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Published online July 16, 2007
Diabetes 56:2579-2588, 2007
DOI: 10.2337/db06-1207
© 2007 by the American Diabetes Association
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Interaction Between Munc13-1 and RIM Is Critical for Glucagon-Like Peptide-1–Mediated Rescue of Exocytotic Defects in Munc13-1–Deficient Pancreatic ß-Cells

Edwin P. Kwan1, Li Xie1, Laura Sheu1, Toshihisa Ohtsuka2, and Herbert Y. Gaisano1

1 Departments of Physiology and Medicine, University of Toronto, Toronto, Ontario, Canada
2 Department of Clinical and Molecular Pathology, University of Toyama, Toyama, Japan

Address correspondence and reprint requests to Dr. Herbert Y. Gaisano, Medical Sciences Bldg., Rm. 7226, 1 King's College Circle, University of Toronto, Toronto, Ontario M5S 1A8, Canada. E-mail: herbert.gaisano{at}utoronto.ca

Abbreviations: 8-pCPT-2'-O-Me-cAMP, 8-(4-chloro-phenylthio)-2'-O-methyladenosine-3',5'-cyclic monophosphate; AUC, area under the curve; BBDC, Banting and Best Diabetes Center; CIHR, Canadian Institutes of Health Research; Cm, membrane capacitance; DAG, diacylglycerol; Epac, exchange protein directly activated by cAMP; F-PIS, first-phase insulin secretion; GLP-1, glucagon-like peptide-1; GSIS, glucose-stimulated insulin secretion; GST, glutathione S-transferase; IBMX, isobutylmethylxanthine; KATP channel, ATP-sensitive K+ channel; KRBH, Krebs-Ringer bicarbonate HEPES buffer; N6-Bnz-cAMP, N6-benzoyladenosine-cAMP; PKA, protein kinase A; RIA, radioimmunoassay; Rim, Rab3A-interacting molecule; RRP, readily releasable pool; SNAP, soluble N-ethylmaleimide–sensitive factor attachment protein; SNARE, SNAP receptor; S-PIS, second-phase insulin secretion; SUR1, sulfonylurea receptor 1


   ABSTRACT
TOP
 ABSTRACT
RESEARCH DESIGN AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
OBJECTIVE—Glucagon-like peptide-1 (GLP-1) rescues insulin secretory deficiency in type 2 diabetes partly via cAMP actions on exchange protein directly activated by cAMP (Epac2) and protein kinase A (PKA)-activated Rab3A-interacting molecule 2 (Rim2). We had reported that haplodeficient Munc13-1+/– mouse islet ß-cells exhibited reduced insulin secretion, causing glucose intolerance. Munc13-1 binds Epac2 and Rim2, but their functional interactions remain unclear.

RESEARCH DESIGN AND METHODS—We used Munc13-1+/– islet ß-cells to examine the functional interactions between Munc13-1 and Epac2 and PKA. GLP-1 stimulation of Munc13-1+/– islets normalized the reduced biphasic insulin secretion by its actions on intact islet cAMP production and normal Epac2 and Rim2 levels.

RESULTS—To determine which exocytotic steps caused by Munc13-1 deficiency are rescued by Epac2 and PKA, we used patch-clamp capacitance measurements, showing that 1) cAMP restored the reduced readily releasable pool (RRP) and partially restored refilling of a releasable pool of vesicles in Munc13-1+/– ß-cells, 2) Epac-selective agonist [8-(4-chloro-phenylthio)-2'-O-methyladenosine-3',5'-cyclic monophosphate] partially restored the reduced RRP and refilling of a releasable pool of vesicles, and 3) PKA blockade by H89 (leaving Epac intact) impaired cAMP ability to restore the RRP and refilling of a releasable pool of vesicles. Conversely, PKA-selective agonist (N6-benzoyladenosine-cAMP) completely restored RRP and partially restored refilling of a releasable pool of vesicles. To determine specific contributions within Epac-Rim2–Munc13-1 interaction sites accounting for cAMP rescue of exocytosis caused by Munc13-1 deficiency, we found that blockade of Rim2–Munc13-1 interaction with Rim-Munc13-1–binding domain peptide abolished cAMP rescue, whereas blockade of Epac-Rim2 interaction with Rim2-PDZ peptide only moderately reduced refilling with little effect on RRP.

CONCLUSIONS—cAMP rescue of priming defects caused by Munc13-1 deficiency via Epac and PKA signaling pathways requires downstream Munc13-1–Rim2 interaction.

