HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

This Article Abstract Full Text (PDF) All Versions of this Article: 542/2/395    most recent 2002.021733v1 Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Ng, Y.-K. Articles by Levitan, E. S. Search for Related Content PubMed PubMed Citation Articles by Ng, Y.-K. Articles by Levitan, E. S.

Journal of Physiology (2002), 542.2, pp. 395-402
© Copyright 2002 The Physiological Society
DOI: 10.1113/jphysiol.2002.021733

Physical mobilization of secretory vesicles facilitates neuropeptide release by nerve growth factor-differentiated PC12 cells

Yuen-Keng Ng, Xinghua Lu and Edwin S. Levitan

	ABSTRACT

Top Abstract Introduction Methods Results Discussion References

It has been speculated that neurosecretion can be enhanced by increasing the motion, and hence, the availability of cytoplasmic secretory vesicles. However, facilitator-induced physical mobilization of secretory vesicles has not been observed directly in living cells, and recent experimental results call this hypothesis into question. Here, high resolution green fluorescent protein (GFP)-based measurements in nerve growth factor-differentiated PC12 cells are used to test whether altering dense core vesicle (DCV) motion affects neuropeptide release. Experiments with mycalolide B and jasplakinolide demonstrate that neuropeptidergic DCV motion at the ends of processes is proportional to F-actin. Furthermore, Ba²⁺ increases DCV mobility without detectably modifying F-actin. Finally, we show that altering DCV motion by changing F-actin or stimulating with Ba²⁺ proportionally changes sustained neuropeptide release. Therefore, increasing DCV mobility facilitates prolonged neuropeptide release.

(Resubmitted 2 April 2002; accepted after revision 14 May 2002)
Corresponding author E. S. Levitan: Department of Pharmacology, E1351 Biomedical Science Tower, University of Pittsburgh, PA 15261, USA. Email: levitan{at}server.pharm.pitt.edu

	INTRODUCTION

Top Abstract Introduction Methods Results Discussion References

It has been speculated that facilitation of release is produced by increasing secretory vesicle mobility. For example, restriction of dense core vesicle (DCV) movement has been proposed to be relevant for F-actin's effect on the initial rate, but not the total extent, of exocytosis by endocrine cells (Vitale et al. 1995; Chowdhury et al. 1999). Likewise, phosphorylation-dependent tethering of small synaptic vesicles via synapsin to F-actin has been suggested to control fast synaptic transmission, although synapsin also affects the release process itself (Hilfiker et al. 1998, 1999). Yet, while the link between F-actin and release is firmly established, live cell imaging of DCVs has not revealed a clear role for increasing DCV motion in enhancing release. Specifically, recent microscopy experiments have found that depolymerizing F-actin reduces DCV movement near the plasma membrane in chromaffin cells and undifferentiated PC12 cells (Lang et al. 2000; Oheim & Stuhmer, 2000) in contradiction to the proposal that F-actin restricts endocrine DCV movement to limit hormone release. Furthermore, depolymerizing F-actin increases fast transmission at central synapses by modifying exocytosis of readily releasable small synaptic vesicles (Morales et al. 2000). Because such vesicles are already docked to active zones, this finding, as well as the endocrine experiments described above, is consistent with known F-actin interactions with phosphoinositides and proteins involved in exocytosis (e.g. Hilfiker et al. 1998; Raucher et al. 2000) instead of a change in cytoplasmic vesicle movement. In addition, increasing small synaptic vesicle motion can lead to decreased synaptic transmission (Betz & Henkel, 1994). Thus, physical mobilization of small synaptic vesicles need not facilitate release. Also, it should be noted that stimulated appearance of secretory vesicles at sites of release could be due to increased capture and utilization of cytoplasmic vesicles rather than increased movement (Zenisek et al. 2000). Given that facilitation by increased vesicle mobility has not been demonstrated directly, recent experimental results indicate that fast neurotransmission and endocrine hormone release may be augmented without increasing vesicle movement.

Yet, controlling vesicle mobility may be very important for neuropeptide release. The cell biology of neuropeptide release is unique; unlike classical transmitters, neuropeptides are released without using active zones, and unlike endocrine hormone release, neuropeptide secretion is supported by DCVs at the ends of processes. Also, unlike small synaptic vesicles and endocrine secretory vesicles, very few neuropeptidergic DCVs are initially docked in native nerve terminals (e.g. Leenders et al. 1999; Karhunen et al. 2001). Live cell imaging suggests that neuropeptide release is accomplished in large part via efficient recruitment of mobile DCVs (Burke et al. 1997; Han et al. 1999b). Yet, the mobile pool is limited (Burke et al. 1997; Han et al. 1999b). Hence, it is conceivable that neuropeptide release is facilitated by increasing DCV motion.

