Okadaic acid disrupts synaptic vesicle trafficking in a ribbon-type synapse

Abstract

Protein phosphorylation plays an essential role in regulating synaptic transmission and plasticity. However, regulation of vesicle trafficking towards and away from the plasma membrane is poorly understood. Furthermore, the extent to which phosphorylation modulates ribbon-type synapses is unknown. Using the phosphatase inhibitor okadaic acid (OA), we investigated the influence of persistent phosphorylation on vesicle cycling in goldfish bipolar cells. We followed uptake of FM1-43 during vesicle recycling in control and OA-treated cells. FM1-43 fluorescence spread to the center of control synaptic terminals after depolarization elicited Ca²⁺ influx. However, OA (1–50 nM) impaired this spatial spread of FM1-43 in a dose-dependent manner. Capacitance measurements revealed that OA (50 nM) did not modify either the amount or kinetics of exocytosis and endocytosis evoked by depolarizing pulses. The extremely low concentrations of OA (1–5 nM) sufficient to observe the inhibition of vesicle mobility implicate phosphatase 2A (PP2A) as a major regulator of vesicle trafficking after endocytosis. These results contrast with those at the neuromuscular junction where OA enhances lateral movement of vesicles between distinct vesicle clusters. Thus, our results suggest that phosphatases regulate vesicle translocation at ribbon synapses in a different manner than conventional active zones.

Cycles of protein phosphorylation and dephosphorylation play a key role in regulating synaptic transmission. Many proteins involved in vesicle trafficking are constitutively phosphorylated and undergo dephosphorylation upon nerve terminal depolarization. Others become phosphorylated in response to calcium influx (reviewed by Turner et al. 1999). Protein kinases may also regulate vesicle fusion by increasing the size of the pool of releasable vesicles or by increasing mobilization of vesicles from the reserve to the readily releasable pool (Llinás et al. 1985; Gillis et al. 1996; Smith et al. 1998; Stevens and Sullivan 1998). Kinases and phosphatases are therefore good candidates as possible regulators of synaptic vesicle cycling (Li and Murthy 2001).

Betz and Henkel (1994) used several phosphatase and kinase inhibitors to investigate the basis of vesicle trafficking in frog motor nerve terminals. Okadaic acid (OA), an inhibitor of phosphatases, was found to disrupt the organization of synaptic vesicles labeled with the fluorescent dye FM1-43. A similar result was observed in hippocampal bouton synapses using CY3-conjugated antibodies as a fluorescent marker for synaptic vesicles (Kraszewski et al. 1995, 1996). These results support the hypothesis that regulated protein phosphorylation and dephosphorylation is crucial for vesicle trafficking and clustering inside nerve terminals.

Goldfish retinal bipolar cell terminals can sustain high release rates and have thus been used to study synaptic vesicle fusion and recycling (Tachibana and Okada 1993; von Gersdorff and Matthews 1997; Rouze and Schwartz 1998; Neves and Lagnado 1999). The large size of the synaptic terminal (> 10 μm) allows direct patch-clamping and confocal microscope fluorescence imaging experiments. These nerve terminals form ribbon-type synapses characterized by the presence of a multilamellar structure (the ribbon itself) with vesicles tethered to it by short filamentous arms (von Gersdorff et al. 1996). The architecture of the ribbon suggests that it functions as a reservoir for a pool of release-ready vesicles (Parsons et al. 1994; von Gersdorff et al. 1996; Lenzi et al. 1999). Following endocytosis, vesicles may either be stored near their original release sites or in a more distal reserve pool from which they may be mobilized according to the intensity of nerve stimulation (Pieribone et al. 1995; Kuromi and Kidokoro 1998; Gomis et al. 1999; Pyle et al. 2000; Richards et al. 2000). Despite the identification of distinct vesicle pools within bipolar cell terminals, the ‘cues’ that govern vesicle reinternalization and trafficking towards and from these vesicle pools remain obscure.

Here we investigate whether OA affects vesicle cycling in a ribbon-type synapse. Bipolar terminals were exposed to OA and stained with FM1-43 to label vesicles undergoing endocytosis (Cochilla et al. 1999). OA restricted fluorescence accumulation to an area near the plasma membrane, even at extremely low concentrations (i.e. 5 nM), but did not affect vesicle exocytosis or endocytosis assayed by capacitance measurements. Our results suggest that PP2A phosphatase activity is necessary for the continuous movement of freshly endocytosed vesicles to the interior of ribbon-type nerve terminals. Furthermore, since multiple bouts of exocytosis occurred in the presence of OA, these results indicate that the mechanisms that govern vesicle mobilization towards the plasma membrane are fundamentally different from those that govern vesicle trafficking away from the plasma membrane in bipolar cells.

