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Vol. 20, Issue 6, 1795-1803, March 15, 2009
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School of Biomedical Sciences, University of Queensland, Brisbane, Queensland, QLD 4072, Australia
Submitted October 21, 2008;
Revised December 19, 2008;
Accepted January 9, 2009
Monitoring Editor: Thomas F.J. Martin
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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; Sudhof, 2004). But although vesicle fusion is the point where content release is initiated it is now recognized that postfusion events can be critical in regulating the kinetics and amount of released vesicle content (Albillos et al., 1997; Graham et al., 2002; Perrais et al., 2004; Vardjan et al., 2007). The mechanisms that control postfusion vesicle behavior are not well understood, but given that they control the release of content they may provide excellent therapeutic targets to up or down-regulate secretory responses in the treatment of disease (Fernandez-Peruchena et al., 2005).
F-Actin and Myosin Involvement
Long-standing observations show that cortical F-actin can influence secretory processes (Burgoyne and Cheek, 1987; Trifaro and Vitale, 1993), but its exact role is not clear (Muallem et al., 1995; Malacombe et al., 2006). For example, it has been suggested that subplasmalemmal F-actin has to be cleared out the way to allow vesicles to dock with the plasma membrane (Giner et al., 2005). It is also possible that F-actin remodelling in some way moves vesicles from within the cell into a position where they might then fuse with the cell surface membrane (Lang et al., 2000) or F-actin may form part of the poststimulation recovery of vesicles (Shupliakov et al., 2002).
The findings of F-actin involvement in secretion further suggest that myosins, a family of molecular motors that bind to actin, may also play a role. There are many classes of mammalian myosins and in conjunction with the F-actin cytoskeleton all use ATP hydrolysis to promote movement or generate force (Hodge and Cope, 2000). Additional studies imply myosin Va (Varadi et al., 2005; Desnos et al., 2007) and myosin VI (Buss et al., 2002) involvement in vesicle movement, but here we focus on the possible role of myosin 2. There is extensive evidence that myosin 2 plays a role in the secretory processes of a variety of cells, including: mast cells (Choi et al., 1994; Ludowyke et al., 2006), natural killer cells (Andzelm et al., 2007), hippocampal (Ryan, 1999) and sensory neurons (Mochida et al., 1994), chromaffin cells (Neco et al., 2004), β cells (Iida et al., 1997; Wilson et al., 2001), exocrine cells (Segawa and Yamashina, 1989; Torgerson and McNiven, 2000; Jerdeva et al., 2005), and oocytes (Becker and Hart, 1999). However, most of these studies rely on measurement of secretory output and therefore lack evidence to show the site(s) of action of F-actin and myosin 2. Also some of these studies use 2,3-butanedione monoxime, as a nonspecific myosin inhibitor, and the clear and dramatic effects of this drug on cell calcium responses (Turvey et al., 2003) cloud interpretation of these data.
More recently, the methods that have been developed to resolve the sequential steps of granule fusion and fission have been applied to the study of the regulation of these processes. With these techniques, accumulating evidence indicates a role for F-actin and myosin 2 in regulating the postfusion behavior of vesicles. In eggs and pancreas, F-actin coats individual vesicles immediately postfusion (Valentijn et al., 2000; Sokac et al., 2003, Turvey and Thorn 2004, Nemoto et al., 2004; Yu and Bement, 2007). In the pancreas, eggs, and chromaffin cells, pharmacological block of actin polymerization influences postfusion vesicle behavior (Sokac et al., 2003; Neco et al., 2004; Nemoto et al., 2004; Larina et al., 2007; Yu and Bement 2007; Doreian et al., 2008) and vesicle content loss (Felmy, 2007). Likewise, inhibition of myosin 2 affects postfusion events, slowing the opening of the fusion pore (Neco et al., 2008; Doreian et al., 2008) and playing a role in promoting vesicle full fusion (Yu and Bement, 2007; Doreian et al., 2008).
