The motor protein myosin Va is involved in multiple successive steps in the development of dense-core vesicles, such as in the membrane remodelling during their maturation, their transport along actin filaments and the regulation of their exocytosis. In the present paper, we summarize the current knowledge on the roles of myosin Va in the different steps of dense-core vesicle biogenesis and exocytosis, and compare findings obtained from different cell types and experimental systems.

Introduction

Hormone-producing cells, for instance those of the adrenal gland, pancreas, pituitary, gastrointestinal tract or the brain, contain DCVs (dense-core vesicles), which are small, spherical organelles packed with hormones and neuropeptides. Although the cells expressing DCVs have a broad range of functions and produce diverse hormones, the biogenesis and exocytosis of their DCVs share a pathway with certain conserved hallmarks. By definition, DCVs contain an electron-dense core consistent with the secreted hormones and neuropeptides, and they undergo regulated exocytosis in response to an appropriate stimulus [1]. It emerges that the F-actin (filamentous actin)-dependent motor protein myosin Va plays roles in the regulation of several key events of this pathway, namely DCV transport, maturation and exocytosis (Figure 1). In the present review, we provide an overview of the up-to-date knowledge of the involvement of myosin Va in these processes and highlight emerging differences between the cell types investigated regarding DCV transport and exocytosis.

Mode of interaction and functional roles of myosin Va on DCVs

Figure 1
Mode of interaction and functional roles of myosin Va on DCVs

The upper panel illustrates the current view of myosin Va recruitment to DCVs by a Rab and a linker protein (Rab effector). Variation of the Rab and/or effectors within and across cell types may lead to functional diversity of myosin Va. The lower panels highlight the major putative functions of myosin Va during the lifetime of a DCV as described so far. These include (i) the maturation process, where small vesicles (open circle) bud from IDCVs (broken arrow) (left), (ii) the F-actin-dependent transport (open arrow) (middle), and (iii) the exocytosis of DCVs (right).

Figure 1
Mode of interaction and functional roles of myosin Va on DCVs

The upper panel illustrates the current view of myosin Va recruitment to DCVs by a Rab and a linker protein (Rab effector). Variation of the Rab and/or effectors within and across cell types may lead to functional diversity of myosin Va. The lower panels highlight the major putative functions of myosin Va during the lifetime of a DCV as described so far. These include (i) the maturation process, where small vesicles (open circle) bud from IDCVs (broken arrow) (left), (ii) the F-actin-dependent transport (open arrow) (middle), and (iii) the exocytosis of DCVs (right).

Maturation of DCVs

A secretory cell contains thousands of DCVs with a half-life of several days. Since maturation of DCVs is comparably fast, with a half-time of, e.g., ~45 min in the neuroendocrine phaeochromocytoma cell line PC12 [2], only a small fraction of the DCVs is immature. Therefore the challenge to monitor IDCVs (immature DCVs) affords specific labelling. This was achieved by a pulse–chase-like labelling of DCVs in PC12 cells based on the expression of DCV marker proteins tagged with green fluorescent protein. This allowed visualizing pools of fluorescent DCVs of a defined age (i.e. maturation status) that could be tracked microscopically [3,4]. Furthermore, the approach led to insights about the functional implications of cytoskeletal elements and motor proteins in IDCV transport and maturation: within a few seconds of biogenesis at the trans-Golgi network, IDCVs move to the cell periphery in a unidirectional, microtubule-dependent manner [3]. An implication of kinesin motor protein involvement in this process has been demonstrated, and the kinesin isoform engaged seems to vary according to the cell type [5,6]. Subsequent to the fast movement towards the cell periphery in PC12 cells, IDCVs remain restricted to the dense cortical F-actin network, where they move in a myosin Va-dependent fashion in all directions while they undergo maturation [3,4]. Knockdown of myosin Va by siRNA (small interfering RNA) or expression of a truncated tail construct which comprises the organelle-binding globular tail, but not the F-actin-binding motor domain (MyoVa-tail) prevents distribution of DCVs into the F-actin-rich cortex [4,7,8]. In other cell types, the early transport events of IDCVs may follow a similar pattern, although this remains to be investigated.

