Human Insulin Vesicle Dynamics During Pulsatile Secretion

Original Article Human Insulin Vesicle Dynamics During Pulsatile Secretion Darren J. Michael,1 Wenyong Xiong,1 Xuehui Geng,2 Peter Drain,2 and Robert H. Chow1

In healthy individuals, plasma insulin levels oscillate in both fasting and fed states. Numerous studies of isolated pancreata and pancreatic islets support the hypothesis that insulin oscillations arise because the underlying rate of insulin secretion also oscillates; yet, insulin secretion has never been observed to oscillate in individual pancreatic ␤-cells. Using expressed fluorescent vesicle cargo proteins and total internal reflection fluorescence (TIRF) microscopy, we demonstrate that glucose stimulates human pancreatic ␤-cells to secrete insulin vesicles in short, coordinated bursts of ⬃70 vesicles each. Randomization tests and spectral analysis confirmed that the temporal patterns of secretion were not random, instead exhibiting alternating periods of secretion and rest, recurring with statistically significant periods of 15– 45 s. Although fluorescent vesicles arrived at the plasma membrane before, during, and after stimulation, their rate of arrival was significantly slower than their rate of secretion, so that their density near the plasma membrane dropped significantly during the cell’s response. To study in greater detail the vesicle dynamics during cyclical bursts of secretion, we applied trains of depolarizations once a minute and performed simultaneous membrane capacitance measurements and TIRF imaging. Surprisingly, young fluorescent insulin vesicles contributed at least half of the vesicles secreted in response to a first train, even though young vesicles were vastly outnumbered by older, nonfluorescent vesicles. For subsequent trains, young insulin vesicles contributed progressively less to total secretion, whereas capacitance measurements revealed that total stimulated secretion did not decrease. These results suggest that in human pancreatic ␤-cells, young vesicles are secreted first, and only then are older vesicles recruited for secretion. Diabetes 56:1277–1288, 2007

From the 1Department of Physiology and Biophysics, Keck School of Medicine, Zilkha Neurogenetic Institute, University of Southern California, Los Angeles, California; and the 2Department of Cell Biology and Physiology, School of Medicine, University of Pittsburgh, Pittsburgh, Pennsylvania. Address correspondence and reprint requests to Robert H. Chow, Department of Physiology and Biophysics, Keck School of Medicine, Zilkha Neurogenetic Institute, University of Southern California, Los Angeles, CA 90089. E-mail: [email protected]. Received for publication 20 March 2006 and accepted in revised form 6 February 2007. Published ahead of print at http://diabetes.diabetesjournals.org on 22 February 2007. DOI: 10.2337/db06-0367. Additional information for this article can be found in an online appendix at http://dx.doi.org/10.2337/db06-0367. D.J.M. and W.X. contributed equally to this work. C-Bright, C-peptide fused to the bright mutant of dsRed fluorescent protein; C-EGFP, C-peptide fused to enhanced green fluorescent protein; C-emGFP, C-peptide fused to emerald green fluorescent protein; FBS, fetal bovine serum; rIAPP-EGFP, rodent isoform of islet amyloid polypeptide fused to enhanced green fluorescent protein; TIRF, total internal reflection fluorescence. © 2007 by the American Diabetes Association. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

DIABETES, VOL. 56, MAY 2007

N

umerous studies (1– 8) suggest that pulsatile insulin secretion is a fundamental property of the healthy pancreas, extending down to the level of single islets and, presumably, to the level of the single ␤-cell. However, no previous study has provided direct evidence in single pancreatic ␤-cells for such coordinated secretion of insulin vesicles. Studies of insulin secretion are especially challenging in primary cultures of pancreatic ␤-cells because the glucosesensitive secretion pathway can be perturbed while cells are prepared and maintained in culture. At present, there is no definitive test for determining the health of cultured pancreatic ␤-cells. Because insulin oscillations degrade during the development of both type 1 (9) and type 2 (10) diabetes, loss of oscillations is likely a sign of unhealthy ␤-cells, and, conversely, pulsatile insulin secretion is likely a key indicator of health for pancreatic ␤-cells. In the work reported here, we studied the dynamics of insulin vesicle trafficking in human pancreatic ␤-cells maintained in primary culture. Because it is known that the function of pancreatic ␤-cells is disturbed by dissociation into individual cells (11), we limited our study to cells that resided within small clusters (5–10 cells). To label insulin vesicles, we transduced the cells with a viral vector encoding a vesicle-targeted fluorescent cargo protein (12). By using a low titer of virus, we were able to find clusters containing only a single cell with fluorescent vesicles, allowing us to study the dynamics of individual insulin vesicles in single pancreatic ␤-cells. We focused in particular on those cells that responded to glucose stimulation with pulsatile insulin secretion, using total internal reflection fluorescence (TIRF) microscopy (13) to image directly individual insulin vesicles that were made fluorescent (14,15). Because a TIRF microscope confines light to a thin optical slice at the base of a cell, it is especially well suited for monitoring the dynamics of membrane-proximal insulin vesicles. Importantly, the population of membrane-proximal vesicles is believed to include those vesicles ready to undergo rapid secretion during the first moments after a stimulus arrives. These “release-ready” or “readily releasable” vesicles (16) have completed many of the steps required for regulated secretion, including docking and priming at the plasma membrane, so that they can fuse within milliseconds after stimulation (17). Detailed functional studies based on electrophysiological (18) and electrochemical (19) approaches have clearly demonstrated in a number of endocrine cell types that the readily releasable pool of vesicles is much smaller than the total cellular pool of vesicles. For example, in rodent pancre1277

1278

*Overall secretion rate ⫽ (total vesicles secreted)/(total imaging time); †burst secretion rate ⫽ (vesicles secreted in pulse)/(pulse duration); ‡maximum secretion rate ⫽ maximum (vesicles secreted in single frame)/(frame duration). For cells stimulated with glucose, the data are compiled from the first 10 min of stimulation. §P ⬍ 0.05, Student’s two-sided t test; 㛳compared to lag for glucose-stimulated rat cells; ¶calculated using the 1st min of secretion after stimulation; #compared to burst secretion rate for glucose-stimulated rat cells; **median value for the distribution of all pulse intervals in all cells. Pulses were identified by a sudden change in the slope of the cumilative secretion plot. See figure legends for statistically significant pulse intervals identified in specific cells. ††Compared to lag for glucose-stimulated human cells; ‡‡compared to arrival rate for glucose-stimulated rat cells. NA, not applicable.

