Two cAMP-dependent pathways differentially regulate exocytosis of large dense-core and small vesicles in mouse β-cells

Authors

  • Hiroyasu Hatakeyama,

    1. 1Division of Biophysics, Center for Disease Biology and Integrative Medicine, Faculty of Medicine, University of Tokyo, Bunkyo-ku, Tokyo, 113-0033 Japan
    2. 2Center for NanoBio Integration, University of Tokyo, Bunkyo-ku, Tokyo, 113-0033 Japan
    3. 3Department of Cell Physiology, National Institute for Physiological Sciences, and Graduate University of Advanced Studies (SOKENDAI), Myodaiji, Okazaki, 444-8585 Japan
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  • Noriko Takahashi,

    1. 1Division of Biophysics, Center for Disease Biology and Integrative Medicine, Faculty of Medicine, University of Tokyo, Bunkyo-ku, Tokyo, 113-0033 Japan
    2. 2Center for NanoBio Integration, University of Tokyo, Bunkyo-ku, Tokyo, 113-0033 Japan
    3. 3Department of Cell Physiology, National Institute for Physiological Sciences, and Graduate University of Advanced Studies (SOKENDAI), Myodaiji, Okazaki, 444-8585 Japan
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  • Takuya Kishimoto,

    1. 1Division of Biophysics, Center for Disease Biology and Integrative Medicine, Faculty of Medicine, University of Tokyo, Bunkyo-ku, Tokyo, 113-0033 Japan
    2. 2Center for NanoBio Integration, University of Tokyo, Bunkyo-ku, Tokyo, 113-0033 Japan
    3. 3Department of Cell Physiology, National Institute for Physiological Sciences, and Graduate University of Advanced Studies (SOKENDAI), Myodaiji, Okazaki, 444-8585 Japan
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  • Tomomi Nemoto,

    1. 3Department of Cell Physiology, National Institute for Physiological Sciences, and Graduate University of Advanced Studies (SOKENDAI), Myodaiji, Okazaki, 444-8585 Japan
    2. 4Section of Information Processing, Center for Brain Experiment, National Institute for Physiological Sciences, and Graduate University of Advanced Studies (SOKENDAI), Myodaiji, Okazaki, 444-8585 Japan
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  • Haruo Kasai

    1. 1Division of Biophysics, Center for Disease Biology and Integrative Medicine, Faculty of Medicine, University of Tokyo, Bunkyo-ku, Tokyo, 113-0033 Japan
    2. 2Center for NanoBio Integration, University of Tokyo, Bunkyo-ku, Tokyo, 113-0033 Japan
    3. 3Department of Cell Physiology, National Institute for Physiological Sciences, and Graduate University of Advanced Studies (SOKENDAI), Myodaiji, Okazaki, 444-8585 Japan
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Corresponding author H. Kasai: Division of Biophysics, Center for Disease Biology and Integrative Medicine, Faculty of Medicine, University of Tokyo, Bunkyo-ku, Tokyo, 113-0033 Japan. Email: hkasai@m.u-tokyo.ac.jp

Abstract

It has been reported that cAMP regulates Ca2+-dependent exocytosis via protein kinase A (PKA) and exchange proteins directly activated by cAMP (Epac) in neurons and secretory cells. It has, however, never been clarified how regulation of Ca2+-dependent exocytosis by cAMP differs depending on the involvement of PKA and Epac, and depending on two types of secretory vesicles, large dense-core vesicles (LVs) and small vesicles (SVs). In this study, we have directly visualized Ca2+-dependent exocytosis of both LVs and SVs with two-photon imaging in mouse pancreatic β-cells. We found that marked exocytosis of SVs occurred with a time constant of 0.3 s, more than three times as fast as LV exocytosis, on stimulation by photolysis of a caged-Ca2+ compound. The diameter of SVs was identified as ∼80 nm with two-photon imaging, which was confirmed by electron-microscopic investigation with photoconversion of diaminobenzidine. Calcium-dependent exocytosis of SVs was potentiated by the cAMP-elevating agent forskolin, and the potentiating effect was unaffected by antagonists of PKA and was mimicked by the Epac-selective agonist 8-(4-chlorophenylthio)-2′-O-methyl cAMP, unlike that on LVs. Moreover, high-glucose stimulation induced massive exocytosis of SVs in addition to LVs, and photolysis of caged cAMP during glucose stimulation caused potentiation of exocytosis with little delay for SVs but with a latency of 5 s for LVs. Thus, Epac and PKA selectively regulate exocytosis of SVs and LVs, respectively, in β-cells, and Epac can regulate exocytosis more rapidly than PKA.

Neurons and secretory cells have two types of secretory vesicles, large dense-core vesicles (LVs) and small vesicles (SVs), both of which undergo Ca2+-dependent exocytosis (Kelly, 1993; Kasai, 1999). It has been reported in many preparations that Ca2+-dependent exocytosis is facilitated by cytosolic cAMP (Takahashi et al. 1999; Tang et al. 2005), and such actions of cAMP are mediated either by protein kinase A (PKA) or by exchange proteins directly activated by cAMP (Epac; Sedej et al. 2005; Seino & Shibasaki, 2005). It has, however, never been clarified how regulation of exocytosis by cAMP differs depending on PKA or Epac, and on the types of vesicles. For example, in pancreatic β-cells, both LVs, containing insulin, and SVs, containing GABA (Thomas-Reetz & De Camilli, 1994), are known to undergo Ca2+-dependent exocytosis (Kanno et al. 2004; MacDonald et al. 2005). Such exocytosis was reported to be facilitated by cAMP using membrane capacitance measurements (Ammäläet al. 1993; Renström et al. 1997; Eliasson et al. 2003). It has, however, been difficult to evaluate the effects of cAMP selective to LVs and SVs, because whole-cell capacitance measurements are not readily able to distinguish between the vesicle types (Takahashi et al. 1997; Braun et al. 2004).