During the phase of movement along cytoskeletal structures, DCVs mature in a highly regulated process composed of several discrete steps, which are completed within a few hours. The observation that movement and maturation occur simultaneously hints at a possible functional link between the two processes. This idea is supported by further observations. During maturation, several IDCVs were shown to fuse homotypically, and their dense cores combined to form a single bigger core [9]. This fusion process requires a set of SNAREs (soluble N-ethylmaleimide-sensitive fusion protein-attachment protein receptors) consisting of syntaxin 6 and synaptotagmin IV that differs from the SNAREs required for exocytosis and provides specificity for this process [10,11]. As maturation proceeds, these SNARE proteins and enzymes involved in cargo processing are removed from the maturing vesicles by clathrin-mediated membrane remodelling [1215]. The exact time window of this remodelling seems to vary according to the cell type or the molecule removed. The removal of synaptotagmin IV, for example, is accomplished within approx. 45 min in AtT-20 cells [14], whereas furin is removed from IDCVs between 12 and 30 min in PC12 cells [3,4]. We demonstrated an involvement of myosin Va in this process by showing that compromising myosin Va inhibits the removal of furin in PC12 cells (T. Kögel, R. Rudolf and H.-H. Gerdes, unpublished work).

Another hallmark of DCV maturation is the acidification of the DCV lumen, which is characterized by a decrease in intraluminal pH from 6.5 in the Golgi apparatus to 5.0–6.0 in mature DCVs. This occurs within 90 min in PC12 cells [16] and is required for the enzymatic processing of pro-hormones to yield bioactive molecules [17] and for the aggregation of cargo [18]. Acidification occurs concomitantly with membrane remodelling, but different experimental manipulations demonstrate that the two processes are separate events: inhibition of ARF (ADP-ribosylation factor)-1 recruitment to IDCVs by brefeldin A or inhibition of myosin Va block the protein removal step, but do not affect acidification-dependent processing of pro-opiomelanocortin [14,19] or secretogranin II (T. Kögel, R. Rudolf and H.-H. Gerdes, unpublished work) respectively. Conversely, the GGA3 (Golgi-associated γ-adaptin ear homology domain ARF-interacting protein 3) clathrin adaptor protein appears to be essential for both protein removal and pro-hormone processing. Knockdown of GGA3 led not only to a retention of the SNARE proteins syntaxin 6 and VAMP4 (vesicle-associated membrane protein 4) in mature DCVs, but also to a decreased pH-dependent activity of pro-hormone convertase 2 [20]. In conclusion, membrane remodelling and luminal processes appear to depend on DCV-bound external factors such as coat and motor proteins. Although myosin Va seems to regulate a maturation step that includes the removal of proteins from DCVs, it does not affect luminal acidification. How exactly myosin Va acts in this remodelling step is unknown, but since it occurs during a critical period where IDCVs undergo active myosin Va-dependent movement within the F-actin cortex in PC12 cells [3,4], we suggest that DCV maturation and F-actin-/myosin Va-dependent transport are closely linked in PC12 cells (Figure 1).

F-actin-dependent movement and exocytosis of DCVs in different cell types

DCVs are stored in the cortical area underlying the PM (plasma membrane) where they await a stimulus that induces fusion with the PM and secretion of the DCV content. According to the biologically diverse roles of DCVs in different cells, the kinetics of this secretory process vary greatly between cell types. In this respect, the F-actin cytoskeleton, the reaction of the F-actin-dependent motor myosin Va on physiological calcium elevation (for a review, see [21]) and, as a result, the mobility of DCVs in the vicinity of the PM preceding exocytosis all play important roles. In the following sections, we give a comparative overview of the modes of myosin Va function on DCV exocytosis in different cell types, which is also summarized in Table 1.

Table 1
Effects of myosin Va manipulation on the exocytosis of DCVs in different cell types
Cell typeExperimental manipulationObservationsReference
Chromaffin cells Anti-myosin Va head antibody Reduced noradrenaline secretion [28
 Anti-myosin Va neck antibody Fewer exocytotic events [29
PC12 cells MyoVa-tail Reduced mobility in the F-actin cortex; fewer exocytotic events [4]; T. Kögel, R. Rudolf and H.-H. Gerdes, unpublished work 
 siRNA against myosin Va More exocytotic events T. Kögel, R. Rudolf and H.-H. Gerdes, unpublished work 
Pancreatic β-cells (MIN6) siRNA against myosin Va Reduced insulin secretion [8
 MyoVa-tail Fewer docked DCVs; fewer exocytotic events [8
Pancreatic β-cells (INS-1E) siRNA against myosin Va Reduced growth hormone secretion [32
 siRNA against myosin Va or Slac-2c-/MyRIP Smaller capacitance increase (=fewer membrane fusion events) [32
 anti-MyoVa-tail antibody Smaller capacitance increase (=fewer membrane fusion events) [32
BON cells MyoVa-tail or siRNA against myosin Va Higher mobility in actin cortex; fewer docked vesicles; fewer long-term docking events; fewer exocytotic events [7
Hippocampal neurons MyoVa-tail More exocytotic events [41
Cell typeExperimental manipulationObservationsReference
Chromaffin cells Anti-myosin Va head antibody Reduced noradrenaline secretion [28
 Anti-myosin Va neck antibody Fewer exocytotic events [29
PC12 cells MyoVa-tail Reduced mobility in the F-actin cortex; fewer exocytotic events [4]; T. Kögel, R. Rudolf and H.-H. Gerdes, unpublished work 
 siRNA against myosin Va More exocytotic events T. Kögel, R. Rudolf and H.-H. Gerdes, unpublished work 
Pancreatic β-cells (MIN6) siRNA against myosin Va Reduced insulin secretion [8
 MyoVa-tail Fewer docked DCVs; fewer exocytotic events [8
Pancreatic β-cells (INS-1E) siRNA against myosin Va Reduced growth hormone secretion [32
 siRNA against myosin Va or Slac-2c-/MyRIP Smaller capacitance increase (=fewer membrane fusion events) [32
 anti-MyoVa-tail antibody Smaller capacitance increase (=fewer membrane fusion events) [32
BON cells MyoVa-tail or siRNA against myosin Va Higher mobility in actin cortex; fewer docked vesicles; fewer long-term docking events; fewer exocytotic events [7
Hippocampal neurons MyoVa-tail More exocytotic events [41