Pancreatic islet ß-cells secrete insulin in a biphasic pattern consisting of a robust first-phase insulin secretion (F-PIS) (5–10 min) triggered by ATP-sensitive K+ (KATP) channel–dependent Ca2+ influx, followed by longer lasting low-level second-phase insulin secretion (S-PIS) (1). F-PIS arises from a small subset of insulin vesicles docked at the plasma membrane and primed for ready release, the readily releasable pool (RRP). S-PIS is restricted by the rate of mobilization and subsequent priming of insulin vesicles to refill the depleted RRP (2). This biphasic insulin release pattern is perturbed in patients with type 2 diabetes, resulting in abolished F-PIS and a blunted S-PIS (3). Glucagon-like peptide-1 (GLP-1) has been shown to partially bypass these insulin secretory defects of type 2 diabetes through cAMP signaling pathways (4), but the molecular mechanism by which GLP-1 restores insulin exocytotic defects in diabetes remains unclear.

SNARE (soluble N-ethylmaleimide–sensitive factor attachment protein [SNAP] receptor) proteins shown to mediate neuroexocytosis (5) also mediate insulin exocytosis (6). Munc13 proteins serve priming functions by assisting in the unfolding of SNARE protein syntaxin to its open conformation, which enables SNARE complex formation (7,8). Munc13 isoforms contain diacylglycerol (DAG)-binding C1 domain (7), which on activation by DAG promotes their translocation from cytosol to the plasma membrane (9) and is required to achieve the full priming activity (7,9,10). Munc13 priming actions lead to increase in size and release probability of the RRP in neurons and ß-cells (10,11). Using Munc13-1–deficient mouse models, Munc13-1 was shown to be the major DAG receptor responsible for potentiation of neurotransmitter (10) and insulin secretion (9,11,12). Remarkably, heterozygous Munc13-1 knockout mice (Munc13-1+/–) exhibited impaired F-PIS and S-PIS, causing glucose intolerance and mimicking type 2 diabetes (11). The reduction of islet level of Munc13-1 in Munc13-1+/– mice is similar to that in type 2 diabetic patients and rodent models (9,13).

cAMP/GLP-1 potentiation of insulin exocytosis has been attributed to an increase in initial size and refilling of the RRP of insulin vesicles (14,15). Whereas molecular substrates acted on by cAMP stimulation in more proximal steps of ß-cell excitation-secretion coupling (i.e., ion channels gating and Ca2+ release) are well studied, cAMP substrates at the level of exocytosis are just now being elucidated (16–19). cAMP/protein kinase A (PKA) activation phosphorylates a number of exocytotic proteins involved in synaptic (20,21) and neuroendocrine secretion (18), the most important of which is Rab3A-interacting molecule (Rim). We hypothesized that Munc13-1 interactions with Rim proteins mediate insulin exocytosis based on the following observations: 1) Rim, enriched in presynaptic active zones (22) as is Munc13-1 (23), is essential for normal probability of neuroexocytosis (24). 2) PKA phosphorylation of Rim is required for Rim-mediated long-term synaptic plasticity (21). 3) Rim binds Munc13-1 at its NH2-terminal domain via its Zn2+ finger region (25,26), which regulates presynaptic recruitment (27) and priming activity (25,26) of Munc13-1. 4) Munc13-1 levels are reduced in Rim-deficient brain (24). Another cAMP exocytic substrate is guanine exchange proteins (exchange proteins directly activated by cAMP [Epacs], also called cAMP-guanine exchange factors [GEFs]) (28). In islet ß-cells, Epac, via PKA-independent mechanisms (18), modulates some proximal steps of insulin secretion, including ion channels (KATP and ClC3 channels) (29,30) and Ca2+ release (17). Epac involvement in more distal steps of insulin exocytosis is via its direct interactions with Rim (18).

The current study is compatible with our hypothesis postulating the interaction of Munc13-1 with PKA-dependent (via Rim) and Epac-dependent signaling in priming insulin exocytosis. Munc13-1+/– mice exhibit abnormal biphasic insulin secretory response mimicking type 2 diabetes, with defects in priming and refilling of a releasable pool of insulin secretory vesicles (11,12). This model presents a unique opportunity to examine the ability of cAMP stimulation to rescue these exocytotic defects and to dissect the independent contribution to this cAMP rescue by Epac or PKA. Remarkably, we find that whereas the ability of Epac to potentiate insulin exocytosis is restricted by the abundance of islet Munc13-1, PKA signaling, in part via Rim2, is more efficacious in overcoming Munc13-1 deficiency-induced exocytotic defects in priming and refilling of a releasable pool of insulin vesicles to restore insulin secretion.


