Involvement of conventional kinesin in glucose-stimulated secretory granule movements and exocytosis in clonal pancreatic ß-cells

Aniko Varadi1, Edward K. Ainscow1, Victoria J. Allan2 and Guy A. Rutter1,*

1 Department of Biochemistry, School of Medical Sciences, University of Bristol, University Walk, Bristol BS8 1TD, UK
2 School of Biological Sciences, University of Manchester, 2.205 Stopford Building, Oxford Road, Manchester M13 9PT, UK

* Author for correspondence (e-mail: g.a.rutter{at}bris.ac.uk)

Accepted 2 August 2002


    Summary
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 Materials and Methods
 Results
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 References
 
Recruitment of secretory vesicles to the cell surface is essential for the sustained secretion of insulin in response to glucose. At present, the molecular motors involved in this movement, and the mechanisms whereby they may be regulated, are undefined. To investigate the role of kinesin family members, we labelled densecore vesicles in clonal ß-cells using an adenovirally expressed, vesicle-targeted green fluorescent protein (phogrin.EGFP), and employed immunoadsorption to obtain highly purified insulin-containing vesicles. Whereas several kinesin family members were expressed in this cell type, only conventional kinesin heavy chain (KHC) was detected in vesicle preparations. Expression of a dominant-negative KHC motor domain (KHCmut) blocked all vesicular movements with velocity >0.4 µm second-1, which demonstrates that kinesin activity was essential for vesicle motility in live ß-cells. Moreover, expression of KHCmut strongly inhibited the sustained, but not acute, stimulation of secretion by glucose. Finally, vesicle movement was stimulated by ATP dose-dependently in permeabilized cells, which suggests that glucose-induced increases in cytosolic [ATP] mediate the effects of the sugar in vivo, by enhancing kinesin activity. These data therefore provide evidence for a novel mechanism whereby glucose may enhance insulin release.

Key words: Kinesin, Insulin, Exocytosis, Glucose, Islet, ß-cell, Pancreas


    Introduction
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 Introduction
 Materials and Methods
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 References
 
Glucose triggers the release of insulin from pancreatic islet ß-cells through a complex sequence of events (Rutter, 2001Go), involving increases in intracellular ATP levels (Malaisse and Sener, 1987Go; Kennedy et al., 1999Go), closure of ATP-sensitive K+ channels (KATP) (AguilarBryan and Bryan, 1999Go) and influx of Ca2+ (Safayhi et al., 1997Go). While these events underlie the acute, or `first phase', of insulin release (Curry et al., 1968Go), KATP-channel independent mechanisms are involved in the second, or sustained phase, of release (Gilon and Henquin, 1992Go; Komatsu et al., 1995Go; Aizawa et al., 1998Go).

The molecular basis of the second phase of secretion is poorly understood (Rorsman et al., 2000Go; Daniel et al., 1999Go). Since only fuel secretagogues such as glucose trigger the sustained phase of release (Rorsman et al., 2000Go), it seems likely that an energy-dependent, presumably ATP-requiring step, is involved in recruiting vesicles from a `reserve' to a `readily releaseable' pool of vesicles (Proks et al., 1996Go; Daniel et al., 1999Go; Rorsman et al., 2000Go).

In previous studies in which we imaged the behaviour of vesicle-targeted enhanced green fluorescent protein (phogrin.EGFP) (Pouli et al., 1998Go; Tsuboi et al., 2000Go), elevated glucose concentrations stimulated both short and longer excursions of vesicles. Montague and colleagues (Montague et al., 1975Go) have demonstrated that the recruitment of insulin-containing vesicles to the plasma membrane may be essential for sustained nutrient-stimulated insulin secretion, and may involve microtubules (see also Pouli et al., 1998Go). By contrast, an important role for microfilaments seems unlikely, since breakdown of microfilaments with clostridium botulinum neurotoxin C2 or cytochalasins E or F enhanced glucose-stimulated release from islets (Li et al., 1994Go).

The above, indirect observations, suggest that kinesin, or kinesin-related motor proteins (KRPs) (Lane and Allan, 1998Go), may be involved in glucose-stimulated movement of insulin-containing vesicles. Kinesins are a family of motor proteins that use ATP hydrolysis to move cargoes along microtubules (MTs) (Goldstein, 1993Go). Kinesin is required for axonal transport in neuronal cells (Rahman et al., 1999Go; Gindhart et al., 1998Go), and recruits vesicles to the release sites of Ca2+-regulated exocytosis in sea urchin embryos (Bi et al., 1997Go).

Conventional kinesin is a heterotetramer of two kinesin heavy chains (KHCs) and two kinesin light chains (KLCs). The KHC head domain is highly conserved among different kinesin-related proteins and is responsible for ATP hydrolysis and force generation (Yang et al., 1990Go). In mice, three conventional kinesin genes (Kif5a, Kif5b and Kif5c) have been identified. Kif5a and Kif5b are the mouse homologues of the human neuronal-KHC and ubiquitous-KHC, respectively (Xia et al., 1998Go), and Kif5b is expressed in primary mouse ß-cells (Meng et al., 1997Go). Furthermore, suppression of Kif5b with antisense oligonucleotides reduced, but did not altogether abolish, glucose-stimulated insulin release from primary ß-cells (Meng et al., 1997Go).

In the present study, we have investigated: (1) the complement of kinesins in clonal ß-cell lines and on dense core insulin secretory vesicles; (2) the role of kinesins in insulin-containing vesicle translocation; (3) the importance of vesicle motility for glucose-stimulated secretion; and (4) the role of ATP in regulating kinesin activity. We demonstrate that sustained insulin release requires kinesin-dependent transport of vesicles to the plasma membrane. Moreover, since vesicle movement could be regulated by increases in ATP concentration over the physiological range in permeabilised cells, these data suggest that kinesin may represent a novel target for regulation by glucose in living ß-cells (Kennedy et al., 1999Go). Regulation of kinesin activity may thus contribute to the KATP channel-independent stimulation of insulin secretion by nutrients (Aizawa et al., 1998Go; Takahashi et al., 1999Go; Seghers et al., 2000Go)