To prove that physical mobilization is important, it is necessary to show that altering DCV dynamics affects neuropeptide release. To date, actin polymerization drugs have been found to produce subtle changes in immature (Rudolf et al. 2001) and mature (Lang et al. 2000; Oheim & Stuhmer, 2000) DCV motion in undifferentiated PC12 cells and chromaffin cells. Those studies may not apply to neuropeptide release because neuronal differentiation alters DCV distribution and dynamics (Ng et al. 2002) as well as cell structure. Furthermore, the effect of changing mobility has not been separated from other consequences of altering F-actin. Therefore, mechanistically diverse treatments for changing DCV motion at the ends of neuronal processes where neuropeptide release occurs are needed to test whether functional mobilization (i.e. enhanced use) of neuropeptidergic DCVs is produced by physical mobilization (i.e. increased DCV motion). Here, GFP-based single particle tracking and release measurements at the ends of processes of neuronally differentiated PC12 cells (Burke et al. 1997; Kaether et al. 1997; Abney et al. 1999; Han et al. 1999b) are employed to demonstrate that stimulating DCV motion by two independent mechanisms facilitates sustained neuropeptide release.

	METHODS

Top Abstract Introduction Methods Results Discussion References

PC12 cells transfected with the GFP-tagged neuropeptide were cultured and neuronally differentiated with 50 ng ml^-1 nerve growth factor on polylysine-coated coverslips, and then imaged as previously described (Burke et al. 1997; Levitan, 1998; Han et al. 1999a,b). Wide field epifluorescence microscopy was performed with a Nikon Diaphot microscope equipped with a 60 1.4 numerical aperture oil immersion objective at 23 °C with an extracellular saline that contained (in mM): 140 NaCl, 5.4 KCl, 5 CaCl₂, 0.8 MgCl₂, 10 glucose, 10 NaHepes, pH 7.5. Stock solutions for mycalolide B and jasplakinolide were made with dimethylsulfoxide. Matched controls contained an equivalent amount of vehicle. Release was evoked by superfusion with a solution in which 100 mM Na⁺ was replaced with K⁺, or by replacement of 100 mM Na⁺ and all Ca²⁺ with K⁺ and Ba²⁺, respectively. Thus, depolarization was continual. Release from PC12 cells was inhibited with N-ethylmaleimide (NEM; Sigma, St Louis, MO, USA) as described previously (Han et al. 1999a).

To limit the number of labelled secretory granules so that movement of individuals could be discerned, an ecdysone-inducible construct of emerald GFP-tagged proANF was transfected into cells along with pVgRXR (Invitrogen). However, to improve the signal over our past experiments with an inducible construct (Han et al. 1999b), emerald GFP-tagged preproANF was subcloned into pEGSH (Stratagene) instead of the Invitrogen expression vector. 1.5 µM muristerone A treatment was initiated 4 h before imaging. Notably, 2 h inductions led to production of a large number of labelled DCVs in the cell body. But these potentially immature DCVs were not transported to the ends of processes. The use of an induction period much longer than the DCV maturation time (Tooze et al. 1991), the apparent ability of cells to select mature DCVs for shipment to release sites and the findings that vesicles induced with this method have release kinetics (Han et al. 1999a) and mobility properties (Y.-K. Ng, W. Han & E. S. Levitan, unpublished results) that are indistinguishable from steady state-labelled vesicles all suggest that functionally mature DCVs were studied.

For analysis of individual DCVs in cultured cells, Inovision (Raleigh, NC, USA) single particle tracking software determined vesicle trajectories based on 10 images acquired at 0.5 Hz as we have shown previously (Burke et al. 1997; Han et al. 1999b). Step sizes were then used to calculate two dimensional diffusion coefficients (Ds) based on the relationship: D = r²/4t, with r being the distance travelled in a time period t. Previous studies have demonstrated that DCV movement conforms to diffusion theory in this preparation; for example, r² vs. time plots are linear (Abney et al. 1999; Han et al. 1999b). Controls with paraformaldehyde-fixed cells demonstrated that the slowest vesicles studied here moved at least an order of magnitude faster than the limit of resolution for the setup. The numbers of measurements referred to in the figures are the number of independently tracked vesicles. The number of cells studied to acquire these trajectories are given in the figure legends.

Release was measured as the decrease in GFP-tagged neuropeptide fluorescence from the ends of processes in accordance with our previous studies (Burke et al. 1997; Levitan, 1998; Han et al. 1999a,b). These measurements utilize the fact that wide field microscopy samples in focus and out of focus fluorescence from an optical section that is much greater in depth than the processes studied here (Levitan, 1998). Furthermore, the validity of this optical method has been verified with biochemical measurements (Burke et al. 1997; Kaether et al. 1997; Han et al. 1999a). Attention was focused on the ends of processes because this is the major site for GFP-tagged neuropeptide accumulation and release in this preparation (e.g. Burke et al. 1997). Furthermore, because whole cells were imaged, it is evident that no significant retrograde transport occurs basally or in response to stimulation. Finally, while we previously reported a Ca²⁺-stimulated brightening of enhanced GFP (EGFP) inside DCVs due to alkalinization (Han et al. 1999a), this effect is not seen with the emerald GFP-based construct used here (data not shown).

