1.56 fF, λ = 0.0183 ± 0.0041 ms−1 , n = 73, r = 0.6772). (B) The increase in Cm is plotted vs. the time integral of ICa (ICa Int). The experimental
points were fitted by a linear regression (y = a + bx; with a = 2.14 ± 0.60, b = 0.296 ± 0.033), n = 42, r = 0.81593, p < 0.0001). (C) The increase in Cm is plotted
vs. Ft /F0 − 1 at the cell border (a = 2.68 ± 0.91, b = 5.24 ± 1.08), n = 41, r = 0.61279, p < 0.0001). There is a clear correlation between Cm and both the measured
fluorescence signal at cell border and the integrated Ca2+ influx.
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F.D. Marengo / Cell Calcium 38 (2005) 87–99
The fluorescence measurements are limited by the resolution of the microscope’s optics, so the Ca2+ concentration
near the cell membrane determined from the Ft /F0 value is
probably a spatial average of the space within 0.5–1 ␮m of the
membrane [7]. This value is likely to underestimate the “true”
Ca2+ signal associated with exocytosis, which is likely to be
confined to a narrow region close to the membrane. In order
to predict the value of the Ca2+ signal sensed by the secretory granules we used a previously developed radial diffusion
model [7], which takes advantage of experimental data to
select appropriate parameters values for endogenous buffers
and diffusion. This model simulates adequately the pattern of
the Ca2+ gradients observed in our patch-clamped cells [7].
We then used the simulations to predict the Ca2+ concentra-
tions existing closer to the plasma membrane. Fig. 3A and B
show the results of simulations for 10, 20, 50, 80 and 100 ms
duration pulses and show the predicted fluorescence signals,
after accounting for blurring of the microscope optics, and
the estimated Ca2+ concentrations. A comparison of Fig. 3A
with Fig. 1A illustrates that the simulated Ft /F0 signals is
a good estimate of the measured gradients (for more details
about the criteria of comparison between measurements and
simulations refer to [7]).
The simulated Ca2+ gradients (Fig. 3B) predict that the
Ca2+ concentration in spaces within 100 nm of cell membrane (the outermost 100 nm shell) increases to over 1 ␮M
for a pulse duration larger than 50 ms. We used a 100 nm
shell as this thickness corresponds to the radius of the chro-
Fig. 3. Analysis of the dependence of exocytosis on the submembrane Ca2+ concentration as estimated with a radial diffusion computer model. (A) Simulated
rhod-2 fluorescence ratio at the end of 10, 30, 50, 80 and 100 ms (in order, from bottom to top) depolarizing pulses represented as the profiles for a section
through the center of the cell. The simulated Ft /F0 signal was blurred using the theoretical point spread function of the microscope [7]. (B) Ca2+ concentrations
obtained from the same simulations shown in (A). (C) Relationship between the measured increase in Cm induced by 10, 30, 50, 80 and 100 ms pulses (the
number of cells included in each point are represented between parenthesis in the figure) and the free [Ca2+ ]i in the outermost 100 nm cytosolic shell, as
predicted by the radial diffusion model (B). This relationship was fitted with a Hill equation. When the model was applied with a shell thickness of 100 nm
(black continuous line), the fitting parameters were Vmax = 22.1 ± 9.1, Kd = 1.35 ± 0.50, n = 2.3 ± 0.6, r = 0.996; with a shell thickness of 50 nm (dotted line),
the fitting parameters were Vmax = 22 ± 27; k = 1.6 ± 1.9; n = 2.2 ± 1.6; r = 0.986; and finally with a shell thickness of 200 nm (dash-dot line), the fitting
parameters were Vmax = 15.7 ± 2.1; Kd = 0.6 ± 0.1; n = 2.8 ± 0.5; r = 0.99332.
F.D. Marengo / Cell Calcium 38 (2005) 87–99
maffin granules [43]. Fig. 3C shows the relationship between
the measured increase in Cm induced by 10, 30, 50, 80 and
100 ms pulses and the Ca2+ concentrations in the outermost
cytosolic shell, as predicted by the radial diffusion model
(Fig. 3B). This plot can be fitted by a Hill type equation with
an “n” value of 2.3 and an affinity (Kd ) of 1.3 ␮M (Fig. 3C).
When the spatial resolution of the simulation was increased or
reduced, changing the shell thickness to 50 or 200 nm, respectively, the fitting parameters were not modified considerably.
