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aj3,*
1Laboratory Neuroendocrinology-Molecular Cell
Physiology, Institute of Pathophysiology, Medical Faculty, Zalo
ka 4,
and 2Celica Biomedical Sciences Center, Stegne 21,
1000 Ljubljana, Slovenia; and 3Departments of
Ophthalmology and Physiology, University of California School of Medicine, San
Francisco, California 94143-0730
Submitted 13 November 2002; accepted in final form 19 March 2003
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ABSTRACT |
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INTRODUCTION |
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). This is in contrast with the millisecond time scale of
action potentials which trigger rapid and transient exocytotic events in
"conventional" neurons. Consequently, photoreceptor and bipolar
synapses are designed to optimize high rates of sustained neurotransmitter
release (reviewed in Juusola et al.
1996; Krizaj and Copenhagen
2002; von Gersdorff
2001). Analysis of molecular components of exocytosis at graded
synapses in the retina revealed that they share many features in common with
conventional synapses (von Kriegstein et
al. 1999). However, in contrast to conventional synapses, graded
synapses can maintain transmitter release for hours, suggesting that they
possess highly efficient mechanisms for vesicle retrieval and/or a virtually
limitless supply of releasable vesicles.
Ultrastructural analysis of synaptic terminals of graded neurons such as
photoreceptors and bipolar cells reveals an elaborate organization dominated
by the synaptic ribbon, a laminar structure whose function is thought to be
fast delivery of vesicles to the active zone
(Gray and Pease 1971;
von Gersdorff 2001). A similar
structure, the "dense body," is found in the synaptic region of
the hair cell (e.g., Safieddine and
Wenthold 1999). It has been suggested that these structures gate
both rapid and sustained phases of neurotransmitter release and thus provide
photoreceptors, bipolar cells, and hair cells with the ability to sustain
vesicle release rates ten to hundreds of times higher compared with those at
conventional synapses (Gray and Pease
1971; Parsons et al.
1994; Rao-Mirotznik et al.
1995; Rieke and Schwartz
1996; von Gersdorff
2001).
Exocytosis at most ribbon synapses is characterized by at least two
kinetically distinct components—a fast component ranging from several
milliseconds to several tens of milliseconds and a slow component in the order
of several hundred milliseconds (Beutner et
al. 2001; Gomis et al.
1999; Moser and Beutner
2000; Neves and Lagnado
1999; Parsons et al.
1994; Rouze and Schwartz
1998; Sakaba et al.
1997; von Gersdorff et al.
1996). Two models have been proposed to account for the fast and
the slow kinetic components of vesicle release at ribbon synapses. The
"sequential model" proposes a sequential activation of separate
vesicle pools. According to this model, a "rapidly releasable
pool" of fusion-ready vesicles is recruited from several rows of
vesicles tethered to the ribbon, whereas a much larger "reserve
pool" may be recruited from the population of nontethered vesicles
within the terminal (Neves and Lagnado
1999; Sakaba et al.
1997; von Gersdorff et al.
1996). The sequential model is supported by the observations in
terminals from teleost bipolar cells in which the rapidly releasable pool was
found to deplete before activation of the reserve pool
(Gomis et al. 1999;
Sakaba et al. 1997;
von Gersdorff et al. 1996),
and in amphibian hair cells, where prolonged rapid exocytosis involves
vesicles from a larger pool than could be in close proximity to the plasma
membrane at active zones (Parsons et al.
1994). Alternatively, the "parallel model" suggests
exocytosis occurs in parallel from spatially separated vesicle pools.
Therefore, according to this model, exocytosis need not occur solely from the
vesicles docked at the ribbon but may occur at active sites elsewhere within
the synaptic terminal. Evidence suggestive of parallel exocytosis in retinal
bipolar cell terminals has been recently provided by direct measurements of
vesicle dynamics using evanescent field microscopy. These experiments showed
that exocytosis of fusion-competent vesicles can occur from sites distal to
the synaptic ribbon (Zenisek et al.