To illustrate changes in movement, three successive images were pseudo-coloured red, green and blue, and then superimposed similarly to Fig. 5 in Han et al. (1999b). If a vesicle is stationary, then the three differently coloured images overlap to generate a white spot. However, a quickly moving vesicle will produce nonoverlapping images to yield multicoloured spot(s).

To visualize F-actin, cells were fixed with 4 % paraformaldehyde, permeabilized with 0.1 % Triton X-100, incubated with 33 nM Texas Red-X phalloidin (Molecular Probes), washed with PBS twice, and then viewed with rhodamine optics either by standard wide field epifluorescence microscopy as described above or with a Leica TCS NT confocal microscope with a 100 1.4 numerical aperture oil immersion objective.

Experimental data were collected from matched cultures to take into account batch-to-batch variations. Statistical significance was measured with Student's t test in experiments with only two conditions. Error bars show standard error of the mean.

	RESULTS

Top Abstract Introduction Methods Results Discussion References

Basal control of neuropeptidergic secretory vesicle motion

As a first step toward testing whether physical mobilization of DCVs affects neuropeptide release, we sought to experimentally manipulate peptidergic DCV motion by perturbing the abundant actin cytoskeleton at the ends of processes where neuropeptidergic DCVs accumulate and undergo exocytosis. Texas Red-X phalloidin labelling of polymerized actin revealed that mycalolide B (Saito et al. 1994) effectively eliminates F-actin at the ends of processes after neuronal differentiation (Fig. 1A). To test whether this depolymerization of actin microfilaments affects DCV motion, a limited number of DCVs were labelled with a GFP-tagged neuropeptide and their motion was monitored with time lapse imaging. As can be seen in the top of Fig. 1B, vehicle treatment does not affect DCV movement (i.e. white vesicles do not change into coloured vesicles). However, F-actin depolymerization enhances the motion of individual DCVs (Fig. 1B, bottom). Analysis of trajectories deduced by single particle tracking revealed that the mean diffusion coefficient for DCVs increases 6-fold in response to mycalolide B (Fig. 1C).

		View larger version [in this window] [in a new window]
Figure 1. Actin depolymerization increases DCV motion A, wide field epifluorescence images of Texas Red-X phalloidin labelling of F-actin. Left, treated with vehicle. Right, treated with 2 µM mycalolide B for 30 min at room temperature. Scale bars indicate 2 µm. B, top, peptidergic vesicles remain nearly immobile for long periods in the presence of vehicle. Three images were acquired every 6 s and RGB colour-coded red, green and blue, respectively, as described in the Methods. With this scheme, stationary vesicles appear white and moving vesicles produce multicoloured spots. Bottom, DCV motion increases after actin depolymerization. Note that mycalolide B treatment stimulates secretory vesicle movement as indicated by the replacement of white vesicle images with coloured vesicle images. Scale bars indicate 2 µm. C, diffusion coefficients of secretory vesicles before (open bars) and after 15 min of treatment (black bars) with vehicle (four cells) or mycalolide B (MB, five cells). Number of vesicle trajectories shown in parentheses. *** P < 0.0005.

If F-actin depolymerization increases DCV motion, then promoting actin polymerization should have an opposite action. Therefore, the effect of jasplakinolide, a membrane permeant compound that acts within minutes to stabilize existing actin microfilaments (Bubb et al. 1994; Cramer, 1999), was examined. We could not assess the impact of jasplakinolide on F-actin because this drug competes with phalloidin for binding (Bubb et al. 1994). Nevertheless, after 10 min of application, jasplakinolide suppresses DCV motion (Fig. 2A). Specifically, the average diffusion coefficient is reduced 4-fold (Fig. 2B). Thus, opposite effects on DCV motion can be produced with mycalolide B and jasplakinolide.

		View larger version [in this window] [in a new window]
Figure 2. Jasplakinolide reduces DCV motion A, sets of images acquired every 6 s were RGB colour-coded as in Fig. 1. Note that DCV movement decreases after exposure to 10 µM jasplakinolide for 10 min as indicated by the increase in white vesicle images in the right panel. Scale bar indicates 2 µm. B, jasplakinolide (Jpk) decreases DCV diffusion coefficients. Number of vesicle trajectories for each measurement shown in parentheses. Data was collected from four cells. *** P < 0.0001.