0.70 ⫾ 0.9 NA 80 ⫾ 6 NA 50** 5 1.7 ⫾ 0.5§†† 0.18 ⫾ 0.03 14.8 ⫾ 3.5 5 63 ⫾ 19 0.08 ⫾ 0.04 18.4 ⫾ 7.8

4.75 ⫾ 0.65 NA 3.79 ⫾ 0.67 6.8 ⫾ 1.3 0.22 ⫾ 0.04¶ 1.00 ⫾ 0.35 0.05 ⫾ 0.02. 0.26 ⫾ 0.08

0.74 ⫾ 0.11 (13) 1,100 ⫾ 150 0.43 ⫾ 0.18 (5)§‡‡ 900 ⫾ 210

0.70 ⫾ 0.10 1,200 ⫾ 50 1,500 ⫾ 300 0.13 ⫾ 0.03 (3) 0.07 ⫾ 0.02 (8) NA 79 ⫾ 4 0.10 ⫾ 0.02¶ 1.29 ⫾ 0.35§# 4.69 ⫾ 1.32 NA 0.02 ⫾ 0.01 0.25 ⫾ 0.09 3.13 ⫾ 1.73 5.5 ⫾ 0.9 NA 46** 0.09 ⫾ 0.03 5.2 ⫾ 1.4 0.02 ⫾ 0.01 10.2 ⫾ 2.7 5 2.0 ⫾ 0.6§㛳 8 54 ⫾ 30

Lag (s) n Simulus type

Rat K⫹ Glucose Human K⫹ Glucose

Overall*

Burst†

Vesicles Number secreted of in bursts Maximum‡ pulses (%) Pulse interval (s) Pulse Pulse mass duration (s) (ves/␮m2)

Secretion rate (ves/min per ␮m2)

TABLE 1. Insulin vesicle secretion kinetics in primary cultures of pancreatic ␤-cells

Arrival rate (ves/h per ␮m2)

Membrane Fluorescent surface vesicle density area (␮m2) (ves/␮m2)

PULSATILE INSULIN SECRETION IN SINGLE ␤-CELLS

atic ␤-cells, just 50 insulin vesicles are thought to be stored in the readily releasable pool (20,21), representing ⬍1% of the ⬃10,000 stored vesicles (22). Once a pancreatic ␤-cell has depleted the vesicles in its readily releasable pool, the rate of secretion drops significantly, even when the secretion stimulus is maintained. On the other hand, if the stimulus is removed and the cell is allowed to rest, the cell will refill its readily releasable pool and will again be capable of a rapid secretory burst. Because a typical pancreatic ␤-cell already contains thousands of insulin vesicles, it is not compelled to synthesize new insulin vesicles to replenish its readily releasable pool. However, it remains unknown which of the thousands of existing vesicles in pancreatic ␤-cells are used to refill the readily releasable pool. Optical methods, such as TIRF, confocal, and multiphoton microscopies (13,23–32), are well suited for studies of vesicle trafficking to the readily releasable pool. According to one previous study in the MIN-6 cell line (27), the readily releasable pool is refilled by vesicles that move from stores deep inside the cell to the plasma membrane. In that study, fluorescent vesicles started to arrive at the plasma membrane soon after a stimulus was applied, and the rate of vesicle arrival at the plasma membrane appeared to be greater for glucose stimulation than for potassium stimulation. As a result, secretion stimulation led the number of membrane-proximal vesicles to increase over time. Similar experiments with the INS-1 cell line, however, did not support these findings, instead suggesting that pancreatic ␤-cells refill their readily releasable pool using insulin vesicles that are already near the plasma membrane (24), so that the number of membrane-proximal fluorescent vesicles did not increase. Although both of these studies of insulin vesicle dynamics in rodent pancreatic ␤-cell lines have played an important role in advancing our understanding of insulin secretion, it is unclear which one more accurately reflects the dynamics of insulin vesicles in human pancreatic ␤-cells. In the current study, we analyze both the temporal patterns of insulin vesicle fusion and the role of insulin vesicles of different ages in refilling the readily releasable pool. RESEARCH DESIGN AND METHODS In the past 4 years, we have received 30 preparations of human islets, all of which were isolated from heart-beating donors who had no evidence of prior type 2 diabetes (see ACKNOWLEDGMENTS). On arrival, these islets were transferred to warm medium (RPMI with 5 mmol/l glucose and 10% fetal bovine serum [FBS] and 5% CO2, 37°C); within 2 h, the islets were dispersed and maintained according to our standard protocols (15). Rat pancreatic ␤-cells were prepared and cultured as previously described (14), using procedures approved by the University of Southern California animal use committee. Cultured cells were used within 5 days. (As described below, we report results of studies using islets harvested from only four human donors [one female and three male subjects, aged 32 ⫾ 6.1 years, BMI 28 ⫾ 3.1 kg/m2] whose pancreatic ␤-cells displayed pulsatile insulin secretion in response to glucose stimulation.) Selection of cells. For inclusion in this study, islets and pancreatic ␤-cells had to satisfy all of the following morphologic and functional criteria. Morphologic criteria. Islets needed to have smooth edges, and the distribution of islet sizes had to span both small and large islets, including at least 10% of islets having diameters ⬎300 ␮m (indicating that the tissue was not over- or underdigested). Moreover, pancreatic ␤-cells derived from islets needed to have smooth membranes, relatively small nuclei, and few large cytoplasmic organelles (⬍1 ␮m). Functional criteria. Pancreatic ␤-cells had to stick to glass cover slips and express fluorescent cargo proteins when infected with adenovirus. Moreover, ⬎80% of cells had to respond to potassium, and more than two cells from the same preparation had to respond vigorously (secrete ⬎30 vesicles) to glucose stimulation. As described below, only four preparations satisfied all of these criteria. DIABETES, VOL. 56, MAY 2007

D.J. MICHAEL AND ASSOCIATES

FIG. 1. Potassium consistently stimulated a rapid, short-duration burst of insulin vesicle fusion (rat). A and B: TIRF micrographs of a single rat pancreatic ␤-cell expressing C-emGFP (12), before (A) and after (B) stimulation with potassium (50 mmol/l). C: Montage showing rapid and complete release of C-emGFP during exocytosis (0.3 s per frame). Full kinetic analysis of this and other fluorescent cargo proteins was published previously (14). D: The number of vesicles fusing during each frame, average values with SE (n ⴝ 5 cells). E: Running sum of the number of secreted vesicles. Vectors and adenovirus. Insulin vesicles were labeled with one of four different fluorescent cargo proteins. Three were made by fusing insulin C-peptide to one of the following fluorescent proteins: emerald green fluorescent protein (C-emGFP), enhanced green fluorescent protein (C-EGFP), or the bright mutant of dsRed fluorescent protein (C-Bright). The fourth was made by fusing the rodent isoform of islet amyloid polypeptide to enhanced green fluorescent protein (rIAPP-EGFP). Construction of adenovirus containing the gene for either C-emGFP or rIAPP-EGFP has been described previously (12,14). To make adenovirus containing C-EGFP and C-Bright, we used standard molecular biological techniques to modify the C-emGFP construct by replacing emerald green fluorescent protein separately with either enhanced green fluorescent protein or the bright mutant of dsRed fluorescent protein. DNA sequences were confirmed using standard sequencing protocols (University of Pittsburgh/University of Southern California Microchemical Core Facility). Once plated, cells were incubated overnight and then transduced with adenovirus (5 ⫻ 107 plaque-forming units/ml in RPMI 1640 with 10% FBS and 5 mmol/l glucose) containing the gene of interest. After inoculation of the cells with adenovirus (4 h), the medium was removed and replaced with fresh medium (RPMI 1640 with 10% FBS and either 5 or 8 mmol/l glucose for human or rat ␤-cells, respectively). Cells were imaged 16 – 48 h later. Solutions and chemicals. Extracellular solution used in imaging experiments was composed of the following (in mmol/l): 136 NaCl, 4.2 KCl, 2.4 CaCl2, 1.2 MgSO4, 1.2 KH2PO4, 1 L-glutamine, 4 glucose, and 10 HEPES (pH 7.4). In the high-potassium stimulating solution, an equal quantity of KCl replaced NaCl to yield a final potassium concentration of 50 mmol/l. In the high-glucose stimulating solution, we added enough glucose to the basal salt solution so that the final concentration of glucose was 8 –20 mmol/l (see figure legends for specific concentrations in each experiment). All solutions were prepared with water from a commercial purification system (Nanopure Infinity; Barnstead Thermolyne, Dubuque, IA). The osmolalities of extracellular and stimulating solutions were matched and fell between 280 and 320 mOsm. All salts for solutions were obtained from Sigma Aldrich (St. Louis, MO) with the highest purity available. TIRF microscopy. For descriptions of our imaging setups for one- and two-color TIRF microscopy, see our previous publications (14,15). ThroughDIABETES, VOL. 56, MAY 2007