2000). Parallel exocytosis has been documented in many other cell
types, including PC12 cells (Kasai et al.
1996), melanotrophs (Kreft et
al. 2003; Poberaj et al.
2002; Rupnik et al.
2000), chromaffin cells (Voets
2000), and pancreatic 
cells
(Takahashi et al. 1997;
reviewed in Kasai 1999).
Ribbon synapses are characterized by their ability to sustain prolonged
periods of elevated [Ca2+]i, which controls
the graded release of the neurotransmitter
(Sakaba et al. 1997;
Tachibana et al. 1993).
Paradoxically, the Ca2+ dependence of release seems to
be different between different classes of ribbon synapses in the retina:
whereas several release mechanisms are thought to operate in bipolar cells,
each at different optimal [Ca2+]i, rod
photoreceptors are thought to operate mostly at
[Ca2+]i < 2 µM
(Lagnado et al. 1996;
Rieke and Schwartz 1996;
Rouze and Schwartz 1998;
von Gersdorff and Matthews
1999). Flash photolysis experiments in bipolar cells suggest that
the [Ca2+]i threshold for fast exocytosis is
high, above approximately 20–50 µM
(Heidelberger et al. 1994;
Mennerick and Matthews 1996;
von Gersdorff et al. 1996).
Bipolar cells thus possess a large rapidly releasable pool activated by a
low-affinity Ca2+ sensor
(Heidelberger et al. 1994; but
see Rouze and Schwartz 1998).
Optical measurements using lipophilic dyes suggest that, in addition to the
high-threshold exocytosis, sustained vesicle cycling in bipolar cells may also
occur at submicromolar [Ca2+]i
(Gomis et al. 1999;
Lagnado et al. 1996;
Rouze and Schwartz 1998).
Bipolar ribbon synapses may thus possess both high- and low-affinity sensors
for exocytosis.
In contrast to bipolar cells, exocytosis in photoreceptors has not been
studied in great detail and the properties of kinetics of release and
Ca2+ dependence of secretion in rods and cones are not
well understood. The secretion responses in rod photoreceptors were shown to
be sustained and proportional to changes in
[Ca2+]i
(Rieke and Schwartz 1996).
Moreover, rod photoreceptors were reported to only operate at relatively low
(0.5 to 2.0 µM) [Ca2+]i
(Rieke and Schwartz 1996).
These findings led to the suggestion that transmitter release in rod
photoreceptors is mainly controlled by residual
[Ca2+]i and operates at high affinity for
Ca2+. It can be predicted, therefore, that exocytosis in
photoreceptors follows changes in global
[Ca2+]i rather than changes within
Ca2+ microdomains. The discrepancy in kinetics and
Ca2+ affinity of vesicle release between photoreceptors
and bipolar cells suggests that the synaptic transmission at the photoreceptor
synapse is fundamentally different from that of the bipolar synapse
(von Gersdorff and Matthews
1999). However, in the original study
(Rieke and Schwartz 1996) the
cells were not studied with the same protocol applied previously to bipolar
cells (Heidelberger et al.
1994). It is therefore not clear whether photoreceptors only
possess high Ca2+ affinity release. To test this
hypothesis experimentally, we re-investigated the kinetics and
Ca2+ dependence of exocytosis in photoreceptors using a
combination of flash photolysis of caged Ca2+ and
capacitance measurements to evoke and monitor changes in
[Ca2+]i and vesicle cycling, respectively.
Our methods were designed to mimic those used in the original studies on
retinal bipolar cells (e.g., Heidelberger
et al. 1994).
We found that kinetics of exocytosis at the photoreceptor synaptic terminal are similar to that reported for retinal bipolar cells, whereas Ca2+-sensitivity threshold is slightly lower, at approximately 15 µM. In a majority of cells tested we found both fast and slow components of exocytosis. Our results therefore suggest that exocytosis in photoreceptors is relatively similar to that observed in bipolar cells with respect to kinetics of vesicle release and its Ca2+ dependence.