Materials and methods

Preparation of goldfish retinal bipolar cells

Bipolar cells were acutely dissociated from goldfish retina according to the protocol described by Heidelberger and Matthews (1992). Briefly, retinas were removed from the eyecups and treated with hyaluronidase (Sigma, 1100 units/mL) followed by Papain digestion (Fluka, 30 units/mL). Three or four pieces of retina were mechanically dissociated with a small-bore glass Pasteur pipette, plated on a glass coverslip, and mounted in a perfusion chamber. Fish Ringer (in mM): NaCl 120, KCl 2.6, MgCl₂ 1.0, HEPES 10, was prepared by omitting CaCl₂ or adding 2.5 mM CaCl₂. Solutions containing 50 mM KCl were prepared by iso-osmotic replacement of NaCl.

Fluorescence imaging with FM1-43

FM1-43 (Betz et al. 1992; Ryum et al. 1993) is a fluorescent tool for monitoring secretion in vivo (reviewed by Guatimosim et al. 1998). Fluorescence images were acquired with a Zeiss water immersion objective (40 × 1.2 NA) in an Axiovert 100 microscope coupled to a Bio-Rad MRC 1024 laser scanning confocal system. A water-cooled argon laser was used to excite the preparation using FITC optics. Bipolar cells were stained with FM1-43 according to the protocol of Lagnado et al. (1996). Dissociated bipolar nerve terminals were perfused with modified Fish Ringer containing FM1-43 (8 μM), 0 mM Ca²⁺ and KCl 50 mM (iso-osmotic replacement of NaCl). After 120 s, bipolar cells were perfused with Fish Ringer containing 2.5 mM Ca²⁺, 8 μM FM1-43, and 50 mM KCl for 120 or 360 s and an image was acquired. Cells pretreated with phosphatase inhibitors were exposed to OA for 30 min before staining with FM1-43.

Capacitance and calcium current measurements

Electrical measurements of calcium currents and membrane capacitance were made using conventional whole-cell patch-clamp recordings (von Gersdorff and Matthews 1997). Capacitance was monitored with the automatic-compensation feature of the computer controlled EPC-9 patch clamp amplifier using the Pulse software emulation of a lock-in amplifier in the ‘Sine + DC’ mode and the XChart software (HEKA, Germany). The lock-in sine wave had a peak-to-peak amplitude of 30 mV at 1000 Hz. The holding potential was −60 mV. The patch pipette solution contained (in mM): 120 Cs-gluconate, 15 TEA-Cl, 25 or 35 HEPES, 2 or 3 MgCl₂, 2 Na₂ATP, 0.5 GTP, 0.5 EGTA at pH 7.2. The external solution contained the following (in mM): 115 NaCl, 2.6 KCl, 1.0 MgCl₂, 2.5 CaCl₂, 12 glucose, 10 HEPES at pH 7.3. OA (25–50 nM) was added to the extracellular solution according to the experimental protocol and 50 nM was included in the patch pipette for experiments with cells pretreated with OA.

Fluorescence imaging analysis

Fluorescence intensity profile analysis

Analyses of the fluorescence distribution pattern in control and OA-treated groups (Figs 1–3 and 6) were performed using the software Image Tools (University of Texas at San Antonio, TX, USA). We performed fluorescence profile analyses to compare spatial fluorescence distribution in control and OA-treated terminals. A line was traced along the same regions of a nerve terminal and fluorescence intensity profiles (256 gray levels) were generated by the software. The graphs of fluorescence against distance (in microns) for each pair of images were plotted using Sigma Plot 5.0.

Fig. 1.

Staining of recycled synaptic vesicles at retinal bipolar cell terminals with the styryl dye FM1-43. DIC image of a goldfish retinal Mb bipolar cell. A bulbous synaptic terminal is connected via a thick axon to a flask-shaped cell soma with short, stout dendrites. Scale bar = 10 μm. (a) Fluorescence image of the same cell after perfusion with Fish Ringer containing 50 mM KCl, 8 μM FM1-43, and 0 mM Ca²⁺. Note the fluorescence only at the plasma membrane. Arrow points to bipolar nerve terminal. (b) Fluorescence image of the same cell obtained after 120 s of perfusion with Fish Ringer containing 50 mM KCl, 8 μM FM1-43 and 2.5 mM Ca²⁺. Note that the fluorescence was internalized during the depolarization in the presence of calcium. Arrow points to bipolar nerve terminal. (c) Same terminal as shown in (a) at higher magnification. Pixel intensity was measured across the horizontal dashed line giving a fluorescence profile that is shown in panel (e). (d) Higher magnification of terminal of panel (b). The same dashed line as in panel (c) was drawn in (d) and fluorescence intensity was measured. Note the conspicuous fluorescence accumulation at the central of the terminal. (e) Fluorescence intensity profile. The dashed line and the continuous line represent the terminal at (d) and (c), respectively. Stained synaptic vesicles are thus able to move from the periphery of the terminal to the center in less than 120 s.