Pancreas
Methods developed to visualize single vesicles during the secretory cycle have shown postfusion F-actin coating of zymogen granules (Nemoto et al., 2004; Turvey and Thorn, 2004). Treatment with the F-actin depolymerizing agent latrunculin decreases granule lifetimes (Nemoto et al., 2004) and closes the fusion pore (Larina et al., 2007). In this article, we show that myosin 2A is located in the apical region and is phosphorylated during cell stimulation. We further show that myosin 2 inhibition does not regulate the number of granule fusion events stimulated by agonist but instead affects the postfusion behavior of the granules, inhibition leading to closure of the fusion pore (here we use the term fusion pore to describe the structural link between the cell membrane and the granule during fusion—some other authors use this term to strictly define the small, nanometer-size pore that forms on initial fusion). Our work is consistent with recent findings in other cell types (Yu and Bement, 2007; Doreian et al., 2008; Neco et al., 2008) and points to myosin 2 as a key player in what is likely to be the complex regulation of postfusion granule behavior that may be important in regulating the release of vesicle content.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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), modified to reduce the time in collagenase and limit mechanical trituration. The resultant preparation was composed mainly of pancreatic lobules and fragments (50–100 cells), which were plated onto poly-L-lysine–coated glass coverslips and used within 3 h of isolation from the animal.
Confocal Imaging
Fixed specimens were imaged using a Zeiss LSM 510 Axioscope confocal laser scanning microscope, with a 63x objective lens (numerical aperture [NA] 1.3), and an optical slice depth of 
1 µm. Images were collected with the appropriate filters and captured in sequential tracks to minimize cross-talk to <2%. Fluorescent probes were from Invitrogen (Carlsbad, CA). All other compounds were from (Sigma-Aldrich, Sydney, Australia). All experiments were performed at least three times. We used MetaMorph software (Molecular Devices, Sunnyvale, CA) for analysis of LSM images and Adobe Photoshop (Adobe Systems, Mountain View, CA) for image processing and presentation.
Indirect Immunofluorescence
Cell preparations were bathed in 2 mg/ml lysine-fixable, fluorescein isothiocyanate (FITC) or tetramethylrhodamine ethyl ester (TMRE) conjugated 3 kDa dextran dyes (Invitrogen, Molecular Probes), stimulated for 5 min with 10 pM cholecystokinin (CCK), fixed in freshly prepared 3.4% paraformaldehyde in PBS for 30 min, and permeablized with 1% Triton-X 100 in PBS for 30 min. Myosin subtypes were detected using goat anti-myosin 2A or 2B antibodies (Sigma-Aldrich) and visualized with rabbit anti-goat Alexa 633 secondary antibody (Invitrogen, Molecular probes). Phospho-myosin distribution in cells was imaged using rabbit anti-phospho-myosin 2 (Ser19) antibody (Cell Signal Technology), and mouse anti-rabbit Alexa 633. F-actin was visualized in fixed and permeabilized tissue with phalloidin Alexa 633 (Invitrogen, Molecular probes), or TRITC-phalloidin (Sigma-Aldrich).
Western Blotting
Pancreatic tissue fragments were treated with myosin inhibitors for 20 min at room temperature before stimulation with acetylcholine (ACh) 10 µM for 1 min followed by atropine (100 µM). Cells were rapidly frozen in liquid nitrogen 4 min later and lysed by 3 freeze-thaw cycles. For time-course studies, cells were not drug treated, but stimulated with acetylcholine (ACh) and atropine, and harvested at various times after this. The lysate was centrifuged to isolate the cytosolic fraction. Samples were boiled in Laemmli buffer and electrophoresed on Mini-Protean II apparatus (Bio-Rad, Hercules, CA) through a 13% polyacrylamide reducing gel before transfer to Hybond-C nitrocellulose membrane (GE Healthcare, Little Chalfont, Buckinghamshire, United Kingdom) by using a Trans Blot semidry transfer cell (Bio-Rad) for Western blotting. The membrane was blocked in 5% bovine serum albumin and then probed with rabbit anti-phospho-myosin 2 antibody, or mouse anti-β-actin as loading control, at 4°C overnight. Bands were detected using anti-rabbit Alexa 680 and anti-mouse Alexa 800 and visualized on an Odyssey 2 fluorescence plate reader (Li-Cor Biosciences, Lincoln, NE). Bands were quantified using MetaMorph and expressed as a ratio of β-actin, and as a percentage of control.