    Materials and Methods
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 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Materials
cDNAs encoding the motor domain (340 amino acids) of rat conventional kinesin heavy chain (KHC340) (the homologue of mouse Kif5b) (Hirokawa, 1998Go), and KHC340 containing a T93N mutation (KHCmut) and fused to a histidine-tag (6His) in vector pET-17b, was kindly provided by R. Cross (Marie Curie Research Institute, Oxted, UK) (Krylyshkina et al., 2002Go). Human growth hormone-containing plasmid pXGH5 was kindly provided by R. Burgoyne (University of Liverpool) (Fisher and Burgoyne, 1999Go). Rabbit-polyconal anti-phogrin antibody raised against the C-terminal domain (amino acids 629-1003) was a kind gift from J. C. Hutton (Barbara Davis Center for Childhood Diabetes, Denver, CO). Cell culture reagents were from Gibco-BRL (Life Science Research, Paisley, UK). Mouse monoclonal anti-green fluorescent protein (GFP) antibody, the human growth hormone (hGH) ELISA Kit, the ECL detection system and all molecular biologicals were obtained from Roche Diagnostics (Lewes, UK). Alexa Fluor 488 or 568 goat anti-mouse, and Alexa Fluor 568 goat anti-guinea pig IgG were from Molecular Probes (Eugene, USA). A rabbit polyclonal antibody that recognises multiple kinesin family members (pan-kinesin), was raised as described (Barroso et al., 2000Go). Rabbit polyclonal anti-ubiquitous kinesin (Niclas et al., 1994Go) was kindly provided by R. Vale (UCSF, San Francisco, CA). Mouse monoclonal anti-human growth hormone and a rabbit polyclonal His-probe antibodies were from Autogen Bioclear UK (Calne, UK). Mouse monoclonal anti-{alpha}-tubulin and anti-dynein (Clone 70.1) antibodies were obtained from Sigma (Poole, UK). A trans-Golgi network protein 38 (TGN38)-specific antibody was kindly provided by G. Banting (University of Bristol, Bristol, UK).

Cell culture
MIN6 and INS-1 pancreatic ß-cells were cultured in DMEM and RPMI 1640 tissue-culture medium supplemented with 15% (v/v) and 10% (v/v) fetal calf serum, respectively, plus penicillin (100 units ml-1) and streptomycin (0.1 mg ml-1) at 37°C in an atmosphere of humidified air (95%) and CO2 (5%) as described previously (Molnar et al., 1995Go). MIN6 cells were used between passages #19 and #35.

Plasmids
KHC (residues 1-340) (KHC340) and KHC340 carying a T93N point mutation (KHCmut) and (6His) cDNA (Krylyshkina et al., 2002Go) was cloned across the XbaI-XhoI sites of pcDNA 3.1(-) and then into the pAdTrack-CMV shuttle vector (He et al., 1998Go) via XbaI-EcoRV sites (Fig. 3A,B Fig. 8A,B). Generation of phogrin.pEGFP.N1 (Pouli, et al., 1998Go) and sub-cloning into a recombinant adenovirus were as described earlier (Tsuboi, et al., 2000Go). Plasmid phogrin. DsRed was constructed by digesting phogrin.EGFP.N1 with AgeI and NotI to remove the EGFP-coding sequence, and replacing it with the AgeI/NotI DsRed-coding fragment from pDsRed-N1 (Clontech). Mitochondrially-targeted DsRed was generated by fusion to the N-terminal 33 amino acids of cytochrome c oxidase subunit VIII (Rizzuto et al., 1989Go). The leader sequence was removed from plasmid pShuttle-CMV.mLuc (Ainscow et al., 2000Go) by digestion with NdeI (which cleaved within the CMV promoter region) and HindIII, and ligated into plasmid pDsRed-N1.



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Fig. 3. Generation and expression of KHCmut conventional kinesin heavy chain construct. (A) cDNA encoding the rat conventional kinesin heavy chain motor domain containing a T93N point mutation at the ATP-binding consensus motif, and a histidine-tag (6His) was cloned into the XbaI-XhoI sites of pcDNA3.1(-). INS-1 cells were transfected, fixed 48 hours later with cold methanol and then co-immunostained with a rabbit polyclonal anti-6His-tag antibody (1:250) (a) and a mouse monoclonal anti {alpha}-tubulin antibody (1:1000) (b) and visualised with Alexa 568 and -488 secondary antibodies, respectively. (c) Overlay of a and b. Overlap appears as yellow. (d-f) High magnification of boxed regions in (a-c). Bars, 2.5 µm (a-c); 1.1 µm (d-f). (B) Cells were co-transfected with KHCmut-pAdTrack-CMV and mitochondrial.DsRed. 48 hours after transfection cells were imaged on a confocal microscope. The KHCmut-expressing cells were identified by exciting EGFP at 488 nm and the DsRed fluorescence of the same cells was visualised by exciting at 568 nm. Typical DsRed in vivo confocal images of mitochondria in INS-1 (a,b) and in HeLa cells (c,d) in KHCmut expressing (a,c) and control cells (b,d) are shown. Scattered lines indicate the position of the plasma membrane obtained as an overlay from the transmitted image of the cell. Bars, 2.5 µm (a-d).

 


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Fig. 8. Expression and effect of KHC340 on vesicle movements and secretion. (A) KHC340 construct was generated as KHCmut described under Fig. 3. (B) INS-1 cells were transfected with the KHC340-pcDNA3.1(-), fixed with cold methanol and then co-immunostained with a rabbit polyclonal anti-6His-tag antibody (1:250) (a) and a mouse monoclonal anti {alpha}-tubulin antibody (1:1000) (b) and visualised with Alexa 568 and -488 secondary antibodies, respectively. (c) Overlay of a and b. Overlap appears as yellow. Bar, 2.5 µm (a-c). (C) Cells were co-transfected with KHC340-pAdTrack-CMV and phogrin.DsRed. Typical 568 nm in vivo confocal images of insulin-containing vesicles (a,c) in KHC340-expressing (a) and control cells (c). Movements of vesicles were imaged and analysed as described in Fig. 5. (b,d) Tracks of granules in a and c. Bars, 2.5 µm (a,c); 1.95 µm (b,d). (D) Effect of KHC340 on glucose-stimulated human growth hormone (hGH) release from MIN6 cells was studied as described in Fig. 7.

 



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Fig. 5. Quantitative analysis of insulin-containing vesicle movements in KHCmut-expressing and control cells. (A) Cells were co-transfected with KHCmut-pAdTrack-CMV and phogrin.DsRed. 48 hours after transfection cells were imaged at a stimulatory glucose concentration (16 mM) on a confocal microscope. The KHCmut-expressing cells were identified by exciting EGFP at 488 nm and the DsRed fluorescence of the same cells was visualised by exciting at 568 nm. Typical 568 nm in vivo confocal images of insulin-containing vesicles (a,c) in KHCmut-expressing (a) and control cells (c) are shown. Images were taken every 5 seconds for 4 minutes or 2 frames/s for 30 seconds (for a total of 60 frames). The movements of vesicles were tracked for 60 frames unless the spot was lost from view. (b,d) Tracks of granules in (a) and (c). Bars, 1 µm (a,c); 0.75 µm (b,d). (B) For quantification of motility, vesicles were randomly selected (20 in each cell) in seven dominant-negative kinesin-expressing (open bars) and seven control cells (closed bars) and the location of granules was determined using the image analysis software MethaMorphTM (see Materials and Methods). The bar diagrams show the probability of vesicles travelling at the indicated velocities. For corresponding movies (Fig. 5Aa,c), see ftp://researcher{at}137.222.66.116/ (long on as `researcher', password c1100cs, directory `kinesin').

