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
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Summary |
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Key words: Kinesin, Insulin, Exocytosis, Glucose, Islet, ß-cell, Pancreas
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Introduction |
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The molecular basis of the second phase of secretion is poorly understood
(Rorsman et al., 2000;
Daniel et al., 1999
). Since
only fuel secretagogues such as glucose trigger the sustained phase of release
(Rorsman et al., 2000
), 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.,
1996
; Daniel et al.,
1999
; Rorsman et al.,
2000
).
In previous studies in which we imaged the behaviour of vesicle-targeted
enhanced green fluorescent protein (phogrin.EGFP)
(Pouli et al., 1998;
Tsuboi et al., 2000
), elevated
glucose concentrations stimulated both short and longer excursions of
vesicles. Montague and colleagues
(Montague et al., 1975
) 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., 1998
). 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., 1994
).
The above, indirect observations, suggest that kinesin, or kinesin-related
motor proteins (KRPs) (Lane and Allan,
1998), 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, 1993
). Kinesin is
required for axonal transport in neuronal cells
(Rahman et al., 1999
;
Gindhart et al., 1998
), and
recruits vesicles to the release sites of Ca2+-regulated exocytosis
in sea urchin embryos (Bi et al.,
1997
).
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.,
1990). 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.,
1998
), and Kif5b is expressed in primary mouse
ß-cells (Meng et al.,
1997
). Furthermore, suppression of Kif5b with antisense
oligonucleotides reduced, but did not altogether abolish, glucose-stimulated
insulin release from primary ß-cells
(Meng et al., 1997
).
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., 1999). Regulation of kinesin activity may thus contribute to
the KATP channel-independent stimulation of insulin secretion by
nutrients (Aizawa et al., 1998
;
Takahashi et al., 1999
;
Seghers et al., 2000
)
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Materials and Methods |
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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., 1995). 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., 2002) was
cloned across the XbaI-XhoI sites of pcDNA 3.1(-) and then
into the pAdTrack-CMV shuttle vector (He
et al., 1998
) via XbaI-EcoRV sites
(Fig. 3A,B
Fig. 8A,B). Generation of
phogrin.pEGFP.N1 (Pouli, et al.,
1998
) and sub-cloning into a recombinant adenovirus were as
described earlier (Tsuboi, et al.,
2000
). 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.,
1989
). The leader sequence was removed from plasmid
pShuttle-CMV.mLuc (Ainscow et al.,
2000
) by digestion with NdeI (which cleaved within the
CMV promoter region) and HindIII, and ligated into plasmid
pDsRed-N1.
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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,
1999) 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., 1998). 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., 1988) 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 2-test (using
Yates's Correction) (Moroney,
1951
) 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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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.,
1985) 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., 2000
). 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., 2000
).
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Results |
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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.
|
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.,
2000) (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., 2000;
Sawin et al., 1992
); and (2)
an anti-ubiquitously expressed) conventional kinesin heavy chain (uKHC)
antibody, which was raised against the less conserved regions of the
-helical coiled-coil domain of conventional KHC
(Niclas et al., 1994
). 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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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--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,
1982) 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.
|
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., 1998) 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.,
1998
). Averaged in a random sample of the whole vesicle population
of single cells, vesicles travelled a significantly shorter distance per unit
time (
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 (
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., 1968) both
INS-1 (Asfari et al., 1992
) and
MIN6 (Ainscowet al., 2000
)
ß-cells display essentially monophasic insulin release in which the early
(KATP-channel-dependent) phase of release
(Asfari et al., 1992
) 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., 1999),
(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., 1999
;
Ainscow and Rutter, 2001
),
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).
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Discussion |
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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,
1995; Krylyshkina et al.,
2002
). 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, 1995
;
Krylyshkina et al., 2002
).
This molecular genetic approach was used in preference to the introduction of
anti-kinesin antibodies (Bi et al.,
1997
), 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, 1995). 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, 1995).
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.,
2002
), as well as in undifferentiated extra-embryonic cells from
KIF5b null mutant mice (Tanaka et al.,
1998
). 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., 2000).
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,
2000
). 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., 1998
)] 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., 1969;
Rorsman et al., 2000
;
Daniel et al., 1999
), 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., 1997
), which
suggests that this feature of regulated exocytosis is similar in both
mammalian cells (INS-1 and MIN6) and those of lower organisms.
|
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., 1997;
Yu et al., 2000
;
Li et al., 1994
) in particular
type II (Wilson et al., 1998
)
or V (Titus, 1997
)], 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., 1997
).
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., 1996
).
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., 1997) and
studies with permeabilized cells (Li et
al., 1994
) 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., 2000
). 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., 1999
;
Detimary et al., 1998
). 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.,
1996
)]. Future studies will be required to explore these
possibilities.
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Acknowledgments |
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References |
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