Chromaffin cells

In chromaffin cells, DCVs at the PM are restricted in mobility, which increases only upon stimulation of exocytosis. Increased DCV mobility is more prominent during the late phase of stimulation [22,23] and is enabled by a combination of F-actin disassembly and maintenance or assembly of some outbound actin filament tracks (for a review, see [24]). Before fusion with the PM, DCVs go through a short immobile docking phase [25]. Using a fluorescent protein that shifts colour from green to red within 16 h, it was shown that chromaffin cells preferentially release younger DCVs, rather than the pre-existing older ones [26]. Chromaffin cells contain a readily releasable pool of DCVs which is released with fast kinetics in the millisecond range upon elevation of cytoplasmic free calcium levels to micromolar concentrations, and a reserve pool which is only released upon sustained stimulation after a delay (for a review, see [27]). An involvement of myosin Va in the regulation of exocytosis was shown by the application of a specific antibody against the head or the neck domain of myosin Va, reducing the amount of regulated exocytosis specifically after sustained stimulation, but not during the initial phase of exocytosis [28,29]. These data point towards a role of myosin Va in recruiting DCVs from the reserve to the readily releasable pool.

PC12 cells

The DCVs of undifferentiated PC12 cells (a rat phaeochromocytoma cell line derived from chromaffin cells) exocytose at a very slow pace (release kinetics of seconds) compared with chromaffin cells (milliseconds), presumably because PC12 cells lack an equivalent of the readily releasable pool (for a review, see [27]). In PC12 cells, DCVs move within the F-actin cortex, and the average movement decreases with the age of the observed DCV pool [3]. At 3 h after biogenesis, approx. 50% of DCVs are immobile and half of these are tethered to F-actin as shown by confocal light microscopy in cells treated with F-actin-depolymerizing agents [3]. A possible involvement of myosin Va in this tethering was suggested by the demonstration that myosin Va is associated with DCVs and that the expression of MyoVa-tail strongly reduced their co-localization with the F-actin-rich cortex and their mobility during their maturation period [4]. Furthermore, expression of MyoVa-tail reduced the amount of exocytosis in PC12 cells (T. Kögel, R. Rudolf and H.-H. Gerdes, unpublished work). Interestingly, down-regulation of myosin Va expression by siRNA increased exocytosis in PC12 cells (T. Kögel, R. Rudolf and H.-H. Gerdes, unpublished work). These data clearly indicate a regulatory role for myosin Va in the exocytosis in PC12 cells.

Pancreatic β-cells

Pancreatic β-cell models such as MIN6 and or INS-1E cells contain a fraction of DCVs, which is immobilized in close proximity to the PM [30,31] and referred to as docked DCVs. This fraction represents the readily releasable pool and shows a fast response in exocytosis upon stimulation with high potassium concentrations [30]. During sustained secretion elicited by high glucose concentrations, new DCVs are recruited from central regions to the periphery of the cell, and this recruitment seems to occur by directed vesicle transport (Figure 2) [8,32]. The involvement of myosin Va in this step was demonstrated by compromising the function of myosin Va in INS-1E cells by RNA silencing or an inhibitory antibody against the cargo-binding site of myosin Va [32]. These manipulations led to a strong reduction in both docked DCVs and exocytotic events under sustained stimulation [32]. In another pancreatic β-cell line, MIN6 β-cells, expression of MyoVa-tail or shRNA (small hairpin RNA) against myosin Va also led to a significant decrease in the number of pre-docked vesicles and to a reduction in DCV release [8]. Taken together, and in agreement with data obtained from chromaffin cells, this points to a role of myosin Va as a classical transport motor that delivers vesicles to release sites by active movement along F-actin.