   RESEARCH DESIGN AND METHODS
TOP
 ABSTRACT
 RESEARCH DESIGN AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
PCR and DNA electrophoresis were performed to determine the genotypes of the Munc13-1+/+ wild-type and Munc13-1+/– knockout mice (10). All experimental procedures are approved by the University of Toronto.

Immunoblotting.
Isolated islets were solubilized in sample buffer (2% SDS) and loaded and separated on SDS-PAGE (Epac2, 30 µg protein, 6% PAGE; Rim2{alpha}, 30 µg protein, 10% PAGE; and ß-actin, 30 µg protein, 10% PAGE). Separated proteins were identified with primary antibodies (goat anti-Epac2 [Santa Cruz Biotechnology, Santa Cruz, CA], 1:100; rabbit anti-Rim2{alpha}, 1:1,000 [Synaptic Systems, Gottingen, Germany]; monoclonal mouse anti–ß-actin, 1:20,000 [Sigma, St. Louis, MO]) and appropriate peroxidase-labeled second antibodies, visualized by chemiluminescence (Pierce).

Islet perifusion secretory assay and cAMP measurements.
Mouse pancreatic islets were isolated as described previously (11), allowed to recover for at least 2 h (11 mmol/l glucose), and then cultured in 2.8 mmol/l glucose (1 h). Batches of ~25 islets were placed in perifusion chambers with a capacity of ~1.3 ml at 37°C and perifused at a flow rate of ~1 ml/min with a Krebs-Ringer bicarbonate HEPES buffer (KRBH; 10 mmol/l HEPES, pH 7.4, and 0.07% BSA). Islets were equilibrated for 30 min in KRBH (2.8 mmol/l glucose) before being subjected to the following stimulation protocols: 1) 2.8 mmol/l glucose for 10 min followed by 16.7 mmol/l glucose plus 10 nmol/l GLP-1 (7-36)-amide (Bachem, Torrance, CA) for 40 min; 2) 2.8 mmol/l glucose for 10 min, then 16.7 mmol/l glucose for 40 min, and a final 16.7 mmol/l glucose plus 10 nmol/l GLP-1 for 40 min; and 3) 2.8 or 16.7 mmol/l glucose, in presence of KCl (30 mmol/l) plus diazoxide (250 µmol/l) as described by Henquin (31). Fractions were collected for insulin determination by radioimmunoassay (RIA) (Linco Research, St. Louis, MO). At the end of each perifusion, islets were collected from the chamber and lysed in acid-ethanol (0 mmol/l HCl in 75% ethanol) for assessment of insulin content. Results are presented as insulin secreted normalized to islet insulin content.

For cAMP measurements, batches of 10–15 islets were preincubated in KRBH (2.8 mmol/l glucose, 1 h, 37°C) with 1 mmol/l isobutylmethylxanthine (IBMX; prevents cAMP degradation by inhibiting cyclic nucleotide phosphodiesterase activity) and then stimulated with glucose (16.7 mmol/l) ± 10 nmol/l GLP-1 (1 h, 37°C). Islet cAMP levels were then determined by RIA (Biomedical Technologies, Stoughton, MA).