Secretory vesicle dynamics controls sustained neuropeptide release

As noted in the introduction, changing F-actin increases the initial rate of release by endocrine cells and hippocampal neurons. Our finding that neuropeptidergic DCV motion can be increased and decreased by depolymerizing and stabilizing F-actin, respectively, has not been observed directly in the aforementioned preparations. Therefore, we analysed how these changes in DCV mobility affect the kinetics of neuropeptide release. Figure 3A shows that depolarization-evoked peptide release, measured as a decrease in normalized initial fluorescence (F₀), is altered by the mycalolide B treatment that promotes vesicle motion. Unlike the results mentioned above, the initial rate of release is not altered. Instead, sustained release seen after the first time point is increased by this treatment. A similar change is seen with release evoked by a Ca²⁺ ionophore (data not shown), indicating that a change in channel activity is not involved. Consistent with this finding, a separate set of experiments that were performed pairwise to control for batch-to-batch variation demonstrated that sustained release is almost completely inhibited by reducing DCV mobility with jasplakinolide (Fig. 3B). Again, this effect is not evident for the initial rate of release. Therefore, changing DCV motion proportionally alters sustained neuropeptide release.

		View larger version [in this window] [in a new window]
Figure 3. F-actin limits sustained neuropeptide release Peptide content at the ends of processes is normalized to initial fluorescence (100 % F0) so that release is evident as a decrease in this quantity. A, F-actin depolymerization with mycalolide B () enhances sustained neuropeptide release evoked by continual depolarization (). n > 10. B, promoting actin polymerization with jasplakinolide () inhibits sustained neuropeptide release evoked by continual depolarization (). n > 10.

Liberation of secretory vesicles by a release facilitator

The mycalolide B and jasplakinolide data indicate that the state of actin polymerization can greatly affect the extent of vesicle mobility. To explore whether this actin reorganization is necessary for large changes in vesicle mobility, Ba²⁺ was employed. Ba²⁺ supports neuropeptide release, possibly more effectively than Ca²⁺ (Sachs et al. 1967; Wayne et al. 1998). Furthermore, this ion facilitates neurotransmission (Zengel & Magleby, 1977, 1980, 1981). Also, Ba²⁺ is not efficiently buffered and so accumulates to greater levels than Ca²⁺, especially in cells with few Ca²⁺ channels.

Figure 4A shows that depolarization-evoked neuropeptide release is greater in the presence of Ba²⁺ than with Ca²⁺. Particularly striking is the fact that this difference is not apparent in the initial rate of release. Rather, sustained release is markedly enhanced. Because this feature was also found with mycalolide B, we examined whether actin depolymerization facilitates release evoked by Ba²⁺. If Ba²⁺-induced facilitation of release does not involve vesicles that interact with F-actin, then its release should be augmented by the facilitation produced by mycalolide B. However, even though Ba²⁺-induced release is not complete, no significant enhancement is produced by F-actin depolymerization (Fig. 4B). A connection between actin-associated DCVs and facilitated release is further supported by experiments with jasplakinolide because this F-actin stabilizing drug inhibits Ba²⁺-evoked neuropeptide release (Fig. 4C). Hence, it appears that Ba²⁺ may access F-actin-sensitive secretory vesicles to enhance neuropeptide secretion.

		View larger version [in this window] [in a new window]
Figure 4. Facilitation of neuropeptide release by Ba2+ A, continual depolarization in the presence Ba2+ () causes more neuropeptide release than in the presence of Ca2+ (). Note that while the initial rate of release is comparable, sustained release is enhanced by depolarization with Ba2+. n > 8. B, 2 µM mycalolide B () does not facilitate Ba2+-evoked release (). n > 12. C, 10 µM jasplakinolide () inhibits Ba2+-facilitated release (). n > 8.

These results might be consistent with the hypothesis that Ba²⁺, like mycalolide B, depolymerizes F-actin to promote vesicle motion and sustained release. This possibility was examined by comparing the abundance of F-actin in control and Ba²⁺-treated cells. Contrary to results with mycalolide B, wide field epifluorescence (data not shown) and confocal detection of Texas Red-X phalloidin indicate that F-actin is abundant after Ba²⁺ stimulation (n = 5) (Fig. 5). We cannot exclude that a change in F-actin was obscured by the fixation necessary for this assay. However, this seems unlikely in view of the detection of depolymerization in response to mycalolide B. Furthermore, the paraformaldehyde solution did not include detergent so that intracellular Ba²⁺ could be retained during fixation. Therefore, the simplest interpretation of the data is that extensive actin depolymerization is not required for facilitation of neuropeptide release.

		View larger version [in this window] [in a new window]
Figure 5. F-actin is unaffected by Ba2+ Confocal images of representative processes that were fixed and stained for F-actin after exposure to control saline or depolarized in the presence of Ba2+. Left panels show Texas Red-X phalloidin labelling of F-actin while right panels show GFP fluorescence from DCVs. Scale bars indicate 2µm.

A potential explanation for these results is that the facilitator lowers the affinity of DCV tethers for F-actin. Such a mechanism would be consistent with enhanced release by actin depolymerization since the tethers could not function in the absence of F-actin. Furthermore, induction of actin polymerization would provide more F-actin for tethering, and hence, inhibit facilitation. This model predicts that Ba²⁺ should increase DCV motion.