To investigate physiological exocytosis, we have developed an approach based on two-photon imaging of secretory preparations immersed in a solution containing highly polar fluorescent tracers (Kasai et al. 2006). Such two-photon extracellular polar-tracer (TEP) imaging has allowed quantification of exocytosis and endocytosis in pancreatic acini (Nemoto et al. 2001; Thorn & Parker, 2005), pancreatic islets (Takahashi et al. 2002; Hatakeyama et al. 2006), adrenal medulla (Kishimoto et al. 2006) and PC12 cells (Kishimoto et al. 2005; Liu et al. 2005). These studies demonstrated that TEP imaging is capable of detecting most exocytic events in intact secretory tissues in a quantitative manner. Moreover, we have developed TEP imaging-based quantification (TEPIQ) analysis, with which it is possible to estimate the diameter of secretory vesicles, even though such vesicles may be smaller than the optical resolution of a two-photon microscope (Kasai et al. 2006). Indeed, we have visualized exocytosis of SVs with a diameter of 55 nm in PC12 cells and shown that these vesicles undergo exocytosis at a rate more than 10 times as fast as that of LVs (Liu et al. 2005).

We have now investigated exocytosis in pancreatic β-cells with TEPIQ analysis. We detected marked Ca2+-dependent exocytosis of SVs with a mean diameter of 80 nm in addition to exocytosis of LVs. The diameter of SVs was confirmed by electron microscopy with photoconversion of diaminobenzidine (DAB). Exocytosis of SVs occurred with a time constant of 0.3 s, whereas that of LVs showed a time constant of > 1 s. Although cAMP markedly potentiated exocytosis of both LVs and SVs, this effect depended on PKA only for LVs and on Epac for SVs. Furthermore, we have applied photolysis of caged cAMP to quantify the speed of cAMP action during high-glucose stimulation, and found that the augmentation of exocytosis by cAMP occurred within a fraction of a second for SVs but with a delay of 5 s for LVs. Thus, we have, for the first time, definitively identified exocytosis of SVs in β-cells, and demonstrated that two cAMP-dependent pathways mediated by Epac and PKA can selectively regulate exocytosis of SVs and LVs, respectively, and that cAMP can regulate exocytosis more rapidly with Epac than with PKA.

Methods

Cell preparations

Eight- to 12-week-old ICR mice (male, Japan SLC; Hamamatsu, Japan) were killed by cervical dislocation. Animal experiments were performed in accordance with the regulations of the Faculty of Medicine, the University of Tokyo, Japan. Pancreatic islets were isolated by collagenase digestion, and small cell clusters (Takahashi et al. 2004; Hatakeyama et al. 2006) or single-cell suspensions were obtained from the islets by trituration (Takahashi et al. 1997). Single β-cells were studied for quantification of kinetics and the extent of SV exocytosis in the experiments shown in Figs 1 and 5 because of their limited diffusion barrier for FM1-43 (Invitrogen, Carlsbad, CA, USA). Islet cell clusters with intact intercellular space were studied for characterization of LV exocytosis in Fig. 2, for estimation of vesicle diameter in Figs 3 and 4, and for stimulation with high glucose in Figs 6 and 7. We studied β-cells in the second layer of islet cell clusters to minimize the possible diffusion barrier imposed by the intercellular space. Cells were cultured for 1–24 h in a humidified atmosphere of 5% CO2/95% air at 37°C in Dulbecco's Modified Eagle's medium (DMEM) containing glucose (1.0 mg ml−1) and supplemented with 10% fetal bovine serum, penicillin (100 μU ml−1) and streptomycin (100 mg ml−1). For experiments, the cells were transferred to a glass-bottomed recording chamber (thickness, 0.1 mm; Matsunami-glass, Osaka, Japan) and immersed in a solution (SolA) containing (mm): 150 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2, 10 Hepes-NaOH (pH 7.4) and 2.8 glucose at 310 mosmol l−1. A SolA-based solution containing 20 mm glucose was prepared by adjusting the osmolarity to 310 mosmol l−1 with deionized water. For some single-cell experiments, we selected only cells with a diameter of > 10 μm so that > 90% of the cells would be expected to be β-cells (Rorsman & Trube, 1986). Forskolin (Sigma, St Louis, MO, USA) was dissolved in DMSO at 10 mm and subsequently diluted in SolA. Myristoylated PKA inhibitor amide 14–22 (PKI; Calbiochem, San Diego, CA, USA), the Rp isomer of adenosine 3′,5′-cyclic monophosphorothioate (Rp-cAMPS, Calbiochem), and 8-(4-chlorophenylthio)-2′-O-methyl cAMP (8-CPT-2′-O-Me-cAMP, Biolog, Bremen, Germany) were dissolved directly in SolA.

Figure 1.

TEP imaging of exocytosis and endocytosis with FM1-43 in isolated β-cells
A, FM1-43 fluorescence (F) images of a cell loaded with NPE AM are shown in panels a1–a4. Photolysis was induced at a time 0 between panels a1 and a2 (UV). The dye was washed out 30 s before panel a4. B, difference images (ΔF) obtained by subtracting the image at rest (a1) from panels a1–a3 in A are shown in panels b1–b3, respectively. C, increase in [Ca2+]i induced by photolysis of NPE in cells loaded with the Ca2+ indicator fura-2FF. D, time course of the change in FM1-43 fluorescence induced by photolysis of NPE during line scanning along the dashed line shown in the inset. Average values from six cells (grey trace) and the single-exponential fit (black line) are shown. E, time course of staining of the plasma membrane of a cell by rapid application of FM1-43. Average values are from 5 cells. F, time course of the change in FM1-43 fluorescence for the entire section of the cell shown in A. The zero level of fluorescence was obtained before application of FM1-43 to the cell. Fluorescence was normalized relative to that of the entire section of the cell before photolysis and is expressed as a percentage of the control value (Fnormalized). G, time courses of the changes in fluorescence in the plasma membrane (PM) region (blue) and in the cytoplasmic (cyt) region (red) of the cell depicted in the inset.

Figure 5.