out the imaging experiments, cells were maintained in a warmed solution (between 35 and 37°C) with continuous exchange (⬃1 ml/min). Pulled glass pipettes (⬃3 ␮m tip diameter) were positioned near cells for transient application of test solutions. Spectral analysis and randomization trials. Periodograms were constructed using standard methods (33). Briefly, to construct a periodogram, each point in an evenly spaced time series of n points was modeled as the sum of sine and cosine terms with amplitudes and frequencies determined according to the following equations:

冘

m⫺1

xi ⫽ A(0) ⫹

兵A共k兲cos共␻ki兲 ⫹ B共k兲sin共␻ki兲其⫹A共m兲cos共␻mi兲, where

k⫽1

m ⫽ n/ 2, ␻k ⫽ ␲*k/m and A(0) ⫽ x៮

冘 n

A共k兲 ⫽ 共2/n兲*

xi cos共␻ki兲

i⫽1

冘 n

B共k兲 ⫽ 共2/n兲*

xi sin共␻ki兲

i⫽1

冘 n

A共m兲 ⫽ 共1/n兲*

xi共⫺1兲i

i⫽1

The magnitude, S(k), for each period in the periodogram is defined as S 2共k兲 ⫽ A2共k兲 ⫹ B 2共k兲 It can be shown that the variance of the original data points partitions among the various periods according to the following equation: 1279


FIG. 2. Insulin vesicles were not secreted at random times in pancreatic ␤-cells stimulated by glucose (rat, 20 mmol/l glucose). A: The number of vesicles fusing during each frame (0.222 s/frame). Each point is from a single frame. B: Running sum of the number of secreted vesicles. C: Two functions, Dⴙ (dashed line) and Dⴚ (solid line), for the observed data (see RESEARCH DESIGN AND METHODS). D: Two functions, Dⴙ (dashed line) and Dⴚ (solid line), after randomly permuting the observed data, i.e., representative result for one randomization trial.

再冘

冎

m⫺1

n

S 2共k兲/ 2 ⫹ A2共m兲 ⫽

k⫽1

冘 n

共x ⫺ x៮ 兲2

i⫽1

The term p(k) is defined by normalizing each of the magnitudes in the periodograms by the sum of the squares about the mean, expressed as

冒冘冒冘 n

p共k兲 ⫽ S 2共k兲

共xi ⫺ x៮ 兲2 for k ⬍ m and

i⫽1

n

p共k兲 ⫽ A2共m兲

共xi ⫺ x៮ 兲2 for

i⫽1

冘 m

k ⫽ m, so that

p共k兲 ⫽ 1.

k⫽1

The p(k) values are then used as the basis for statistical testing of significance in randomization trials. However, before determining whether the magnitude of any of the observed periods was statistically different from the expected distribution of p(k) values, we first determined the likelihood that the overall pattern was significantly different from that expected for a random series. For this step, we used the Kolmogorov-Smirnov test (33) as follows:

冘 j

␮j ⫽

k⫽1

冒冘 m⫺1

S 2共k兲

S 2共k兲

k⫽1

D⫹ ⫽ max 兵j/共m ⫺ 1兲 ⫺ ␮j其 D⫺ ⫽ max 兵␮j ⫺ 共 j ⫺ 1兲/共m ⫺ 1兲其 D ⫽ max 兵D⫹, D⫺其 The significance levels of D for the observed data were determined by randomization trials. In each trial, the observed data were randomly permuted, and the value of D was calculated. At the end of the N ⫺ 1 trials, the 1280

value of D for the observed data was grouped with all of the values of D from the randomization trials. The significance level of D was then calculated as 100 ⫻ rank/N, where rank refers to the rank of D for the observed data within the set of N values for D. If the significance level of D for the observed data was less than 5%, then we tested individual periods for significance using a similar randomization method. However, when testing individual periods, the statistically significant level was adjusted by the Bonferonni correction (␣ ⫽ 5%/number of periods tested) (33). In some cases, the data were first compressed using a simple nonoverlapping, box-car summation. In these cases, secretion data from X continuous frames were summed, where X represents the compression factor (see RESULTS for typical values of X). All calculations were performed using scripts written for R (34), a freely distributed variant of the statistical programming language S⫹. To verify our scripts, we reproduced the data presented in Table 11.5 of Manly (33). To calculate the overall pulse interval reported in Table 1, we included data from all cells that showed pulsatile secretion (i.e., all cells that passed the first part of our two-part test), even those in which we could not identify a significant period of oscillation. Bursts were identified by sharp changes in the cumulative secretion plots. The distribution of pulse intervals was skewed to the right, so we report the median value for this parameter (see Table 1). All other values are reported as the means ⫾ SE, unless otherwise stated. Parameters in Table 1 were compared using Student’s t test with the Welch modification to the degrees of freedom. Calcium measurements. At ⬃24 – 48 h after cells were infected with C-Bright adenovirus, they were incubated for 30 min at room temperature in external solution containing 1 ␮mol/l of the AM (acetoxymethyl) ester form of the calcium indicator Oregon Green 488 BAPTA-1 (Invitrogen, Carlsbad, CA). Thereafter, the cells were washed once with dye-free external solution and then kept in dye-free solution for 30 – 60 min at 37°C. To measure insulin vesicle secretion and calcium changes simultaneously, images were collected from the red and green channels of our two-color microscope (15). Green (543 nm) light from an He-Ne laser was configured for TIRF excitation of insulin vesicles containing C-Bright, whereas blue light (488 nm) from an Ar⫹ laser was configured for wide-field excitation of the cytoplasmic calcium indicator. Fluorescence images from blue and green excitation were captured separately. To minimize photobleaching of the calcium indicator, we used an asymmetric acquisition scheme: nine images with green excitation light were collected for every one image with blue excitation light. The overall acquisiDIABETES, VOL. 56, MAY 2007