Assessment of Fusion Pore Dynamics (Figure 5 and 6)
Freshly prepared cells were placed on poly-L-lysine-coated coverslips and incubated with lysine-fixable FITC-dextran (2 mg/ml) at 37°C. Lysine-fixable TMRE-dextran (4 mg/ml) was added either along with the FITC-dextran or at various times after the start of the ACh stimulation. After stimulation, cells were fixed in fresh 3.4% paraformaldehyde in phosphate-buffered saline (PBS) for 30 min. To determine the TMRE/FITC (red/green) ratio, fluorescence intensity was measured in a region of interest (diameter, 0.5 µm) in granule and luminal areas. The TMRE/FITC ratio in the granules were normalized to the ratios within the adjacent lumens (see Larina et al., 2007 for details). All graphs were produced in Excel (Microsoft, Redmond, WA).
Live-Cell Two-Photon Imaging (Figure 7)
We used a custom-made, video-rate, two-photon microscope using a Sapphire-Ti laser (Thorn et al., 2004), with a 60x oil immersion objective (NA 1.42; Olympus, Tokyo, Japan), providing an axial resolution (full width, half maximum) of 1 µm. We imaged exocytic events by using sulforhodamine B (SRB, 20 µg ml–1; Sigma-Aldrich) and FITC as a membrane-impermeant fluorescent extracellular markers excited by femtosecond laser pulses at 800 nm, with fluorescence emission detected at 450–510 nm (FITC) and 550–700 nm (SRB).
Images (resolution of 10 pixels/µm; average of 15 video frames) were analyzed with MetaMorph. An epifluorescent mercury light source provided high-intensity light to photobleach the FITC extracellular dye in an 30-µm diameter field at the image plane. Exocytotic event kinetics was measured from regions of interest (0.78 µm2; 100 pixels) over granules. Traces were rejected if extensive movement was observed. All data are shown as mean ± SEM.
Neurite Outgrowth Assay
We tested the efficacy of Y27632 in an independent assay to validate our results on the epithelial cells. Y27632 has been well characterized to stimulate differentiation and neurite outgrowth in PC12 cells (Zhang et al., 2005). PC12 cells were plated onto six-well plates at 10 cells/well and allowed to adhere. Cells were cultured with 0, 10, 30, or 50 µM Y26732 or 100 ng ml–1 of neurite growth factor for 48 h. Cells were then imaged by light microscopy, and the number of cells with greater than one neurite was counted. Y27632 caused neurite growth in PC12 cells in a dose-related manner, comparable at 50 µM with neurite growth factor (see Supplemental Data).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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). Here, we wanted to examine the specific distribution of myosin 2 isoforms. The myosin 2 isoforms A and B were immunolocalized in isolated mouse pancreatic acinar cell clusters that were costained with phalloidin to reveal the F-actin distribution. In addition, the cells were bathed in lysine-fixable fluorescent dye (FITC), which, after fixation, labels the acinar lumen (Larina and Thorn 2005). Figure 1 shows the dotted appearance of myosin 2B in the perinuclear and basal domain but not present in the apical region. In contrast myosin 2A (Figure 1) colocalized extensively with both the luminal FITC marker and with phalloidin staining indicating that it was predominantly, although not exclusively, localized to the apical region. Our work now shows that the apical myosin isoform is likely myosin 2A and is therefore in a position to regulate the exocytic release of digestive enzymes because this occurs exclusively at the apical plasma membrane.
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, Torgerson and McNiven 2000), physiological levels of stimulation in these cells results in a rapid and prolonged increase in myosin 2 light chain phosphorylation. The persistence of myosin phosphorylation, despite the transient stimulation, suggests that phosphatase activity in this system may be weak.