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METHODS |
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Rod and cone photoreceptor cells from tiger salamander (Ambystoma
tigrinum) retina were acutely dissociated by enzymatic dissociation and
mechanical trituration, as previously described
(Krizaj and Copenhagen 1998).
Larval-stage tiger salamander retinas were dissected and incubated on a shaker
in calcium-depleted saline with papain (7 U/ml; Worthington, Freehold, NJ) for
25 min and triturated with a BSA-coated Pasteur pipette. The outer segments of
many rods were shorn during the isolation procedure, resulting in the absence
of the dark current and hyperpolarization of their membrane potentials. Cell
isolation was performed at room light; therefore, all cells that kept their
outer segments were completely light adapted. Cells were kept at 4°C in
80% L-15 medium supplemented with 10 mM HEPES, 20 mM glucose, 1 mM pyruvic
acid, 1 mg/ml bovine serum albumin (BSA), and 1 µl/ml liquid media
supplement containing transferrin and selenium (Sigma, St. Louis, MO). Cells
were seeded onto acid-cleaned glass coverslips coated with IgG and/or IgM
(Jackson ImmunoResearch, West Grove, PA) and the Sal-1 antibody (a kind gift
from Dr. Peter MacLeish; MacLeish et al.
1983).
Immunocytochemistry and confocal microscopy
For immunocytochemical detection of synaptotagmin I, enzymatically
dissociated photoreceptor cells were washed with phosphatebuffered saline
(PBS) and then fixed for 15 min in 4% paraformaldehyde in PBS. Cells were kept
in fixative containing 0.1% of Triton X-100 for 10 min and washed four times
with PBS. Nonspecific staining was reduced by incubating cells in 3% BSA and
10% normal goat serum in PBS. Cells were then incubated with primary
antibodies for 2 h at 37°C. We used mouse anti-synaptotagmin I monoclonal
antibody (Leveque et al. 1992;
Takahashi et al. 1991) diluted
1:2,000 in PBS containing 3% BSA. Cells were then washed and incubated in PBS
containing Alexafluor 546 labeled anti-mouse secondary antibodies (1:2,000;
Molecular Probes, Oregon, USA) and 3% BSA for 45 min. Light Antifade Kit
(Molecular Probes) was used for mounting. Cells were monitored with a confocal
microscope (Zeiss, LSM 510, objective x63, NA = 1.4). Alexafluor 546 was
excited with a He/Ne (543 nm) laser. Emission signals were filtered using a LP
560-nm filter.
Membrane capacitance measurements
Compensated membrane capacitance (Cm) measurements were used
(Lindau and Neher 1988;
Neher and Marty 1982;
Zorec et al. 1991), employing
a SWAM IIB patch-clamp/lock-in amplifier (Celica, Ljubljana, Slovenia),
operating at 1.6 kHz lock-in frequency.
On establishment of the whole-cell configuration, Cm and Ga (access conductance) were compensated by Cslow and Ga compensation controls. A sine voltage of 314 mV (peak-to-peak) was applied. The phase-angle setting was determined by a 1-pF calibration pulse and by monitoring the projection of this pulse from the C (signal proportional to Cm) to the G output of the lock-in amplifier (asterisk in Fig. 2). These two signals were stored unfiltered (C-DAT4 recorder, Cygnus, USA) for off-line analysis. Simultaneously, we recorded filtered (300 Hz, 4-pole Bessel) C and G signals, the fluorescence intensity from a C660 photon counter (Thorn EMI, UK), and membrane current (0–10 Hz, low-pass). PhoCal program (LSR, UK) was used to acquire signals every 5 ms. For high temporal resolution measurements of Cm, the records on DAT were played back and a 10-s epoch of the signal enveloping each flash was digitized at 50 kHz using a computer disk recorder (CDR) program (J. Dempster, Strathclyde Electrophysiology Software). Signals were digitally filtered at 1 kHz (2-way 150th order FIR filter, Math Works MATLAB) and resampled at 10 kHz. The pipette solution contained the following (in mM): 88 KCl, 8 TEACl, 32 KOH/HEPES, 1.6 Na2ATP, 1.6 MgCl2, 4.0 K4-NP-EGTA, 2.9 CaCl2, 0.5 furaptra, pH 7.6. The bath contained the following (in mM): 105.4 NaCl, 4 KCl, 1.6 MgCl2, 0.4 NaH2PO4, 4 NaHCO3, 8 Na HEPES, 8 D-glucose, 1.4 CaCl2, pH 7.6.