Patch-clamp membrane capacitance measurements.
Single cells were obtained by dispersing mouse pancreatic islets (11). The intracellular pipette solution contained 125 mmol/l K-glutamate, 10 mmol/l KCl, 10 mmol/l NaCl, 1 mmol/l MgCl2, 5 mmol/l HEPES, 0.05 mmol/l EGTA, and 3 mmol/l MgATP, pH 7.1. For cAMP dialysis experiments, 0.1 mmol/l cAMP, 10 µmol/l 8-(4-chloro-phenylthio)-2'-O-methyladenosine-3',5'-cyclic monophosphate (8-pCPT-2'-O-Me-cAMP) (Biolog, Hayward, CA), or 100 µmol/l N6-benzoyladenosine-cAMP (N6-Bnz-cAMP) (Biolog) was included in the intracellular solution. For Ca2+ infusion experiments, the same pipette solution plus or minus 0.1 mmol/l cAMP was used except that EGTA was changed to 10 mmol/l, and 9 mmol/l CaCl2 was added, resulting in a free Ca2+ concentration estimated as 1.5 µmol/l (32). For dialysis experiments with glutathione S-transferase (GST) fusion proteins, 1 µmol/l GST–Rim2{alpha}-PDZ or GST–Rim2{alpha}-PDZ(AAA) (gifts from S. Seino, Division of Cellular and Molecular Medicine, Kobe University Graduate School of Medicine, Kobe, Japan) was included in the intracellular solution. Extracellular solution consisted of 138 mmol/l NaCl, 5.6 mmol/l KCl, 1.2 mmol/l MgCl2, 2.6 mmol/l CaCl2, 5 mmol/l HEPES, and 5 mmol/l D-glucose, pH 7.4. For H89 experiments, ß-cells were pretreated with 3 µmol/l H89 (Sigma) for ~10 min, and this H89 concentration was maintained in the extracellular solution during recordings. Cell membrane capacitance (Cm) was estimated by Lindau-Neher technique (33), implementing the "Sine + DC" feature of the Lock-in module (40 mV peak-to-peak and a frequency of 500 Hz for depolarization-evoked exocytosis) in whole-cell configuration. Recordings were conducted using EPC10 patch clamp amplifier, Pulse, and X-Chart softwares (HEKA Electronik, Lambrecht, Germany). Exocytic events were elicited by a train of eight 500-ms depolarization pulses (1-Hz stimulation frequency) from –70 to 0 mV. For Ca2+ infusion experiments, a 40-mV peak-to-peak 800-Hz sine wave about the holding potential (–70 mV) was applied (32). All recordings were performed at 30°C.

Statistical analysis.
Results are expressed as means ± SE where appropriate. Statistical comparisons were performed by t test in which an acceptable level of significance was considered at P < 0.05.


   RESULTS
TOP
 ABSTRACT
 RESEARCH DESIGN AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
GLP-1 stimulation normalizes insulin secretory deficiencies in islets from Munc13-1+/– mice.
Using islet perifusion, we examined biphasic glucose-stimulated insulin secretion (GSIS) in islets from Munc13-1+/+ and Munc13-1+/– mice. Glucose (16.7 mmol/l) stimulated a biphasic secretory pattern in both groups of islets (Fig. 1A). However, Munc13-1+/– islets displayed a markedly lower level of insulin secretion in both phases compared with Munc13-1+/+ islets, culminating in 37% reduction in F-PIS (3.42 ± 0.33 vs. 2.17 ± 0.32) and 45% reduction in S-PIS (10.51 ± 1.12 vs. 5.79 ± 0.56) as quantified by area under the curve (AUC) analysis (Fig. 1A and B). Remarkably, subsequent addition of 10 nmol/l GLP-1 potentiated S-PIS in Munc13-1+/– islets to the same extent as Munc13-1+/+ islets (Fig. 1A). In a second protocol, GLP-1 potentiated F-PIS and S-PIS in Munc13-1+/+ and Munc13-1+/– islets to the same extent (Fig. 1C).


Figure 1
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  		FIG. 1. GLP-1 stimulation rescues impaired GSIS response in perifused islets from Munc13-1+/– mice. A: Groups of ~25 islets from Munc13-1+/+ and Munc13-1+/– mice were perifused with agonists as indicated. Insulin released was normalized to total insulin content. B: Means ± SE of AUCs of insulin levels during islet perifusion from A. F-PIS insulin secretion was quantified from 11–19 min and S-PIS from 19–50 min from A. Data are means ± SE of seven determinations in five separate experiments in A to B. C: Islets from Munc13-1+/+ and Munc13-1+/– mice were perifused with agonists as indicated. Data are means ± SE of five determinations in five separate experiments in C. D: Islets from Munc13-1+/+ and Munc13-1+/– mice were perifused with agonists as indicated. E: A mean ± SE of AUC of insulin levels during islet perifusion from D. Data are means ± SE of six determinations for each condition in two separate experiments in D to E. F: cAMP production was evoked from groups of 10–15 islets from Munc13-1+/+ and Munc13-1+/– mice by incubating them for 1 h at 16.7 mmol/l glucose + 1 mmol/l IBMX, with or without 10 nmol/l GLP-1 (7-36)-amide. Data are means ± SE of five to six determinations in two separate experiments. *P < 0.05 for Munc13-1+/– vs. Munc13-1+/+.