 


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Fig. 7. Effect of KHCmut on glucose-stimulated human growth hormone (hGH) release from MIN6 and INS-1 cells. Cells were co-transfected with 0.5 µg hGH-encoding plasmid pXGH5 together with 1 µg KHCmut-pAdTrack-CMV or the corresponding empty vector pAdTrack-CMV. (A) For immunocytochemistry, cells were fixed and co-stained with guinea-pig ployclonal anti-insulin (1:500) and mouse monoclonal anti-hGH (1:150) antibodies and then visualised with Alexa 488 goat anti-guinea pig and Alexa 568 goat anti-mouse secondary antibodies. (a) Alexa 488 and (b) Alexa 568 fluorescence (488 and 568 nm excitations, respectively). (c) Overlay of a and b. Overlap appears as yellow. (d-f) High magnification of boxed regions in a-c. Bars, 2.5 µm (a-c); or 0.4 µm (d-f). (B,C) For hGH assay, cells were cultured for 2 days in complete growth medium and then starved in DMEM medium containing 3 mM glucose 12 hours before stimulation. INS-1 (B) or MIN6 (C) cells were incubated first in the presence of 3 mM glucose for 20 minutes and then at 16 mM or 30 mM glucose for 20 or 90 minutes. Following stimulation, the cells were lysed in 0.5% Triton X-100 and assayed for total hGH using a colorimetric sandwich ELISA method. hGH release was expressed as a percentage of the total hGH and was compared with values obtained with the empty vector (pAdTrack-CMV) at 3 mM glucose. Number of transfections for each condition are indicated in the columns. **P<0.001 compared with basal (empty vector at 3 mM glucose).

 
Adenoviral infection and immunoadsorption of phogrin.EGFP-containing vesicles
INS-1 or MIN6 cells were infected with the recombinant phogrin.EGFP adenoviral construct at a multiplicity of 30-100 viral particles/cell, for 1 hour. Cells were subsequently used 24-48 hour post-infection when >95% of cells were infected. Cells were scraped into ice-cold buffer A [10 mM morpholeno-sulphonic acid (MOPS), 260 mM sucrose, (pH 6.5) 1 µM phenylmethylsulfonyl fluoride (PMSF), 5 µg ml-1 aprotinin, 5 µg ml-1 leupeptin] then homogenised with a teflon homogeniser and centrifuged at 500 g for 1 minute. The post-nuclear supernatant was further centrifuged at 2400 g for 10 minutes at 4°C and the pellet was resuspended in buffer B (50 mM Hepes, 1 mM EDTA, 150 mM NaCl, 1 µM PMSF, 5 µg ml-1 aprotinin, 5 µg ml-1 leupeptin) to a concentration of 1-2 mg ml-1. 100-200 µg of this homogenate was pre-cleared with 100 µl of packed Protein-A sepharose in buffer B overnight and then centrifuged at 14,000 g for 3 seconds; the supernatant was then transferred to a fresh tube. Vesicles carrying phogrin.EGFP were specifically adsorbed using a mouse monoclonal anti-EGFP antibody. Anti-EGFP antibody (20 µl) was first incubated with 50 µl Protein-A sepharose in buffer B overnight, and the beads washed three times with buffer B. Pre-cleared samples (150-250 µl) were added to the antibody-bound beads and incubation continued for a further 24 hours at 4°C. Samples were centrifuged at 500 g for 30 seconds, and the immunoadsorbed vesicles were washed four times with buffer B and then analysed by SDS-PAGE and immunoblotting (Varadi et al., 1996Go). The following control experiments were performed: immunoadsorption with (1) a rabbit polyclonal anti-phogrin antibody that crossreacts with the cytosolic C-terminal domain of phogrin; (2) mouse monoclonal anti-SREBP antibody (sterol response element binding protein-1 is not associated with dense core secretory vesicles) (AzzoutMarniche et al., 2000Go); (3) rabbit pre-immunsera and (4) without any IgGs but in presence of Protein-A sepharose. The presence of kinesin and dynein in the immunoabsobed vesicle preparation was detected using a polyclonal-pan-kinesin antibody (1:1000), a rabbit polyclonal anti-ubiquitous conventional kinesin (uKHC) antibody (1:2000) and a mouse monoclonal anti-dynein antibody (1:1000). Immunostaining was revealed with horseradish-peroxidase conjugated anti-rabbit IgG (1:160,000) and anti-mouse IgG (1:40,000) using an enhanced chemiluminescence (ECL) detection system.

Transient transfection and assay of human growth hormone (hGH) release
INS-1 and MIN6 cells were seeded at a density of 4-6x105/ml on 24-mm-diameter poly-L-lysine-coated coverslips, and cultured overnight. Cells were co-transfected with 0.5 µg hGH-encoding plasmid pXGH5 (Fisher and Burgoyne, 1999Go) together with 1 µg phogrin-pcDNA3 or the corresponding empty vector (pcDNA3); KHCmut-pAdTrack-CMV or the corresponding empty vector (pAdTrack-CMV); KHC340-pAdTrack-CMV or the corresponding empty vector (pAdTrack-CMV) using 10 µg/ml lipofectamine in Optimem I medium for 4 hours. The cells were cultured for 48 hours in complete growth medium (containing 25 mM glucose) which was then replaced with a 3 mM glucose-containing DMEM medium 12 hours prior to stimulation. For assay of hGH release, cells were washed three times in Krebs-Ringer-Hepes-Bicarbonate (KRH) buffer comprising 140 mM NaCl, 3.6 mM KCl, 0.5 mM NaH2PO4, 0.5 mM MgSO4, 2.0 mM NaHCO3, 3 mM glucose, 10 mM Hepes (pH 7.4) and 1.0 mM CaCl2 equilibrated with O2/CO2 (95:5, v/v) at 37°C. Cells were stimulated by incubating with 1 ml KRH buffer containing 3 mM glucose at 37°C. After 20 minutes, 0.5 ml of supernatant was removed and replaced with a high glucose-containing KRH buffer (37°C) to obtain a final glucose concentration of 16 mM or 30 mM. Cells were stimulated for 20 minutes at 37°C then 0.5 ml medium was removed and replaced with KRH buffer containing 16 mM or 30 mM glucose and further incubated for an additional 70 minutes (90 minute time point). At the end of the incubation, the supernatants were removed and the cells lysed in 600 µl of 0.5% (v/v) Triton X-100 (15 minutes at 22°C). The samples were collected and assayed for total hGH content. hGH assay was carried out using a colorimetric sandwich ELISA method according to the manufacturer's instructions (Roche Diagnostics).