Pharmacology of Ca2+-dependent exocytosis of SVs in β-cells
A, time courses of FM1-43 fluorescence after photolysis of NPE in representative β-cells either maintained under control conditions or pretreated with forskolin (2 μm) for 10 min. B, cells were exposed (or not) to PKI (5 μm) or Rp-cAMPS (200 μm) for 30 min before incubation for 10 min with forskolin (2 μm) or 8-CPT-2′-O-Me-cAMP (10 μm), as indicated. They were then stimulated by NPE photolysis, and the diffuse increase in FM1-43 fluorescence was measured. Circles represent data obtained from individual cells, with columns and error bars indicating means ±s.e.m. The dashed line shows the control level (NPE photolysis alone). Statistical analyses were performed with the Kruskal–Wallis test (P < 0.001) followed by the Mann–Whitney U test for comparison with control values. ***P < 0.001.

Figure 2.

Kinetics of LV exocytosis in β-cells present within islet cell clusters as revealed by TEP imaging of FM1-43 and SRB
A and B, time courses of the changes in fluorescence of FM1-43 and SRB associated with a discrete exocytic event appearing in the centre of the images. Time 0 was set as the onset of the increase in FM1-43 fluorescence. Fluorescence intensities of FM1-43 and SRB were converted to membrane area and volume of vesicles by TEPIQ analysis of ΔS and ΔV, respectively. Arrow in B indicates the time difference between the onsets of the FM1-43 and SRB signals. C and D, cumulative latency distributions of LV exocytic events induced by photolysis of NPE. The time scale in C is expanded in D. Latency of exocytic events was estimated with either SRB (black) or FM1-43 (blue) in 4 cells and is expressed as events per cell. τ, time constant.

Figure 3.

Vesicle diameters estimated by TEPIQ analysis
A, TEP image for SRB in an islet cell cluster. B, difference TEP images for SRB (upper) and FM1-43 fluorescence (lower) obtained before and after NPE photolysis at time 0. A discrete exocytic event is indicated by the white arrow. The circular region (h) indicates the area containing the discrete exocytic event, whereas the boxed region (i) contains only a diffuse increase in fluorescence. C, a ΔFSRBFFM1-43 ratio image obtained 6.33 s after UV irradiation for the cell shown in B. A pseudocolour coding, displayed below, is used to represent the diameter of vesicles estimated by TEPIQ analysis of ΔVS. D, time courses of the changes in ΔV and ΔS for the circular region (h) shown in B. Double-headed arrows p1 and p2 denote the time periods before and after the appearance of the discrete fluorescent spot, respectively. E, time courses of changes in vesicle diameter obtained from TEPIQ analysis of the data in D. F and G, time courses of the changes in ΔV and ΔS (F) and in ΔVS-TEPIQ diameter of vesicles (G) for the boxed region (i) in B.

Figure 4.

Ultrastructural identification of endocytic vesicles in β-cells
Endocytic vesicles were examined by electron microscopy in cells loaded with FM1-43FX. Photoconversion of DAB was induced by fluorescence of FM1-43FX remaining after extensive washout. Some FM1-43FX molecules remained in the plasma membrane (PM) despite such washout, resulting in its staining with DAB (long filled arrows). A, a cluster of cells immersed in a solution containing FM1-43FX for 30 min at rest. Open arrows, open arrowheads and filled arrowheads indicate LVs, constitutive endocytic vesicles, and lysosomes or endosomes, respectively. B, a cluster of cells immersed in a solution containing FM1-43FX for 90 s without stimulation. M, mitochondria. C, a cluster of cells that was immersed in a solution containing FM1-43FX for 1 min before photolysis of NPE and was fixed with glutaraldehyde within ∼15 s after photolysis. Short filled arrows indicate small endocytic vesicles. The external scale bar (500 nm) applies to all panels with the exception of the insets in A and C, which show magnified images of DAB-positive endocytic vesicles and for which the associated scale bars represent 50 nm.

Figure 6.

Induction of SV exocytosis by high-glucose stimulation
A, time course of the increase in [Ca2+]i induced by a high concentration (20 mm) of glucose in a cluster of islet cells loaded with the Ca2+ indicator fura-4F. The high-glucose solution was applied at time 0. B, time course of the increase in FM1-43 fluorescence in a cluster of islet cells exposed to 20 mm (black) or 2.8 mm glucose (blue) at time 0. FM1-43 was applied to the cell for 1 min before stimulation. C, SRB and SRB/FM1-43 ratio images of a cluster of islet cells stimulated with high glucose for 200 s. Diameters of vesicles are represented by the pseudocolour coding indicated in Fig. 3C.

Figure 7.

Potentiation of high-glucose-induced exocytosis by photolysis of caged cAMP
A, time course of the increase in [Ca2+]i induced by application of a high concentration (20 mm) of glucose at time 0 in a cluster of islet cells loaded with the Ca2+ indicator fluo-5F. Photolysis of caged cAMP (UV) was induced 200 s after glucose stimulation. B, cumulative events of LV exocytosis (revealed by discrete SRB fluorescent spots) before and after photolysis of caged cAMP performed at time 0, 200 s after the onset of stimulation with high glucose. Straight lines represent linear regression lines for two portions of the black trace. Red circles show data obtained from cells pretreated with Rp-cAMPS (200 μm) for 30 min before uncaging of cAMP. C, averaged time courses of FM1-43 fluorescence before and after UV irradiation at time 0 in control cells (n= 4, thin line) or in cells loaded with caged cAMP (n= 5, thick line). The cells were exposed to high glucose 200 s before UV irradiation. Bleaching of FM1-43 fluorescence is evident in the control cells (arrowhead). D, corrected time course for the effect of uncaging of cAMP on high-glucose-stimulated SV exocytosis obtained by subtracting the control trace from the experimental trace shown in C. E, time course of SV exocytosis in cells incubated for 30 min in the absence (control) or presence of Rp-cAMPS (200 μm) before exposure to high glucose and photolysis of caged cAMP.