 
To more rigorously determine which phase(s) of insulin secretion GLP-1 was rescued, we used a well-established protocol (31) of depolarizing concentrations of KCl (30 mmol/l) plus KATP-channel opener diazoxide (250 µmol/l) in the presence of low glucose (2.8 mmol/l) to isolate F-PIS (triggering phase) or in the presence of high-concentration glucose (16.7 mmol/l) to isolate S-PIS (amplifying phase). In low glucose conditions (KCl plus diazoxide), Munc13-1+/– islets displayed a markedly lower level of F-PIS compared with Munc13-1+/+ islets (Fig. 1D), culminating in 38% reduction (13.30 ± 1.16 vs. 8.93 ± 1.08) as quantified by AUC analysis (Fig. 1E). In high glucose conditions (KCl plus diazoxide), Munc13-1+/– islets displayed a lower level of S-PIS compared with Munc13-1+/+ islets (Fig. 1D), culminating in 31% reduction (31.56 ± 2.66 vs. 21.77 ± 1.37) (Fig. 1E). GLP-1 was therefore able to rescue both F-PIS and S-PIS in Munc13-1+/– islets (Fig. 1A and C). The relatively greater reduction in S-PIS in glucose alone (Fig. 1B) versus S-PIS in KCl plus diazoxide and high glucose (Fig. 1E) in Munc13-1+/– islets indicates a permissive role of the triggering phase in the amplifying phase in GSIS (31). Of note, this normalization of insulin secretory defects in Munc13-1+/+ islets is not due to any alteration of islet cAMP accumulation, which was similar between Munc13-1+/+ and Munc13-1+/– islets in either GSIS or GLP-1 stimulation conditions (Fig. 1F).

cAMP stimulation rescues exocytotic defects in Munc13-1+/– islet ß-cells by enhancing the size of RRP and accelerating refilling of a releasable pool of vesicles.
We examined the specific pool of insulin vesicles acted on by GLP-1 stimulation to normalize biphasic insulin secretion in Munc13-1+/– islets (Fig. 1) by Cm measurements of single ß-cells. Cm changes elicited by the first two pulses approximate the number of vesicles released from and therefore the size of the RRP of primed fusion-ready vesicles. Subsequent pulses estimate the rate of refilling or mobilization of insulin vesicles from reserve pool(s) to RRP. The size of RRP and rate of refilling of a releasable pool of vesicles have been shown to correlate favorably with F-PIS and S-PIS from whole pancreatic islets (1).

Using this strategy of Cm measurements of single ß-cells, we observed that depolarization-evoked insulin exocytosis was severely reduced in Munc13-1+/– compared with Munc13-1+/+ ß-cells (Fig. 2A–C), consistent with our previous report (11). The size of RRP of vesicles ({Sigma}{Delta}Cm1st-2ndpulse) was reduced by 52% in Munc13-1+/– ß-cells (2.79 ± 0.15 vs. 5.77 ± 0.87 femto Farad per pico Farad [fF/pF] in Munc13-1+/+ ß-cells; Fig. 2D). Rate of refilling of a releasable pool of vesicles ({Sigma}{Delta}Cm3rd-8thpulse) was reduced by 54% in Munc13-1+/– ß-cells (3.82 ± 0.72 vs. 8.36 ± 1.04 fF/pF in Munc13-1+/+ ß-cells; Fig. 2D). No changes was observed in whole-cell currents evoked during the depolarization pulses, suggesting that the interactions between the ß-cell Kv channels and exocytotic SNARE proteins were unaltered in the Munc13-1+/– ß-cells (11). Like GLP-1, intracellular dialysis of 100 µmol/l cAMP to maximally activate cAMP signaling (29) greatly enhanced insulin exocytosis in both Munc13-1+/+ and Munc13-1+/– ß-cells (Fig. 3A–C). Remarkably, increase in RRP size ({Sigma}{Delta}Cm1st-2ndpulse) by cAMP activation in Munc13-1+/– ß-cells (11.60 ± 1.88 fF/pF) was very similar to Munc13-1+/+ ß-cells (13.96 ± 1.84 fF/pF) (Fig. 3D). The increased rate of refilling of a releasable pool of vesicles ({Sigma}{Delta}Cm3rd-8thpulse) by cAMP activation in Munc13-1+/– ß-cells (31.64 ± 6.02 fF/pF) was only mildly reduced by 28% (P = 0.11) compared with Munc13-1+/+ ß-cells (44.21 ± 4.67 fF/pF) (Fig. 3D). Similarly, the normalized exocytosis in Munc13-1+/– ß-cells is not due to altered whole-cell currents as explained above. The full rescue of RRP in Munc13-1+/– ß-cells by maximal cAMP stimulation is commensurate to GLP-1–potentiated F-PIS in the Munc13-1+/– islet perifusion study (Fig. 1). However, the less-effective rescue of refilling of a releasable pool of vesicles by cAMP in Munc13-1+/– ß-cells is in contrast to the normalized S-PIS.