FIG. 3. Glucose stimulated short-duration bursts of insulin vesicle secretion (rat and human). A and C: Rat cells (20 mmol/l glucose). B and D: Human cells (12 mmol/l glucose). tion rate was ⬃2.5 Hz. Cross talk between fluorophores was minimal, and no effort was made to correct signals. Capacitance measurements. For capacitance measurements, pancreatic ␤-cells were bath perfused at 1 ml/min flow rate in external solution containing (in mmol/l): 138 NaCl, 5.6 KCl, 1.2 MgCl2, 2.6 CaCl2, 4 glucose, 10 HEPES, and 10 tetraethylammonium-Cl. Patch pipettes had initial resistances of 3– 4 M⍀ when filled with internal solution containing the following (in mmol/l): 10 NaCl, 145 Cs-glutamate, 10 HEPES, 1 MgCl2, 2 Mg-ATP, and 0.3 Na-GTP. Whole-cell recordings were made using the EPC-9 patch-clamp amplifier and Pulse software (Heka, Lambrecht/Pfalz, Germany). For capacitance measurements, we used the “Sine⫹dc” mode of the “Lock-in” extension in Pulse software. In brief, a 1-kHz frequency sine-wave voltage, with a 40-mV peak-to-peak amplitude, was superimposed on the ⫺70-mV holding potential. Stimulation trains consisted of eight voltage pulses, with each pulse depolarizing the cell membrane from ⫺70 to 0 mV for 500 ms; pulses in the train were separated by 100 ms. Trains were repeated approximately every minute. Capacitance recordings and TIRF imaging were synchronized through transistor-transistor logic signals. Calculating diffusion coefficients. To calculate the diffusion coefficients of single insulin vesicles, we first tracked the positions of single insulin vesicles using commercially available tracking software (Metamorph; Molecular Devices, Sunnyvale, CA). For a vesicle tracked for n frames, we then calculated the vesicle’s net displacement over i frames, where i ⫽ 1 to n ⫺ 1, using frame j as the starting frame, where j ⫽ 1 to n ⫺ i. For each i, we calculated the average of the square of the vesicle’s net displacement. The mean square displacement was then plotted as a function of time, and the data were fitted using a weighted linear regression; points were weighted proportionally with respect to the number of points that contributed to the average. The slope of the fit is directly proportional to the two-dimensional diffusion coefficient (D2 ⫽ slope/4, where D2 is the two-dimensional diffusion coefficient). Calculations and curve-fittings were performed using scripts written in R.

RESULTS

We prepared primary cultures of pancreatic ␤-cells from rat and human islets and transduced them with a low titer of a viral vector encoding a fluorescently tagged vesicle cargo protein (see below for specific constructs). Within 48 h, we imaged individual fluorescent vesicles with a TIRF microscope, a microscope that selectively illuminates membrane-proximal vesicles at the base of the cell (13). Each of the cells we selected for study resided within DIABETES, VOL. 56, MAY 2007

a small cluster of cells. Typically, a single transduced cell contained hundreds of fluorescent vesicles near the plasma membrane, which were scattered about at a modest density (⬃0.7 fluorescent vesicles per ␮m2) (see Table 1). In resting cells, most of the membrane-proximal fluorescent vesicles moved only slightly, jostling about a central point (two-dimensional diffusion coefficient of 4.9 ⫾ 0.5 ⫻ 10⫺12 cm2/s, n ⫽ 78 vesicles, eight human pancreatic ␤-cells) (35,36). We then characterized secretion by stimulating cells with solutions containing elevated concentrations of potassium or glucose. Potassium consistently stimulated a rapid and short burst of insulin vesicle fusion. Figure 1A shows a rat pancreatic ␤-cell expressing C-emGFP, a probe that consistently leaves insulin vesicles rapidly and completely during a single round of exocytosis (12,14,15). When this cell was transiently stimulated with potassium, it responded quickly (⬍5 s) (Table 1, “lag”) by secreting ⬃50 vesicles, all fusing with the plasma membrane within a span of 10 s (Fig. 1B–E). We repeated this experiment for a number of rat and human pancreatic ␤-cells (Table 1) and found that potassium consistently stimulated a single short-duration, large-amplitude burst of insulin vesicle fusion. Rat and human pancreatic ␤-cells secreted insulin vesicles at similar rates (see Table 1, which summarizes characteristics of the burst). Note that with a TIRF microscope, we are imaging only the footprint of a cell adhering to a glass coverslip—a portion of the plasma membrane encompassing ⬍50% of the cell’s total membrane surface area (for a precise estimate, see online Appendix 1 [available at http://dx.doi. org/10.2337/db06-0367]). Previous work has shown that vesicles fuse with the plasma membrane not only where cells contact the glass surface, but also where cells contact each other (15). If we assume that the rate of secretion is uniform over the entire cell, then extrapola1281


FIG. 4. Bursts of insulin vesicle secretion coincide with transient increases in intracellular calcium concentration (human, 12 mmol/l glucose). A: The number of vesicles labeled with C-Bright that fused during each frame (0.4 s/frame). B: Running sum of the number of secreted vesicles. C: Integrated green fluorescence (Oregon Green 488 BAPTA-1) in a region of interest encompassing the entire cell. Fluor, fluorescence.

tion predicts that the number of vesicles secreted by a potassium-stimulated human pancreatic ␤-cell is ⬃200, which represents the size of the readily releasable pool. In rats the estimate of the readily releasable pool size is ⬃90. Glucose-stimulated small-amplitude bursts of insulin vesicle fusion. Next, we investigated insulin vesicle exocytosis stimulated by glucose. Fewer rat and human pancreatic ␤-cells responded to glucose than to potassium (for human pancreatic ␤-cells, 45 of 49 cells responded to potassium, and 12 of 42 cells responded to glucose), and several features of the glucose-stimulated response differed from the potassium-stimulated response. First, when cells responded to glucose, there was a significant delay (Table 1, “lag”), in contrast to the immediate response to potassium stimulation. The delay is expected because glucose stimulates secretion through a pathway that is more complicated than the one activated by potassium. Once stimulated, cells secreted more slowly in response to glucose than to potassium (Table 1). Finally, whereas cells responded to potassium with a single, short-duration burst of secretion, they responded to glucose with widely varying temporal patterns of vesicle fusion. In general, the temporal patterns of secretion were not random and included some cases in which vesicles fused in well-separated bursts, consisting of 10 –15 vesicles (in 1282