Immunocytochemistry and confocal microscopy
Cells were co-transfected with 1 µg plasmid DNA encoding KHCmut or KHC340 and phogrin.DsRed or mitochondrial.DsRed. Immunocytochemistry was performed as described earlier (Pouli et al., 1998Go). Images were captured on a Leica TCS-NT confocal laser-scanning microscope attached to a DM IRBETM epifluorescence microscope using a x63 PL Apo 1.4 NA oil-immersion objective (Leica, Heidelberg, Germany).

Live cell confocal imaging and image analysis
Prior to imaging, cells were incubated in KHR buffer containing a stimulatory concentration of glucose (16 mM or 30 mM) for 5 minutes at 37°C. Cells co-expressing (1) phogrin.DsRed and KHCmut-pAdTrack-CMV or the empty vector (pAdTrack-CMV); or (2) mitochondrial-DsRed and KHCmut-pAdTrack-CMV or the empty vector (pAdTrack-CMV) were identified by exciting EGFP at 488 nm and using FITC emission filters. The DsRed fluorescence of the same cells was visualised by exciting at 568 nm and using TRITC (tetramethylrhodamine isothiocyanate) filters for fluorescence emission.

To study the effect of various cytosolic ATP concentrations on vesicle movement, phogrin.EGFP expressing cells were permeabilised for 1 minute at 20°C with 20 µM digitonin in intracellular buffer containing 140 mM KCl, 10 mM NaCl, 1 mM K2HPO4, 2 mM Na-succinate, 20 mM Hepes, 0.5 mM EGTA, 0.27 mM CaCl2, 0.3-4.0 mM MgCl2 and 5.5 mM glucose, pH 7.05. Concentrations of free Ca2+ and Mg2+ were calculated using `Metlig' software (Rutter et al., 1988Go) to be 0.2 µM and 0.26 mM, respectively.

Images were acquired every 5 seconds for 4 minutes (giving a total of 60 frames) using a Leica TCS-NT confocal laser-scanning microscope. Alternatively, to provide greater temporal resolution, cells were imaged on an UltraVIEWTM Live Cell Confocal Imaging system (PerkinElmer Life Sciences, Boston, MA). In the latter case, images were acquired at a rate of 2 frames second-1 for 30 seconds (60 frames total). The movements of 20 randomly-selected vesicles or mitochondria, that were present in the first image of each recorded sequence, were tracked in each cell using the image analysis software MetaMorphTM (Universal Imaging, West Chester, PA). Fluorescent spots representing DsRed-labeled granules or mitochondria were tracked for 60 frames unless the spot was lost from view or coalesced with another spot. The software displayed the first image in a sequence, and was then directed by mouse click to the granule/mitochondrial structure of interest. The program provided, for each granule tracked, a table of x and y coordinates as function of time. The speed of a given single vesicle or mitochondrion was calculated (µm/s) for images taken on the Leica TCS-NT confocal laser-scanning microscope at the magnification values given. For images acquired on the UltraVIEWTM system, the magnification parameters were set at a constant level, and velocity is given in arbitrary units (Fig. 9B). Differences between the behaviour of vesicles and mitochondria in control and dominant-negative kinesin-expressing cells were assessed by a {chi}2-test (using Yates's Correction) (Moroney, 1951Go) on histograms generated from the velocity data (Fig. 5B; Fig. 6B). The statistical data for one experimental condition was usually obtained from 6-7 cells.



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Fig. 9. ATP dependence of insulin-containing vesicle movements. (A) Cells transfected with phogrin.EGFP were permeabilised using 20 µM digitonin in intracellular buffer. Vesicle movement was viewed on an Ultra VIEWTM Live Cell Confocal Imaging system (see Materials and Methods). Typical 488 nm in vivo confocal images of insulin-containing vesicles (a,c,e,g) in INS-1 cells are shown. Images were taken every 0.5 seconds for 30 seconds (for a total of 60 frames). The location of vesicles was determined using the image analysis software MethaMorphTM. The movements of vesicles were tracked for 60 frames unless the spot was lost from view. (b,d,f,h) Tracks of granules in a, c, e and g. Bars, 2.5 µm (a,c,e,g); 3.25 µm (b,d,f,h). (B) During imaging the magnification parameters were set at a constant level and the velocity is given in arbitary units. Differences between the behaviour of vesicles at various ATP concentrations were assessed by a {chi}2-test (using Yates's correction) on histograms generated from the distance moved data. The inset shows the probability of vesicles moving >2.8 arbitrary units in the presence of 1.0 mM (filled bar) and 5.0 mM (open bar) ATP. For URL address to movies, see legend to Fig. 5.

 


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Fig. 6. Quantitative analysis of mitochondrial movements in KHCmut-expressing and control cells. Cells were co-transfected with KHCmut-pAdTrack-CMV and mitochondrial.DsRed. Movement of mitochondria was analysed as described in Fig. 5. Typical 568 nm in vivo confocal images of mitochondria (a,c) in KHCmut-expressing (a) and control cells (c). (b,d) Tracks of granules in (a) and (c). Bars, 1 µm (a-d). (B) The bar diagrams show the probability of mitochondria travelling at the indicated velocities. For URL address to movies, see legend to Fig. 5.

 

Measurement of intracellular free Ca2+ concentration [Ca2+]i and NAD(P)H
Changes in [Ca2+]i were measured at 37°C with entrapped Fura-2 (Grynkiewicz et al., 1985Go) using a Leica DM-IRBI inverted microscope (x40 objective) and a Hamamatsu C4742-995 charge-coupled device camera driven by OpenLabTM software (Improvision, Coventry, UK) (Ainscow et al., 2000Go). Cells transfected with KHCmut or the empty vector were loaded with 5 µM Fura-2/AM and 0.1% Pluronic F-127 (BASF, Mount Olive, NJ) for 40 minutes in KRH buffer initially containing 3 mM glucose. Autofluorescence due to NAD(P)H was measured as previously described (Ainscow et al., 2000Go).


    Results
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 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Immunoadsorption of phogrin.EGFP-containing vesicles
EGFP was targeted to the limiting membrane of large dense core insulin secretory vesicles (LDCVs) by fusion at the C-terminus of the trans-membrane spanning LDCV protein, phogrin [phosphatase on the granules of insulinoma cells (Wasmeier and Hutton, 1996Go)], to produce chimaeric phogrin.EGFP (Pouli et al., 1998Go). Confirming the correct targeting of phogrin.EGFP to LDCVs, >95% of the expressed protein was localised to insulin-containing structures in both INS-1 (Pouli et al., 1998Go; Tsuboi et al., 2000Go) and MIN6 cells, and no phogrin.EGFP-positive/insulin-negative structures were revealed in >100 single cells examined.