Figure 2
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  		FIG. 2. Exocytosis in single islet ß-cells of Munc13-1+/+ and Munc13-1+/– mice. Changes in cell Cm ({Delta}Cm) were measured from single ß-cells using a train of eight depolarizing pulses (500 ms in duration) from –70 to 0 mV. A: Representative recordings of {Delta}Cm normalized to basal cell Cm (fF/pF) from Munc13-1+/+ and Munc13-1+/– ß-cells evoked by depolarization alone (top). Whole-cell currents normalized to basal Cm (pA/pF) evoked during first and eighth pulse were also shown (bottom). Note that Cm changes do not acutely change current density, which remains the same during the train of pulses in both groups of ß-cells. B: The corresponding summaries of {Delta}Cm normalized to basal cell Cm (fF/pF) (n = 8–10 cells). C: The mean incremental changes in Cm (i.e., {Delta}Cmn – {Delta}Cmn-1) evoked by the individual pulses in B. D: The size of RRP of insulin vesicles ({Sigma}{Delta}Cm1st-2ndpulse) and the rate of refilling of a releasable pool of vesicles ({Sigma}{Delta}Cm3rd-8thpulse) (n = 8–10 cells). Data are means ± SE. *P < 0.05 for Munc13-1+/– vs. Munc13-1+/+.

 

Figure 3
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  		FIG. 3. cAMP potentiation of insulin exocytosis in single islet ß-cells of Munc13-1+/+ and Munc13-1+/– mice. Single ß-cells were subjected to the same depolarization protocol as in Fig. 2, and {Delta}Cm was determined. A: Representative recordings of {Delta}Cm (fF/pF) (top) and whole-cell currents (pA/pF) (bottom) of ß-cells when evoked in presence of dialyzed 100 µmol/l cAMP. B: Summaries of changes in {Delta}Cm (fF/pF) (n = 9–10 cells). C: Mean {Delta}Cmn – {Delta}Cmn-1 evoked by the individual pulses in B. D: The size of RRP of insulin vesicles ({Sigma}{Delta}Cm1st-2ndpulse) and rate of refilling of a releasable pool of vesicles ({Sigma}{Delta}Cm3rd-8thpulse) (n = 9–10 cells). E: Representative recordings of {Delta}Cm (pF) of ß-cells evoked by intracellular infusion of a Ca2+-EGTA buffer with a free [Ca2+] of 1.5 µmol/l ± 0.1 mmol/l cAMP. F: Summarizes the rates of Cm increase (dC/dt) determined 0–30 or 60–120 s after establishment of whole-cell configuration (n = 8–11 cells). Data are means ± SE. *P < 0.05 for Munc13-1+/– vs. Munc13-1+/+.

 
Of note, depolarization protocols have some inherent limitations. First, Ca2+ channels induced to open by the depolarization protocols might be affected by either the Munc13-1 deficiency or the cAMP rescue. Second, Munc13-1 deficiency might alter the coupling of insulin vesicles to Ca2+ channels or influence Ca2+-induced Ca2+ release. Any of these secretory components could have been acted on by cAMP to rescue exocytosis (Fig. 2) in a manner independent of the residual Munc13-1. To negate the contributions from these confounding possibilities, we "bypassed" the plasma membrane Ca2+ channel by infusing into ß-cells a constant stimulatory concentration of Ca2+ (1.5 µmol/l) to induce Ca2+-evoked exocytosis (Fig. 3E). Analysis of the rate of change of Cm (cAMP free) at a 60- to 120-s period, which approximates the refilling rate of a releasable pool of vesicles, showed a reduction of 41% in Munc13-1+/– ß-cells (3.32 ± 0.22 vs. 5.67 ± 0.51 fF · s–1 in Munc13-1+/+ ß-cells; Fig. 3F). This magnitude of reduction is similar to that observed with the serial depolarization (54%) study (Fig. 3A–D). However, when we calculated the rate of change of Cm during the first 30 s, which approximates the RRP size, we saw a reduction of 27% in Munc13-1+/– ß-cells (6.70 ± 0.53 vs. 9.24 ± 0.85 fF · s–1 in Munc13-1+/+ ß-cells; Fig. 3F). This reduction is milder than that seen when exocytosis was induced by serial depolarization (52%) (Fig. 3A–D), suggesting a reduction in Ca2+ sensitivity. These results indicate that while Munc13-1 deficiency caused exocytotic defects by reducing the initial size and refilling of the RRP, the Munc13-1 deficiency also seemed to have caused some reduction in the sensitivity of exocytosis to Ca2+, the latter also contributing to the priming defects. Remarkably, in presence of cAMP, Cm increase in Munc13-1+/– ß-cells was completely normalized to that in Munc13-1+/+ ß-cells (14.84 ± 0.89 vs. 14.95 ± 0.80 fF · s–1 in Munc13-1+/+ ß-cells; Fig. 3E and F) during the first 30 s and was only mildly reduced by 22% in Munc13-1+/– ß-cells (10.30 ± 0.88 vs. 13.27 ± 1.49 fF · s–1 in Munc13-1+/+ ß-cells; Fig. 3E and F) during the 60- to 120-s period, indicating that cAMP rescues the exocytotic defects caused by the Munc13-1 deficiency.