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All recordings were made at room temperature. The salts were obtained from
Sigma. Cells were voltage clamped at a holding potential of –50 mV. The
average cell capacitance was 13.2 ± 0.83 pF (n = 23; mean
± SE). Recordings were made with pipette resistances between 1 and 4
M
(measured in KCl-rich solution), giving access conductance of more
than 80 nS (117 ± 14.2 nS; mean ± SE). The pipette and bath
solutions were of similar osmolarity (within 5%) measured by freezing point
depression (Camlab, Cambridge, UK). Na2ATP was included in the
pipette solution since it has a major role in preparing synaptic vesicles for
fusion (Heidelberger 1998).
Under our experimental conditions, endocytosis was not regularly observed,
probably as a result of the washout of a critical component (e.g.,
Parsons et al. 1994). We
therefore focused on the exocytotic component. Moreover, although several UV
flashes were applied during any given experiment, the second flash always
elicited a smaller capacitance jump, presumably because of vesicle depletion
and blocked endocytosis. Only the first exocytotic responses to photolysis
flashes were analyzed in this study.
Flash photolysis and [Ca2+]i measurements
The Ca2+ cage NP-EGTA (Molecular Probes) was used to
elevate [Ca2+]i by flash photolysis
(Ellis-Davies and Kaplan 1994).
A 1-ms UV flash from an Xe arc flash lamp
(Rapp and Güth 1988) was
delivered to cells through a x40 fluor oil immersion objective of a
Nikon Diaphot microscope. The same optical pathway as in flash photolysis was
used to illuminate the fluorescent [Ca2+] indicator
furaptra (Molecular Probes). A combination of two dichroic mirrors was used.
The first one was a 390-nm dichroic positioned at 45°, which passed
through the 420-nm light for furaptra excitation from a Xe arc lamp and
reflected the light below 390 nm for NP-EGTA flash photolysis from a Xe arc
flash lamp. The second 430-nm dichroic reflected both lights through the
objective to the cell under experiment and allowed only furaptra fluorescent
light to pass back to the photomultiplier through a 510-nm barrier filter.
[Ca2+]i measurements were performed as
described by Carter and Ogden
(1994). The equation used in
calculation is
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;
Krizaj et al. 2003). |
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RESULTS |
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We photolyzed the caged Ca2+ compound NP-EGTA to
rapidly increase cytosolic [Ca2+] in a spatially uniform
manner in the photoreceptor terminals
(Neher and Zucker 1993). Only
cells with well-preserved morphology of the synaptic terminal were examined
physiologically (Fig. 1). In
several cells, the synaptic terminals were resorbed following isolation and
these cells were avoided in single-cell capacitance measurements described in
this study.