Detecting such an effect induced by a strong secretagogue is complicated by the fact that rapidly diffusing DCVs are preferentially depleted to support neuropeptide release (Han et al. 1999b). Hence, the very same DCVs that are mobilized would be depleted by the release process. Therefore, the effect of Ba²⁺ on DCV translocation was examined after inhibiting exocytosis with NEM. As can be seen in Fig. 6A, depolarization in the presence of Ba²⁺ promotes DCV motion. This implies that mobilization is not an indirect consequence of release. Particle tracking measurements show that the mean DCV diffusion coefficient is increased 4-fold by Ba²⁺ (Fig. 6B). In contrast, no liberation is produced by incubation with saline. Most importantly, this result implies that physical mobilization of DCVs, with or without extensive F-actin depolymerization, enhances sustained neuropeptide release.

		View larger version [in this window] [in a new window]
Figure 6. Ba2+ increases secretory vesicle motion A, sets of images acquired over 6 s in NEM-treated cells either before (left) or after depolarization with Ba2+ for 15 min (right) were RGB colour-coded. Scale bar indicates 2 µm. B, change in DCV diffusion coefficients in NEM-treated cells that were exposed for 15 min to control saline (six cells) or depolarized in the presence of Ba2+ (eight cells). Number of vesicle trajectories is shown in parentheses. *** P < 0.0001.

	DISCUSSION

Top Abstract Introduction Methods Results Discussion References

The goal of this study was to investigate whether changes in DCV motion affect neuropeptide release. The hypothesis that functional mobilization of secretory vesicles can be produced by physical mobilization is appealing because it provides a simple mechanism for promoting availability of secretory vesicles at sites of release. Yet proving this hypothesis has been hindered historically by the inability to follow secretory vesicle movement in living cells. Thus, the speculation that physical mobilization of vesicles is important for facilitation has been based on indirect experiments that can now be explained by alternative mechanisms (see Introduction). However, direct imaging of GFP-labelled neuropeptidergic DCVs led to the detection of a large fraction of immobile cytoplasmic reserve DCVs at the ends of neuronal processes (Burke et al. 1997; Han et al. 1999b). This suggested that physical mobilization could have a significant effect on neuropeptide release. Hence, we examined the effect of altering DCV motion from neuronal processes to demonstrate the dependence of neuropeptide release on DCV dynamics. Specifically, we perturbed an endogenous mechanism that hinders DCVs near sites of exocytosis, and then demonstrated parallel effects on DCV motion at the ends of processes and sustained neuropeptide release. Second, a facilitator of sustained release was identified and shown to act by a distinct mechanism to promote DCV movement. Therefore, the simplest interpretation of the experimental results is that sustained neuropeptide release is facilitated specifically by enhancing DCV motion.

These results raise the issue of whether this process is important for physiological regulation of neuropeptide secretion that is significant for controlling behaviour. In this study, DCV mobility was altered with the use of pharmacological treatments. We also examined the effect of depolarization in the presence of NEM. Although Ca²⁺ influx occurs under such conditions (Han et al. 1999a), no change in DCV motion was detected (X. Lu & E. S. Levitan, unpublished results). Furthermore, facilitation of release, which is indicative of DCV mobilization, was not observed by prolonged dialysis with an elevated Ca²⁺ solution, application of a Ca²⁺ ionophore or by photolyzing caged Ca²⁺ (unpublished results). These findings could reflect that Ca²⁺ cannot enhance DCV dynamics in PC12 cells (i.e. Ca²⁺ is not a physiological DCV mobility enhancer in this system), or that this preparation requires an intracellular concentration that is attained more easily with Ba²⁺, which is not efficiently buffered. The latter conclusion is supported by experiments with transgenic Drosophila that express the same GFP-tagged neuropeptide used in this study (Rao et al. 2001). We have found that depolarization acts via Ca²⁺ to increase neuropeptidergic vesicle motion in larval neuromuscular junction synaptic boutons (Levitan et al. 2002). Thus, a physiological messenger controls DCV dynamics in vivo. The results presented here suggest that physical mobilization of DCVs occurs to facilitate neuropeptide release.

The difference between Ba²⁺ and Ca²⁺ evoked neuropeptide release has not been studied previously in detail. Although Ba²⁺ was known to be a very effective secretagogue, this could have been attributed to K⁺ channel block that induces depolarization that in turn opens voltage-gated Ca²⁺ channels. However, our finding that Ba²⁺ elicits more sustained release cannot be attributed to an effect on membrane potential because both ions were applied in the presence of elevated K⁺ to induce tonic depolarization. Instead, detailed analysis suggests that the increase in vesicle diffusion can fully account for the facilitation by Ba²⁺ (X. Lu, Y.-K. Ng, W. Han, M. J. Saxton, D. Axelrod & E. S. Levitan, unpublished results). Thus, the difference in Ca²⁺ and Ba²⁺ evoked neuropeptide release can be explained by dissimilar control of DCV mobility without invoking use of separate exocytotic machinery.

