Impaired maturation of large dense-core vesicles in muted-deficient adrenal chromaffin cells

Large dense-core vesicles (LDCVs), which are formed at the trans-Golgi network (TGN), are key organelles for secretion of hormones and neuropeptides in specialized secretory cells, including neuroendocrine and non-neuroendocrine cells. LDCVs are characterized as a type of lysosome-related organelle (LRO) (Wei and Li, 2013). However, the molecular players and mechanisms governing the biogenesis of LDCVs are not fully understood. During LDCV biogenesis, sorting machineries are important to ensure proper cargo assembly into mature and condensed vesicles (Kim et al., 2006; Tooze et al., 2001). Two hypotheses have been proposed for cargo sorting into LDCVs: sorting for entry and sorting for retention (Arvan and Castle, 1998). Prevention of cargo mis-sorting into lysosomes for degradation is also important during LDCV biogenesis (Yu et al., 2011).

Previous studies have suggested that the granin family plays an important role in LDCV biogenesis (Kim et al., 2001), but the machinery for granin sorting into LDCVs is largely unknown. The adaptor protein-3 (AP-3) complex is likely involved in granin sorting into immature LDCVs. In AP-3-deficient chromaffin cells, LDCVs contain less granins, chromogranin A (CgA, also known as CHGA) and secretogranin II (SgII, also known as SCG2), and exhibit enlarged size, decreased number and an increase in releasing quantal size (i.e. the amount of transmitter released per vesicle) (Asensio et al., 2010; Grabner et al., 2006). AP-3 and the HOPS complex interact physically and might act in the same pathway in regulating endosomal trafficking (Angers and Merz, 2009; Rehling et al., 1999; Wilkin et al., 2008). Knockdown of VPS41 (a HOPS subunit) in PC12 cells leads to similar defects of LDCVs as in AP-3-deficient cells (Asensio et al., 2013). Similarly, the LDCVs in adrenal chromaffin cells from mice deficient in dysbindin [a subunit of biogenesis of lysosome-related organelles complex-1 (BLOC-1)] are reduced in number and increased in vesicle size, leading to reduced secretion events and increased quantal size (Chen et al., 2008). These findings suggest that AP-3, HOPS and BLOC-1 are likely involved in the sorting of cargoes into immature LDCVs (selective import) or the sorting of unnecessary molecules out of immature LDCVs (selective export). However, the underlying mechanisms of how these trafficking complexes control the import and export processes of LDCV proteins are very limited.

The BLOC-1 complex consists of eight subunits: muted, pallidin, dysbindin, cappuccino (also known as BLOC1S5, BLOC1S6, DTNBP1 and BLOC1S4, respectively), snapin, BLOS1 (also known as BLOC1S1 or GCN5L1), BLOS2 (also known as BLOC1S2) and BLOS3 (also known as BLOC1S3) (Ciciotte et al., 2003; Falcón-Pérez et al., 2002; Gwynn et al., 2004; Li et al., 2003; Starcevic and Dell'Angelica, 2004). KXD1 has been identified as an additional subunit of BLOC-1 (Hayes et al., 2011; John Peter et al., 2013; Yang et al., 2012). Current understanding of the functions of BLOC-1 are mainly focused on cargo trafficking from endosomes to lysosomes (for a review, see Wei and Li, 2013) and endosomal maturation (John Peter et al., 2013). BLOC-1 deficiency causes Hermansky–Pudlak syndrome (HPS) in mice and humans, which is characterized by LRO dysfunction (Wei and Li, 2013). The KXD1-knockout mice (Yang et al., 2012) exhibit mild defects in LROs compared with other typical HPS mouse mutants of BLOC-1 such as the sdy (dysbindin deficiency), mu (muted deficiency) and pa (pallidin deficiency) mice, whereas snapin- and BLOS1-knockout mice are embryonic lethal and exhibit impaired autophagy (Cai et al., 2010; Scott et al., 2014; Tian et al., 2005; Zhang et al., 2014). This suggests that the deficiency of different BLOC-1 subunit leads to variable phenotypes on LROs. Comparison of the features of LRO defects in different BLOC-1 mouse mutants might reveal the underlying mechanisms in their involvement of LRO biogenesis.