TEP imaging and TEPIQ analysis

TEP imaging was performed with an inverted microscope (IX70, Olympus, Tokyo, Japan) and a laser scanner (FV300, Olympus) equipped with a water-immersion objective lens (UPlanApo60xW/IR, numerical aperture of 1.2) and a mode-locked femtosecond-pulse laser (Tsunami, Spectra Physics, Mountain View, CA, USA) as described by Kasai et al. (2005). The laser power at the specimen was 3–10 mW, and two-photon excitation was performed at 830 or 850 nm. All fluorophores and caged compounds were obtained from Invitrogen with the exception that fura-2FF was from TEF Laboratories (Austin, TX, USA). The fluorescence of FM1-43, FM1-43FX, fura-4F, fluo-5F, or fura-2FF was measured at 400–550 nm, and that of sulforhodamine B (SRB) at 570–700 nm. Fluorescence was detected with a photomultiplier in the microscope, and images were acquired every 0.3–5.0 s at a plane 5–20 μm from the glass coverslip. The 12-bit images were analysed and colour-coded with ‘fall’ or custom-made look-up tables of the image acquisition and analysis software, either Fluoview of the FV300 microscope or IPLab Spectrum (BD Biosciences Bioimaging, Rockville, MD, USA). All experimental procedures were performed under illumination with yellow light (FL40S-Y-F, National, Osaka, Japan) to prevent unintentional photolysis of caged compounds. In the experiment shown in Fig. 1E, FM1-43 was added rapidly (< 5 ms) to the cells with a piezoelectric device (ASB003A, NEC TOKIN, Tokyo, Japan). TEPIQ analysis was performed as described, with the effeciency of focal illumination pxy(0) = 0.57 and conversion coeffecient mC= 0.28 (Kasai et al. 2005). Fluorescence intensities of solutions containing FM1-43 [20 μm, dissolved in 40 mm 3-((3-cholamidopropyl)-dimethylammonio)-1-propanesulphonate (CHAPS)] and SRB (0.4 mm) were measured daily and remained constant for the entire experimental period of 1.5 years, indicating that the laser and photomultipliers were stable during this time. Imaging experiments were performed at room temperature (24–25°C).

Photolysis of caged compounds and monitoring of [Ca2+]i

The acetoxymethyl (AM) ester forms of fura-4F, fura-2FF, fluo-5F, and o-nitrophenyl-EGTA (NPE) were dissolved in DMSO at 2–10 mm. Cells were incubated for 30 min at 37°C in serum-free DMEM containing 10 μm fura-4F AM or fura-2FF AM, 10–25 μm NPE AM, 0.03% Cremophor EL (Sigma) and 0.1% bovine serum albumin (BSA) and were then washed with SolA. Cell-permeable 4,5-dimethoxy-2-nitrobenzyl (DMNB)-caged cAMP was dissolved in DMSO at 50 mm and loaded into cells by incubation for at least 30 min at 37°C in serum-free DMEM containing 100 μm DMNB-caged cAMP and 0.1% BSA. Changes in [Ca2+]i in cells subjected to photolysis of DMNB-caged cAMP were monitored with fluo-5F in cells that had also been loaded with 10 μm fluo-5F AM in DMEM containing 0.03% Pluronic F-127 (Invitrogen). Photolysis of caged compounds was induced with a mercury lamp (U-ULS100HG, Olympus) through a 360 nm bandpass filter. The radiation of the mercury lamp was gated with an electric shutter (IX-ESU, Olympus) with an opening duration of 10–500 ms.

Increases in [Ca2+]i induced by photolysis of NPE or high glucose were monitored as the decrease in fluorescence intensity of fura-2FF or fura-4F and were calculated as:

display math

where Kd, Fmax and Fmin represent the dissociation constant for the interaction between the Ca2+ indicator and Ca2+, the maximal fluorescence intensity and the minimal fluorescence intensity, respectively. In vivo calibration of fura-2FF was performed as previously described (Nemoto et al. 2004; Takahashi et al. 2004; Oshima et al. 2005). For fura-2FF we estimated Fmin/Fmax to be 0.1, and Kd is 31 μm. For fura-4F, we estimated Fmin/Fmax to be 0.46, and Kd is 1.16 μm (Wokosin et al. 2004). Increases in [Ca2+]i in cells loaded with caged cAMP were monitored as the increase in fluorescence intensity of fluo-5F and calculated as:

display math

For fluo-5F, we estimated Fmax/Fmin to be 3.5, and Kd is 2.3 μm.

Photoconversion of DAB and electron microscopy

Staining of islet cell clusters with FM1-43FX, an aldehyde-fixable analog of FM1-43, and subsequent photoconversion of DAB and electron-microscopic analysis were performed as previously described (Kishimoto et al. 2005; Liu et al. 2005), with slight modifications. Photolysis of NPE was induced during TEP imaging to identify the responsive cells. The cells were plated in a recording chamber thickly coated with a collagen gel (MatriGel, BD Biosciences) in order to prevent cell slippage during subsequent manoeuvres. The recording chamber (0.2 ml) was subjected to superfusion (0.1 ml s−1) with 100 mm sodium cacodylate buffer (pH 7.4, Nacalai Tesque, Kyoto, Japan) containing 2% glutaraldehyde within ∼15 s after photolysis, and superfusion was continued for at least 1 min. Cells were exposed to the fixative overnight at 4°C and were then washed first for 1 h with 100 mm glycine in phosphate-buffered saline (PBS, pH 7.4, Sigma) and then for 5 min with 100 mm NH4Cl at room temperature. After an additional brief wash with PBS (pH 7.9), the cells were incubated for 20 min with PBS containing DAB (1 mg ml−1) at pH 7.9. Fluorescence excitation (100 W mercury lamp, 475 nm) was then performed for 30 min with the cells in the freshly prepared DAB solution. The cells were subsequently washed overnight with 100 mm sodium cacodylate buffer, exposed to 2% osmium tetroxide for 90 min, dehydrated with a graded series of ethanol solutions, and embedded in EPON812 (TAAB, Reading, UK). After incubation for 2 days at 60°C, the coverslip was removed from the EPON812 with hydrogen fluoride, and silver–gold thin sections (∼80 nm) were cut horizontally relative to the surface of the coverslip and mounted on copper grids for observation with an electron microscope (JEM-1200EX, JEOL, Tokyo, Japan).