Our results pose the question of whether increasing secretory vesicle mobility is important for enhancing classical neurotransmission. It is notable that increasing DCV motion specifically alters sustained release. This suggests that the total releasable pool was expanded by making more DCVs available for refilling the readily releasable pool. Such a 'back filling' mechanism is unlikely to affect initial release by endocrine cells with an abundance of docked DCVs and synapses with active zones. In such preparations, reported increases in utilization of the readily releasable pool by F-actin depolymerization may reflect F-actin interactions with phosphoinositides and proteins (e.g. Hilfiker et al. 1998; Raucher et al. 2000) that influence handling of docked vesicles. However, it is conceivable that sustained release that requires continual refilling of the readily releasable pool becomes limited by the availability of mobile vesicles that must be recruited at docking sites. Indeed, physical mobilization of small synaptic vesicles may be important for the facilitating effects of depolymerizing F-actin and Ba²⁺ on amphibian neuromuscular junctions (Zengel & Magleby, 1977, 1980, 1981; Wang et al. 1996) and the Ba²⁺-induced increased in the number of recycling vesicles in hippocampal synapses (Fernandez-Alfonso et al. 2001). Yet, until facilitator-induced physical mobilization of small synaptic vesicles is detected directly, a role for changing vesicle mobility in facilitation is most compelling for neuropeptides because (a) their release can occur over long time periods, (b) few neuropeptidergic vesicles are docked initially, (c) the refilling of the readily releasable pool relies on reserve vesicles since neuropeptidergic vesicles cannot be regenerated by local endocytosis, (d) neuropeptidergic vesicles are held in reserve by immobilization, and (e) sustained release is proportional to DCV mobility.