The mu mutant mouse, which carries a spontaneous mutation on the Muted gene (Bloc1s5), abolishes the expression of the muted protein and affects the biogenesis of several LROs such as melanosomes and platelet-dense granules (Zhang et al., 2002). In our pilot study, we observed that mu mice had a different profile of morphological changes in LDCVs compared to sdy mice (Chen et al., 2008) or snapin-knockout mice (Pan et al., 2009; Tian et al., 2005). In comparison with additional HPS mouse mutants, we propose that the muted protein is involved in the export of the granin CgA during LDCV biogenesis.

To investigate whether LDCVs are affected in mu mice (mu/mu), we analyzed the morphological changes of LDCVs from adrenal gland of controls (mu/+) and mu mice by electron microscopy. The gross ultrastructure of mu chromaffin cells was similar to that in the control mice (Fig. 1A). The round granular limits were visible and dense cores were condensed in mu chromaffin cells. In control cells, many granules had a loosely packaged dense-core and a spacious halo around the dense core. However, in mu cells, the halos were less spacious and the dense cores occupied more of the space in the whole vesicles (Fig. 1A). We then examined the number and size of LDCVs in sections of randomly selected cells from two different animals for each genotype. There was no significant difference in the total number of vesicles per square micrometer between control and mu chromaffin cells (Fig. 1B). For vesicle size analysis, all round granules with a well-preserved membrane, a halo and an electron-dense round core were included. The average increase of the size (diameter) of LDCVs in mu chromaffin cells was ∼6% (Fig. 1C). Furthermore, the electron-dense core size (diameter) was significantly enlarged by ∼17.6% in mu chromaffin cells (Fig. 1D). Hence, the ratio of dense core diameter compared to vesicle diameter was increased by ∼10% in mu mice compared with control mice (Fig. 1E). Similarly, we observed enlarged vesicle size and dense core size in another type of LDCV in islet α-cells, which contain CgA and glucagon (Portela-Gomes and Stridsberg, 2001), in the mu mice (supplementary material Fig. S1). These results suggest that loss of the muted protein likely affects LDCV morphology in a conserved mechanism.

CgA is a major constituent of LDCVs, acting as an assembly factor to drive secretory granule biogenesis (Kim et al., 2001). To investigate the molecular mechanism of the morphological defects of mu LDCVs, we measured the steady-state protein levels of CgA in adrenal glands in mu mice. We found a significant increase (1.54-fold) of full-length CgA (80 kDa) in mu mice compared to wild-type (WT, con) mice (Fig. 2A,B). We quantified the amount of an intermediate CgA-derived peptide (∼23 kDa) and there was likewise an increase in mu mice (Fig. 2C,D), which is in agreement with the increase of full-length CgA. This suggests that the increased CgA in mu mice is unlikely to be caused by the inefficiency of CgA cleavage. In contrast, the protein levels of another granin, SgII, were unaltered in mu mice compared to control mice (Fig. 2E,F). We confirmed the increase of CgA in muted-knockdown (KD) PC12 cells (Fig. 2G). Consistently, the intensities of CgA in muted-KD PC12 cells was significantly increased (Fig. 2H,I).

As the total number of LDCVs in mu cells did not change, we reasoned that the increased CgA might result from impaired export from immature LDCVs or from mislocalized non-LDCV organelles. We monitored the distribution of CgA by western blotting after equilibrium sedimentation through a 5–50% OptiPrep gradient. We found an almost complete absence of CgA of mu adrenal glands in the lighter fractions 3 and 4, which correspond to the clathrin heavy chain, compared with in wild-type mice. In contrast, the distribution of SgII was similar between mu and WT mice (Fig. 3A). These results suggest that the muted protein might facilitate the removal or export of CgA from immature secretory granules (ISGs) via clathrin-coated vesicles (Orci et al., 1985a; Tooze and Tooze, 1986).