Insulin-containing vesicles were prepared from MIN6 and INS-1 cells by immunoadsorption of the phogrin.EGFP chimaera (see Materials and Methods) using either anti-phogrin (Fig. 1a-e, P) or anti-EGFP antibodies (Fig. 1a-e, GFP). In homogenates (Hom.) from phogrin.EGFP-virus-infected cells, both antibodies bound to a diffuse band migrating with a molecular mass of 82-86 kDa, corresponding well to the expected size of the chimaera (Fig. 1a,b, Hom.). This band was completely absent in homogenates from non-infected cells (not shown). As expected, anti-phogrin antibody also crossreacted with endogenous phogrin with molecular weight 60-64 kDa (Fig. 1a, Hom.). Since the anti-EGFP antibody bound to the phogrin.EGFP chimaera exclusively, this antibody was routinely used for immunoadsorption.



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Fig. 1. Immunoadsorption of phogrin.EGFP-containing vesicles from INS-1 cell homogenates. Cells were infected with the phogrin.EGFP adenoviral construct and homogenised 24 hours after infection (Hom.). Phogrin.EGFP-containing vesicles were then immunoadsorbed using a polyclonal anti-phogrin antibody (P), a monoclonal anti-EGFP antibody (GFP), or an irrelevant monoclonal anti-sterol response element binding protein 1 (SREBP1) antibody (Cont). The immunoadsorbed vesicles were analysed by 7.5-15% SDS-PAGE and immunoblotting. The blots were probed with: (a) an anti-phogrin antibody; (b) an anti-EGFP antibody; (c) an anti-insulin antibody; (d) an anti-glycerol phosphate dehydrogenase (mGPDH) antibody to detect mitochondrial contamination; or (e) an anti-manose-6-phosphate receptor (M6PR) antibody for identifying lysosomal contamination. Molecular weight markers are indicated on the left and arrows show the position of phogrin.EGFP (a,b).

 

Following immunoadsorption with the anti-EGFP antibody, a single 82-86 kDa band was labelled in the resulting immunoprecipitated fraction with either anti-phogrin or anti-EGFP antibodies (Fig. 1a,b, GFP). No detectable phogrin.EGFP chimaera was observed in control samples in which immunoadsorption performed with an irrelevant monoclonal antibody (raised to sterol response element binding protein 1; SREBP1) (AzzoutMarniche et al., 2000Go) (Fig. 1a,b, Cont.), or in the presence of Protein-A sepharose without IgGs, not shown) with either antibody. Furthermore, the immunoadsorbed samples showed intense immunostaining for insulin (Fig. 1c, P, GFP). These samples were next tested for the presence of the most likely contaminating organelles, mitochondria and lysosomes. Neither mitochondrial glycerol phosphate dehydrogenase- (mGPDH) nor lysosomal anti-mannose-6-phosphate receptor (M6PR)-specific antibodies revealed detectable amounts of these marker proteins in the immunoadsorbed samples, which revealed that the immunoadsorption protocol provided a preparation of insulin-containing vesicles of high purity, (Fig. 1d,e, P, GFP), while both antigens were abundant in cell homogenates (Fig. 1d,e, Hom.).

Conventional kinesin is associated with insulin-containing granules in ß-cell lines
Expression of conventional kinesin in MIN6 or INS-1 cells, and its association with phogrin.EGFP-containing vesicles, were examined by immunoblot analysis of cell homogenates and purified secretory vesicles, respectively (Fig. 2A). Two kinesin-specific antibodies were used: (1) a pan-kinesin antibody, raised against a conserved region of the motor domain, and which crossreacts with conventional kinesin heavy chain as well as with a range of kinesin-related proteins (Barroso et al., 2000Go; Sawin et al., 1992Go); and (2) an anti-ubiquitously expressed) conventional kinesin heavy chain (uKHC) antibody, which was raised against the less conserved regions of the {alpha}-helical coiled-coil domain of conventional KHC (Niclas et al., 1994Go). The pankinesin antibody crossreacted with a protein migrating with molecular mass ~120 kDa in the immunoadsorbed samples without labelling any other kinesin or kinesin-related protein (Fig. 2A, left panel, GFP). However, this antibody labelled several other, presumably kinesin-related proteins, in INS-1 cell homogenates (Fig. 2A, left panel, Hom.). The anti-uKHC antibody also crossreacted with the 120 kDa protein in INS-1 cell homogenate (Fig. 2A, middle panel, Hom.) and immunoadsorbed samples (Fig. 2A, middle panel, GFP). Immunocytochemistry of INS-1 cells with the uKHC antibody also revealed, in agreement with the above data, that a small proportion of conventional kinesin (10-20% of total) is associated with insulin-containing vesicles (Fig. 2B).



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Fig. 2. Conventional kinesin heavy chain (KHC) is associated with insulin granules in ß-cells. (A) INS-1 cells were homogenised (Hom.) and the phogrin.EGFP-containing vesicles were immunoadsorbed with a monoclonal anti-EGFP antibody (GFP) or with a monoclonal anti-SREBP antibody (Cont.) (for details, see Materials and Methods). Immunoblots were probed with a polyclonal-pan-kinesin antibody (left panel), a rabbit polyclonal anti-ubiquitous conventional KHC antibody (uKHC) (middle panel) or a monoclonal anti-dynein antibody (right panel). 15 µg protein was loaded from the cell homogenates and 2.5 µg from purified kinesin from pig brain (Kin); the protein content of the immunoadsorbed samples was not determined. Note that the immunoreactive band of ~60 kDa detected by the anti-dynein antibody in the immunoadsobed sample is corresponds to IgG. (B) INS-1 cells were co-immunostained with a rabbit polyclonal anti-uKHC (1:200) and a guinea pig polyclonal anti-insulin (1:500) antibody and visualised with an Alexa 488 goat anti-rabbit (1:500) and an Alexa 568 goat anti-guinea pig (1:500) secondary antibody. (a) Alexa 488 fluorescence (488 nm excitation). (b) Alexa 568 fluorescence (568 nm excitation). (c) Overlay of a and b. Overlap appears as yellow. (d-f) High magnification of boxed regions in (a-c). Bars, 5 µm (a-c); 0.5 µm (d-f).

 

To confirm that the relatively small proportion of total conventional kinesin that was associated with vesicles (Fig. 2A, left and middle panels, GFP) did not result from contamination with cytosol, we screened the vesicle fraction with an antibody to an abundant cytosolic protein, dynein. While dynein immunoreactivity was abundant in the cell homogenate (Fig. 2A, right panel, Hom.), this immunoreactivity was undetectable in the vesicle preparation (Fig. 2A, right panel, GFP) (note that the strongly labelled ~60 kDa band corresponds to IgG).

Dominant-negative-acting kinesin (KHCmut) does not affect the sub-cellular localisation of membrane-bound organelles
cDNA encoding the motor domain of rat KHC, carrying a point-mutation (T93N) and a 6His-tag (KHCmut), was introduced into INS-1 or MIN6 cells alone or with cDNA encoding EGFP (on the same plasmid) (Fig. 3A,B). The sub-cellular localisation of the KHCmut protein in INS-1 cells was studied by immunocytochemistry using an anti-6His-tag and an anti-{alpha}-tubulin antibodies (Fig. 3A,a-f). As expected, anti-6His-tag antibody showed a filamentous staining pattern (Fig. 3A,a,d) and most of the over-expressed KHCmut (>90%) localized to microtubules (Fig. 3A,c,f).