Fig. 3.

The OA effect on vesicle recycling is dose-dependent. Brightness through different regions across bipolar nerve terminals. Data from line profiles were plotted and grouped in three regions: two at the cell periphery (0–2 μm and 8–10 μm) and one region at the central zone (4–6 μm) for OA-treated nerve terminals. White bars correspond to mean ± SEM of pixels intensity after perfusion with Fish Ringer with 50 mM KCl, 8 μM FM1-43 and 0 mM Ca²⁺. Black bars corresponds to mean ± SEM of pixels intensity after perfusion with Fish Ringer with 50 mM KCl, 8 μM FM1-43 and 2.5 mM Ca²⁺. The results presented are from five cells pretreated with 0.1 nM OA (a), four cells with 1.0 nM (b), three cells with 5.0 nM (c), seven cells with 25 nM (d) and six cells with 50 nM (e).

Fig. 6.

OA impairs vesicle movement towards the center of terminals. (a) Representative fluorescence intensity profile of control bipolar cells (n = 4) that were perfused with Fish Ringer containing 50 mM KCl, 8 μM FM1-43 and 0 mM Ca²⁺ for 2 min. The continuous line represents the fluorescence intensity profile measured through the line drawn across the nerve terminal. The dashed line represents the fluorescence intensity profile across the same nerve terminal after perfusion with Fish Ringer containing 50 mM KCl, 8 μM FM1-43 and 2.5 mM Ca²⁺ for 6 min. Note the typical fluorescence spreading towards the central zone of the terminal. (b) Bipolar cells were pretreated with OA 25 nM for 30 min and exposed to Fish Ringer containing 50 mM KCl, 8 μM FM1-43 and 0 mM Ca²⁺ for 2 min. The continuous line represents the fluorescence intensity profile measured through the line drawn across a representative nerve terminal (n = 7). The dashed line represents the fluorescence intensity profile across the same nerve terminal after perfusion with Fish Ringer containing 50 mM KCl, 8 μM FM1-43 and 2.5 mM Ca²⁺ for six minutes. Note the much smaller increase in fluorescence at the central zone. (c and d) Brightness through different regions across bipolar nerve terminals. Data from line profiles were plotted and grouped in three regions: two at the cell periphery (0–2 μm and 8–10 μm) and one region at the central zone (4–6 μm) for control and OA-treated nerve terminals. White bars correspond to mean ± SEM of pixels intensity after perfusion with Fish Ringer containing 50 mM KCl, 8 μM FM1-43 and 0 mM Ca²⁺ (four and seven cells for control and OA, respectively). Black bars corresponds to mean ± SEM of pixel intensity after 6 min of perfusion with Fish Ringer containing 50 mM KCl, 8 μM FM1-43 and 2.5 mM Ca²⁺ (four and seven cells for control and OA, respectively).

Fluorescence intensity measurements of regions labeled by FM1-43 in different conditions

Fluorescence measurements described in Fig. 5 were performed using the software Metamorph Imaging System 3.6 (Universal Imaging Corporation). We tested if fluorescence changes near the plasma membrane were an artifact due to repeated perfusions or reflected a specific effect of OA. Cells were stimulated with Fish Ringer lacking Ca²⁺ or where Ca²⁺ was equimolarly replaced by Mg²⁺ (i.e. total of 2.5 mM Mg²⁺). An image (image 1) was saved after 120 s of perfusion in Fish Ringer containing 8 μM FM1-43, 50 mM KCl, 0 mM Ca²⁺ (image 1, Figs 5a–c). A second image (image 2) was then obtained after 240 s of perfusion with Fish Ringer containing 8 μM FM1-43, 50 mM KCl and 0 mM Ca²⁺ (Fig. 5d), 2.5 mM Ca²⁺ (Fig. 5e) or where Ca²⁺ was replaced with 2.5 mM Mg²⁺ (Fig. 5f). Images were automatically thresholded (30–255 gray levels range) and a region of interest was saved for image 1. The same treatment was then applied for image 2. The total number of pixels contained in the thresholded region of image 2 was subtracted from image 1. The resulting pixel values were averaged with Excell and the graph was plotted using Sigma Plot 5.0.

Fig. 5.