Differential localization and mobility of DCVs under resting and stimulated conditions in endocrine cells compared with neurons

Figure 2
Differential localization and mobility of DCVs under resting and stimulated conditions in endocrine cells compared with neurons

In endocrine cells, a fraction of DCVs is typically immobilized close to the PM at rest. Upon stimulation, this fraction undergoes exocytosis. Inhibition of myosin Va mostly reduces cortical restriction, mobility and exocytosis of DCV. In neurons, DCVs are highly mobile at rest. Mobile DCVs exocytose preferentially. Inhibition of myosin Va enhances exocytosis. The differences depicted between endocrine and neuronal cells require further validation. Black circles, intracellular DCVs; grey circles, exocytosing DCVs; arrowheads indicate DCV movement; grey lines, F-actin; black lines, membrane.

Figure 2
Differential localization and mobility of DCVs under resting and stimulated conditions in endocrine cells compared with neurons

In endocrine cells, a fraction of DCVs is typically immobilized close to the PM at rest. Upon stimulation, this fraction undergoes exocytosis. Inhibition of myosin Va mostly reduces cortical restriction, mobility and exocytosis of DCV. In neurons, DCVs are highly mobile at rest. Mobile DCVs exocytose preferentially. Inhibition of myosin Va enhances exocytosis. The differences depicted between endocrine and neuronal cells require further validation. Black circles, intracellular DCVs; grey circles, exocytosing DCVs; arrowheads indicate DCV movement; grey lines, F-actin; black lines, membrane.

BON cells

BON cells, derived from a human pancreatic carcinoid tumour, are endocrine-like cells that share functional similarities with intestinal enterochromaffin cells. Analysis of BON cells by evanescent wave microscopy showed that a fraction of DCVs is tethered in close proximity to the PM and does not undergo significant movements [7,33,34]. However, in contrast with chromaffin cells, stimulation of BON cells for regulated secretion causes fusion of DCVs with the PM without increasing the motility of DCVs [7]. Rather, the data indicate a ~20 nm step for the transfer of DCVs from a tethered non-fusogenic state to a docked fusogenic state upon stimulation [7]. Both expression of MyoVa-tail and down-regulation of myosin Va expression by siRNA, contrasting with the situation in PC12 cells, led to a reduced secretory response of these cells [7]. This decrease was paralleled by a reduced number of immobile DCVs at the PM [7]. Furthermore, it was shown that myosin Va silencing reduces the number of long-lasting, but not short-lasting, docking periods rather than reducing the transport of DCVs to release sites [7]. This suggests a role for myosin Va in the docking of DCVs to the PM [7]. The idea of such a role is corroborated by in vitro experiments that show binding of myosin Va to the SNARE complex: at elevated calcium concentrations, myosin Va binds to syntaxin 1A directly [29]. Prevention of this interaction strongly reduced the exocytotic response in chromaffin cells, mainly in the sustained phase of release [29]. For enterochromaffin cells, the current view therefore indicates that myosin Va is implicated in the tethering and docking of DCVs from a reserve pool.

Neurons

Finally, for DCVs in the growth cones of hippocampal neurons and neuronal growth factor-differentiated PC12 cells, mobility and exocytosis is different from all above-mentioned cell types. First, there is no evidence of a noticeable pool of pre-docked DCVs [3537]. Instead, neuronal DCV movement is dominated by directed motion, presumably along microtubule or F-actin tracks [35], and it is a mobile DCV fraction that preferentially undergoes exocytosis [35,37] (Figure 2). Secondly, the exocytotic response of DCVs is very slow, and this delay may be caused by the absence of a docked or readily releasable pool of DCVs in neurons and PC12 cells [35,38,39]. In neurons, MyoVa-tail expression inhibited retrograde axonal DCV transport, suggesting that it may act as a classical vesicle-transport motor [40]. In contrast with all other cell types investigated, the expression of MyoVa-tail enhanced the secretory response of DCVs in cultured hippocampal neurons [41]. A similar effect was seen when F-actin was depolymerized, suggesting that myosin Va may tether DCVs to actin filaments at rest and release them upon stimulation so as to allow fusion with the PM [41].