The density of DAB-positive vesicles (in μm−2) was converted into the number of vesicles per cell by multiplying by Hr3/3, where r represents the mean radius of β-cells (6 μm; Takahashi et al. 1997) and H is a correction factor (0.535) to account for the effect of thickness of the section (s= 80 nm); H is equal to s/(s+d), where d is vesicle diameter (70 nm), as previously described (Liu et al. 2005). The number of LVs undergoing exocytosis is predicted as 0.009 μm−2, assuming that the number of LV exocytic events is 20 per cell within 15s after photolysis of NPE, the thickness of the thin sections is 80 nm, and the diameters of LVs and β-cells are 350 nm and 12 μm, respectively; the corresponding value for SVs determined by electron microscopy of thin sections is ∼0.6 μm−2 (see Results).

Statistical analysis

Statistical analyses were performed as indicated in text or figure legends. A P value of < 0.05 was considered statistically significant.

Results

Ca2+-dependent exocytosis of SVs in β-cells

We performed two-photon imaging of individual mouse β-cells immersed in a solution containing the polar fluorescent tracer FM1-43 in order to detect exocytosis (Fig. 1A and B), as previously described for PC12 cells (Liu et al. 2005). The FM1-43 stains the outer leaflet of the plasma membrane, which it does not permeate (Betz & Bewick, 1992). Exocytosis was induced by photolysis of the caged-Ca2+ compound NPE, which was loaded into the cells in the form of an AM ester. Photolysis of NPE resulted in a rapid increase in [Ca2+]i, which was estimated with the Ca2+ indicator fura-2FF to be > 10 μm (Fig. 1C). Photolysis of NPE also induced a rapid and diffuse increase (ΔF) in FM1-43 fluorescence (Fig. 1A and B) of 20 ± 9% (mean ±s.d., n= 26) with a time constant of 0.32 ± 0.18 s (n= 6; Fig. 1D). These kinetics reflect those of the increase in membrane area, given that staining of the plasma membrane with FM1-43 occurred more rapidly, with a time constant of 0.05 ± 0.01 s (n= 5), when FM1-43 was quickly applied to β-cells with the use of a piezoelectric device (Fig. 1E).

Most vesicle membrane inserted into the plasma membrane by exocytosis appeared to be subsequently internalized by endocytosis. The increase in diffuse fluorescence was thus largely unaffected by washout of the tracer 30 s after stimulation (Fwash, 36 ± 13%, n= 5; Fig. 1F). Although the remaining fluorescence signal was slightly larger than the increase induced by stimulation (ΔF), this difference is likely to reflect the fact that washout of FM1-43 from the plasma membrane is incomplete in β-cells (residual fluorescence intensity of 9 ± 3%, n= 5). Given that such incomplete washout of FM1-43 is a potential source of error, we were not able to determine accurately the amount of membrane taken up by endocytosis. We detected, however, an increase in FM1-43 fluorescence in the cytoplasm (Fig. 1G), indicating that a substantial number of endocytic vesicles underwent translocation from the plasma membrane into the cytoplasm.

TEP imaging with the fluid polar-tracer SRB is able to detect exocytosis of LVs in mouse β-cells within islet cell clusters as discrete fluorescent spots (Fig. 2A and B). These LVs have a diameter of 0.35 μm (Kasai et al. 2005), and their exocytosis occurred with a time constant of > 1 s (Fig. 2C). Staining of LVs with FM1-43 often preceded that with SRB by ∼0.3 s (Fig. 2A and B), probably as a result of the slow opening of the fusion pore in β-cells (Takahashi et al. 2002). We therefore measured the time course of LV exocytosis based on the onset of the FM1-43 signal that precedes the SRB signal (Fig. 2C and D). Although LV exocytosis detected with FM1-43 was slightly faster than that detected with SRB, the overall time courses visualized with the two tracers were almost identical. We therefore concluded that exocytosis of LVs is too slow to account for the rapid increase in diffuse FM1-43 staining induced by NPE photolysis (Fig. 1D) and that the diffuse exocytic events detected by FM1-43 must be mediated mostly by vesicles other than LVs, if not all (see Figs 3 and 6).

We next estimated the diameter of the vesicles responsible for the diffuse FM1-43 staining by TEPIQ analysis. TEPIQ analysis can estimate the diameters of exocytic vesicles based on the fluorescence intensities of SRB and/or FM1-43 (Kasai et al. 2005, 2006). There are three types of TEPIQ analysis: ΔV-, ΔS- and ΔVS-TEPIQ analysis, respectively. In particular, ΔVS-TEPIQ analysis uses the fluorescence intensities of both SRB and FM1-43, and can estimate the diameter of a population of vesicles even when single exocytic events are not resolved. Given that ΔVS-TEPIQ analysis requires staining with both SRB and FM1-43, we studied regions of the cell facing the intercellular space of the islet (Fig. 3A), where the background level of SRB fluorescence is low as a result of the narrowness of the intercellular space (see Fig. 4). We found diffuse increases in FM1-43 fluorescence in islet cell clusters (Fig. 3B) similar to those observed in single β-cells (Fig. 1A and B). The baseline-subtracted images of SRB and FM1-43 fluorescence indicated that the increase in the diffuse signal occurred before the appearance of discrete signals (Fig. 3B). The ratio between SRB and FM1-43 images allows estimation of the distribution of vesicle diameters (Fig. 3C). The diameters of the vesicles responsible for the discrete spots of fluorescence were estimated from the increases in the fluorescence of SRB (ΔV-TEPIQ analysis) or FM1-43 (ΔS-TEPIQ analysis) as 0.39 ± 0.055 (n= 12) and 0.37 ± 0.074 μm (n= 12), respectively (Fig. 3D and E), consistent with our previous results (Kasai et al. 2005). The discrete signals thus reflect the exocytosis of individual LVs. The diameter of vesicles responsible for the diffuse signal apparent immediately after NPE photolysis was estimated as 0.079 ± 0.028 μm (n= 11; Fig. 3DG) by TEPIQ analysis of ΔVS. The diffuse signal in β-cells thus reflects exocytosis of many SVs, as does that in PC12 cells (Liu et al. 2005). No change in the diameter of the small vesicles was detected between 0.2 and 10 s after exocytosis (Fig. 3G), suggesting that their diameter remained the same until they underwent endocytosis.