OA causes fluorescence accumulation near the plasma membrane. (a–f) Representative goldfish bipolar terminals of control (a,c,d,f) and pretreated with OA (b,e). Bipolar cells were exposed to Fish Ringer 0 mM Ca²⁺ containing 50 mM KCl, 8 μM FM1-43 (Fig. 5a–c) and an image was saved after 2 min of perfusion (image 1). Another image (image 2) was saved after an additional two minutes perfusion with the following solutions: Fish Ringer containing 50 mM KCl, 8 μM FM1-43 with 0 mM Ca²⁺ (5d) or 2.5 mM Ca²⁺ (Fig. 5e) or 2.5 mM Mg²⁺ (Fig. 5f). Each pair of images for each cell was automatically given thresholds using the software Metamorph and the total number of pixels contained in the threshold region in image 2 was subtracted from image 1. (g) Bars represent variations of the thresholded region for each experimental condition tested. Black bars are mean ± SEM for cells that were not exposed to OA (10 cells for 0 Ca²⁺ and seven cells for 2.5 mM Mg²⁺). The white bar is mean ± SEM of six cells that were treated with OA (50 nM) and exposed to 2.5 mM Ca²⁺.

Results

Okadaic acid interferes with the pattern of FM1-43 staining in bipolar cell terminals

Betz and Henkel (1994) observed at the frog neuromuscular junction (NMJ) that in nerve terminals labeled with FM1-43 OA interfered with the formation of synaptic vesicle clusters. OA also appeared to unmask a powerful lateral translocation process that caused vesicles to move from one active zone cluster to another. We performed experiments to investigate if sustained phosphorylation induced by OA causes spatial changes in the distribution of FM1-43 staining of goldfish retinal bipolar cells that contain ribbon-type active zones.

Figure 1(a) shows a typical confocal image (from a total of 12 cells) from a bipolar cell that was exposed to FM1-43 in the absence of Ca²⁺. Note that only the plasma membrane was stained with the fluorescent dye. Activity dependent staining of synaptic vesicles was then elicited by perfusion with 50 mM KCl Fish Ringer containing Ca²⁺ (2.5 mM) and FM1-43 (Fig. 1b). This treatment depolarizes the cells to about 0 mV and triggers Ca²⁺ influx (data not shown). After 120 s, the fluorescence moved from the plasma membrane towards the center of the terminal. A profile analyses (see Material and methods) of bipolar terminals exposed to FM1-43 in the absence of Ca²⁺ shows fluorescence only in the nerve terminal periphery, which corresponds to staining of the plasma membrane (Figs 1c and e, continuous line). When Ca²⁺ was added to the perfusion medium, fluorescence was spread throughout the terminal during the 120-s perfusion period (Figs 1d and e, dashed line). This staining pattern was in good agreement with previous studies performed by Lagnado et al. (1996) and Rouze and Schwartz (1998). Indeed, Rouze and Schwartz (1998) showed that endocytosed vesicles quickly diffuse throughout the bipolar nerve terminal cytoplasm within about 70 s.

Terminals pretreated with 50 nM OA for 30 min presented a strikingly different staining pattern from that observed in control terminals. Figure 2 shows a representative cell (n = 6) that was exposed to FM1-43 before (Fig. 2a) and 120 s after (Fig. 2b) addition of 2.5 mM Ca²⁺ to the perfusion medium. There was little change in the overall fluorescence from the plasma membrane to the interior and center of the terminal in response to calcium influx (compare Fig. 2c with 2d). Only the edges of the terminal presented slightly more staining after Ca²⁺ influx (compare dashed and continuous lines in Fig. 2e). However, the spread of FM1-43 into the middle of the nerve terminal, a pattern that was so prominent under control conditions (see Fig. 1d), was not detected in any of the OA (50 nM) treated terminals examined (n = 6). OA thus had a dramatic effect on the ability of dye to spread throughout the terminal.

Fig. 2.

Okadaic acid (OA) interferes with the pattern of FM1-43 staining of bipolar cell terminals. Goldfish bipolar cells were treated with OA (50 nM) and stained with the dye FM1-43. (a) Fluorescence image of a bipolar cell after perfusion with Fish Ringer containing 50 mM KCl, 8 μM FM1-43 and 0 mM Ca²⁺. Note the fluorescence only at the plasma membrane. Arrow points to bipolar nerve terminal. (b) Fluorescence image obtained after perfusion with Fish Ringer containing 50 mM KCl, 8 μM FM1-43 and 2.5 mM Ca²⁺. Note that the fluorescence did not spread to the center of the terminal during the depolarization in the presence of calcium. The arrow points to the bipolar nerve terminal. (c) Expanded view of the same terminal as shown in Fig. 2(a). (d) Same terminal as shown in Fig. 2(b). (e) Fluorescence intensity profile. The dashed line and continuous line represent the fluorescence intensity profile for (c) and (d), respectively.