We then tested whether the increased amount of CgA in mu mice was localized to LDCVs or mislocalized to other organelles. Immature and mature LDCVs have been considered as the major prohormone conversion sites (Koshimizu et al., 2010; Orci et al., 1985b). In this regard, the processed CgA peptides (60 kDa and 23 kDa) are mainly concentrated in LDCVs. We observed that these peptides were enriched in fractions 7–10, which represent the distribution of LDCVs (Fig. 3A). We then quantified the total amount of CgA in these LDCV-enriched fractions and our results showed that the LDCV-CgA proportion in mu mice was increased compared with the WT mice (Fig. 3B). Consistent with this, we quantified the intensities of CgA in cytosolic individual granules that are defined as LDCVs (Walter et al., 2014) and showed that the CgA intensity in the LDCV proportion was increased in muted-KD PC12 cells compared with the control cells (Fig. 3C).

To confirm that loss of muted protein leads to the increase in CgA, we transfected EGFP–muted into the stable muted-knockdown PC12 cells (Fig. 4A). We quantified the intensities of total CgA and the CgA in individual LDCVs in the transfected cells and showed that both total CgA and the LDCV-CgA proportion were significantly decreased in the EGFP–muted-expressing cells compared with the EGFP-transfected cells (Fig. 4B,C). We verified the expression of the muted protein in these cells (Fig. 4D). These results suggest that the increased LDCV-CgA as well as total CgA can be rescued by the expression of functional muted protein.

We then investigated whether CgA is mislocalized to other non-LDCV organelles in the mu cells. We observed that CgA was largely retained in the SgII-positive LDCVs in both the muted-KD cells and control cells, with a partial colocalization with LAMP1, which is a marker of both lysosomes and LDCVs. We did not observe apparent mislocalization of CgA to other organelles including the endoplasmic reticulum (ER) (SEC61B), TGN (STX6), early endosomes (EEA1) and late endosomes (Rab7) in the muted-KD cells when compared to control cells (Fig. 5), suggesting that its subcellular localization is unaltered.

Taken together, in the absence of muted protein, the proportion of CgA which is sorted to constitutive secretory vesicles or lysosomes might be trapped in ISGs, and thus lead to the increase of CgA in LDCVs. As CgA has the ability to aggregate to form electron-dense cores, the increased CgA content in mu chromaffin cells likely leads to the increased size of electron-dense cores as well as the increased size of LDCVs.

We tested whether the size change in LDCV in mu cells would affect the size of the readily releasable pool (RRP) and the release frequency. In neuroendocrine cells, the RRP generally represents the population of fusion-competent vesicles, and the immediately releasable pool (IRP) corresponds to a subpool of the RRP that is closely associated with Ca²⁺ channels (Voets et al., 1999). Whole-cell patch-clamp was applied to single chromaffin cells. Average responses obtained from control and mu mice showed that secretory response was drastically reduced in mu chromaffin cells as shown by the reduced IRP and RRP size (Fig. 6A–D). These results indicate that lack of muted protein leads to a smaller RRP size and results in a reduction of vesicular secretion in chromaffin cells.

The reduction in RRP size or vesicle release could result from the reduction of docked vesicles when LDCV maturation is impaired, as evident by the enlargement of LDCVs in mu cells. We measured the number of docked vesicles (0–100 nm from plasma membrane) in the mu adrenal chromaffin cells. Indeed, the docked vesicles in mu cells were significantly reduced compared to the control mice (Fig. 6E,F).

We then investigated that whether the docked vesicles release with normal kinetics. We recorded the kinetics of each releasing spike by carbon-fiber electrodes. We found no significant changes in half-height duration (HHD), rise time (RT) and quantal size (Q) of amperometric spikes between control and mu mice (supplementary material Fig. S2). This suggests that the docked vesicles might represent a normal population of LDCVs that have normal fusion and release kinetics. This population might not be affected by the muted protein.

To explore whether the docking or fusion machinery is affected by the loss of muted protein, we determined the steady state levels of the synaptic proteins in the mu adrenal glands. Western blotting analyses confirmed the expression of the muted protein in adrenal cells. As expected, no band was observed in the mu adrenal glands. We did not find apparent changes in the steady-state levels of a panel of synaptic proteins including SNARE proteins (SNAP25, syntaxin-1 and VAMP2), synaptophysin (SYP) and Munc18 between controls and mu mice (supplementary material Fig. S3A). These results suggest that the smaller RRP pool does not result from changes in the steady-state levels of these well-known docking or fusion proteins.