While “n” was maintained between 2 and 3, Kd was kept
close to 1 ␮M (for details, please refer to figure legend). We
should note that these Kd values might be underestimated due
to vesicle pool depletion during longer depolarization pulses
(see Section 3). However, adding to the fitting an additional
point corresponding to the Ca2+ concentration predicted for
200 ms pulses (1.72 ␮M) and the associated capacitance measurement did not change these results (Vmax = 26.1 ± 8.3;
Kd = 1.6 ± 0.46; n = 2.1 ± 0.4). These results suggest that
depolarization-induced exocytosis occurs with a Ca2+ affinity in the 1–2 ␮M range. These values are consistent with
the estimates of the Ca2+ sensitivity obtained in experiments using intracellular dialysis of Ca2+ and release of
Ca2+ from internal stores to stimulate exocytosis [3], but
they indicate a slightly higher Ca2+ affinity of the fusion
mechanism in comparison with previous studies using photolyisis of caged Ca2+ compounds [4,27]. It is necessary to
consider that in our experiments, we estimate the sensitivity to Ca2+ of a group of vesicles responsive to single short
depolarizations, while photolysis experiments measures the
response of the whole ready releasable pool. In reference
to that it is interesting to mention a recent paper describing
the presence of a pool of vesicles in bovine chromaffin cells
with higher sensitivity to Ca2+ than classic ready releasable
pool [44].
3.2. Cellular Ca2+ and exocytosis in response to
repetitive depolarizing stimuli
Fig. 4 shows data from a typical experiment where a cell
was stimulated with a train of depolarizing pulses (10 pulses
at 2 Hz). The Ca2+ gradient after the first few pulses is similar
to those shown for short pulses in Fig. 1, although the gradient was superimposed on a larger globally elevated Ca2+
remaining from previous pulses. However, after five pulses
the gradient is no longer clear because there is also an important fluorescence increase in the cell interior. After 10 pulses,
the increase in fluorescence at the center of the cell was larger
than at the periphery. The results in this cell were typical of the
Ca2+ response observed during repetitive stimulation, which
we have characterized in detail elsewhere [20].
Like the Ca2+ distribution, the pattern of exocytosis
evolves during repetitive stimulation as illustrated by the Cm
changes shown in Fig. 4B. During the first few pulses of
a train, there is synchronous exocytosis during the pulse,
similar to that seen for single pulses in Fig. 1B, but not
significantly afterwards. This exocytotic response decreased
93
Fig. 4. Spatial distribution of Ca2+ signal and pattern of exocytosis in
response to repetitive depolarizing stimuli. Sequence of dynamic ratio
(Ft /F0 ) images (A) and exocytosis (B) during repetitive (train) stimulation
(pulses of +20 mV and 50 ms duration; 500 ms between stimuli). Ratios
were obtained from images taken before the beginning train and at the end
the pulse indicated. In addition to the fast increase in Cm that occurs during
each pulse (synchronous exocytosis), repetitive stimulation also induced a
slow increase between pulses (asynchronous exocytosis). Calcium currents
(green traces) and stimulation pulses (blue traces) are shown below.
slightly during the first three or four pulses (Fig. 6), presumably due to depletion of the IRP. However, later during the train a slower phase of Cm increase was observed
between pulses, in addition to the synchronous exocytosis.
We refer to this slow increase between pulses as asynchronous
exocytosis. Similar patterns of exocytosis have previously
been reported in adrenal chromaffin cells and other neuroendocrine cells [3,8,23–25]. This asynchronous exocytosis is
still observed between depolarizing pulses when the submembrane Ca2+ gradient within <1 ␮m of the cell membrane
will have dissipated [7,20]. Therefore, exocytosis during this
slow phase appears to be triggered by a relatively low and sustained Ca2+ concentration. Although there is some variation
between experiments, this asynchronous phase usually starts
between the fifth and the seventh pulse of the trains, as in the
example in Fig. 4B (for average data refer to Figs. 5 and 6).