Electron microscopy of SV exocytosis and endocytosis in β-cells

The conclusions of TEPIQ analysis were supported by electron microscopy of islet cell clusters in which DAB was photoconverted by fluorescence of an aldehyde-fixable analogue of FM1-43, FM1-43FX (Henkel et al. 1996; Harata et al. 2001; Brumback et al. 2004). As a control, we first labelled the constitutive endocytic pathway by immersing cells in FM1-43FX for 30 min without stimulation. Such treatment revealed many small DAB-positive vesicles as well as endosome- and lysosome-like structures (Fig. 4A), as observed in PC12 cells (Liu et al. 2005). In contrast, no staining of intracellular organelles was detected in cells exposed to FM1-43FX for only 90 s before fixation (Fig. 4B).

Many small DAB-positive vesicles were apparent in cells fixed within 15 s after stimulation by NPE photolysis during TEP imaging (Fig. 4C). While some DAB-positive vesicles were still docked at the plasma membrane, many were scattered in the cytoplasm (Fig. 4C), consistent with the results obtained with TEP imaging (Fig. 1G). The diameter of the stained vesicles was 0.070 ± 0.011 μm (n= 41), similar to the value estimated by ΔVS-TEPIQ analysis. The number of DAB-positive vesicles was 0.6 ± 0.3 μm−2, corresponding to a total of 4000 ± 1900 vesicles per cell (see Methods) or to 13 ± 6% of the original area of the plasma membrane. These results are consistent with the observed diffuse fluorescence increase of 20 ± 9% and with the fact that most of the membrane added to the cell surface by exocytosis was captured by endocytosis (Fig. 1F). We thus demonstrated the occurrence of marked exocytosis of SVs in β-cells with both TEPIQ analysis and electron microscopy. Staining of docked LVs was never seen, unlike PC12 cells (Kishimoto et al. 2005), confirming that the diffuse signal was not mediated by transient opening of the fusion pore of LVs. We considered the diffuse FM1-43 signal to reflect selectively the exocytosis of SVs in the following experiments.

Exocytic images of LVs were rarely captured by electron microscopy, even though exocytosis of LVs was detected by TEP imaging. This discrepancy is, however, consistent with our estimation of the rate of LV exocytic events as 20 per cell within 15 s after photolysis of NPE (Fig. 2C), which predicts the number of LVs undergoing exocytosis in the thin sections required for electron microscopy to be 0.009 μm−2 (see Methods). This value is only 1.5% of the corresponding value for SVs (0.6 μm−2). It would thus be expected to be difficult to detect LV exocytosis in β-cells by electron microscopy (Orci et al. 1973). The fact that SV exocytosis was readily identified (Fig. 4C) underscores our conclusion that the extent of SV exocytosis is far greater than that of LV exocytosis in β-cells.

Regulation of Ca2+-dependent SV exocytosis by cAMP

We next examined the effect of cAMP on the Ca2+-dependent exocytosis of SVs. Forskolin, an activator of adenylate cyclase that increases the cytosolic concentration of cAMP, increased by 39 ± 15% (n= 28; P < 0.001, Mann–Whitney U test) the extent of SV exocytosis induced by photolysis of NPE and detected by measurement of the diffuse FM1-43 fluorescence signal (Fig. 5A). The potentiation of SV exocytosis by cAMP was not blocked by antagonists of PKA, including PKI and Rp-cAMPS (Fig. 5B). Certain PKA-independent cAMP signalling is mediated by guanine nucleotide exchange factors that are directly activated by cAMP (Epac) and which are selectively activated by 8-CPT-2′-O-Me-cAMP at 10 μm (Enserink et al. 2002). We found that 8-CPT-2′-O-Me-cAMP at 10 μm mimicked the effect of forskolin on SV exocytosis (Fig. 5B), suggesting that the augmentation of Ca2+-dependent exocytosis of SVs by cAMP is dependent on Epac, but not on PKA.

These properties of SV exocytosis contrast with those of LV exocytosis characterized in our previous study (Hatakeyama et al. 2006). First, Ca2+-dependent exocytosis of LVs was significantly enhanced by forskolin at a high glucose concentration (20 mm) but not at the low glucose concentration (2.8 mm) at which SV exocytosis was shown here to be potentiated by forskolin. Second, the effect of forskolin was inhibited by PKI. Third, 8-CPT-2′-O-Me-cAMP at 10 μm did not increase the extent of LV exocytosis at either concentration of glucose. These observations thus indicate that Epac and PKA selectively regulate Ca2+-dependent exocytosis of SVs and LVs, respectively, in β-cells.

Regulation by cAMP of SV or LV exocytosis induced by high glucose

Exocytosis of SVs was also induced by exposure of β-cells to a high glucose concentration (20 mm). High-glucose stimulation, which increased [Ca2+]i with a latency of ∼100 s (Fig. 6A), thus also induced an increase in diffuse FM1-43 fluorescence in synchrony with the increase in [Ca2+]i (Fig. 6B). TEPIQ analysis of ΔVS revealed that the vesicles responsible for this exocytosis were SVs with a diameter of 0.075 ± 0.016 μm (n= 10; Fig. 6C). No LV exocytic events were detected in the image shown in Fig. 6C.