Compared with the characteristics of LDCVs in dysbindin-1 deficient sdy mice, mu mice have a different profile of LDCV phenotypes (supplementary material Table S1) although both dysbindin-1 and muted reside in the same BLOC-1 complex. This suggests that dysbindin-1 and muted play different roles in cargo sorting for LDCV biogenesis. Notably, vesicle number is reduced in sdy mice but is normal in mu mice. The reduction of LDCV number in AP-3- or HOPS-deficient chromaffin cells might lead to the decrease of CgA and SgII as shown in supplementary material Table S1. We confirmed the reduction of CgA and SgII in pe (β3A AP-3 deficiency) and bf (VPS33a HOPS deficiency) mice compared with their wild-type controls. In sdy mice, CgA was decreased and that there was also a reduction in the levels of SgII although this was not significant (Fig. 7). In contrast, the CgA steady-state level was increased but SgII was normal and the number of LDCVs was not changed in mu mice (Figs 1, 2). This suggests that the increase in CgA in mu mice is not due to the increase in LDCV number. This further supports our hypothesis that the increase of CgA in mu cells is caused by the retention of CgA in immature LDCVs. Muted protein plays an important role in the export of CgA, but has no or little effect on SgII.

Here, we characterized the defects in LDCV morphology (enlarged size and reduced docking) by examining electron microscopy images of mu adrenal chromaffin cells. In the absence of muted protein, the CgA content was increased, which could be an explanation for the enlarged sizes of LDCVs and dense cores (Kim et al., 2001). The increase of CgA does not result from an increased vesicle number or impaired processing, rather it is likely caused by the failure of CgA export from the vesicles when muted protein is lacking. In general, an enlarged size of LDCV represents immature LDCVs (Kim et al., 2006). Consequently, failure of CgA export might impair LDCV maturation and docking and finally vesicle release (supplementary material Fig. S3B). These findings support the sorting-for-retention model and provide molecular insights into the sorting machinery of LDCV and its pathology.

Cargo sorting into LDCVs is a very complicated and highly regulated process. It has been postulated that there different LDCV populations exist, which might have different constituents (Duncan et al., 2003; Grabner et al., 2005; Watanabe et al., 1991). The segregation of CgA and SgII in different populations implicates that there exist different sorting machineries for these granins (Watanabe et al., 1991). As shown in Fig. 5C, there were some granules that CgA and SgII did not colocalize in control cells, further supporting that separate CgA-positive vesicles and SgII-positive vesicles exist, although a large number of the granules contain both CgA and SgII. Furthermore, our results showed an increase in CgA but a normal level of SgII, and a distributional shift in CgA, but not SgII, in the mu adrenal glands. This indicates that the muted protein is involved in the sorting of CgA, but not SgII. This also explains why vesicle docking and release events exist and why the releasing kinetics is normal (supplementary material Fig. S2) in the mu chromaffin cells given that the secretion of SgII-positive LDCV population is normal.

Control of vesicle number and size could be a reflection of the complexity of multiple regulators. In BLOC-1, loss of dysbindin-1 (sdy) and loss of muted (mu) both lead to an enlargement of LDCV size, but the LDCV number is unchanged in mu mice, in contrast to reduced LDCV numbers in sdy mice (Chen et al., 2008). In other LROs, such as melanosomes and platelet-dense granules, both mu and sdy mice show reduced vesicle numbers, indicating the biogenesis of LROs utilizes shared mechanisms in endo-lysosomal trafficking (Swank et al., 1998; Wei and Li, 2013). Taken together, mounting evidence indicates that each LRO has unique biosynthetic and cargo-sorting pathways, although they might share some common sorting mechanisms. By contrast, dysbindin is known to bind to a variety of proteins belonging to the dystrophin protein complex, the WAVE-2 complex and others (Ghiani and Dell'Angelica, 2011; Li et al., 2007). This suggests that different interacting partners of BLOC-1 subunits are operative (Li et al., 2007). In addition, dysbindin-1 has distinct isoforms that might function in different pathways (Talbot et al., 2009). We have recently shown that dysbindin-1C isoform functions in a BLOC-1-independent pathway in regulating autophagy and neuronal survival (Wang et al., 2014; Yuan et al., 2015). The altered ultrastructure of hippocampal synaptic vesicles might cause reduced neurotransmission and thereafter the schizophrenia-like symptoms observed in sdy mice (Chen et al., 2008; Feng et al., 2008; Wang et al., 2014). However, except for DTNBP1 and BLOC1S3, genes encoding other BLOC-1 subunits have no significant association with schizophrenia although an epistatic interaction between DTNBP1 and MUTED might exist (Morris et al., 2008). Currently, there is no support for MUTED as a susceptibility gene for schizophrenia (Gerrish et al., 2009). Thus, the different profiles of releasing vesicle defects in sdy and mu mice might have different mechanisms and consequences.