Epac activation alone is incapable of fully rescuing the exocytotic defects in Munc13-1+/– islet ß-cells.
cAMP/GLP-1–mediated potentiation of insulin secretion has been attributed to its actions on two major cAMP receptors, cAMP/Epac and cAMP/PKA (18). We hypothesized that these two cAMP receptors play a distinct role in rescuing the insulin exocytotic defects in Munc13-1+/– ß-cells. We measured islet expressions of Epac2 and Rim2 (a major PKA substrate), which were identical between Munc13-1+/– and Munc13-1+/+ mice (Fig. 4A). These cAMP receptors are therefore equally intact in both Munc13-1+/+ and Munc13-1+/– ß-cells for cAMP to exert its full actions.


Figure 4
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  		FIG. 4. Epac-mediated (PKA-independent) potentiation of insulin exocytosis in single-islet ß-cells of Munc13-1+/+ and Munc13-1+/– mice. A: Western blots of Rim2 and Epac2 proteins from pancreatic islets of Munc13-1+/+ and Munc13-1+/– mice. Thirty micrograms protein per lane was loaded. Note that expression levels of Rim2 and Epac2 are comparable between Munc13-1+/+ and Munc13-1+/– islets (densitometry of Munc13-1+/– islets relative to that of Munc13-1+/+ islets: ß-actin, 77%; Rim2, 86%; Epac2, 115%). Single ß-cells were subjected to the same depolarization protocol as in Fig. 2, and {Delta}Cm was determined. B: Summaries of changes in {Delta}Cm (fF/pF) of ß-cells when evoked in presence of dialyzed 10 µmol/l 8-pCPT-2'-O-Me-cAMP (n = 15–20 cells). Representative Cm recordings and whole-cell current traces in are shown in Supplemental Fig. 1A. C: Mean {Delta}Cmn – {Delta}Cmn-1 evoked by individual pulses in B. D: The size of RRP of insulin vesicles ({Sigma}{Delta}Cm1st-2ndpulse) and rate of refilling of a releasable pool of vesicles ({Sigma}{Delta}Cm3rd-8thpulse) (n = 15–20 cells). Data are means ± SE. *P < 0.05 for Munc13-1+/– vs. Munc13-1+/+.

 
We proceeded to examine the independent actions of Epac signaling. We performed intracellular dialysis of 10 µmol/l 8-pCPT-2'-O-Me-cAMP, a selective agonist for Epac but not PKA (17). Although 8-pCPT-2'-O-Me-cAMP enhanced depolarization-evoked exocytosis (cAMP-free) in both groups of ß-cells (Fig. 4B and C), Epac-potentiated exocytosis was less than that potentiated by maximal cAMP stimulation (Fig. 3A–D), in keeping with previous reports (28,29). More importantly, absolute (Fig. 4B) and incremental (Fig. 4C) increases in Cm by serial depolarization were lower in every single pulse in Munc13-1+/– ß-cells compared with Munc13-1+/+ ß-cells, culminating in 43% reduction in size of RRP ({Sigma}{Delta}Cm1st-2ndpulse) in Munc13-1+/– ß-cells (5.48 ± 0.50 vs. 9.68 ± 0.97 fF/pF in Munc13-1+/+ ß-cells) and a remarkably similar reduction of 41% in rate of refilling of a releasable pool of vesicles ({Sigma}{Delta}Cm3rd-8thpulse, 9.81 ± 1.35 vs. 16.51 ± 1.47 fF/pF in Munc13-1+/+ ß-cells; Fig. 4D). These results suggest that although Epac signaling pathway is functional and contributes to cAMP-mediated rescue of exocytic defects in Munc13-1+/– ß-cells, Epac signaling alone is insufficient in fully restoring the insulin exocytotic defects caused by Munc13-1 deficiency, implying that Munc13-1 abundance is required for full potentiating effects of Epac activation. This led us to postulate that PKA signaling would also contribute to cAMP-induced rescue of exocytosis caused by Munc13-1 deficiency.