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Figure 2 shows a
Cm response of a rod photoreceptor to a flash of UV light. The
flash discharge increased [Ca2+]i to 56 µM
in <10 ms (the time resolution limit of fluorescence recording system;
Fig. 2, arrow), whereupon
[Ca2+]i returned to the baseline with a time
constant of around 5 s. Following flash photolysis and
[Ca2+]i increase within the terminal, an
exponential increase in Cm to the final amplitude of 1.86 pF
(labeled A in Fig. 2)
was observed. A small change in the real part of admittance signal (G) was
observed following flash photolysis (G,
Fig. 2). The change in G was
transient and was not correlated to the increase in Cm. It may be
attributed to calcium-induced activation of ion channels
(Maricq and Korenbrot 1988;
Moriondo et al. 2001) or/and
may reflect fusion pore conductance of many synchronously exocytosed vesicles
(Lindau 1991). The time course
of the Cm response in this cell was well described by a single
exponential function with a time constant of 1.19 s. Assuming a vesicle
diameter of 30 nm and assuming the specific membrane capacitance of 8
fF/µm2, a single-vesicle capacitance is approximately 23 aF
(von Gersdorff et al. 1996;
Zenisek et al. 2000).
Therefore the amplitude of 1.86 pF corresponds to the fusion of approximately
80,000 vesicles with the membrane of the photoreceptor. The number of
exocytosed vesicles is at least one order of magnitude larger than the pool of
rapidly releasable vesicles docked at approximately 60 ribbons in the teleost
Mb1 bipolar cell (von Gersdorff et al.
1996; von Gersdorff and
Matthews 1999).
We measured the maximal amplitude of Cm elevation after stimulation with flash photolysis of NP-EGTA. Only Cm responses to the first photolysis flash were analyzed in any cell (see METHODS). The evoked maximal [Ca2+]i varied between 5 and 60 µM. The Cm amplitude exhibited a sigmoidal dependence on [Ca2+]i with the half-maximal response at approximately 30 µM for both rods and cones (Fig. 3). The threshold of [Ca2+]i at which exocytosis was observed was at 10–15 µM [Ca2+]i, whereas maximal Cm amplitude was observed at approximately 50 µM [Ca2+]i.
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To quantify the kinetics of Cm responses, both single- and double-exponential functions were fitted to the rising phase of each Cm trace. Capacitance responses were best described by either a single exponential (39% of 18 photoreceptors studied) or a double-exponential time course to the peak (61%). The goodness of fit was evaluated by squared correlation coefficients (coefficients of determination) and by visual inspection. The two types of responses are illustrated in Fig. 4 for two cones. The response of the cell depicted in Fig. 4A was best fit with a single exponential function with a time constant of 1.19 s. The majority of cells exhibited two visually distinct time courses. These were best fit with a double exponential function. One such cell, shown in Fig. 4B, had a time constant for the rapid phase of the Cm response of 94.1 ms, whereas the time constant for the slow phase was 1.77 s. The range of time constants for the rapid exocytic response was from 0.5 to 135.0 ms and for the slow exocytic response in the biphasic Cm traces was from 0.02 to 8.40 s. No significant differences in the kinetics of Cm responses were observed between rods and cones (Fig. 3). The Ca2+ dependence of rate constants appeared as a step-like function, with an all-or-none response beyond the threshold [Ca2+]i of 10–15 µM (Fig. 5C). The slopes of the linear fit of logarithmized rapid, slow, and single time constants versus [Ca2+]i did not differ significantly from zero.
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Figure 5A shows the estimated amplitudes of exponential functions obtained in fitting monophasic and biphasic exponential functions to the experimentally recorded flash-evoked changes in Cm. Mean amplitudes of rapid and slow components of double-exponential exocytotic responses were not significantly different (0.84 ± 0.35 and 0.82 ± 0.20 pF, respectively; n = 11 cells, mean ± SE). Likewise, the mean amplitude of the single exponential response in Cm was not significantly different (0.70 ± 0.23 pF; n = 7) from either the mean rapid or the mean slow components. The mean rate constants of single- or double-exponential curves in Cm are graphed in Fig. 5B. Mean rate constant of the rapid exocytosis was 53 times faster than the mean rate constant for the slow component of Cm response (420 ± 168 and 7.85 ± 5.02 s–1, respectively). The mean rate constant of single exponential exocytotic response in Cm (17.5 ± 12.4 s–1) was not significantly different from the mean rate constant of the slow component of double-exponential exocytotic response. Corresponding mean time constants for rapid, slow, and single exponential response are 24.7 ± 13.7 ms, 1.59 ± 0.75 s, and 0.51 ± 0.17 s, respectively. These results indicate that the majority of cells responded to stimulation with a biphasic exocytotic response. This may indicate that in these photoreceptor cells two kinetically distinct readily releasable pools of vesicles coexist.