the cell footprint) that all fused within a span of 15 s (Figs. 2 and 3 and Table 1). Extrapolated to the entire cell (as was performed above for potassium stimulation), we estimate that each glucose-stimulated burst consists of ⬃70 vesicles in human ␤-cells and 30 vesicles in rat ␤-cells. As expected, the bursts of insulin vesicle fusion coincided with transient increases in intracellular calcium (Fig. 4). A two-step statistical analysis revealed that these bursts of secretion were separated by significant periods ranging from 15 to 45 s (Figs. 2 and 5). See RESEARCH DESIGN AND METHODS for details of the analysis, and see the legend for Fig. 5 for exact values of significant periods. Our analysis also revealed statistically significant periods in cells in which the pulses of vesicle fusion were not obvious on initial inspection of the raw data (Fig. 6). Changes in the density of fluorescent membraneproximal insulin vesicles. Before, during, and after stimulation with either potassium or glucose, new fluorescent insulin vesicles arrived at the plasma membrane at ⬃1 vesicle/␮m2 per h—much more slowly than the rate at which fluorescent vesicles were secreted (Table 1). Thus, during vigorous stimulation, there was a net decrease in the number of fluorescent vesicles near the plasma membrane (Fig. 7). A role for older insulin vesicles in refilling the readily releasable pool. We next investigated the kinetics of refilling the readily releasable pool of vesicles. For these experiments, we monitored secretion from individual cells by simultaneous application of two different techniques: TIRF imaging and patch-clamp measurements of membrane capacitance. TIRF imaging selectively reports exocytosis of vesicles containing fluorescent cargo protein (age ⬍48 h, the time since viral transduction). In contrast, membrane capacitance measurements monitor changes in the membrane surface area, which increases on addition of membrane from any vesicle, whether it contains fluorescent cargo (age ⬍48 h) or not (age ⬎48 h). To stimulate secretion, we applied a train of membranedepolarizing voltage steps to cells whose plasma membrane was voltage-clamped in the whole-cell configuration. Depolarization is known to activate voltage-gated calcium channels and to trigger increases in membrane capacitance (37), which should be accompanied by exocytosis of fluorescent vesicles. Indeed, as seen in Fig. 8, individual depolarizations elicit a capacitance step, and during a train of depolarizations, the size of the capacitance steps progressively diminishes until there is no change for the last few stimuli. This indicates that the readily releasable pool of vesicles is exhausted at the end of the train. To calculate the contribution of fluorescent vesicles to overall secretion during a train of depolarizations, we counted the number of vesicles that disappeared and assumed that each vesicle contributed 2 fF of membrane capacitance (estimates range from 1 to 3.8 fF per vesicle) (38 – 40). We also assumed that the rate of secretion of fluorescent vesicles in the footprint is representative of the rate across the entire surface of the cell. With these assumptions, we estimate that fluorescent vesicles contributed at least half of the vesicles secreted in response to the first train of depolarizing voltage steps. (If we were to use the estimate of 3.5 fF per vesicle, a large estimate of unitary capacitance [41], then all secretion for the first train would have been of new fluorescent vesicles.) After the first train of depolarizations, we allowed the cells to rest for ⬃1 min and then applied a second train. As shown in Fig. 8, cells had largely restored their ability to DIABETES, VOL. 56, MAY 2007


FIG. 5. Spectral analysis and randomization tests identified statistically significant periods between bursts of insulin vesicle secretion (human, 12 mmol/l glucose). All time series have been compressed so that each point represents ⬃1 s of data. Results are from a representative cell. A: Time series showing the number of vesicles fusing per second. B: Time series showing the running sum for the number of secreted vesicles. C: Periodograms for the data shown in A. D: Significance levels of the observed p(k) values relative to 50,000 simulations. The dotted line shows the relevant significance level after Bonferroni’s correction (␣ ⴝ 5%/272). *Statistically significant periods (19.3 and 41.8 s).

secrete, as judged by the reappearance of capacitance changes stimulated by a second train of depolarizing voltage steps. However, the fraction of secretion attributed to fluorescent vesicles decreased significantly for the second train compared with the first train (Fig. 8 and supplemental Appendix 2). In one exceptionally stable cell, we were able to apply four separate trains of stimuli (consecutive trains separated by 1 min), and we observed that the fraction of secretion attributed to fluorescent vesicles decreased progressively for succeeding trains (Fig. 8B, lower panel). These results demonstrate directly that human pancreatic ␤-cells contain both young (fluorescent) and old (nonfluorescent) vesicles in their readily releasable pools. Remarkably, the ␤-cells first secrete their young (fluorescent) vesicles, and only later use their older (nonfluorescent) vesicles to refill their pool of secretion-competent vesicles. DISCUSSION

In the current study, we have characterized the dynamics of the functional vesicle pools in individual human pancreatic ␤-cells. Our work was made possible because we had access to a number of human islet preparations, including islets that were of transplant quality—prepared for use in treating patients with type 1 diabetes (42). In addition, we developed a strategy to study single cells located in small cell clusters because previous work suggests that isolated ␤-cells function differently from ␤-cells found within islets (11). We transduced cells using a low titer of virus, and we chose for study only those clusters in which one cell, at most, was infected. In addition, we calculated rates of vesicle arrival and secretion, normalized to a square micron of surface area, so that we could obtain whole-cell estimates for vesicle behavior by multiplyDIABETES, VOL. 56, MAY 2007

ing the normalized rates by the total surface area of cells, derived from membrane capacitance measurements. Although our study includes only a small number of cells, the cells we selected for analysis displayed a number of features suggesting that they were of exceptionally good health. Most importantly, they displayed oscillations of insulin vesicle fusion that coincided with oscillations in intracellular calcium levels. Additional features that corroborated the good health of the cells included their ability to express fluorescent cargo protein when transduced with adenovirus, their tendency to flatten when maintained in culture, and their ability to refill the readily releasable pool of vesicles rapidly when stimulated to secrete (see below). Screening for such cellular characteristics may prove useful as a means of verifying the health of islets for transplantation. Size of the readily releasable pool of vesicles in human pancreatic ␤-cells. Using potassium stimulation, we obtained estimates of the size of the readily releasable pool of vesicles in pancreatic ␤-cells. In human ␤-cells, potassium stimulation rapidly triggered a single, shortduration (⬃10 s) burst of exocytosis consisting of ⬃55 vesicles per cell footprint, which extrapolates to ⬃200 vesicles for the entire cell. This estimate of the readily releasable pool in human ␤-cells is similar to previous estimates of the readily releasable pool in rodent ␤-cells (22). Indeed, in parallel experiments, we measured the readily releasable pool of vesicles in rat ␤-cells to be ⬃90 vesicles for the entire cell. These estimates may be slight underestimates because they are based on imaging fluorescent insulin vesicles, which are a subset of all insulin vesicles. However, as we note below, most insulin vesicles undergoing exocytosis for a first stimulus are young fluo1283


FIG. 6. Glucose stimulated several surges of insulin vesicle fusion: compressed data (human, 8 mmol/l glucose). The data shown in A are compressed fourfold. A: The number of vesicles fusing during each second. B: Running sum of the number of secreted vesicles. Inset shows the initial response, which includes three clear surges in secretion. C: Periodogram for the data shown in A. D: Significance level of the observed p(k) values relative to 105,000 simulations. The dotted line shows the relevant significance level after Bonferroni’s correction (␣ ⴝ 5%/287). *Statistically significant periods (64.5, 72.6, and 580.9 s). Note: It is unclear whether the significance of the longest period is meaningful because there is only one full cycle for this period in our data set.