Several bipolar cells were pretreated with OA at different concentrations (0.1–50 nM), stained with FM1-43 and imaged. We divided bipolar terminals in three regions according to the distance from the membrane edges: periphery (0–2 μm and 8–10 μm) and central zone (4–6 μm). At concentrations above 1.0 nM OA [1.0 nM (n = 4 cells), 5.0 nM (n = 3 cells), 25 nM (n = 7 cells), 50 nM (n = 6 cells), OA interfered with the spread of FM1-43 to the center of the terminals and caused fluorescence to accumulate near the plasma membrane (Figs 3b–e). At a lower concentration (0.1 nM, n = 4, Fig. 3a), bipolar cells displayed an FM1-43 staining pattern similar to that obtained in control nerve terminals (i.e. fluorescence was present in the middle of the terminal).

Capacitance measurements indicate that OA does not affect synaptic vesicle exocytosis or endocytosis

One interpretation for the above results is that synaptic vesicle fusion or retrieval is blocked. To determine the effect of OA on synaptic vesicle exoendocytosis, bipolar cell terminals were treated with 25 nM OA for 30 min and capacitance measurements were performed. Terminals were voltage-clamped in the whole-cell mode and subjected to several 200-ms, 1-s or 5-s depolarizations to 0 mV. This triggered Ca²⁺-mediated exocytosis and subsequent endocytosis. In addition, the patch pipette contained 50 nM OA to avoid any possible washout of the effects of OA via whole-cell dialysis of the terminals. Capacitance measurements showed that OA treatment did not affect the rate of endocytosis (Fig. 4, Table 1). For both OA treated and control terminals, 200 ms depolarizations were followed immediately by endocytosis which proceeded with an average time constant of about 2 s (Figs 4a and c, Table 1). These time constants are in close agreement with previously reported rates of endocytosis following 200 ms depolarizations in the goldfish bipolar cell terminals (von Gersdorff and Matthews 1997). Capacitance jumps and endocytosis in control and OA treated cells could be elicited several times within the first few minutes after patch pipette break-in. Because FM dye measurements of endocytosis were performed following longer potassium induced depolarizations, we also tested 1 and 5-s depolarizing pulses. Again, OA-treated terminals exhibited the same rates of endocytosis as untreated terminals (Figs 4b and d, Table 1). Following 1- and 5-s depolarizations, fast endocytosis was delayed in both control and OA-treated terminals (Fig. 4f), possibly due to inhibition of endocytosis by elevated calcium levels and/or continued exocytosis (von Gersdorff and Matthews 1997; Rouze and Schwartz 1998; Neves and Lagnado 1999; Heidelberger 2001). This delay before onset of fast endocytosis after 1- and 5-s depolarizations was unchanged by OA treatment (see Table 1). These cells also exhibited undelayed, fast endocytosis after 200-ms pulses when 1- or 5-s pulses were alternated with 200 ms pulses (data not shown). The same average rate of fast endocytosis for all pulse durations (Fig. 4f) was observed in control and OA-treated cells. Moreover, OA treatment did not significantly affect the amplitude or kinetics of the calcium current at the bipolar cell terminal (Fig. 4e and Table 1). If anything, the amplitude of the Ca²⁺ currents was slightly larger on average with OA pretreatment, but this was a small effect (two-sample t-test p > 0.05). Depolarization of Fluo-3-AM labeled bipolar cells (n = 8 for control and n = 5 for OA-treated cells) in the presence and absence of OA led to equal changes in intracellular Ca²⁺ in both conditions (not shown). Thus Ca²⁺ channels were not altered significantly by OA treatment.

Fig. 4.

Capacitance and Ca²⁺ current. OA does not affect synaptic vesicle exocytosis or endocytosis. Isolated presynaptic terminals were voltage-clamped in the whole-cell mode for capacitance recordings. OA (25 nM) was applied for 30 min prior to capacitance measurements for treated cells and 50 nM OA was included in the patch pipette. (a–d) Typical membrane capacitance (C_m) recordings (dots) from four presynaptic terminals reflecting endocytosis after two depolarizing pulses of 200 ms (a,c) and one depolarizing pulse of 5 s (b,d) for both OA treated (c,d) and control (a,b) terminals. The depolarization was from −60 mV to 0 mV. The solid lines are best-fit single exponentials. (a) and (c) were baseline corrected to account for a very slow stimulus-independent drift in the resting capacitance. Recordings were taken within the first minute after whole-cell break-in. The time scale is the same for panels (a–d). (e) Typical Ca²⁺ currents (I_Ca) and membrane capacitance jump (C_m) recorded from one OA-treated and one control terminal during a 200-ms pulse. Capacitance recordings during the sinusiodal voltage command were briefly interrupted for a depolarizing pulse to elicit Ca²⁺ influx and transmitter release (see Methods). I_Ca was recorded in this fashion for each pulse during capacitance recordings. (f) Average time constants of endocytosis following 200 ms, 1 s and 5 s depolarizing pulses from −60–0 mV for OA-treated and control terminals. Error bars reflect SEM.

Table 1.