	REFERENCES

Top Abstract Introduction Methods Results Discussion References

ABNEY, J. R., MELIZA, C. D., CUTLER, B., KINGMA, M., LOCHNER, J. E. & SCALETTAR, B. A. (1999). Real-time imaging of the dynamics of secretory granules in growth cones. Biophysical Journal 77, 2887-2895	[Abstract/Full Text]
BETZ, W. J. & HENKEL, A. W. (1994). Okadaic acid disrupts clusters of synaptic vesicles in frog motor nerve terminals. Journal of Cell Biology 124, 843-854	[Abstract]
BUBB, M. R., SENDEROWICZ, A. M., SAUSVILLE, E. A., DUNCAN, K. L. & KORN, E. D. (1994). Jasplakinolide, a cytotoxic natural product, induces actin polymerization and competitively inhibits the binding of phalloidin to F-actin. Journal of Biological Chemistry 269, 14869-14871	[Abstract]
BURKE, N., HAN, W., LI, D., TAKIMOTO, K., WATKINS, S. C. & LEVITAN, E. S. (1997). Neuronal peptide release is limited by secretory granule mobility. Neuron 19, 1095-1102	[Medline]
CHOWDHURY, H. H., POPOFF, M. R. & ZOREC, R. (1999). Actin cytoskeleton depolymerization with Clostridium spiroforme toxin enhances the secretory activity of rat melanotrophs. Journal of Physiology 521, 389-395	[Abstract/Full Text]
CRAMER, L. P. (1999). Role of actin-filament disassembly in lamellipodium protrusion in motile cells revealed using the drug jasplakinolide. Current Biology 9, 1095-1105	[Medline]
FERNANDEZ-ALFONSO, T., SANKARANARAYANAN, S. & RYAN, T. A. (2001). Barium converts nonrecycling synaptic vesicles to recycling synaptic vesicles in CNS synapses. Society for Neuroscience Abstracts 27, 386
HAN, W., LI, D., STOUT, A. K., TAKIMOTO, K. & LEVITAN, E. S. (1999a). Ca2+-induced deprotonation of peptide hormones inside secretory vesicles in preparation for release. Journal of Neuroscience 19, 900-905	[Abstract/Full Text]
HAN, W., NG, Y-K., AXELROD, D. & LEVITAN, E. S. (1999b). Neuropeptide release by efficient recruitment of diffusing cytoplasmic secretory vesicles. Proceedings of the National Academy of Sciences of the USA 96, 14577-14582	[Abstract/Full Text]
HILFIKER, S., PIERBONE, V. A., CZERNIK, A. J., KAO, H.-T., AUGUSTINE, G. J. & GREENGARD, P. (1999). Synapsins as regulators of neurotransmitter release. Philosophical Transactions of the Royal Society B 354, 269-279
HILFIKER S., SCHWEIZER, F. E., KAO, H. T., CZERNIK, A. J., GREENGARD, P. & AUGUSTINE, G. J. (1998). Two sites of action for synapsin domain E in regulating neurotransmitter release. Nature Neuroscience 1, 29-35	[Medline]
KAETHER, C., SALM, T., GLOMBIK, M., ALMERS, W. & GERDES, H. H. (1997). Targeting of green fluorescent protein to neuroendocrine secretory granules: a new tool for real time studies of regulated protein secretion. European Journal of Cell Biology 74, 133-142	[Medline]
KARHUNEN, T., VILIM, F. S., ALEXEEVA, V., WEISS, K. R. & CHURCH, P. J. (2001). Targeting of peptidergic vesicles in cotransmitting terminals. Journal of Neuroscience 21, 1-5
LANG, T., WACKER, I., WUNDERLICH, I., ROHRBACH, A., GIESE, G., SOLDATI, T. & ALMERS, W. (2000). Role of actin cortex in the subplasmalemmal transport of secretory granules in PC-12 cells. Biophysical Journal 78, 2863-2877	[Abstract/Full Text]
LEENDERS, A. G., SCHOLTEN, G., WIEGANT, V. M., DA SILVA, F. H. & GHIJSEN, W. E. (1999). Activity-dependent neurotransmitter release kinetics: correlation with changes in morphological distributions of small and large vesicles in central nerve terminals. European Journal of Neuroscience 11, 4269-4277	[Medline]
LEVITAN, E. S. (1998). Studying neuronal peptide release and secretory granule dynamics with GFP. Methods 16, 182-187	[Medline]
LEVITAN, E. S., TULLY, A. & DEITCHER, D. L. (2002). Stimulation induces physical mobilization of neuropeptidergic secretory vesicles in living synapses. Biophysical Journal 82, 279a
MORALES, M., COLICOS, M. A. & GODA, Y. (2000). Actin-dependent regulation of neurotransmitter release at central synapses. Neuron 27, 539-550	[Medline]
NG, Y.-K., LU, X., WATKINS, S. C., ELLIS-DAVIES, G. C. R. & LEVITAN, E. S. (2002). NGF changes the cellular organization of regulated peptide release by PC12 cells. Journal of Neuroscience 22, 3890-3897	[Abstract/Full Text]
OHEIM, M. & STUHMER, W. (2000). Tracking chromaffin granules on their way through the actin cortex. European Biophysics Journal 29, 67-89	[Medline]
RAO, S., LANG, C., LEVITAN, E. S. & DEITCHER, D. L. (2001). Visualization of neuropeptide expression, transport, and exocytosis in Drosophila melanogaster. Journal of Neurobiology 49, 159-172	[Medline]
RAUCHER, D., STAUFFER, T., CHEN, W., SHEN, K., GUO, S., YORK, J. D., SHEETZ, M. P. & MEYER, T. (2000). Phosphatidylinositol 4,5-bisphosphate functions as a second messenger that regulates cytoskeleton-plasma membrane adhesion. Cell 100, 221-228	[Medline]
RUDOLF, R., SALM, T., RUSTOM, A. & GERDES, H. H. (2001). Dynamics of immature secretory granules: role of cytoskeletal elements during transport, cortical restriction, and F-actin-dependent tethering. Molecular and Cellular Biology 12, 1353-1365
SACHS, H., SHARE, L., OSINCHAK, J. & CARPI, A. (1967). Capacity of the neurohypophysis to release vasopressin. Endocrinology 81, 755-770	[Medline]
SAITO, S., WATABE, S., OZAKI, H., FUSETANI, N. & KARAKI, H. (1994). Mycalolide B, a novel actin depolymerizing agent. Journal of Biological Chemistry 269, 29710-29714	[Abstract]
TOOZE, S. A., FLATMARK, T., TOOZE, J. & HUTTNER, W. B. (1991). Characterization of the immature secretory granule, an intermediate in granule biogenesis. Journal of Cell Biology 115, 1491-1503	[Abstract]
VITALE, M. L., SEWARD, E. P. & TRIFARO, J. M. (1995). Chromaffin cell cortical actin network dynamics control the size of the release-ready vesicle pool and the initial rate of exocytosis. Neuron 14, 353-363	[Medline]
WANG, X. H., ZHENG, J. Q. & POO, M. M. (1996). Effects of cytochalasin treatment on short-term synaptic plasticity at developing neuromuscular junctions in frogs. Journal of Physiology 491, 187-195	[Abstract]
WAYNE, N. L., KIM, J. & LEE, E. (1998). Prolonged hormone secretion from neuroendocrine cells of Aplysia is independent of extracellular calcium. Journal of Neuroendocrinology 10, 529-537	[Medline]
ZENGEL, J. E. & MAGLEBY, K. L. (1977). Transmitter release during repetitive stimulation: selective changes produced by Sr2+ and Ba2+. Science 197, 67-69
ZENGEL, J. E. & MAGLEBY, K. L. (1980). Differential effects of Ba2+, Sr2+, and Ca2+ on stimulation-induced changes in transmitter release at the frog neuromuscular junction. Journal of General Physiology 76, 175-211	[Abstract]
ZENGEL, J. E. & MAGLEBY, K. L. (1981). Changes in miniature endplate potential frequency during repetitive nerve stimulation in the presence of Ca2+, Ba2+, and Sr2+ at the frog neuromuscular junction. Journal of General Physiology 77, 503-529	[Abstract]
ZENISEK, D., STEYER, J. A. & ALMERS, W. (2000). Transport, capture and exocytosis of single synaptic vesicles at active zones. Nature 406, 849-854	[Medline]

Acknowledgements

This research was supported by NIH grant R01 NS32385 to E.S.L.