Since kinesin is thought to be involved in membrane transport to the plus-end of microtubules (anterograde transport), we determined whether over-expression of mutant kinesin caused a generalised blockade of membrane transport in the cells. Mitochondria, which have been suggested to move along MTs in other cell types (Ball and Singer, 1982Go) were visualised by monitoring the fluorescence of mitochondrially targetted DsRed (see Materials and Methods). As shown in Fig. 3, mitochondria in INS-1 or MIN6 cells transfected with KHCmut (Fig. 3Ba), empty vector (Fig. 3Bd) or KHC340 (not shown), were widely distributed throughout the cytoplasm in each case. In marked contrast, however, KHCmut overexpression in HeLa cells, to levels comparable with those obtained in INS1 or MIN6 cells, induced the collapse of mitochondrial structure to the perinuclear region (Fig. 3Bc,d).

We next determined the effect of KHCmut expression on the distribution of LDCVs as well as proximal elements of the secretory pathway. Insulin-containing vesicles were visualised after cell fixation with an anti-insulin antibody (Fig. 4a-f) or, in live cells, by expressing a phogrin. DsRed chimaera (Fig. 5A). No evident difference in the intensity or pattern of insulin staining was apparent in cells transfected with either mutant kinesin or empty vector, suggesting that the synthesis and targeting of insulin was not impaired by inhibition of kinesin function. Moreover, the distribution of phogrin.DsRed-labelled secretory vesicles was unaffected in live cells by expression of mutant kinesin (Fig. 5A), although we occasionally (in 5-10% of cells) observed accumulation of vesicles towards the centre of mutant-kinesin-transfected cells expressing very high levels of KHCmut (not shown). Stained with an anti-TGN38 antibody, perinuclear localisation of the trans-Golgi network was observed in both KHCmut-expressing cells and non-transfected cells (Fig. 4g-i). Thus, the assembly and position of the Golgi apparatus was unaffected by introduction of KHCmut.



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Fig. 4. Localisation of insulin and Golgi apparatus in KHCmut-expressing (a-c, g-i; arrow) and control (d-f, g-i) ß-cells. Cells were transfected with the KHCmut-pAdTrack-CMV construct (A). Downstream of the multiple cloning site, the shuttle vector carries cDNA for EGFP driven by a distinct second CMV promoter. 48 hours after transfection the cells were fixed and probed with (a-f) a guinea-pig anti-insulin (1:500) or (g-i) a mouse monoclonal anti-TGN38 antibody (1:100) and then visualised with the appropriate Alexa 568 secondary antibodies. (a,d,g) Transmitted light images. (b,e,h) Alexa 568 fluorescence (568 nm excitation). (c,f,i) Intrinsic EGFP fluorescence (488 nm excitation). Bars, 2.5 µm (a-i).

 

Dominant-negative kinesin blocks glucose-stimulated excursions of insulin-containing granules but has no effect on mitochondrial motion
To study the effect of a mutant kinesin on insulin-containing vesicle movement in real time, ß-cells were transfected with cDNAs encoding phogrin.DsRed in the presence or absence of an expression construct encoding mutant kinesin. Importantly, expression of phogrin.DsRed had no impact on either the early or late phases of glucose-stimulated hGH secretion (not shown) demonstrating that this construct is unlikely to perturb vesicle recruitment to sites of exocytosis (not shown).

To quantitate the impact of the mutant kinesin on secretory granule movement (Fig. 5), live cell imaging was performed first using confocal microscopy. As previously described (Pouli et al., 1998Go) in the presence of stimulatory glucose concentrations >5 mM) vesicles displayed both short oscillatory movements (<1-2 vesicle diameters) but also longer excursions (several microns). The latter excursions were completely absent at low glucose concentrations (Pouli et al., 1998Go). Averaged in a random sample of the whole vesicle population of single cells, vesicles travelled a significantly shorter distance per unit time ({chi}2=P<0.1%) in mutant kinesin-transfected cells than in control cells (Fig. 5Aa-d). Thus, the majority of vesicles (85%) in the mutant kinesin-expressing cells moved with velocities of 0.2 µm second-1 or less, and no vesicle moved more than 0.4 µm second-1 (Fig. 5B). By contrast, 38% of vesicles in control cells moved 0.2 µm second-1, 27% travelled 0.4 µm second-1, and 35% of the vesicles (versus 0% after introduction of KHCmut) moved more than 0.4 µm second-1 (Fig. 5B). By contrast, mitochondrial movement (which was insensitive to glucose concentration; E.K.A. and G.A.R., unpublished) was not significantly affected ({chi}2=P>5%) by the expression of the mutant kinesin (Fig. 6A,B). These data suggest that, in the clonal ß-cell lines examined here, conventional kinesin plays an important role in the activitation of secretory vesicle movement by elevated glucose concentrations.

Conventional kinesin is important for the substained phase of glucose-stimulated secretion from ß-cells
We next investigated how the reduced vesicle motility, caused by the inhibition of kinesin function, may affect glucose-stimulated insulin secretion. In contrast to whole islets (Curry et al., 1968Go) both INS-1 (Asfari et al., 1992Go) and MIN6 (Ainscowet al., 2000Go) ß-cells display essentially monophasic insulin release in which the early (KATP-channel-dependent) phase of release (Asfari et al., 1992Go) is followed by sustained release of the hormone with no clear nadir between the two phases. To distinguish the effects of kinesin on the first and sustained phases of hormone release in these two cellular models, we co-transfected cells with cDNAs encoding hGH and dominant-negative kinesin (see Materials and Methods), and monitored the release of hGH after either 20 or 90 minutes. hGH released during these two different periods was then taken as a guide to the two phases of insulin release. As revealed by immunocytochemistry (Fig. 7A), expressed hGH was correctly targeted into dense core vesicles of the regulated secretory pathway, and KHCmut was efficiently introduced into all cells expressing hGH. Thus, of 150 cells examined that expressed EGFP, all were positive for the 6His-tag of KHCmut (not shown). Furthermore, the majority of cells (135 of 150 from three independent experiments) showed positive staining for both hGH and 6His-tag (not shown). Providing further confirmation that introduction of KHCmut did not inhibit the correct targeting of hGH (and presumably insulin) into dense core vesicles, the rate of release of hGH was low under basal conditions (2.3±0.15% and 2.7±0.23% for control and KHCmut-transfected INS-1 cells, respectively, at 3 mM glucose; Fig. 7B). In this cell line, expression of the dominant-negative kinesin strongly reduced the stimulation by 30 mM glucose of hGH release measured after a 90 minute incubation (2.89±0.32 versus 19.62±0.29-fold, for KHCmut-transfected and control cells, respectively; P<0.001 for the effect of KHCmut) but was without effect on secretion stimulated by 30 mM glucose after a 20 minute incubation (Fig. 7B).