Synaptotagmin I is localized to vesicles in the synaptic terminal of photoreceptors
Synaptotagmin I is a synaptic vesicle protein suggested to be involved in
neurotransmitter release at conventional synapses
(Geppert et al. 1994;
Schiavo et al. 1997). This
ubiquitous vesicle protein has a relatively low affinity for
Ca2+ (Davis et al.
1999) and is thought to support fast, "synchronous"
transmitter release (Geppert et al.
1994; Goda and Stevens
1994; Kreft et al.
2003; Schiavo et al.
1997; Voets et al.
2001). Expression of synaptotagmin I in salamander rods and cones
provides additional confirmatory evidence for our observation that these cells
are capable of fast exocytosis.
Figure 1 (left) shows a Nomarski image of an isolated rod photoreceptor. The cell consists of the inner segment with a cell body and a prominent synaptic terminal. The outer segment was shorn from this cell during the dissociation procedure. The cell was immunolabeled with the antibody against synaptotagmin I. As shown in Fig. 1 (right), synaptotagmin I is selectively expressed in the synaptic terminal, consistent with a large pool of vesicles being preserved under our experimental conditions. Similar results were obtained with SV2, another marker for vesicles within photoreceptor terminals (data not shown). This finding indicates that rapid exocytosis, observed in salamander photoreceptors, may be subserved by molecular machinery similar to that found in conventional synapses. Moreover, the observation of robust presence of synaptotagmin- and SV2-positive terminals is consistent with our physiological measurements of secretory function of the photoreceptor terminal in dissociated cells.
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DISCUSSION |
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).
These findings (Rieke and Schwartz
1996) suggested that transmitter release from rod photoreceptors
is fundamentally different from that at the bipolar cell terminal
(Heidelberger et al. 1994;
Sakaba et al. 1997).
In our experiments, measurements of Cm combined with flash
photolysis of caged Ca2+ revealed that the majority of
cells (61%) respond to flash photolysis with an exocytotic response consisting
of two kinetic components (Fig.
4). These two components were observed in both rods and cones
(Fig. 3). The fast response was
characterized by a mean time constant of 24 ms, ranging from 0.5 to 135 ms.
The vesicles comprising the fast exocytotic pool are likely to include those
already docked at active sites and tethered to the ribbon, as shown in studies
with bipolar cells (Heidelberger et al.
1994; Mennerick and Matthews
1996; Neves and Lagnado
1999; Sakaba et al.
1997; von Gersdorff et al.
1996). The capacitance jump corresponding to the rapidly
releasable pool in teleost bipolar cells is approximately 150 fF, whereas the
mean size of the fast component in salamander photoreceptors was 840 fF,
corresponding to release of approximately 34,000 vesicles. This is similar to
what has been observed in mammalian cochlear hair cells (approximately 40,000
vesicles; Beutner et al. 2001)
but is much higher than the estimates in bipolar cells (about 6,000 vesicles;
von Gersdorff et al. 1996). A
photolysis flash could trigger exocytosis of several tens of thousands of
vesicles in both fast and slow components, suggesting that the exocytosis
draws vesicles from a larger pool than could exist in close proximity to the
synaptic ribbon. For example, a ribbon in a mammalian rod may tether about 770
vesicles (Rao-Mirotznik et al.