Effect of OA treatment on exo-endocytosis

	OA (200 ms) (n = 11)	Control (200 ms) (n = 13)	OA (1 s) (n = 5)	Control (1 s) (n = 6)	OA (5 s) (n = 6)	Control (5 s) (n = 4)
τ endo (s)	2.07 (± 0.61)	2.13 (± 0.69)	4.72 (± 1.41)	4.83 (± 0.33)	8.8 (± 4.1)	9.8 (± 3.7)
Cm jump (fF)	181 (± 64)	176 (± 45)	133 (± 45)	171 (± 50)	246 (± 56)	266 (± 65)
Peak ICa2+ (pA)	324 (± 103)	294 (± 102)	278 (± 57)	244 (± 52)	259 (± 68)	191 (± 35)
Delay before fast endo (s)	0	0	5.7 (± 1.4)	5.9 (± 2.5)	14.6 (± 3.3)	10.3 (± 1.1)

OA treatment did not alter the average time constant of endocytosis, average capacitance jumps, or average calcium currents for depolarizing pulses of 200 ms, 1 s or 5 s. Values are reported as average ± SD. There was no statistical difference at any pulse duration between OA treated and control terminals for these parameters (Student’s t-test; p < 0.05). Data reflect the fastest capacitance response and corresponding calcium current measured within the first two minutes after whole-cell break-in for each terminal.

Exocytosis (capacitance jumps) was also unaffected by OA treatment (Table 1), and the amount of membrane exocytosed under control and OA-treated conditions matched closely with values previously reported in this preparation (von Gersdorff and Matthews 1997). The recovery from capacitance saturation, or recovery from vesicle pool depletion, was complete within 20–30 s in control and OA treated cells. When two depolarizing pulses of 200 ms were given 20 s apart (Fig. 4a and b), the second C_m jump measured as a fraction of the first C_m jump was 0.83 ± 0.037 (mean ± SEM; n = 10) in control and 0.87 ± 0.036 (n = 8) in OA (no significant difference; unpaired t-test, p > 0.1). Typically, exocytosis and endocytosis run down in goldfish bipolar cells during the course of prolonged recordings under whole-cell dialysis. While this run down was observed during long-lasting recordings, there was no difference in the persistence of exo-endocytotic responses between OA-treated and control terminals (data not shown). Capacitance measurements thus show no detectable effect of OA on the amount or the kinetics of exocytosis and endocytosis.

OA causes FM1-43 fluorescence to accumulate near the plasma membrane

Because capacitance recordings indicated that OA does not block membrane fusion or fission, it seems improbable that dye accumulation near the plasma membrane occurs due to inhibition of exo- or endocytosis. One explanation for this result would be that dye accumulation in the periphery of the nerve terminal occurs spontaneously during repetitive perfusions with FM1-43. An alternative explanation would be that OA induces sustained phosphorylation of the recycling or mobilization machinery and impairs vesicle cycling. Therefore, after stimulation, this prolonged phosphorylation would prevent the movement of FM1-43 stained vesicles towards the interior of the terminal. To address these issues bipolar cells were exposed to FM1-43 in Fish Ringer with KCl (50 mM) and 0 mM Ca²⁺. After 120 s, an image of the terminal was obtained (image 1, Fig. 5a–c) and the same cell was submitted to another round of perfusion with this solution followed by re-imaging after 120 s (image 2, Fig. 5d–f). Subtracting image 2 from image 1 indicates whether dye continues to accumulate near the plasma membrane in the absence of endocytosis, i.e. during a condition where dye is not supposed to be internalized. Figure 5(g) (first bar) shows that there was a small increase in the number of fluorescent pixels at the periphery of the cell (approximately 500 pixels) during repetitive perfusions with Fish Ringer with FM1-43, 50 mM KCl and 0 mM Ca²⁺. A similar increase in fluorescence was detected when calcium was replaced by 2.5 mM Mg²⁺ (approximately 600 pixels, Fig. 5g, third bar) to keep the divalent ion concentration constant. However, OA-treated terminals showed a threefold increase in fluorescent area (Fig. 5g, second bar) when compared to these control experiments (Ca²⁺ 0 mM or total Mg²⁺ 2.5 mM). This observation support the idea that in OA treated bipolar cell terminals (Fig. 5g) vesicle internalizes FM1-43 but remain confined near the plasma membrane.