Fig. 5 (black line) shows this typical pattern of Cm change,
with synchronous and asynchronous phases of exocytosis
(data pooled from 12 different cells). The appearance of the
asynchronous exocytosis also seems to coincide with the start
of an apparent potentiation (facilitation) of the synchronous
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F.D. Marengo / Cell Calcium 38 (2005) 87–99
Fig. 5. Different patterns of exocytotic response were obtained during repetitive stimulation. The experiments were divided in three groups according
to the pattern of exocytosis and each group was averaged point by point: the
first group (black) contains most of the cells and shows the same characteristics as the example in Fig. 4. A second group (light gray) was characterized
by large capacitance changes during the early pulses (especially the first)
and by the asynchronous Cm changes appearing earlier (in the third pulse it
is clear). The three cells of the second group had an unstable ICa record and
showed signs of Ca2+ overload (a steady increase in fluorescence throughout
the experiment). And finally a third group (dark gray) of cells that showed
neither potentiation of synchronous component nor an asynchronous Cm
increase.
component (Figs. 4B and 5, black line). The total exocytosis caused by repetitive stimulation is much larger than
the largest values seen for single pulses, suggesting that the
pool of vesicles prepared for fusion has been replenished by
mobilization from another pool. Here, we use vesicle mobilization to indicate the availability of more fusion-prepared
vesicles and make no claims to the mechanism, which might
involve biochemical priming, vesicle translocation or other
processes.
Most of the cells stimulated with repetitive depolarizing
pulses displayed a pattern of exocytosis similar to that shown
in Fig. 4. However, some cells showed different patterns of
synchronous and asynchronous exocytosis. We divided our
results into three groups based on these patterns. The first, and
largest group showed the pattern that we have just described
as typical behavior (Fig. 5, black line, n = 12). A second group
only showed synchronous exocytosis and lacked both the
asynchronous phase and the potentiation of the synchronous
component (Fig. 5, dark gray line, n = 3). The cells in the second group had a total capacitance increase of 14.8 ± 3.6 fF
(measured as the average between 1 and 2 s after the end
of the train). This value is close to the asymptotic value of
16 fF calculated for single pulses (Fig. 2A). The fluorescence
ratio images at the end of the 10th pulse for this second group
showed a normal spatial pattern of Ca2+ increase, and the spatially averaged Ft /F0 was 2.35 ± 0.16, which was similar to
the values obtained for the first group (2.70 ± 0.25). Therefore, we cannot attribute the different secretion pattern to
changes in Ca2+ signal. A possible explanation could be some
sort of impediment for refilling the immediately releasable
pool of vesicles. Two measurements included in this group
(in a total of three) were obtained a long time (about 30 min)
after whole cell was established. In addition the three cells of
this group had low-series resistances (<9 M). We can speculate that a diffusive factor necessary to induce facilitation
of synchronous exocytosis and appearance of asynchronous
exocytosis was washed away. The third pattern was characterized by large synchronous Cm changes occurring during
the first part of the train and by an early appearance of the
asynchronous Cm changes (Fig. 5, light gray line, n = 3). In
this latter group, there was an increase in the resting rhod-2
fluorescence during the course of the experiment. Resting
fluorescence increased 132 ± 11% over a period of 200 s,
measured from 6 min after the establishment of whole cell
configuration. This increase continued for at least 15 min
without further stimulation. We also noticed that the Ca2+
current was unstable in these cells, suggesting that there may
be some “leaky” Ca2+ entry. In fact, we found that these
cells exhibited a rather large leak current (−15.0 ± 1.5 pA,
n = 3) at the normal holding potential of −70 mV compared
to the group with the “typical” Cm behavior (−2.9 ± 0.9 pA,
n = 12). All this data together suggest that this third group
was subjected to a steady increase in resting Ca2+ . On this
basis, we investigated how increasing resting cytosolic Ca2+
may affect secretion.