rescent insulin vesicles, so our estimates are likely to be fairly accurate. Pulsatile secretion of insulin vesicles. Despite the preponderance of evidence suggesting that individual pancreatic ␤-cells should secrete insulin in discrete pulses or bursts, there has been little direct evidence of such pulsatile insulin secretion at the level of the single cell and single vesicle. In fact, most previous studies of insulin vesicle fusion at the single-cell level reported that pancreatic ␤-cells respond to glucose by secreting insulin vesicles at a relatively steady rate (27,30,32). Here, we have shown directly that individual pancreatic ␤-cells can and do secrete insulin vesicles in short-duration, small-amplitude bursts that recur regularly and that consist of ⬃70 vesicles per burst for human ␤-cells and ⬃30 vesicles per burst for rat ␤-cells (extrapolated in both cases to the entire cell). As noted above, these estimates may be slight underestimates. Furthermore, individual glucose-stimulated bursts do not fully deplete the readily releasable pool. Instead, they release only about a third of the vesicles in the readily releasable pool. Our observations reveal the single-cell contribution to the oscillations of insulin secretion observed at higher levels of tissue organization. Intracellular calcium concentration is a key regulator of insulin vesicle secretion (17,43), and in previous publications, oscillations in intracellular calcium concentration have been recorded in both intact islets (44) and in single pancreatic ␤-cells (45). In isolated ␤-cells from mice, three classes of intracellular calcium oscillation have been described, with periods ranging from tens of seconds to a few minutes (46). As we have shown (Fig. 4), bursts of 1284

insulin vesicle fusion coincide with transient increases in intracellular calcium concentration, and the periods separating the bursts fit within the previously reported range for periods of oscillations in intracellular calcium concentration (46). Not all studies of clonal ␤-cells (36) and isolated pancreatic islets (30) have shown that intracellular calcium levels oscillate in response to glucose stimulation. Rather, some studies have reported that intracellular calcium concentration increased suddenly and remained elevated. Although it is not clear why only certain cells responded to glucose with easily identified oscillations, the loss of pulsatile insulin secretion observed during the development of both type 1 (9) and type 2 (10) diabetes suggests that the cells and tissue responding with oscillations are the more healthy cells. Refilling the readily releasable pool in human pancreatic ␤-cells. We characterized refilling of the readily releasable pool in primary cultures of human and rat pancreatic ␤-cells. Contrary to what was reported previously in pancreatic beta tumor cell lines (27), we did not observe a dramatic increase in the rate of fluorescent vesicle arrival at the plasma membrane during a human ␤-cell’s response to either potassium or glucose. In fact, we even observed a net decrease in the overall number of fluorescent vesicles at the plasma membrane because the rates of secretion exceeded by at least an order of magnitude the rates of arrival of new fluorescent insulin vesicles (Table 1 and Fig. 7). Although secretion stimulation did not dramatically enhance the arrival of fluorescent vesicles at the plasma membrane, we did observe the appearance of fluorescent DIABETES, VOL. 56, MAY 2007


FIG. 7. Density of membrane-proximal fluorescent insulin vesicles decreases during potassium and glucose stimulation (human). A: The percentage of membrane-proximal fluorescent insulin vesicles remaining in glucose-stimulated cells. Solid lines are from the same five human cells reported in Table 1. Broken line shows average values with SE. B: Montage showing images of a single cell during glucose-stimulated response. Times indicated are relative to the moment that glucose was first applied. C: The percentage of membrane-proximal fluorescent insulin vesicles remaining in potassium-stimulated cells. Solid lines are from eight representative cells. The heavy band shows average values with SE. D: Montage showing images of a single cell during potassium-stimulated response. Note: For A and C, 100% ⴝ initial density.

vesicles at the plasma membrane, with an arrival rate of ⬃1 vesicle/␮m2 per h. This apparently slow rate of arrival is actually more than sufficient to account for the daily rate of insulin secretion at a typical human pancreatic ␤-cell— over a 24-h period, 24,000 vesicles would be expected to reach the plasma membrane of a single human ␤-cell (1 vesicle/␮m2 per h ⫻ 1,000 ␮m2 per cell ⫻ 24 h). (Note, however, that this rate may be an overestimate of the true rate of vesicle delivery at the membrane because some vesicles appear not to dock firmly and may actually undergo repeated appearances at the membrane.) In comparison, the average daily insulin secretion in healthy humans has been reported (47) to be ⬃75 pmol/min, corresponding to secretion of ⬃120 insulin vesicles per ␤-cell per day (assuming 106 islets per human pancreas (42), 103 ␤-cells per human islet (37) and 5 ⫻ 105 insulin molecules per insulin vesicle (48). The second observation that explains why fluorescent vesicles are not needed for rapid resupply of the readily releasable pool is that nonfluorescent vesicles are capable of rapid priming to join the readily releasable pool. We were able to study this process by combining TIRF microscopy and membrane capacitance measurements (Fig. 8). For a first train of depolarizations, at least half of the vesicles contributing to secretion were fluorescent—surprising, given that the fluorescent vesicles were vastly outnumbered by nonfluorescent vesicles. This observation DIABETES, VOL. 56, MAY 2007

corroborates the finding in another type of endocrine cell that new vesicles are preferentially secreted (49). After allowing a minute for replenishment of the readily releasable pool, a second train used to probe the makeup of the readily releasable pool showed that a significant fraction of the resupplied vesicles were nonfluorescent, or older, vesicles. This result implies that, over short time spans (on the order of several minutes), pancreatic ␤-cells preferentially secrete their young (labeled) vesicles (⬃50% of the vesicles secreted in response to the initial train of depolarizations) (see supplemental Appendix 2) and then recruit older (unlabeled) vesicles to refill the readily releasable pool of insulin vesicles. Over longer time spans, especially between meals when secretion stimulation is much diminished, we expect that new (fluorescent) vesicles will repopulate the membrane-proximal pool, arriving there shortly after they bud from the Golgi body (as has been suggested for other types of endocrine cells) (50). The rapid and directed movement of new vesicles from the Golgi body to the plasma membrane is distinct from the relatively slow and random movement of vesicles once they are near the plasma membrane (50). Of note, the average diffusion coefficient of insulin vesicles in clonal ␤-cells (36) is approximately two orders of magnitude greater than the average diffusion coefficient we report here (4.9 ⫾ 0.5 ⫻ 10⫺12 cm2/s) in primary cultures of 1285