1995). This would correspond to about 18 fF. Thus even if taking
into account that salamander ribbons can accommodate a larger number of
vesicles than mammalian rods and that a single salamander rod terminal may
possess multiple ribbons, it is unlikely that the capacitance jump observed
during the fast exocytic response derives solely from the pool of vesicles
tethered to the ribbon and docked at the active site. The amplitudes of
exocytic responses in rods and cones were not significantly different (not
shown).
We also found that a smaller percentage of examined cells lacked the faster
component of exocytosis. It is possible that cells with a single exocytotic
(slow) phase lack the rapid phase due to the cell isolation procedure.
Whatever the cause underlying the differences between the slow and fast
response, the absence of the latter during the slow exocytotic response argues
against the model in which the rapid and slow components in photoreceptor
Cm occur by sequential coupling of two pools of vesicles
(Neher and Zucker 1993;
Thomas et al. 1993;
Xu et al. 1998). As is the
case with the fast pool, the size of the slow sustained pool is likely to be
larger than the combined surface area of the vesicles tethered to the ribbon,
suggesting an additional contribution from fusion competent vesicles distal to
the ribbon. Zenisek et al.
(2000) reported that vesicle
exocytosis in fish bipolar cells may occur at sites away from the active zone
and thus ribbons may not be absolutely required for docking and exocytosis at
graded synapses. Similarly, electron microscopic analysis of salamander
photoreceptor terminals suggests that exocytosis may occur at the ribbons as
well as at basal junctions, specializations of the synaptic terminal which
contact postsynaptic processes in the absence of associated synaptic ribbons
(Lasansky 1973). Several kinds
of basal junctions have been observed in salamander rods and cones
(Lasansky 1973); likewise,
recent studies have shown that synaptic vesicles in frog saccular cells may be
docked at the presynaptic membrane at some distance from the active zones
(Lenzi et al. 1999). A
parsimonious explanation for the two phases observed in salamander
photoreceptors may be that they occur via parallel vesicle release processes
at ribbon and at extra-ribbon sites. Changes in Cm may also reflect
fusion of subcellular compartments other than vesicles (for example,
lysosomes, endosomes, granules, etc.). Using capacitance measurements on
single cells, we cannot distinguish the relative contributions of these
compartments to the total Cm increase. Furthermore, large changes
in Cm may be due to multivesicular exocytosis.
If photoreceptors operated only using asynchronous release, they would be
expected to lack synaptotagmin I, known to be a major vesicle component
underlying fast synchronous exocytosis
(Geppert et al. 1994).
Synaptotagmin I binds multiple Ca2+ ions
(Fernandez-Chacon et al. 2001;
Ubach et al. 1998), has a
relatively low affinity for Ca2+
(Davis et al. 1999) and is
essential for fast, Ca2+-triggered exocytosis in
conventional synapses (Geppert et al.
1994). We therefore examined the subcellular distribution of
synaptotagmin I in photoreceptors, using immunocytochemistry and confocal
microscopy. The anti-synaptotagmin I antibody strongly labeled synaptic
terminals in both rods and cones, consistent with the presence of fast,
synchronous vesicle release in these cells. The expression of synaptotagmin I
in salamander photoreceptors is consistent with the recent report by von
Kriegstein et al. (1999) in
which a prominent synaptotagmin I signal was observed in the outer retina of
cattle and supports the idea that conventional and ribbon synapses share many
molecular components of the exocytotic machine
(Ullrich and Sudhof 1994;
von Kriegstein et al. 1999).
For example, other elements of the soluble N-ethylmaleimide-sensitive
factor attachment protein receptor complex found in photoreceptors include
synaptosome-associated protein of 25 kDa and synaptobrevin
(Sherry et al. 2001;
von Kriegstein et al.
1999).
[Ca2+]i above 15 µM was required to
trigger secretion in photoreceptors with a half-saturating response at
approximately 30 µM (Fig.