CgA has been implicated in playing multiple roles in the endocrine, cardiovascular and nervous systems through the regulation of multiple DCV biogenesis in different tissues. Our results have shown similar defects in both the adrenal chromaffin LDCV and the LDCV in pancreatic islet α-cells (supplementary material Fig. S1), suggesting that they might share a conserved mechanism in CgA sorting. CgA knockout in mice leads to decreased size and number of LDCVs in chromaffin cells, as well as hypertension in these animals (Mahapatra et al., 2005). In our case, loss of muted protein causes an increase in CgA and impairs LDCV release frequency from the chromaffin cells. Whether CgA secretion is reduced in vivo in mu mice, or whether there is a link to hypertension or other LDCV phenotypes in this mutant requires further detailed investigation.

The mu mutant mice and control CHMU/Le mice (wild-type, WT) were as described previously (Zhang et al., 2002). Other mouse mutants and their controls including pearl (pe) and buff (bf) in the C57BL/6J (B6) background, sandy (sdy) in the DBA/2J (DBA) background were are summarized Li et al. (Li et al., 2004). They were originally obtained from the Jackson Laboratory (Bar Harbor, Maine, USA), were transferred to Dr Richard T. Swank's laboratory at Roswell Park Cancer Institute (Buffalo, New York, USA) and are bred in the animal facility of the Institute of Genetics and Developmental Biology, Chinese Academy of Sciences. All animal experiments were performed under permission from the Institutional Animal Care and Use Committee and the Peking University Committee on Ethics in the Care and Use of Laboratory Animals. The genotypes of wild type (+/+), heterozygous (mu/+) and homozygous (mu/mu) mice were identified by a PCR method based on the mutation in the Muted gene (Zhang et al., 2002). Age-matched wild-type CHMU/Le mice and heterozygous mice were used as background controls.

The polyclonal muted antibody was generated in New Zealand white rabbits against a His-tagged fusion protein corresponding to the mouse full-length muted protein (GenBank RefSeq NM_139063). The mouse monoclonal antibodies against Syntaxin 6 (#ab12370), the rabbit polyclonal CgA (#ab45179), VAMP2 (#ab3347) and SEC61B (#ab78276) were obtained from Abcam (Cambridge, UK). The rabbit polyclonal Rab7 (#9367) and clathrin heavy chain (#4796) antibodies were from Cell Signaling Technology (Danvers, MA). The rabbit polyclonal SgII antibody (#K55101R) was from Meridian Life Science (Memphis, TN). The mouse monoclonal β-actin antibody (#A5441) was purchased from Sigma-Aldrich (St. Louis, MO). The mouse monoclonal EEA1 (#610457), Munc-18 (#610337) and the rat monoclonal LAMP1 (#553792) antibodies were from BD Biosciences (San Diego, CA). Mouse monoclonal SYP (#sc-17750), syntaxin 1 (#sc-12736), SNAP25 (#sc-20038), GFP (#sc-9996) and goat polyclonal CgA (#sc-1488) antibodies were purchased from Santa Cruz Biotechnology (Dallas, TX).