Fig. 6 shows the results of experiments where the resting Ca2+ concentration was increased by raising the pipette
internal solution Ca2+ concentration to 300 nM (225 ␮M total
Ca2+ plus 300 ␮M EGTA). In these experiments, we waited
at least 6 min before stimulating the cell to be sure that
the cell interior was fully dialyzed with the high-Ca2+ solution. Fig. 6A compares these “high Ca2+ ” experiments (light
gray line) with the group of control experiments exhibiting the typical behavior (black line). The high-Ca2+ cells
showed an exocytotic response that was characterized by
large synchronous Cm increases during the first few depolarizing pulses and early appearance of the asynchronous
phase. This pattern was similar to the “leaky” cells in Fig. 5
(light gray line). We further analyzed the pattern of exocytosis by measuring the synchronous and asynchronous changes
in capacitance as a function of pulse number. This analysis is summarized in Fig. 6B and C for both control (black)
and high-Ca2+ experiments (light gray). Under control conditions, the synchronous exocytosis was slightly depressed
after the second to the fourth pulse presumably due to vesicle depletion, but becomes facilitated after the fifth pulse
(Fig. 6B) at the same time that the asynchronous exocytosis was observed (Fig. 6C). In the experiments with elevated
Ca2+ in the pipette, the synchronous increase in Cm was
considerably larger during the first five pulses of a train in
comparison with the controls (Fig. 6B), and the asynchronous
component appeared after the second pulse (Fig. 6C). Note
that the synchronous and asynchronous components of exocytosis refer to exocytosis occurring during and following a
depolarizing pulse, respectively, and should not be considered
F.D. Marengo / Cell Calcium 38 (2005) 87–99
95
Fig. 6. Raising the pipette Ca2+ concentration potentiated synchronous exocytosis and induced earlier onset of asynchronous exocytosis., The figures are depicted
in a counterclockwise manner. (A) Raising the free Ca2+ concentration in the pipette solution to 300 nM (light gray) induced a significantly larger exocytotic
response compared to the control experiments (black). The temporal average of all points following the capacitance jump for each pulse is significantly bigger
in the cells dialyzed with 300 nM Ca2+ (* p < 0.05; ** p < 0.01; *** p < 0.001). The figure shows an averaged record obtained from 6 and 12 cells, respectively
(all points for each interval between pulses were included). (B) Analysis of synchronous exocytosis during each pulse in a train of 10 pulses (50 ms, −70
to +20 mV) at 2 Hz in control (black) and 300 nM Ca2+ -dialyzed cells (light gray). The values are averages calculated from the same experiments as shown
in (A). The values obtained in cells dialyzed with 300 nM Ca2+ are significantly larger than the corresponding control during the first five pulses (Student’s
t-test). (C) Analysis of asynchronous exocytosis measured in the 450 ms interval after each pulse during repetitive stimulation in control (black) and 300 nM
Ca2+ -dialyzed cells (light gray). Asynchronous exocytosis increases after the sixth pulse in the controls, whereas there is an earlier appearance (after second
pulse) of asynchronous component in the high-Ca2+ conditions (Student’s t-test). (D) Asynchronous component plotted against cumulative ICa integral during
train obtained for three different cells.
independent processes. The synchronous changes induced
by pulses later in the train will include a component due to
the asynchronous phase of earlier pulses, but we have not
attempted to correct for this crossover. Analysis of the Cm
changes measured for single 50 ms pulses also show a significant increase in the synchronous Cm response in the cells
dialyzed with 300 nM free Ca2+ solution (22.87 ± 4.17 fF,
n = 5) compared to 10.26 ± 1.95 for control (n = 20, p < 0.01),
suggesting that the immediately releasable pool is larger
under these conditions.
Other authors had previously associated the appearance
of facilitation and slow delayed components of exocytosis with accumulation of residual Ca2+ [3]. Horrigan and
Bookman [21] pointed out that a slow and delayed component of exocytosis only appears after stimuli of single or
repetitive stimulation when the resulting calcium currents
were big (approximately between 200 and 300 pA). Addition-
ally, Seward and Nowycky [8] reported the appearance of a
delayed secretory phase initiated after several pulses during a
train, which was activated after a specific “threshold” amount
of Ca2+ had entered to the cell. Another interesting observation is that exogenously added ␣-SNAPs stimulate exocytosis
when Ca2+ is in the 100–300 nM range [45]. Since SNAPs
are believed to increase secretion by increasing the size of the
most readily releasable pools of chromaffin granules [46],
this is a possible mechanism whereby a small increase in
basal Ca2+ could increase both the synchronous and asynchronous components of exocytosis. The observation that
increasing the pulse duration during repetitive stimulation
results in an earlier appearance of asynchronous exocytosis
is also consistent with exocytosis being induced by a residual
Ca2+ mechanism [8].