3). This Ca2+ dependence of release is
similar to that previously reported in retinal bipolar cells. Dialysis of
bipolar cell terminals with solutions containing buffered amounts of elevated
calcium concentration and flash photolysis experiments demonstrated that
exocytosis in these cells requires [Ca2+]i
above approximately 20–50 µM
(Heidelberger et al. 1994;
von Gersdorff and Matthews
1994). Our finding therefore suggests that exocytosis in
photoreceptors may be triggered within microdomains of high-calcium
concentration expected near open calcium channels
(Chad and Eckert 1984;
Neher 1998) rather than by the
relatively low [Ca2+]i measured in cells at
dark potentials (approximately 300 nM at approximately –40 mV;
Krizaj et al. 1999). This view
is also supported by the observation that a high density of voltage-gated
calcium channels is often found at the active zone close to the synaptic
ribbon (Morgans 2001;
Morgans et al. 1998;
Nachman-Clewner et al. 1999;
Raviola and Raviola 1982;
reviewed by Lenzi and von Gersdorff
2001). The technique we used is not sensitive enough to resolve
fusion of single synaptic vesicles (e.g.,
Kreft and Zorec 1997).
Similarly, exocytotic rates of ≤2,000 vesicles/s measured in bipolar cells
using optical dyes (Lagnado et al.
1996; Rouze and Schwartz
1998) may have been missed during capacitance measurements by
Heidelberger et al. (1994)
(cf. Beutner et al. 2001).
Rieke and Schwartz (1996)
reported that photoreceptor exocytosis operates at high (0.5–1.0 µM)
Ca2+ affinity. We do not know the reason for the
observed differences in our study and in the work by Rieke and Schwartz
(1996), but different
experimental approaches may account for this. For example, Rieke and Schwartz
(1996) adjusted the intensity
and duration of the photolysis flash so that the time course of
[Ca2+]i changes approximated the cell voltage
during the light response. The long rise times (on the order of seconds) and
long decay times of [Ca2+]i following flash
photolysis may have contributed to favoring a component of release with high
Ca2+ affinity. However, the time course of
Ca2+ decay was much slower from the time course expected
from the synaptic terminal (Morgans et al.
1998; D. Krizaj, unpublished observations) and known properties of
synaptic transfer at the photoreceptor synapse
(Schnapf and Copenhagen 1982).
It is possible that photoreceptors possess both fast release and slow
asynchronous release at high and low Ca2+ affinity.
Synaptotagmin I may be the Ca2+ sensor required for the
fast release, whereas slow release may be supported by at present unknown
proteins or different isoforms of synaptotagmins. More than 12 isoforms of
synaptotagmins are present in the brain in addition to synaptotagmin I and may
participate in regulating secretion at higher Ca2+
affinities (Sugita et al.
2002). For example, synaptotagmin sIV, VI, VII, VIII, and IX have
been recently identified at the "ribbon" synapse of the hair cell
(Safieddine and Wenthold
1999).
In summary, our study revealed that at least two separate pools of secretory vesicles are present in rod and cone photoreceptors and that salamander photoreceptors possess a large pool of fusion-ready vesicles activated by a low-affinity Ca2+ sensor similar to the bipolar cells. Our results suggest that graded release in photoreceptors and bipolar cells may be regulated by similar molecular pathways.
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ACKNOWLEDGMENTS |
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This work was supported by Ministry of Education, Science, and Sport of the
Republic of Slovenia Grant J3-2344-7421-00 to M. Kreft, National Eye Institute
Grant EY-32918 to D. Kriaj, Wheeler Center for Neurobiology of
Addiction to D. Kriaj, University of California San Francisco Academic
Senate Individual Investigator Award to D. Kriaj, P3 521 381 to R.
Zorec, and the European Union Grant QLG3 2001–2004 to R. Zorec.
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FOOTNOTES |
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* M. Kreft and D. Kriaj contributed equally to this paper. 
Address for reprint requests: R. Zorec, Celica Biomedical Sciences Center, Stegne 21, 1000 Ljubljana, Slovenia (E-mail: Robert.Zorec{at}mf.uni-lj.si).
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