We then examined the effects of cAMP on exocytosis of LVs and SVs by photolysis of caged cAMP during glucose stimulation. Cells were loaded with the cell-permeable DMNB-caged cAMP analogue (100 μm), the photolysis of which was triggered after the glucose-induced increase in [Ca2+]i. We chose fluo-5F to measure [Ca2+]i for these experiments because UV-induced uncaging of cAMP did not result in bleaching of this Ca2+ indicator. Photolysis of caged cAMP did not affect [Ca2+]i in our experimental conditions (n= 5; Fig. 7A), but it potentiated exocytosis of LVs with a latency of 4.8 ± 2.6 s (n= 19; Fig. 7B). This latter effect of photolysis of caged cAMP was prevented by Rp-cAMPS (n= 6; Fig. 7B), consistent with the notion that the effect of cAMP on LV exocytosis is mediated by PKA. Photolysis of caged cAMP did not induce LV exocytosis in the absence of glucose stimulation (n= 3; data not shown).

Photolysis of caged cAMP also potentiated high-glucose-stimulated exocytosis of SVs (Fig. 7C). Ultraviolet irradiation in the absence of caged cAMP had no such effect but revealed bleaching of FM1-43 (Fig. 7C). We therefore corrected for the effect of bleaching by subtraction of the trace obtained in the absence of caged cAMP (Fig. 7D), revealing that the photolysis-induced augmentation of SV exocytosis occurred with little delay. A linear regression analysis of the effect of cAMP between 0 and 20 s suggested that such latency was 0.3 s at most (n= 6; 95% upper confidence limit, 2.4 s). The potentiating effect of the uncaging of cAMP was not substantially altered by Rp-cAMPS (n= 6; Fig. 7E), consistent with the notion that the effect of cAMP on SV exocytosis is mediated by Epac. Photolysis of caged cAMP did not induce SV exocytosis in a low-glucose solution (n= 3; data not shown). These data thus suggested that cAMP regulates Ca2+-dependent exocytosis more rapidly through Epac than it does through PKA.

Discussion

Differential regulation of exocytosis of LVs and SVs by cAMP

We found that two distinct cAMP pathways, mediated by either PKA or Epac, selectively regulate exocytosis of LVs or SVs, respectively, in β-cells. In addition to the pharmacological distinctions presented, two important differences between the two pathways, consistent with their molecular mechanisms, were apparent. First, augmentation of LV exocytosis by cAMP required a high concentration of glucose, whereas that of SV exocytosis did not, possibly because of the requirement for a high cytosolic concentration of ATP for the phosphorylation reaction in the PKA pathway (Birnbaumer et al. 1969; Ho & Sutherland, 1975). Epac is a guanine nucleotide exchange factor and so may be less dependent on ATP concentration. Second, potentiation of SV exocytosis by cAMP was faster than that of LV exocytosis. The latency for augmentation of LV exocytosis (∼5 s) may reflect the time required for phosphorylation by activated PKA. In contrast, the action of Epac may be faster because it requires only nucleotide exchange, which occurs within a fraction of a second (John et al. 1990; Itzen et al. 2007). Epac has been proposed to regulate exocytosis through direct binding to Rim2 (Ozaki et al. 2000). Rim proteins are putative effectors of Rab3 and are thought to serve as Rab3-dependent regulators of synaptic vesicle fusion (Wang et al. 1997), a role that they may also play in exocytosis of SVs in β-cells.

Small vesicle exocytosis in β-cells

Using TEP imaging, we have here demonstrated that massive exocytosis of SVs precedes that of LVs in β-cells stimulated with photolysis of NPE, consistent with a previous study based on amperometric and membrane capacitance measurements (Takahashi et al. 1997). TEP imaging has also revealed that the rapid capacitance increase was mediated by exocytosis of SVs in PC12 cells (Liu et al. 2005). Similar rapid capacitance increases were reported in mast cells (Kirillova et al. 1993), fibroblasts (Coorssen et al. 1996; Ninomiya et al. 1996), pancreatic acinar cells (Ito et al. 1997) and adrenal chromaffin cells (Ninomiya et al. 1997; Haller et al. 1998). Thus, mammalian cells may commonly possess numerous SVs which can undergo rapid Ca2+-dependent exocytosis.

We found massive exocytosis of SVs in β-cells even during glucose stimulation. If we assume that the diameters of SVs and β-cells are 80 nm and 12 μm, respectively, an increase in FM1-43 fluorescence of 3% per cell min−1 represents exocytosis of 675 SVs per cell min−1, a rate that is more than 100-fold greater than that of 6.4 LVs per cell min−1 previously determined for LV exocytosis (Hatakeyama et al. 2006). Exocytosis of SVs in β-cells is therefore likely to play an important functional role. Although SVs in β-cells contain GABA (Thomas-Reetz & De Camilli, 1994), fewer than 100 quantal GABA-induced currents were detected after photolysis of a caged-Ca2+ compound in β-cells expressing recombinant GABAA receptors (Braun et al. 2004), whereas we detected exocytosis of ∼4500 SVs in response to such photolysis by TEPIQ analysis (20% increase in membrane area) and 4000 endocytic vesicles by electron microscopy. These observations suggest that GABA is present in, at most, only ∼1% of SVs in β-cells, a situation similar to that in PC12 cells, in which only a small proportion of SVs contain detectable acetylcholine (Ninomiya et al. 1997; Liu et al. 2005). Furthermore, although the number of SV exocytic events is large, the release of transmitter from SVs is likely to be inefficient, given that the volume of these vesicles is only ∼1%[(80/350)3] of that of LVs and that the total volume secreted by SVs (0.18 μm3 per cell min−1) is similar to that secreted by LVs (0.14 μm3 per cell min−1). The action of neurotransmitters released from SVs is likely to be transient and limited to the site of release, owing to both the small volume of the vesicles and the low affinity of neurotransmitter receptors relative to that of hormone receptors.