FIG. 8. Pancreatic ␤-cells first secrete young insulin vesicles and then refill the readily releasable pool using older vesicles (human). A: Stimulus protocol (upper panel), representative calcium currents (middle panel), and membrane capacitance changes (lower panel). B (top panel): Fluorescent vesicles secreted per capacitance change (in fF) for two consecutive stimulations. ***P < 0.05, Student’s t test for paired means, n ⴝ 6 cells. B (bottom panel): Fluorescent vesicles secreted per capacitance change (in fF) for four consecutive stimulations (n ⴝ 1 cell). C–J: TIRF images from a representative pancreatic ␤-cell. C: Before first stimulation. D: After first stimulation. E: False color overlap of C (green) and D (red). Secreted vesicles appear green. Newly arriving vesicles appear red. Unchanged vesicles appear yellow. F–H: Same cell, same pattern of images as C–E, but now for second stimulation. I: False-color overlay of D (green) and F (red). J: Same image as I, with three lines added, indicating where the line traces were taken to make the traces in K. K: Three line traces showing spatial overlap of green and red fluorescence in J. Note: The absence of red vesicles in all panels reflects the slow rate of arrival of new fluorescent vesicles.

human pancreatic ␤-cells. On the other hand, the value we report agrees with previous studies of mobility for large dense-core vesicles in other types of endocrine cells (35,51,52). The greater mobility of insulin vesicles in clonal cells might account for the higher rate of vesicle arrival on glucose or potassium stimulation in comparison with what we observed in primary cultures of human pancreatic ␤-cells. An ultrastructural study of fixed pancreatic ␤-cells suggested that the density of membrane-proximal insulin vesicles increases by ⬃50% after 40 min of glucose stimulation (21). It is important to note that electron microscopes visualize all insulin vesicles and cannot distinguish 1286

young from old vesicles. The insulin vesicles studied by TIRF microscopy and transient expression of a fluorescent cargo protein are young (⬍48 h old) and represent a subset of the insulin vesicles studied by the electron microscopy approach. Thus, the results from electron microscopy, TIRF microscopy, and membrane capacitance measurements taken together imply that large numbers of older insulin vesicles move closer to the plasma membrane in a glucose-stimulated pancreatic ␤-cell after the cell first secretes its youngest vesicles. Summary. In summary, we have demonstrated for the first time that glucose stimulates some pancreatic ␤-cells to secrete insulin vesicles in discrete bursts that recur DIABETES, VOL. 56, MAY 2007


regularly. Furthermore, we report that ␤-cells preferentially secrete first their youngest insulin vesicles and then recruit older insulin vesicles for rapid release. ACKNOWLEDGMENTS

Funding for this work was provided by generous gifts from the family of Betty Smith Rose and by grants from the U.S. National Institutes of Health provided to D.J.M. (DK10181) and R.H.C. (DK60623) and from the American Heart Association to R.H.C. We thank Robert Ritzel for helpful comments on this manuscript. Human islets for basic research were made available by the Islet Cell Resource Centers (Washington University, University of Pennsylvania, University of Minnesota, and City of Hope Medical Center [Duarte, CA]) as well as Emory University (Atlanta, GA). We thank all members of the Islet Cell Resource Centers for their time and efforts. REFERENCES 1. Stagner JI, Samols E, Weir GC: Sustained oscillations of insulin, glucagon, and somatostatin from the isolated canine pancreas during exposure to a constant glucose concentration. J Clin Invest 65:939 – 942, 1980 2. Polonsky KS, Given BD, Van Cauter E: Twenty-four-hour profiles and pulsatile patterns of insulin secretion in normal and obese subjects. J Clin Invest 81:442– 448, 1988 3. Porksen N: The in vivo regulation of pulsatile insulin secretion. Diabetologia 45:3–20, 2002 4. Lang DA, Matthews DR, Peto J, Turner RC: Cyclic oscillations of basal plasma glucose and insulin concentrations in human beings. N Engl J Med 301:1023–1027, 1979 5. Goodner CJ, Walike BC, Koerker DJ, Ensinck JW, Brown AC, Chideckel EW, Palmer J, Kalnasy L: Insulin, glucagon, and glucose exhibit synchronous, sustained oscillations in fasting monkeys. Science 195: 177–179, 1977 6. Porksen N, Hollingdal M, Juhl C, Butler P, Veldhuis JD, Schmitz O: Pulsatile insulin secretion: detection, regulation, and role in diabetes. Diabetes 51:S245–S254, 2002 7. Ritzel RA, Hansen JB, Veldhuis JD, Butler PC: Induction of beta-cell rest by a kir6.2/sur1-selective k-atp-channel opener preserves beta-cell insulin stores and insulin secretion in human islets cultured at high (11 mm) glucose. J Clin Endocrinol Metab 89:795– 805, 2004 8. Ritzel RA, Veldhuis JD, Butler PC: Glucose stimulates pulsatile insulin secretion from human pancreatic islets by increasing secretory burst mass: dose-response relationships. J Clin Endocrinol Metab 88:742– 747, 2003 9. O’meara NM, Sturis J, Herold KC, Ostrega DM, Polonsky KS: Alterations in the patterns of insulin secretion before and after diagnosis of IDDM. Diabetes Care 18:568 –571, 1995 10. Lang DA, Matthews DR, Burnett M, Turner RC: Brief, irregular oscillations of basal plasma insulin and glucose concentrations in diabetic man. Diabetes 30:435– 439, 1981 11. Bertram R, Previte J, Sherman A, Kinard TA, Satin LS: The phantom burster model for pancreatic beta-cells. Biophys J 79:2880 –2892, 2000 12. Watkins S, Geng X, Li L, Papworth G, Robbins PD, Drain P: Imaging secretory vesicles by fluorescent protein insertion in propeptide rather than mature secreted peptide. Traffic 3:461– 471, 2002 13. Axelrod D: Selective imaging of surface fluorescence with very high aperture microscope objectives. J Biomed Opt 6:6 –13, 2001 14. Michael DJ, Geng X, Cawley NX, Loh YP, Rhodes CJ, Drain P, Chow RH: Fluorescent cargo proteins in pancreatic beta cells: design determines secretion kinetics at exocytosis. Biophys J 87:L3–L5, 2004 15. Michael DJ, Haataja L, Ritzel RA, Chow RH: Pancreatic beta cells secrete insulin in fast- and slow-release forms. Diabetes 55:600 – 607, 2006 16. Heinemann C, Vonruden L, Chow RH, Neher E: A two-step model of secretion control in neuroendocrine cells. Pflueg Arch Eur J Physiol 424:105–112, 1993 17. Jahn R, Lang T, Sudhof TC: Membrane fusion. Cell 112:519 –533, 2003 18. Voets T: Dissection of three Ca2⫹-dependent steps leading to secretion in chromaffin cells from mouse adrenal slices. Neuron 28:537–545, 2000 19. Chow RH, Klingauf J, Heinemann C, Zucker RS, Neher E: Mechanisms DIABETES, VOL. 56, MAY 2007