Authors' present addresses

Y.-K. Ng: Department of Neurobiology, Duke University, Durham, NC, USA.

X. Lu: Department of Medicine, University of Pittsburgh, PA, USA.

This article has been cited by other articles:

				T. Pangrsic, M. Potokar, M. Stenovec, M. Kreft, E. Fabbretti, A. Nistri, E. Pryazhnikov, L. Khiroug, R. Giniatullin, and R. Zorec Exocytotic Release of ATP from Cultured Astrocytes J. Biol. Chem., September 28, 2007; 282(39): 28749 - 28758. [Abstract] [Full Text] [PDF]

				D. Shakiryanova, M. K. Klose, Y. Zhou, T. Gu, D. L. Deitcher, H. L. Atwood, R. S. Hewes, and E. S. Levitan Presynaptic Ryanodine Receptor-Activated Calmodulin Kinase II Increases Vesicle Mobility and Potentiates Neuropeptide Release J. Neurosci., July 18, 2007; 27(29): 7799 - 7806. [Abstract] [Full Text] [PDF]

				S. Speese, M. Petrie, K. Schuske, M. Ailion, K. Ann, K. Iwasaki, E. M. Jorgensen, and T. F. J. Martin UNC-31 (CAPS) Is Required for Dense-Core Vesicle But Not Synaptic Vesicle Exocytosis in Caenorhabditis elegans J. Neurosci., June 6, 2007; 27(23): 6150 - 6162. [Abstract] [Full Text] [PDF]

				S. Huet, E. Karatekin, V. S. Tran, I. Fanget, S. Cribier, and J.-P. Henry Analysis of Transient Behavior in Complex Trajectories: Application to Secretory Vesicle Dynamics Biophys. J., November 1, 2006; 91(9): 3542 - 3559. [Abstract] [Full Text] [PDF]

				M. W. Allersma, M. A. Bittner, D. Axelrod, and R. W. Holz Motion Matters: Secretory Granule Motion Adjacent to the Plasma Membrane and Exocytosis Mol. Biol. Cell, May 1, 2006; 17(5): 2424 - 2438. [Abstract] [Full Text] [PDF]

				B. A. Scalettar How Neurosecretory Vesicles Release Their Cargo Neuroscientist, April 1, 2006; 12(2): 164 - 176. [Abstract] [PDF]

				Y.-S. Park, D.-J. Jun, E.-M. Hur, S.-K. Lee, B.-S. Suh, and K.-T. Kim Activity-Dependent Potentiation of Large Dense-Core Vesicle Release Modulated by Mitogen-Activated Protein Kinase/Extracellularly Regulated Kinase Signaling Endocrinology, March 1, 2006; 147(3): 1349 - 1356. [Abstract] [Full Text] [PDF]

				M. A. Bittner and R. W. Holz Phosphatidylinositol-4,5-bisphosphate: Actin Dynamics and the Regulation of ATP-Dependent and -Independent Secretion Mol. Pharmacol., April 1, 2005; 67(4): 1089 - 1098. [Abstract] [Full Text] [PDF]

				M. A. Silverman, S. Johnson, D. Gurkins, M. Farmer, J. E. Lochner, P. Rosa, and B. A. Scalettar Mechanisms of Transport and Exocytosis of Dense-Core Granules Containing Tissue Plasminogen Activator in Developing Hippocampal Neurons J. Neurosci., March 23, 2005; 25(12): 3095 - 3106. [Abstract] [Full Text] [PDF]

				X. Xin, F. Ferraro, N. Back, B. A. Eipper, and R. E. Mains Cdk5 and Trio modulate endocrine cell exocytosis J. Cell Sci., September 15, 2004; 117(20): 4739 - 4748. [Abstract] [Full Text] [PDF]

				M. Oheim A Deeper Look into Single-Secretory Vesicle Dynamics Biophys. J., September 1, 2004; 87(3): 1403 - 1405. [Full Text] [PDF]

				Y.-K. Ng, X. Lu, A. Gulacsi, W. Han, M. J. Saxton, and E. S. Levitan Unexpected Mobility Variation among Individual Secretory Vesicles Produces an Apparent Refractory Neuropeptide Pool Biophys. J., June 1, 2003; 84(6): 4127 - 4134. [Abstract] [Full Text] [PDF]

This Article Abstract Full Text (PDF) All Versions of this Article: 542/2/395    most recent 2002.021733v1 Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Ng, Y.-K. Articles by Levitan, E. S. Search for Related Content PubMed PubMed Citation Articles by Ng, Y.-K. Articles by Levitan, E. S.