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) and investigated its upstream regulation by using ML-9, an inhibitor of myosin light chain kinase (MLCK), the major regulator of myosin phosphorylation, or Y27632, a Rho-kinase inhibitor. The latter is expected to affect phosphorylation of myosin either directly through Rho-kinase or indirectly via an enhancement of phosphatase activity (Amano et al., 1996). Cells were drug treated for 20 min and then stimulated for 1 min with 10 µM ACh followed by 100 µM atropine. Cells were lysed by rapid freeze-thaw and processed for SDS-polyacrylamide gel electrophoresis and Western blotting by using ser-19 phospho-myosin 2 antibody and β-actin loading control. Bands were quantified and corrected for β-actin and expressed as a percentage of no-drug controls. Figure 3 shows that ML-9 treatment (30 µM) significantly decreased myosin phosphorylation to 82.6% compared with untreated controls (p < 0.05). In contrast, neither (–)-blebbistatin nor Y27632 (50 µM) significantly changed phospho-myosin levels. For Y27632, we were expecting to see a change due to its direct or indirect action. To confirm that the compound was bioactive, we used Y27632 in a neurite growth assay in PC12 cells (Zhang et al., 2005). Here, it has been shown that Y27632 is as effective as neurite growth factor in inducing neurite outgrowth (Zhang et al., 2005), a finding we confirm (see Supplemental Data). We conclude that the negative effects of Y27632 on myosin 2 phosphorylation in the pancreatic acinar cells are truly indicative of low levels of Rho-kinase activity.
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). Figure 4 shows that without CCK there is a "spontaneous" count of approximately one granule per cell; something we have observed before and presumably due to neuronal and cell-to-cell regulatory mechanisms still present in the tissue fragments (Thorn et al., 2004). CCK stimulation leads to a threefold increase in the numbers of fused granules per cell. This increase is not affected by either the myosin inhibitor, (–)-blebbistatin (50 µM) or by the myosin light chain kinase inhibitor ML-9 (30 µM). Thus, in acinar cells, inhibition of myosin 2 does not alter the number of granules secreted per cell in response to CCK stimulation. We conclude that the exocytic fusion process itself is not affected by myosin 2 inhibition. Because studies have shown that myosin inhibition can significantly reduce the amount of enzyme secreted from acinar cells (Mizuno et al., 2000; Torgerson and McNiven, 2000), our data place myosin 2 as a postfusion regulator of secretion possibly through a control of the granule behavior at the plasma membrane.
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). Adding a second dye, 5 min after stimulation probes whether the fusion pore is open: if open, the dye enters then the granules are labeled; if closed, the second dye cannot enter and the first dye is trapped. Quantification of the ratio of second (TMRE) to first dye (FITC) therefore gives us a measure of fusion pore behavior, which we have used previously to show that granule fusion is often transient (Larina et al., 2007). Figure 5A shows that pretreatment with (–)-blebbistatin or ML-9 before cell stimulation (with 10 pM CCK) increased the prevalence of granules with a low second to first dye ratio. To analyze the data, the TMRE/FITC fluorescence intensity within a region drawn over each granule, normalized to the dye ratio in the lumen, was calculated. If the fusion pore is open, then there will be almost equal amounts of FITC and TMRE present, and their ratio is nearly 1. Values <0.4 indicated that less than half of the amount of dye 2 compared with dye 1 was in the granule, and the fusion pore was deemed closed. Analysis of >50 cell clusters from 15 of these images from six experiments showed that untreated cells had a fusion pore closure rate of under 20% at 5 min after stimulation (Figure 5B). Treatment with the myosin 2 inhibitor blebbistatin, however, resulted in 58% closed granules 5 min after stimulation (p < 0.001, compared with controls; n = 3 preparations). The (+)-blebbistatin inactive enantomer was used as a control in this experiment (with no effect). We conclude that myosin 2 inhibition leads to a marked increase in the number of closed fusion pores at 5 min after stimulation, suggesting early fusion pore closure. We wanted to examine the functional upstream regulation of myosin 2 by MLCK because in Western blot experiments (Figure 3), we show that ML-9 decreases phosphorylation. Cell fragments were treated with ML-9 and examined for fusion pore closure at 5 min after stimulation using the two-dye fixed cell technique as described. As with (–)-blebbistatin treatment, there was a marked increase in the number of granules with closed fusion pores in ML-9–treated cells (Figure 6). In similar experiments, pretreatment of tissue fragments with the Rho-kinase inhibitor Y27632 had no significant effect on the number of closed fusion pores at 5 min after stimulation (data not shown). These data suggest the Ca2+/calmodulin MLCK pathway primarily regulates myosin 2 phosphorylation in these cells, with little role for Rho-kinase.