OA interferes with synaptic vesicles movement within bipolar cell terminals during sustained depolarization

We next examined whether a longer period of calcium influx (i.e. 360 s instead of 120 s) would allow vesicles to move from the periphery to the center of OA-treated terminals (Fig. 6). Again, control terminals and OA-treated terminals presented a distinct fluorescence line profile (compare Fig. 4a and b for a typical pair of cells). These results were quantified in Fig. 4(c and d) for several cells. In control bipolar cell terminals, the fluorescence measured at the central zone (Fig. 4c, second bars) increased from 23.079 ± 9 (mean ± SEM) to 183.25 ± 26 gray levels, after 360 s of perfusion with High KCl Fish Ringer with 2.5 mM Ca²⁺ and FM1-43. In contrast, when the same procedure was performed in OA treated nerve terminals, the pixel intensity at the central zone changed from 20.43 ± 4 (mean ± SEM) to 79.76 ± 10 gray levels (Fig. 4d). Thus, the fluorescence increase at the center in control terminals was almost threefold the value obtained for OA-treated nerve terminals. This result suggests that prolonged calcium influx could not prevent or overcome the effect of OA on vesicle mobilization and that vesicles had their movement restricted even for longer periods of time.

Discussion

Little is known about how synaptic vesicles are mobilized towards the plasma membrane following a bout of exocytosis, although this process seems to be regulated by Ca²⁺ and PKC (reviewed by Neher 1998). Even less is known about what directs vesicle traffic following endocytosis or membrane retrieval (Koenig and Ikeda 1996; Cochilla et al. 1999; Li and Murthy 2001). Here we have identified one step of the synaptic vesicle recycling pathway, namely the trafficking of vesicles away from the plasma membrane after endocytosis, that is sensitive to very low doses of OA. In addition, the finding that vesicles can still move towards the membrane for release in the presence of OA implies that the trafficking of vesicles towards the plasma membrane is controlled by different mechanisms from those that control trafficking away from the plasma membrane. We have thus ‘broken up’ the vesicle recycling process into two distinct steps: an OA-insensitive step (mobilization towards the plasma membrane), and an OA-sensitive step, that returns vesicles to the center of nerve terminals.

Serine/threonine protein phosphatases can be classified into protein phosphatase 1 (PP1) and PP2 (with subtypes PP2 A, B and C), and PP4, PP5, PP6 and PP7 (reviewed by Herzig and Neuman 2000). Analyses of phosphatase tissue distribution shows that PP1 and PP2 are particularly rich in neurons. Although there are several results that relate protein phosphorylation and dephosphorylation to plasticity and endocytosis in conventional synapses (reviewed by Llinás 1999; Cremona and De Camilli 2001), little is known about this regulation in ribbon-type synapses.

Okadaic acid is a polyether fatty acid first isolated from the black marine sponge Hallicondria okadaii, but is also produced by several types of dinoflagellates (marine plankton). It is a major toxic component associated with diarrhetic seafood poisoning (Cohen et al. 1990) due to its accumulation in the midgut of mussels, clams and scallops feeding on dinoflagellates. It is a potent phosphatase inhibitor of PP1 and PP2A with half maximal inhibition ranging from 50 to 300 nM for PP1 and 0.1–2 nM for PP2A (Bialojan and Takai 1988; Herzig and Neuman 2000). It also inhibits PP2B to a lesser extent (concentrations higher than 1 μM). In our experiments, bipolar cell nerve terminals exposed to OA (1–50 nM) had a distinct FM1-43 staining pattern when compared to untreated terminals. Because very low doses of OA (1 or 5 nM) interfered with vesicle recycling, it is likely that PP2A plays an important role in regulating vesicle trafficking following endocytosis at this ribbon-type synapse.

Phosphatase inhibitors such as OA induce changes in the activity of many intracellular targets, including ion channels. For example, studies performed in central and peripheral nervous system neurons suggest that inhibition of phosphatases by OA may increase Ca²⁺ current amplitude through high-voltage activated calcium channels (Herzig and Neuman 2000). To test whether OA interfered with Ca²⁺ channel function of bipolar cells, we measured Ca²⁺ currents electrophysiologically and intracellular Ca²⁺ concentration changes with the fluorescent dye Fluo-3 (data not shown). Using these two independent techniques, we detected no difference in Ca²⁺ channel function between control or OA-treated terminals (Fig. 4e for Ca²⁺ current). Therefore, altered Ca²⁺ channel function is not responsible for the effect of OA.