determining the time course of secretion in neuroendocrine cells. Neuron 16:369 –376, 1996 20. Olofsson CS, Gopel SO, Barg S, Galvanovskis J, Ma XS, Salehi A, Rorsman P, Eliasson L: Fast insulin secretion reflects exocytosis of docked granules in mouse pancreatic b-cells. Pflueg Arch Eur J Physiol 444:43–51, 2002 21. Straub SG, Shanmugam G, Sharp GW: Stimulation of insulin release by glucose is associated with an increase in the number of docked granules in the beta-cells of rat pancreatic islets. Diabetes 53:3179 –3183, 2004 22. Barg S, Eliasson L, Renstrom E, Rorsman P: A subset of 50 secretory granules in close contact with L-type Ca2⫹ channels accounts for firstphase insulin secretion in mouse beta-cells. Diabetes 51:S74 –S82, 2002 23. Allersma MW, Bittner MA, Axelrod D, Holz RW: Motion matters: secretory granule motion adjacent to the plasma membrane and exocytosis. Mol Biol Cell 17:2424 –2438, 2006 24. Barg S, Olofsson CS, Schriever-Abeln J, Wendt A, Gebre-Medhin S, Renstrom E, Rorsman P: Delay between fusion pore opening and peptide release from large dense-core vesicles in neuroendocrine cells. Neuron 33:287–299, 2002 25. Leung YM, Sheu L, Kwan E, Wang G, Tsushima R, Gaisano H: Visualization of sequential exocytosis in rat pancreatic islet beta cells. Biochem Biophys Res Commun 292:980 –986, 2002 26. Ma L, Bindokas VP, Kuznetsov A, Rhodes C, Hays L, Edwardson JM, Ueda K, Steiner DF, Philipson LH: Direct imaging shows that insulin granule exocytosis occurs by complete vesicle fusion. Proc Natl Acad Sci U S A 101:9266 –9271, 2004 27. Ohara-Imaizumi M, Nakamichi Y, Tanaka T, Ishida H, Nagamatsu S: Imaging exocytosis of single insulin secretory granules with evanescent wave microscopy: distinct behavior of granule motion in biphasic insulin release. J Biol Chem 277:3805–3808, 2002 28. Qian WJ, Aspinwall CA, Battiste MA, Kennedy RT: Detection of secretion from single pancreatic beta-cells using extracellular fluorogenic reactions and confocal fluorescence microscopy. Anal Chem 72:711–717, 2000 29. Qian WJ, Gee KR, Kennedy RT: Imaging of Zn2⫹ release from pancreatic beta-cells at the level of single exocytotic events. Anal Chem 75:3468 – 3475, 2003 30. Takahashi N, Kishimoto T, Nemoto T, Kadowaki T, Kasai H: Fusion pore dynamics and insulin granule exocytosis in the pancreatic islet. Science 297:1349 –1352, 2002 31. Taraska JW, Perrais D, Ohara-Imaizumi M, Nagamatsu S, Almers W: Secretory granules are recaptured largely intact after stimulated exocytosis in cultured endocrine cells. Proc Natl Acad Sci U S A 100:2070 –2075, 2003 32. Tsuboi T, Rutter GA: Multiple forms of “kiss-and-run” exocytosis revealed by evanescent wave microscopy. Curr Biol 13:563–567, 2003 33. Manly BJ: Randomization, Bootstrap and Monte Carlo Methods in Biology. Boca Raton, FL, CRC Press, 1991 34. R Development Core Team: R: A Language and Environment for Statistical Computing. Vienna, Austria, 2005. Available online from http:// www.R-project.org 35. Steyer JA, Almers W: Tracking single secretory granules in live chromaffin cells by evanescent-field fluorescence microscopy. Biophys J 76:2262– 2271, 1999 36. Ivarsson R, Obermuller S, Rutter GA, Galvanovskis J, Renstrom E: Temperature-sensitive random insulin granule diffusion is a prerequisite for recruiting granules for release. Traffic 5:750 –762, 2004 37. Rorsman P, Renstrom E: Insulin granule dynamics in pancreatic beta cells. Diabetologia 46:1029 –1045, 2003 38. Ammala C, Eliasson L, Bokvist K, Larsson O, Ashcroft FM, Rorsman P: Exocytosis elicited by action potentials and voltage-clamp calcium currents in individual mouse pancreatic B-cells. J Physiol 472:665– 688, 1993 39. MacDonald PE, Braun M, Galvanovskis J, Rorsman P: Release of small transmitters through kiss-and-run fusion pores in rat pancreatic beta cells. Cell Metab 4:283–290, 2006 40. MacDonald PE, Obermuller S, Vikman J, Galvanovskis J, Rorsman P, Eliasson L: Regulated exocytosis and kiss-and-run of synaptic-like microvesicles in INS-1 and primary rat beta-cells. Diabetes 54:736 –743, 2005 41. Kanno T, Ma XS, Barg S, Eliasson L, Galvanovskis J, Gopel S, Larsson M, Renstrom E, Rorsman P: Large dense-core vesicle exocytosis in pancreatic beta-cells monitored by capacitance measurements. Methods 33:302–311, 2004 42. Lakey JRT, Burridge PW, Shapiro AMJ: Technical aspects of islet preparation and transplantation. Transpl Int 16:613– 632, 2003 43. Jackson MB, Chapman ER: Fusion pores and fusion machines in Ca2⫹triggered exocytosis. Annu Rev Biophys Biomol Struct 35:135–160, 2006 44. Gilon P, Henquin JC: Influence of membrane potential changes on cytoplasmic Ca2⫹ concentration in an electrically excitable cell, the insulinsecreting pancreatic b-cell. J Biol Chem 267:20713–20720, 1992 1287


45. Grapengiesser E, Gylfe E, Hellman B: Glucose-induced oscillations of cytoplasmic Ca2⫹ in the pancreatic beta-cell. Biochem Biophys Res Commun 151:1299 –1304, 1988 46. Grapengiesser E, Gylfe E, Hellman B: Three types of cytoplasmic Ca2⫹ oscillations in stimulated pancreatic beta-cells. Arch Biochem Biophys 268:404 – 407, 1989 47. Boden G, Ruiz J, Kim CJ, Chen XH: Effects of prolonged glucose infusion on insulin secretion, clearance, and action in normal subjects. Am J Physiol 33:E251–E258, 1996 48. Huang L, Shen H, Atkinson MA, Kennedy RT: Detection of exocytosis at individual pancreatic beta cells by amperometry at a chemically modified microelectrode. Proc Natl Acad Sci U S A 92:9608 –9612, 1995

1288

49. Duncan RR, Greaves J, Wiegand UK, Matskevich I, Bodammer G, Apps DK, Shipston MJ, Chow RH: Functional and spatial segregation of secretory vesicle pools according to vesicle age. Nature 422:176 –180, 2003 50. Rudolf R, Salm T, Rustom A, Gerdes HH: Dynamics of immature secretory granules: role of cytoskeletal elements during transport, cortical restriction, and f-actin-dependent tethering. Mol Biol Cell 12:1353–1365, 2001 51. Johns LM, Levitan ES, Shelden EA, Holz RW, Axelrod D: Restriction of secretory granule motion near the plasma membrane of chromaffin cells. J Cell Biol 153:177–190, 2001 52. Oheim M, Loerke D, Stuhmer W, Chow RH: Multiple stimulation dependent processes regulate the size of the releasable pool of vesicles. Eur Biophys J 28:91–101, 1999

DIABETES, VOL. 56, MAY 2007