Adrenal glands from control and mu mice were homogenized in 50 mM Tris-HCl pH 7.4, 150 mM NaCl, 1% Triton X-100, and a protease inhibitor cocktail (Sigma-Aldrich). The extracts were centrifuged at 13,000 g for 15 min to remove nuclei and cell debris, equal amounts of homogenates were separated on 12% SDS-PAGE gels and transferred onto polyvinylidene difluoride (PVDF, Millipore) membranes. The membranes were immunoblotted with specific antibodies and visualized by ECL (Pierce, Rockford, IL). Protein bands intensities were quantified by NIH Image J after normalization to β-actin.

Adrenal glands from WT and mu mice were homogenized in 300 µl HB buffer (250 mM sucrose, 20 mM Tris-HCl, 1 mM EDTA, pH 7.4) plus a protein inhibitor cocktail (Sigma-Aldrich). A post-nuclear supernatant (PNS) was generated by centrifugation at 1000 g for 10 min. The PNS was layered onto a linear 5–50% linear Optiprep gradient (Axis-Shield, Norway) formed in HB buffer using a Gradient Master (Biocomp, USA). Gradients were spun at 30,000 rpm for 15–16 hours in an SW41 rotor (Beckman, USA) at 4°C. Fourteen equal fractions (900 µl each) were collected from the top of the gradient and subjected to acetone precipitation. The precipitated protein (fractions 3–13) was analyzed by SDS-PAGE and immunoblotting.

PC12 cells were cultured at 37°C with 5% CO₂ in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% horse serum and 5% fetal bovine serum (Gibco-BRL, Gaithersburg, MD). Short hairpin RNAs (shRNA) against rat Muted (GenBank RefSeq NM_001107347) were designed by the siRNA target finder (Ambion, Austin, TX). Corresponding DNAs were synthesized and incorporated into the BamHI and HindIII sites of pSilencer 5.1-H1 Retro (Ambion) vector. The target sequence for Muted was 5′-AAGGATCTTGGAGAAATTCAT-3′ and for control was 5′-AATTCTCCGAACGTGTCACGT-3′. These vectors were transfected into PC12 cells using Lipofectamine^TM 2000 (Invitrogen, Carlsbad, CA) following the manufacturer's instructions. After 24 hours, the transfected cell clone expressing the vector was selected by 5 µg/ml of puromycin (Invivogen). For the rescue experiment, the Muted cDNA (NM_139063.1) was amplified by PCR and cloned in-frame into pEGFP-C2 using the XhoI and BamHI sites. Cells were transfected by using the Neon transfection system (Life Technologies, Carlsbad, CA) according to the manufacturer's instruction. Transfected cells were seeded onto poly-D-lysine-coated glass coverslips and then processed for immunostaining 72 hours after transfection.

Chromaffin cells were isolated as previously described (Sørensen et al., 2003) with minimal modifications. Adrenal glands were dissected from 6- to 8-week-old heterozygous mu/+ or mu mice, placed in filtered Locke's buffer (in mM: 154 NaCl, 5.6 KCl, 5.0 HEPES, 3.6 NaHCO₃, and 5.6 glucose, pH 7.4). Fat tissue and adrenal cortex was removed from adrenal medulla under a surgical microscope. The medullae were incubated in 0.2 ml equilibrated (95% O₂∶5% CO₂) papain solution (40 units/ml in Locke's buffer; the papain was from Sigma-Aldrich) at 37°C for 1 hour with gentle shaking. The enzyme solution was discarded and replaced with 500 µl culture medium (DMEM supplemented with 10% fetal calf serum, 100 IU/ml penicillin and 100 µg/ml streptomycin). After washing two times in 500 µl culture medium, the medullae were triturated gently through a 10-µl pipette tip. 50 µl cell suspension was plated on poly-D-lysine-coated glass-bottomed dishes to allow the cells to settle for 30 min at 37°C before supplementing with 2 ml culture medium. The cells were cultured at 37°C in 5% CO₂ and used within 3 days.