In summary, it appears that myosin Va commonly acts in conjunction with the F-actin cytoskeleton to control DCV exocytosis. The effect of this control differs among cell types and probably depends on the biological function of the substances released. Myosin Va could restrict reserve pool DCVs to F-actin under resting conditions and move DCVs along F-actin when required either at rest or during stimulation, depending on the cell type. After docking to the PM, myosin Va may be separated from DCVs [42] or from F-actin in order to allow DCVs to undergo exocytosis.

Myosin Va-interaction partners

The cell-type-specific impact of myosin Va manipulation favours the view of a regulatory rather than a direct role of myosin Va in exocytosis, which must depend on additional, cell-type-specific, factors. Important factors are the structure and dynamics of the F-actin cortex at rest and during stimulation that are variable between cell types and have a great impact on exocytosis. Other important factors are regulatory proteins that affect myosin Va function that may be differentially expressed. Moreover, myosin Va itself exists in different splice variants that affect the vesicle-binding domain and thereby the choice of regulatory proteins. The alternatively spliced exon F is, for example, expressed in melanocytes and recognized by the vesicle protein melanophilin/Slac [Slp (synaptogmin-like) lacking C2 domains] 2a [43]. This interaction is regulated by a protein of the RabGTPase family, Rab27a [44], which is also found on insulin granules and regulates exocytosis [45]. In neurons, myosin Va does not express exon F [46], and therefore another protein combination should link myosin Va to DCVs in neurons. In PC12 cells, Rab27 and Rab3 are targeted to DCVs and are necessary for the correct localization of DCVs [47]. The linker proteins that connect the Rab proteins with myosin Va in this cell type are not known, but likely candidates are proteins related to melanophilin/Slac2a, namely long proteins of the Slp or Slac type that bind myosin Va [32,43], cytoskeletal elements [48], cell signalling molecules and the exocytotic machinery (for a review, see [49]). Particularly, the Rab3- or Rab27-interacting proteins rabphilin, RIM2 (Rab3-interacting molecule 2), MyRIP (myosin VIIA- and Rab-interacting protein), granuphilin and Noc2 were shown to affect exocytosis (for a review, see [49]). An antibody that prevents the interaction of myosin Va with MyRIP/Slac2c was shown to inhibit the secretory response of INS-1E cells [32]. Furthermore, cells of both Noc2- and Rab3D-knockout mice have abnormally large DCVs [50,51]. As DCV size is likely to be determined during DCV biogenesis or maturation, these observations make Noc2, together with Rab3D, prime candidates in the search for determinants of myosin Va-regulated DCV biogenesis and maturation. Together, different myosin Va-interaction partners may specify the myosin Va function in different cell types and at different developmental stages of DCVs (Figure 1).

Conclusions

In summary, current data support the model that myosin Va is essential for the maintenance, mobilization and exocytosis of DCVs in many cell types, where it plays diverse roles: myosin Va regulates the maturation of DCVs and it can either perform F-actin-dependent movements or restrict DCV movement in resting cells (Figure 1). During stimulated exocytosis, myosin Va transports vesicles towards release sites and can replenish releasable DCVs from the reserve pool. Finally, it may be involved directly in the docking of DCVs to SNARE components in the PM. Increasing evidence points to proteins of the Rab family (Rab3 and Rab27) and of differentially expressed Rab effectors as potential regulators of the function of myosin Va in a manner that corresponds to the cell-type-specific needs during DCV maturation and exocytosis.

Molecular Mechanisms in Exocytosis and Endocytosis: 7th Junior Academics Meeting, an Independent Meeting held at University of Edinburgh, Edinburgh, U.K., 5–7 April 2009. Organized and Edited by Rolly Wiegand (Edinburgh, U.K.).

Abbreviations

     
  • ARF

    ADP-ribosylation factor

  •  
  • DCV

    dense-core vesicle

  •  
  • F-actin

    filamentous actin

  •  
  • IDCV

    immature DCV

  •  
  • MyoVa-tail

    a truncated tail construct of myosin Va comprising the organelle-binding globular tail, but not the F-actin-binding motor domain

  •  
  • GGA3

    Golgi-associated γ-adaptin ear homology domain ARF-interacting protein 3

  •  
  • MyRIP

    myosin VIIA- and Rab-interacting protein

  •  
  • PM

    plasma membrane

  •  
  • siRNA

    small interfering RNA

  •  
  • Slp

    synaptotagmin-like

  •  
  • Slac

    Slp lacking C2 domains

  •  
  • SNARE

    soluble N-ethylmaleimide-sensitive fusion protein-attachment protein receptor

Funding

T.K., C.M.B. and H.-H.G. thank the University of Bergen and the Norwegian Research Council for their generous support.

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