When phosphatase activity is impaired by OA, fluorescence accumulates only at the edges of the bipolar cell terminal. This staining pattern is in clear contrast to the diffusely stained control terminals (compare Figs 1 and 2; see also Fig. 5a–c from Rouze and Schwartz 1998) and suggests that OA may disrupt either exocytosis or endocytosis, or both. Trying to release the fluorescence accumulated in OA-treated terminals would be the most direct way to test this paradigm. Lagnado et al. (1996) were able to follow destaining of bipolar terminals that were previously stained with FM1-43. However, Rouze and Schwartz (1998) were not able to see destaining of bipolar terminals during depolarization with KCl. One explanation is that vesicles stained with FM1-43 during depolarization (10 000–20 000 vesicles; see Rouze and Schwartz 1998) mix with almost one million non-labeled vesicles that are distributed in bipolar terminals (von Gersdorff et al. 1996). Therefore, to address if OA interfered with synaptic fusion and/or retrieval we then performed membrane capacitance measurements to independently track synaptic vesicle exocytosis and endocytosis. Capacitance measurements revealed that these two processes are unaffected by OA treatment (Fig. 4 and Table 1). These experiments show that this phosphatase inhibitor did not interfere with exo-endocytosis, suggesting that PP2A does not modulate these processes in bipolar cell terminals. This hypothesis is also supported by the observation that stimulation-dependent dephosphorylation of proteins involved in endocytosis is mediated by another phosphatase: calcineurin (phosphatase 2B; Liu et al. 1994; Bauerfeind et al. 1997; Lai et al. 1999). Indeed, if OA blocked exocytosis we would expect labeling with FM1-43 to be similar to that observed in terminals depolarized in the absence of Ca²⁺. This was clearly not the case (Fig. 5). Thus, we favor the hypothesis that movement of freshly endocytosed vesicles towards the interior of the terminal is inhibited by OA.

It is interesting to compare our results with those obtained by Betz and Henkel (1994) in the frog NMJ. Studying terminals labeled with FM1-43 and treated with OA (1–5 μM for 30 min) they observed a greatly facilitated ‘lateral movement’ of vesicles between active zones and this enhanced mobility disrupted the formation of vesicle clusters near active zones. However, the concentrations of OA used (1–5 μM) did not permit a specification of the particular phosphatase type(s) involved in this process. EM micrographs of goldfish bipolar cell nerve terminals show that ribbons have a halo of about 110 tethered vesicles, but the ribbon complex itself is not surrounded by a tight cluster of vesicles organized in similar manner as the active zones of the frog NMJ. Bipolar cell terminals have a spherical shape when isolated and contain a large number of synaptic vesicles (480 000–910 000), the vast majority of which are uniformly distributed throughout the cytoplasm. In fact, only a small fraction of the total vesicle population is found attached to the ribbons (< 1%; von Gersdorff et al. 1996) in isolated bipolar terminals. This makes it very difficult to identify clusters of FM1-43 labeled synaptic vesicles in this preparation with confocal microscopy (see, however, Zenisek et al. 2000). Our results obtained with OA-treated bipolar cells thus contrast with those of Betz and Henkel (1994) since we observed an inhibition of vesicle mobility from the plasma membrane towards the center of the synaptic terminal. Our finding that dye accumulates near the edges of the plasma membrane in the presence of OA indicates that phosphatases participate in vesicle translocation at ribbon-type synapses but in a very different manner from their role in conventional CNS (Kraszewski et al. 1996) and peripheral synapses (Betz and Henkel 1994). Interestingly, Li and Murthy (2001) have also shown recently that staurosporin, a kinase inhibitor, affects postendocytic vesicle movement.

An alternative interpretation for our results is that during OA treatment some vesicles were unable to internalize the dye because endocytosis became extremely fast (Pyle et al. 2000) or switched to a ‘kiss-and-run’ mode of exo-endocytosis (Stevens and Williams 2000). Such interpretation would require that a significant amount of vesicles were unlabeled and competed with the labeled ones, thus decreasing the fluorescence in the terminal. We feel this to be very unlikely because these invisible (or unlabeled) vesicles would have also to be undetected by the capacitance measurements, which were unaltered by OA treatment. Therefore, if a significant amount of endocytosis shifted to an extremely fast ‘kiss-and-run’ mode during the depolarization of the terminal (with OA treatment) then we should have recorded a decrease in the capacitance jump. Alternatively, if endocytosis becomes accelerated after the depolarization of the terminal (with OA treatment) then the decay rate of the capacitance back to baseline should have sped up. Neither of these effects was observed with OA treatment, even when the OA concentration was as high as 50 nM inside and outside the patch pipette.

Given that OA seems to impair postendocytic vesicle movement it is interesting that OA did not affect the terminal’s ability to engage in repeated bouts of exocytosis (see Table 1). One possibility is that the large reservoir of vesicles in the bipolar cell terminal provides a safety margin that allows these cells to function even if part of their synaptic vesicle recycling pathway is severely compromised. In conclusion, our studies indicate that phosphorylation via PP2A plays an important role in regulating the mobility and transport of recently internalized vesicles within ribbon-containing synaptic terminals. Furthermore, they show that recruitment of vesicles towards the plasma membrane for exocytosis is regulated by a different mechanism from that used to regulate translocation of vesicles away from the plasma membrane after endocytosis. These two legs of the synaptic vesicle cycle are thus differentially regulated.

Abbreviations used

NMJ

neuromuscular junction

OA

okadaic acid

PP

phosphatase