Trafficking of proteins and lipids between the plasma membrane and endosomes is another possible function of SV exo-endocytosis. Small vesicles are likely to contribute to trafficking more efficiently than they do to secretion as a result of their large surface-to-volume ratio. Membrane area added to the plasma membrane by SV exocytosis was ∼3% per cell min−1 during glucose stimulation, a rate about five times as large as that for LV exocytosis (0.54% per cell min−1). Small vesicles may play a similar role in ‘non-secretory’ cells that exhibit a substantial amount of SV exocytosis (Steinhardt et al. 1994; Borgonovo et al. 2002; McNeil & Steinhardt, 2003). The fusion pore of SVs in β-cells appears to expand to an extent that allows lipids and proteins to diffuse along its lateral wall. This conclusion is based on our findings that ΔVS-TEPIQ analysis and electron microscopy yielded similar values for SV diameter, indicating that SVs were stained not only with FM1-43 but also with SRB and that the fusion pore is > 1.4 nm in diameter. Such fusion pores were shown to be lipidic and to allow diffusion of lipids and proteins in β-cells (Takahashi et al. 2002).

Trafficking of molecules by SV exocytosis precedes LV exocytosis, given that the actions of both Ca2+ and cAMP were faster for SVs than for LVs. Exocytosis of SVs may thus precondition the plasma membrane for exocytosis and endocytosis of LVs. For example, a sialylated form of the neural cell adhesion molecule NCAM (PSA-NCAM) is expressed specifically in β-cells and is mobilized to the cell surface in an activity-dependent manner (Bernard-Kargar et al. 2001; Kiss et al. 1994). Moreover, surface expression of PSA-NCAM correlates with glucose-stimulated insulin secretion (Bernard-Kargar et al. 2001). In pancreatic islets, NCAM is thought to contribute to maintenance of cell–cell interactions and is required for normal turnover of secretory granules (Langley et al. 1989; Esni et al. 1999). Increased surface expression of PSA-NCAM might therefore facilitate contact between β-cells and other islet cells in order to preserve islet integrity in the face of secretion of reactive substances stored in insulin granules, for example, insulin, Zn2+ ions, protons, ATP, GABA, carboxypeptidase E and islet amyloid polypeptide (Hutton et al. 1983; Hutton, 1989; Gammelsaeter et al. 2004). Impairment of such preconditioning might result in islet dysfunction.

We have examined exocytosis and fusion pores in the plasma membrane of β-cells facing the intercellular space within islets of Langerhans, which is the physiological site of exocytosis. This is important because the properties of fusion pores are markedly affected by experimental conditions. For instance, membrane surface tension plays a critical role in fusion pore expansion (Monck & Fernandez, 1996). Capacitance measurements in the cell-attached configuration can resolve the fusion pore of small vesicles; however, they should therefore be interpreted with caution (MacDonald et al. 2006), because the patch membrane is mechanically distorted. In addition, it is not possible to estimate fusion pore diameter by capacitance measurements without assuming other pore dimensions, such as length, about which limited information is available. Capacitance measurements also do not allow estimation of the diameter of fusion pores larger than 2 nm for SVs (Kasai et al. 2006) and tend to underestimate fusion pore size. In contrast, TEPIQ analysis allows measurement of the size of fusion pores without assumptions of pore geometry and is able to estimate pore sizes as large as 20 nm (Kishimoto et al. 2006).

Our TEP imaging is likely to detect most exocytic events of LVs and SVs in the plane of focus in β-cells. The number of LV exocytic events (6.4 per cell min−1) detected by TEP imaging was consistent with that estimated by radioimmunoassay during glucose stimulation (Hatakeyama et al. 2006). Rapid increases in FM1-43 fluorescence intensity, which were attributed to SVs by TEPIQ analysis of ΔVS, amounted to 20% of the original area of the plasma membrane. Capacitance measurement estimated the increase in the membrane area to be only ∼5% (Takahashi et al. 1997), probably because of concurrent endocytosis. The contribution of SVs to the rapid capacitance increase in β-cells was also supported by the observation that such increases were not associated with amperometric detection of LV exocytosis (Takahashi et al. 1997). Such amperometric signals were sensitive to inhibitors of PKA (Takahashi et al. 1999), in line with our results for LVs. In contrast, the rapid capacitance increase was found to be resistant to Rp-cAMPS and PKI (Renström et al. 1997), and 8-CPT-2′-O-Me-cAMP potentiated the rapid component of capacitance increases during trains of depolarization (Eliasson et al. 2003), consistent with our results for SVs.

Conclusion

In summary, exocytosis of SVs occurs more rapidly than that of LVs during stimulation with cytosolic Ca2+ and cAMP, and does so even at lower glucose concentrations in the blood. Therefore, SV exocytosis probably precedes LV exocytosis in most physiological conditions. Exocytosis of SVs may precondition the plasma membrane for subsequent insulin granule exocytosis, which may otherwise be harmful to the islet. Although it is currently unknown why Epac selectively regulates SVs but not LVs in β-cells, such two distinct pathways of cAMP were reported to be involved in exocytosis in melanotrophs (Sedej et al. 2005), therefore it is possible that similar mechanisms play roles in cAMP-dependent functions in various types of cells. The more rapid control of SV exocytosis by Epac compared with control of LV exocytosis by PKA is also consistent with the putative role of Epac in regulation of synaptic vesicle exocytosis (Sakaba & Neher, 2001; Kaneko & Takahashi, 2004), which may be rapidly modulated by various neurotransmitters that affect adenylate cyclase activity. In any case, we have found that two effectors of cAMP (Epac and PKA) control exocytosis with different rates. Similar mechanisms may operate for other cAMP-dependent cellular functions.

Appendix

Acknowledgements

We thank Y. Hara, M. Yoshida and N. Takahashi for technical assistance. This study was supported by Grants-in-Aid from the Ministry of Education, Culture, Sports, Science, and Technology of Japan and from the Japanese Society for the Promotion of Science, as well as by research grants from the Human Frontier Science Program Organization and Takeda Science Foundation.

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