Intracellular Ca2+ release is not significant under our
experimental conditions [7,20], and therefore, intracellular
96
F.D. Marengo / Cell Calcium 38 (2005) 87–99
Ca2+ concentration increase during a train should be mainly
a consequence of Ca2+ entry through Ca2+ channels [20]. If
residual Ca2+ follows ICa and the delayed exocytosis during
a train follows residual Ca2+ , it is expected that asynchronous
exocytosis should follow cumulative ICa integral [8]. When
the asynchronous component was plotted versus cumulative
ICa integral during trains, we noted that asynchronous exocy-
tosis appeared after certain amount of calcium enters into
the cell (Fig. 6D). For a total of 11 individual cells analyzed, 8 cells showed asynchronous components that started
between 30 and 60 pC. This is equivalent to a range of
1.5–3.10−10 ␮mols of Ca2+ ions entering the cell. For an
average cell of 6.5 ␮m cellular radio, assuming an accessible
cytosolic volume of 60%, and using a total buffer capac-
Fig. 7. Kinetic model of exocytosis with independent Ca2+ sensor with four Ca2+ -binding sites. This model is a three-pool (four-state) sequential model of
exocytosis, with a small immediately releasable pool (IRP, pool C), a pool for quickly replenishing the IRP (pool B), and a larger reserve (or depot, pool A) pool.
The rate constant for fusion (γ) is modulated by the fraction of the Ca2+ sensor with all four sites occupied (see Section 2). (A and B) The Ca2+ input functions
(in the outermost 100 nm cytosolic shell) for repetitive (ten 50 ms pulses at 2 Hz) and single-pulse (200 ms duration) stimuli are represented in black solid line,
and was estimated using a radial diffusion model [7,20]. (C) Simulation of capacitance response during repetitive stimulation (ten 50 ms pulses at 2 Hz). The
pool sizes were A = 5000 fF, B = 680 fF and C = 16 fF. The sizes of A and B were those determined by Heinmann et al. [37] for Ca2+ dialysis data and the size
of C is the size of the immediately releasable pool determined in Fig. 2. The rate constants used were k1 = k−1 = 0 s−1 (k1 = 0.009 s−1 , k−1 = 0.01375 s−1 gave
virtually identical results), k2 = 0.2 s−1 , k−2 = 6.8 s−1 , γ = 1000 s−1 , α = 8 ␮M−1 s−1 and β = 20 s−1 to give a Kd of 1.5 ␮M. These parameters gave a reasonable
approximation of the measured capacitance changes (e.g. see Fig. 4) (D). Simulation of capacitance response during single-pulse stimulation (200 ms duration).
Using the parameters shown for repetitive stimulation (in C) does not result in saturable exocytosis (black trace). The rate of refilling of the IRP (k2 ) must
be slowed k2 = 0.02 s−1 to achieve saturable responses for single-pulse stimulation (light gray traces), which are shown with several different fusion rates
(γ = 1, 2 and 5 ms−1 ). (E and F) To achieve saturation for single pulses and sufficient mobilization for repetitive stimulation, the mobilization must be made
Ca2+ -dependent. (E) Shows the capacitance responses to repetitive stimuli when the refilling of the IRP is Ca2+ -insensitve and slow enough to achieve saturation
for single pulses (light gray) and when it was made Ca2+ -sensitive with a single site high-affinity Ca2+ -binding sensor (Kd = 300 nM, dark gray) and a two site
lower affinity sensor (Kd = 900 nM, black). The analysis of asynchronous exocytosis for these three simulations, performed as in Fig. 6C is represented in the
inset. (F) Simulation for single pulses using same parameters as for repetitive stimulation (see E).
F.D. Marengo / Cell Calcium 38 (2005) 87–99
ity (endogenous plus exogenous buffers) of 1600 [7], these
numbers are equivalent to an increase in cytosolic Ca2+ of
160–320 nM over resting concentration.
Our results suggest that a moderate increased basal Ca2+
concentration, as it is produced by residual accumulation during a train, is responsible of vesicle mobilization to most
readily releasable pools, and in particular to immediately
releasable pool. From flash photolysis experiments, IRP was
defined as a sub-pool of vesicles included in the ready
releasable pool but closer to Ca2+ channels. In a recent paper,
Becherer et al. [26] using evanescent field imaging, show that
elevated Ca2+ reduced the distance between docked vesicles
and Ca2+ entry sites. These authors suggested a mechanism
for facilitation in which vesicles are moved closer to Ca2+
channels.
3.3. Models of exocytosis
Our results have raised questions regarding the Ca2+ affinity of the fusion step and the possible involvement of other
Ca2+ -dependent steps in vesicle mobilization. To explore
these questions further, we used computer simulations based
on kinetic models of Ca2+ -dependent exocytosis. A previous kinetic model [37] consisting of two pools of vesicles
(reserve and ready releasable pools) may be extended to add
an immediately releasable pool (see Section 2). The rate constant for fusion was made sensitive to a Ca2+ sensor with
multiple-binding sites (see Section 2).