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). Tissue fragments bathed in extracellular FITC dye were stimulated with CCK (10 pM), allowing dye to enter exocytic cells. Then, at each cycle of photobleaching, the fluorescence of the dye within the granule decreases, with fluorescence recovery indicating that the fusion pore is open and fresh dye can enter, whereas lack of recovery indicating fusion pore closure. Photobleaching CCK-stimulated cells every 55 s, we observed fluorescence recovery in most granules (n = 33) over a period of 10 min after fusion (mean pore lifetime 10.44 ± 0.44 min) (Figure 7A). In contrast, after 20-min blebbistatin pretreatment (50 µM; n = 16) followed by CCK stimulation, fluorescence recovery in fused granules was seen in the first few cycles of photobleaching but at later cycles recovery was not seen, indicating premature closure of the fusion pore (mean pore lifetime 4.86 ± 0.43 min) (Figure 7B). To analyze the data we arbitrarily defined a slope (fitted in Excel by a simple linear regression) of <5% fluorescence recovery per minute as indicative of lack of recovery. The summarized data (Figure 7C) show that the modal values for closure of the fusion pore in control cells is 11 min compared with 4 min after blebbistatin (p < 0.001). We conclude that these methods of probing fusion pore behavior show that inhibition of myosin 2 decreases the fusion pore lifetimes.
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; Turvey and Thorn, 2004; Nemoto et al., 2004). Phalloidin staining of the cell clusters treated with (–)-blebbistatin (50 µM), ML-9 (30 µM), or Y27632 (30 µM), however, did not affect the appearance of F-actin–coated granules (Figure 8A), nor the number of actin-coated granules (Figure 8B). There were n >200 granules per group, and the difference between no drug and the treatment groups did not reach statistical significance. This data are consistent with the hypothesis that granule F-actin coating is initiated by actin nucleation probably under the control of Cdc42 (Nemoto et al., 2004; Yu and Bement, 2007) and suggests that myosin 2 is specifically necessary to maintain fusion pore opening.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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F-Actin and Myosin 2 Play a Postfusion Role during Exocytosis
The exact role of F-actin postfusion is still not clear, but evidence suggests there are two distinct sites of action: one at the fusion pore and the other in coating fused granules. In the first, F-actin dynamics regulate the behavior of the fusion pore. We have directly shown that F-actin disruption with latrunculin closes the fusion pore (Larina et al., 2007). Consistent with this, Felmy (2007) shows that the expression of actin and actin associated proteins, expected to affect F-actin dynamics, influence the release of granule content. And finally, granule collapse, presumably reflecting loss of fusion pore integrity can be enhanced by F-actin disruption (Sokac et al., 2003; Doreian et al., 2008). The second site of actin involvement leads to the F-actin coating of fused granules in eggs and acinar cells (Segawa and Yamashina, 1989, Nemoto et al., 2004; Sokac et al., 2003; Turvey and Thorn, 2004). This coating might be specific to these cell types or it might reflect difficulties in visualizing the smaller secretory granules in other cells types. Previous findings show that this coating requires actin nucleation (Sokac et al., 2003; Nemoto et al., 2004) and our work, showing myosin 2 activity is not part of the mechanism of F-actin coating of the granule (Figure 8), is consistent with this. The functional significance of F-actin coating is not known (Malacombe et al., 2006). It is apparently not specifically required to maintain granule integrity because fused granules are still observed after drug treatments that specifically prevent F-actin coating (Nemoto et al., 2004; Yu and Bement, 2007). Yu and Bement (2007) have evidence that it might be part of a contractile process, squeezing content out, an idea first put forward by Segawa and Yamashina (1989). Furthermore, F-actin coating might be an early step in the endocytic recovery of granule membrane as suggested from work in neurons (Shupliakov et al., 2002).