Similar to INS-1 cells, the rate of release of hGH from MIN6 cells at basal (3 mM) [glucose] was low, and nearly identical in empty vector and dominant-negative kinesin-transfected cells (1.26±0.07 versus 1.17±0.13% of total cellular hGH; Fig. 7C). However, after a 20 minute incubation at 16 mM glucose, hGH secretion was stimulated more than fivefold in both groups (Fig. 7C), with no significant difference between the two (6.99±0.297 versus 8.53±1.87-fold above the basal release rate for control and KHCmut-transfected cells, respectively; Fig. 7C). Strikingly, after a 90 minute incubation, glucose-stimulated hGH release from dominant-negative kinesin-expressing MIN6 cells was almost abolished (3.35±0.24 versus 40.61±0.18-fold, for KHCmut-transfected and control cells, respectively; P<0.001; Fig. 7C).

To determine whether the effect of KHCmut was due specifically to a dominant-negative action of this protein on endogenous KHC we next overexpressed wild- type KHC340, which did not carry the T93N point mutation (Fig. 8A), and studied its effect on vesicle movements and secretion. This protein showed homogenous cytosolic staining in ß-cells (Fig. 8A) and, as expected, did not strongly localise to the microtubules (Fig. 8A). KHC340 expression had no effect on vesicle movements (Fig. 8B) or hGH release (Fig. 8C).

To determine whether the effects of mutant kinesin may be the indirect result of an alteration in glucose metabolism or intracellular Ca2+ handling, changes in intracellular NAD(P)H or [Ca2+] in response to KCl or prolonged glucose stimulation (90 minutes) were also monitored. No difference was observed between dominant-negative kinesin- and empty vector-expressing cells (data not shown).

Cytosolic ATP stimulates vesicle movements within the physiological concentration range
Since kinesin activity is ATP-dependent, we next explored the possibility that this motor protein may permit vesicle movement to be regulated by changes in cytosolic ATP concentration. We therefore studied the ATP dependence of vesicle movements in permeabilised INS-1 cells (see Materials and Methods). In the complete absence of cytosolic ATP, virtually all vesicle movements stopped (Fig. 9Aa,b, 9B). Changing the MgATP concentration to 0.1 mM (Fig. 9Ac,d, 9B) significantly (P<0.01) increased the velocity of vesicles. Thus, whereas only 2.5% of vesicles moved more than 0.8 arbitrary distance units frame-1 in the absence of ATP, versus 61.8% at 0.1 mM ATP. By contrast, in the presence of 1 mM MgATP, (which corresponds to the resting cytosolic [ATP] in ß-cells) (Kennedy et al., 1999Go), (Fig. 9Ae,f, 9B) the movement of vesicles significantly increased (P<0.01 with respect to zero MgATP) and 17.7% of vesicles moved with a velocity of over 1.6 arbitrary distance units frame-1 at 0.1 mM ATP and 39% at 1 mM ATP. Addition of 5 mM MgATP, a concentration in the range of [ATP] reached in intact ß-cells after elevation of [glucose] from 3 mM to 30 mM (Kennedy et al., 1999Go; Ainscow and Rutter, 2001Go), further increased the velocity of vesicles. Thus, 12.8% of vesicles moved with a velocity of over 2.8 arbitrary distance units frame-1 at 1 mM ATP, whereas 21.7% of vesicles achieved this velocity at 5 mM ATP (P<0.01; Fig. 9Ag,h, 9B).


    Discussion
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 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Conventional kinesin is the principal (+) end-directed regulated secretory vesicle motor protein in two clonal ß-cell lines
Here we have developed a method by which targeted EGFP can be used for the immunoadsorption and isolation of insulin-containing vesicles. This has allowed us to identify motor proteins that may be responsible for the anterograde translocation of insulin-containing vesicles to the cell surface when glucose concentrations rise. This new method provided a highly pure vesicle preparation that was essentially free of mitochondrial and lysosomal contamination (Fig. 1a-e). Conventional KHC was abundant in these preparations (Fig. 2), consistent with earlier studies that showed expression of Kif5b conventional KHC in RIN-m38 and primary mouse ß-cells (Balczon et al., 1992Go; Meng et al., 1997Go). By contrast, other members of the kinesin family, clearly present in MIN6 and INS-1 cells revealed by western analysis of whole cell homogenates (Fig. 2A), were completely absent from the vesicle preparation. These data therefore strongly implicate conventional kinesin as the motor protein responsible for anterograde transport of vesicles during glucose-stimulated exocytosis of insulin (Fig. 2).

Conventional kinesin is required for stimulated vesicle movement
To explore the functional importance of vesicle-associated kinesin, described above, we overexpressed the motor domain of rat conventional KHC containing a T93N mutation in the catalytic (ATP-binding) site. This construct has previously been shown to function as a dominant-negative inhibitor of kinesin function (Nakata and Hirokawa, 1995Go; Krylyshkina et al., 2002Go). Thus, it has been shown that KHCmut, which is a `rigor' kinesin, binds tightly to but rarely detaches from MTs, and does not support MT motility in other cell types (Nakata and Hirokawa, 1995Go; Krylyshkina et al., 2002Go). This molecular genetic approach was used in preference to the introduction of anti-kinesin antibodies (Bi et al., 1997Go), since it permitted parallel studies of both vesicle movement (Fig. 5) and exocytosis from cell populations (Fig. 7).

Interestingly, KHCmut acted specifically to block glucose-induced, but not un-stimulated, movement of vesicles (Fig. 5). Thus, KHCmut had no effect on the distribution of vesicles under basal conditions, indicating that transport of vesicles to their location within the cell prior to stimulation is independent of vesicle-type. Moreover, a generalised blockade of MT transport was not observed after KHCmut expression as indicated by the observations that: (1) the localisation of mitochondria, Golgi apparatus and insulin-containing vesicles were all similar in mutant kinesin-transfected and control cells (Fig. 3B, Fig. 4, Fig. 6); and (2) the synthesis and storage of co-transfected hGH was unaltered (Fig. 7). Taken together, these observations revealed that KHCmut interferes specifically with the binding of active kinesin to secretory vesicles and their movement in response to elevated glucose concentrations. Interestingly, an earlier study showed that the rigor kinesin specifically blocked anterograde transport of lysosomes in mouse fibroblast L cells (Nakata and Hirokawa, 1995Go). Thus a similar, kinesin-dependent mechanism, may be involved in the long range transport of both dense core secretory vesicles and lysosomes, although the former would appear to occur only after cell stimulation.