We investigated how our capacitance measurements are
simulated by this exocytosis model using the estimated Ca2+
concentration in the 100 nm outer shell, determined by the
Ca2+ radial diffusion model, as Ca2+ input function (Fig. 7A
and B). Based on our measured size of 16 fF for the IRP
(Fig. 2) and the estimate of a Kd of 1–2 ␮M for the Ca2+ sensor (Fig. 3), we simulated satisfactorily the Cm measurements
during repetitive stimulation using a Kd of 1.5 ␮M (Fig. 7C).
However, simulations using these parameters yielded a poor
fit for single-pulse data and did not show saturation with
pulses longer than 100 ms (Fig. 7D, black line). To reach
saturation of the capacitance change during single pulses and
to obtain good-quality simulations for trains the rate constant for refilling the IRP (k2 ) had to be decreased (Fig. 7D)
and made Ca2+ -dependent (Fig. 7E and F). Fig. 7E and F
shows simulations using a calcium sensor for k2 with a singlebinding site and affinity of 0.3 ␮M (dark gray traces) and
with a sensor with two sites and an affinity of 0.9 ␮M (black
traces). The three-pool model described above gives a fairly
good description of the Ca2+ -dependence of depolarizationinduced exocytosis for single pulses and for repetitive stimulation. However, this model underestimates exocytosis for
pulses less than 50 ms long. A possible explanation for this
discrepancy is that there may be some co-distribution of Ca2+
channels and chromaffin granules [47]. In support of this
hypothesis, it was observed that for short depolarizing stimuli the Ca2+ entry is not uniform [7,6] and exocytosis occurred
preferentially at these “Ca2+ hotspots” [48]. A higher density
97
of Ca2+ channels would give a larger local Ca2+ concentration at these sites leading to more exocytosis, while repetitive
stimulation leads to a more uniform Ca2+ increase around the
cell perimeter.
The IRP is a fraction of the readily releasable pool of
vesicles that are preferentially associated with Ca2+ channels
[22,42]. Thus, flash photolysis causes release from the whole
readily releasable pool, whereas depolarization only releases
from a fraction of this pool. With such a model, the Ca2+ dependent mobilization of vesicles to the IRP may not be a
Ca2+ -dependent reaction per se. Instead, the IRP is the fraction of vesicles suitably located to see a high enough Ca2+
signal and the apparent vesicle mobilization comes about by
changes in the spatial distribution of the Ca2+ concentration,
so that a larger fraction of the readily releasable pool is sensitive to depolarization-induced Ca2+ entry. An alternative
hypothesis proposes a real Ca2+ -dependent mobilization of
granules to a site closer to Ca2+ channels. In support to this
possibility, Becherer et al. [26], using evanescent field imaging, described the mobilization of docked vesicles to a spatial
dominium closer to Ca2+ entry sites.
4. General conclusions
We have used simultaneous measurement of Ca2+ gradients and exocytosis to study the Ca2+ dependence of
depolarization-induced exocytosis during single pulse and
repetitive stimuli. The Ca2+ measurements were complemented by computer modeling to enable us to make predictions of the Ca2+ signal immediately beneath the cell
membrane that is sensed by the secretory apparatus. From
these data, we conclude that the Ca2+ sensor for fusion has
an affinity of 1–2 ␮M and three or more Ca2+ -binding sites.
This Kd estimation has an additional support from the simulations of the secretion process, which requires also a high
affinity sensor. We used the data to examine kinetic models
of exocytosis and conclude that the data are consistent with
Ca2+ -dependent fusion from a small immediately releasable
pool that can be refilled by Ca2+ -dependent mobilization of
vesicles.
Acknowledgements
The work presented in this manuscript was mostly carried out in Dr. Jonathan R. Monck’s laboratory, Department
of Physiology, UCLA. I specially thank Dr. Jonathan R.
Monck for his intellectual and material contribution, which
was indispensable for the completion of this work. I also
thank Dr. Bernard Ribalet and Dr. Lidia Szczupak for the
critical reading of the manuscript. This work was supported
by grant GM54340 (to Dr. J. Monck) from the National Institutes of Health (USA) and by grant PICT 05-11661 (to Dr. F.
Marengo) from SCYT-ANPCYT (Argentina).
98
F.D. Marengo / Cell Calcium 38 (2005) 87–99
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