There are some contradictions in this work on F-actin that probably reflect differences in the consequences of drug actions in different cell types, as well as possible different functions for the preexisting cortical F-actin and newly formed F-actin. For example, in some studies treatment with agents to disrupt F-actin formation destabilizes the fused granules as measured by a favoring of granule collapse (Sokac et al., 2004, Doreian et al., 2008) and an increase in the measured quantal size of granule content release (Doreian et al., 2008). In contrast, Yu and Bement (2007) show that mild F-actin disruption (with cytochalasin D) "traps" fused granules at the plasma membrane. Consistent with this, in acinar cells that are known to have a particularly robust preexisting F-actin cytoskeleton, resistant to drug disruption (Turvey et al., 2005), latrunculin treatment leads to the long-term retention of fused granules (Nemoto et al., 2004). We might tentatively conclude that treatments that disrupt the preexisting F-actin cytoskeleton destabilize granules leading to collapse, whereas with more subtle treatments, or in cells with a particularly extensive F-actin cortex, granule integrity is maintained but behaviors such as fusion pore opening are affected.
Our data place myosin 2 as necessary for the maintained opening of the fusion pore. This is consistent with other reports of myosin 2 inhibition leading to a slowing of fusion pore expansion (Doreian et al., 2008; Neco et al., 2008) and, of course, does not exclude an additional role for myosin in providing a contractile force for expulsion of granule content (Segawa and Yamashina, 1989; Yu and Bement, 2007). To maintain fusion pore opening myosin 2 is likely to be part of a machinery that is linked to the pore membrane and linked to the surrounding F-actin cytoskeleton providing forces to pull (or maintain) pore opening.
Other Potential Regulators of the Fusion Pore
The indication that myosin 2 is a regulator of the fusion pore, places it as potential component in a fusion pore macromolecular complex. A range of other factors such as dynamin (Graham et al., 2002; Tsuboi et al., 2004), complexin (Archer et al., 2002), calcium (Alés et al., 1999; Haller et al., 2001; Elhamdani et al., 2006; Llobet et al., 2008), syntaxin (Wang et al., 2003), and amphiphysin (Llobet et al., 2008) have also been shown to affect pore dynamics. Although a coherent view of the composition and regulation of the fusion pore complex has yet to emerge, it is likely, given this diversity of possible control pathways, that its regulation may subserve genuine physiological functions.
Possible Role of Fusion Pore Dynamics in Regulating Granule Content Release
The regulation of granule content release is an exciting possible function for the control of fusion pore behavior. It has been shown that different exogenous content can be differentially released from granules (Michael et al., 2004; Felmy 2007). In dense-core granules, ATP content has been shown to be released rapidly and preferentially compared with peptide content (Obermuller et al., 2005). Bauer et al. (2004) and Perrais et al. (2004) show that protein content can be retained after a cycle of fusion and fission and recycled back into the cell. These studies are consistent with either fusion pore size or dynamics regulating content release. Furthermore, Felmy (2007) shows that molecular weight is not the sole determinant of content release kinetics, indicating that, in addition to pore size and dynamics as potential regulators of release, the time course of intragranule matrix dissociation may also play a role.
Role of F-Actin and Myosin 2 in the Physiology of Secretion in Acinar Cells
Torgerson and McNiven (2000) originally described a decrease in enzyme secretion from acinar cells treated with myosin inhibitors. Our work now suggests that at least in part this is due to the premature closure of the fusion pore. We might imagine this would have two consequences. First, it could directly limit the release of content from the fused granule. This seems unlikely because we have shown that content release is very rapid, with the majority of content lost within a few seconds (Thorn and Parker, 2005). Second, and more likely, the closure of the fusion pore could prevent the release of content from secondary granules, fused to the first as a result of compound exocytosis (Nemoto et al., 2001). In this way, granule content would equilibrate within the fused granules but could not escape to the outside.
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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ACKNOWLEDGMENTS |
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Footnotes |
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Present address: * Princess Alexandra Hospital, Level 4, R Wing, Diamantina Institute, Ipswich Rd., Woolloongabba QLD 4102, Australia. 
Address correspondence to: Peter Thorn (p.thorn{at}uq.edu.au)
Abbreviations used: ACh, acetylcholine; CCK, cholecystokinin; FITC, fluorescein isothiocyanate; MLCK, myosin light chain kinase; TMRE, tertramethylrhodamine ethyl ester; SRB, sulforhodamine B.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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