Role of kinesins in mitochondrial distribution and movement in ß-cells
In common with our own findings in pancreatic ß-cells, the rigor kinesin did not affect the localisation of Golgi apparatus, mitochondria or lysosomes in mouse fibroblast L cells (Nakata and Hirokawa, 1995Go). In contrast, KHCmut caused a complete collapse of mitochondrial structure in HeLa cells (Fig. 3B), a phenomenon previously observed in Xenopus and fish fibroblasts (Krylyshkina et al., 2002Go), as well as in undifferentiated extra-embryonic cells from KIF5b null mutant mice (Tanaka et al., 1998Go). Thus, it would appear that in some cell types, including fibroblasts, normal mitochondrial distribution requires active kinesin and, in particular, KIF5b-driven movement along MTs. However, it seems likely that kinesins play little or no role in the overall distribution of mitochondria in smaller mammalian cells, including ß-cells.

We observed relatively slow short-range movements of the mitochondria in ß-cells (Fig. 6), which can probably be explained in large part by Brownian (thermal) motions (Margineantu et al., 2000Go). Thus, the velocities we measured for mitochondrial movements (Fig. 6.) were in the range previously described for mitochondrial motions after the suppression of longer excursions in axons (Ligon and Steward, 2000Go). KHCmut expression had no effect on these movements which occurred with the same range of velocities as those of secretory vesicles at low glucose concentrations [data not shown and (Pouli et al., 1998Go)] or after introduction of KHCmut. However, longer range movements of mitochondria [which were very rarely observed in the time frame (~5 minutes) of the experiments described here], but which may be important for determining the distribution of mitochondria in the cell, would appear not to be driven primarily by kinesin motors in the islet ß-cells.

Importance of conventional kinesin for glucose-stimulated insulin secretion
Remarkably, the dramatically reduced motility of vesicles in KHCmut-expressing cells (Fig. 5) did not affect basal secretion or the initial phase of glucose-stimulated growth hormone release in INS-1 or MIN6 cells (Fig. 7). However, the later (sustained) phase of secretion was completely inhibited by KHCmut in both cell types. These results are most simply interpreted in terms of the two-compartment model for insulin release (Grodsky et al., 1969Go; Rorsman et al., 2000Go; Daniel et al., 1999Go), as illustrated in Fig. 10. Since the initial phase of secretion is shown here to be independent of kinesin-driven vesicle transport (Fig. 7B, Fig. 10), the vesicles involved must have passed through recruitment steps and be very close to, or functionally docked at, the plasma membrane. By contrast, the requirement for conventional kinesin function for sustained secretion (Fig. 7B), indicates that this process involves the recruitment of vesicles from an intracellular site away from the membrane (Fig. 7B). Blockade of kinesin function with a monoclonal antibody against the kinesin motor domain (SUK4) inhibited only the slow (second) phase of Ca2+-regulated exocytosis in wounded sea urchins eggs (Bi et al., 1997Go), which suggests that this feature of regulated exocytosis is similar in both mammalian cells (INS-1 and MIN6) and those of lower organisms.



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Fig. 10. Role of kinesin-mediated vesicle translocation in glucose-stimulated secretion. Typical biphasic insulin secretion from pancreatic islet ß-cells when glucose concentration is elevated to 16 mM (upper panel) and the corresponding glucose-stimulated vesicle movements (lower panel) are shown. (1) At sub-stimulatory [glucose], a small proportion (~5% of total) (Rorsman et al., 2000Go) of granules is docked and immediately available for release (releasable pool). However, most of the granules are situated some distance from the plasma membrane in a `reserve pool' and need to be chemically modified or physically translocated to release sites. (2) Shortly after elevation of [glucose], granules that are already docked undergo exocytosis and transient stimulation of insulin secretion is observed. (3) Kinesin-dependent vesicle recruitment. Note that the nadir in the rate of secretion between end of exocytosis of vesicles in the releaseable pool (2), and the activation of kinesin-dependent vesicle recruitment (3) is postulated to be longer in islets than cell lines, where the two phases of secretion are not readily distinguished. (4) ATP-dependent mobilisation of granules from the reserve pool via kinesin-mediated transport. This stage represents the sustained release of insulin. (5) KHCmut blocks the transport of granules from the reverse pool by irreversibly binding to microtubules (Nakata and Hirokawa, 1995Go; Krylyshkina et al., 2002Go) and inhibits the second phase of insulin release. KHC, kinesin heavy chain; KHCmut, motor domain of kinesin heavy chain containing a T93N point mutation; KLC, kinesin light chain; MT, microtubule.

 

While the present studies highlight an important role for kinesin in secretory vesicle movement, it should be noted that they do not exclude a role for other motor proteins, including myosin [(Iida et al., 1997Go; Yu et al., 2000Go; Li et al., 1994Go) in particular type II (Wilson et al., 1998Go) or V (Titus, 1997Go)], perhaps at a later step in vesicle transport to the plasma membrane. Thus, inhibition of myosin ATPase activity with 2,3-butanedione monoxime significantly decreases secretion from sea urchin embryonic cells (Bi et al., 1997Go). Unfortunately, the effect of this inhibitor could not be tested in ß-cells in the present study since it also inhibits Ca2+-channels under some conditions (Byron et al., 1996Go).

Role of vesicle-associated kinesin as a potential ATP sensor in glucose stimulated insulin release
Analysis of capacitance changes during the release of caged ATP (Eliasson et al., 1997Go) and studies with permeabilized cells (Li et al., 1994Go) have demonstrated that Ca2+-induced exocytosis in ß-cells is highly dependent on access to cytoplasmic ATP. This requirement is largely confined to the second phase of insulin release (Rorsman et al., 2000Go). Our new data (Fig. 9.) suggest that kinesin-driven vesicle transport, which requires the hydrolysis of ATP, may be one site at which glucose-induced increases in [ATP] could act to enhance the second phase of insulin secretion. Thus, vesicle movement was (1) entirely dependent on [ATP] (Fig. 9); and (2) was stimulated by [ATP] increases over the range observed during challenge of ß-cells with glucose (Kennedy et al., 1999Go; Detimary et al., 1998Go). An important question is whether, in the ß-cell, kinesin may be a direct target of glucose-triggered increases in [ATP] (acting at the catalytic site of the ATPase domain) or an indirect target of the nucleotide [e.g. via a protein kinase (Hisatomi et al., 1996Go)]. Future studies will be required to explore these possibilities.


    Acknowledgments
 
This work was supported by project grants from the Biotechnology and Biological Sciences Research Council (BBSRC), the Medical Research Council (UK), The Wellcome Trust, Diabetes UK, the Human Frontiers Science Program, the European Union and the UK Joint Infrastructure Fund (Office of Science and Technology & Wellcome Trust). V.J.A. was supported by a Senior Fellowship from the Lister Institute and E.K.A. was supported by an MRC Research Training Fellowship. We thank the Medical Research Council for providing an Infrastructure Award and Joint Research Equipment Initiative Grant to establish the School of Medical Sciences Cell Imaging Facility, and Mark Jepson and Alan Leard for their assistance.


    References
 Top
 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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