PC12 cells were plated on poly-D-lysine-coated glass coverslips for 24 hours. Cells were fixed with 4% paraformaldehyde for 20 min at room temperature, permeabilized for 15 min with 0.4% Triton X-100 in PBS, blocked for 30 min with 1% BSA in PBS, then incubated for 16 hours at 4°C with indicated primary antibodies in PBS containing 1% BSA. Cells were subsequently washed and incubated for 1 hour with a Alexa-Fluor-594-conjugated donkey anti-mouse IgG (1∶2000; Invitrogen), Alexa-Fluor-594-conjugated donkey anti-rabbit IgG (1∶2000; Invitrogen), Alexa-Fluor-488-conjugated donkey anti-goat IgG (1∶2000; Invitrogen), Alexa-Fluor-488-conjugated goat anti-rat IgG (1∶2000; Invitrogen) or Alexa-Fluor-488-conjugated donkey anti-rabbit IgG (1∶2000; Invitrogen) in PBS containing 1% BSA. Confocal images were acquired using a 100× oil objective with NA 1.40 on a Nikon confocal microscope (ECLIPSE Ti-C2, Japan). Images were obtained using the NIS-Elements AR 3.2 software provided by Nikon. For quantification of the fluorescence intensity, the fluorescence signal was obtained from the entire cell, the nucleus was subtracted and image was analyzed with NIH Image J software.

Electron microscopy was carried out as previously described (Chen et al., 2008). Briefly, adrenal glands removed from homozygous mu mice and heterozygous mu/+ littermates (8 weeks old) were fixed with 3% glutaraldehyde in 0.1 M phosphate buffer (pH 7.4) overnight and were postfixed with 1% OsO₄ for 2 hours at 4°C. Tissue samples were then dehydrated in a graded series of ethanol and were embedded in Epon812. Ultra-thin sections (∼60 nm) were collected and stained with uranyl acetate and lead citrate. The sections were examined on a FEI Tecnai 20 electron microscope and analyzed with NIH Image J. At least 100 digital images were taken from randomly selected chromaffin cells of two different animals for both genotypes. Numerical density of the secretory granules was calculated as the number of morphologically identifiable LDCVs per section divided by the chromaffin cell cytoplasmic area. Vesicles with a round membrane and an electron-dense core were included in the diameter analysis. Diameters of vesicles and electron-dense cores were determined by direct measurement. The observer was blind to the genotypes.

The islets from age-matched mu/+ and mu/mu male mice were fixed in 2% paraformaldehyde and 2.5% glutaraldehyde in 0.1 M phosphate buffer (pH 7.4) overnight and were post-fixed with 1% OsO₄ for 2 hours at 4°C. Tissue samples were then dehydrated in a graded series of acetone and were embedded in Spurr (SPI-Chem, West Chester, PA). Ultra-thin sections (∼70 nm) were collected and stained with lead citrate. The sections were examined on an electron microscope (JEM 1400; JEOL, Japan) at 80 kV. Digital images were taken from randomly selected α-cells of two different animals for both genotypes.

Whole-cell recordings were performed using an EPC-9 amplifier and Pulse software (HEKA, Lambrecht, Germany). A 800 Hz, 20 mV peak-to-peak sinusoid stimulus was applied about a direct current holding potential of 80 mV. The intracellular solution contained (in mM): Cs-glutamate, 145; NaCl, 8; MgCl₂, 1; Mg-ATP, 2; Na₂-GTP, 0.3; and Cs-HEPES, 10 (pH 7.2). The extracellular solution contained (in mM) NaCl, 140; KCl, 2.8; MgCl₂, 1; CaCl₂, 2; HEPES, 10; and glucose, 2 (pH 7.4). Experiments were performed at room temperature.

We prepared adrenal medulla slices and performed whole-cell recordings as described previously (Chen et al., 2008) by using an EPC9/2 amplifier and Pulse software (HEKA Elektronik, Lambrecht/Pfalz, Germany). For analysis of the kinetic properties of amperometric spikes, including the half-height duration (HHD), the rise time (RT) and quantal size (Q), only events that were >5 s.d. of the noise were used (Zhou and Misler, 1996). Data analysis was carried out using Igor software (WaveMetrix, USA) with a custom-made macro program (Zhou and Misler, 1996).

All values in the text are shown as mean±s.e.m. Student's t-test was used to compare means of two groups. Differences of P<0.05 were considered statistically significant.

We are in debt to Dr. Richard T. Swank for his generous providing of the HPS mutant mice used for this study and for his proofreading of this manuscript. We thank Ms. Lei Sun for her help in making electron microscopy samples and taking electron microscopy images.