PKA activation alone contributes to cAMP-induced rescue of exocytotic defects in Munc13-1+/– islet ß-cells.
To examine the independent contribution of PKA signaling in rescuing exocytosis in Munc13-1+/– ß-cells, we blocked Epac signaling by dialyzing into ß-cells 1 µmol/l GST-Rim2PDZ peptide while simultaneously including 100 µmol/l cAMP in the pipette solution to activate the PKA signaling pathway. GST-Rim2PDZ directly interacts with Epac2 in a manner that abolished Epac2-Rim2 binding, which is essential for Epac to potentiate exocytosis (18). Cm changes elicited by the first four depolarization pulses were similar between Munc13-1+/– and Munc13-1+/+ ß-cells (Fig. 5A and B), including similar sizes of RRP ({Sigma}{Delta}Cm1st-2ndpulse) between Munc13-1+/– (9.67 ± 1.02 fF/pF) and Munc13-1+/+ (11.89 ± 1.26 fF/pF) ß-cells (Fig. 5C), indicating little effect by Epac blockade on RRP size. Cm changes elicited by subsequent four pulses were lower in Munc13-1+/– ß-cells (Fig. 5A and B), such that overall refilling of a releasable pool of vesicles encompassing the third to eighth pulse ({Sigma}{Delta}Cm3rd-8thpulse) remained impaired by 30% in Munc13-1+/– ß-cells (19.16 ± 2.74 vs. 27.22 ± 2.24 fF/pF in Munc13-1+/+ ß-cells; Fig. 5C). These patterns are rather similar to those effected by maximal cAMP activation (Fig. 3). As control for Rim2PDZ, 1 µmol/l GST-Rim2PDZ(AAA), a mutant that cannot bind Epac2 (28), did not affect cAMP-mediated potentiation of exocytosis (online appendix Supplemental Fig. 1C [available at http://dx.doi.org/10.2337/db06-1207]). To confirm that the inhibitory effects of GST-Rim2PDZ were attributable to blockade of Epac signaling, we showed that GST-Rim2PDZ completely inhibited the potentiating effects of 8-pCPT-2'-O-Me-cAMP (Fig. 5D–F).


Figure 5
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  		FIG. 5. Assessment of blockade of Epac-mediated potentiation of insulin exocytosis in single islet ß-cells of Munc13-1+/+ and Munc13-1+/– mice. Single ß-cells were subjected to the same depolarization protocol as in Fig. 2, and {Delta}Cm was determined. Summaries of changes in {Delta}Cm (fF/pF) of ß-cells when evoked in presence of dialyzed 100 µmol/l cAMP plus 1 µmol/l GST-Rim2PDZ peptide (n = 12–13 cells) (A) or 10 µmol/l 8-pCPT-2'-O-Me-cAMP plus 1 µmol/l GST-Rim2PDZ peptide (n = 6–7 cells) (D). Representative Cm recordings and whole-cell current traces are shown in Supplemental Fig. 1B and D. B and E: Mean {Delta}Cmn – {Delta}Cmn-1 evoked by individual pulses in A and D, respectively. C and F: The size of RRP of insulin vesicles ({Sigma}{Delta}Cm1st-2ndpulse) and the rate of refilling of a releasable pool of vesicles ({Sigma}{Delta}Cm3rd-8thpulse) in A and B (n = 12–13 cells) and D and E (n = 6–7 cells), respectively. D: Summaries of changes in {Delta}Cm (fF/pF) of ß-cells in C (n = 6–7 cells). Data are means ± SE. *P < 0.05 for Munc13-1+/– vs. Munc13-1+/+.

 
Because the PKA signaling pathway was discriminated only indirectly by inhibiting Epac signaling in Fig. 5, we examined direct PKA activation by dialyzing 100 µmol/l N6-Bnz-cAMP, an agonist selective for PKA but not Epac (34). N6-Bnz-cAMP stimulation increased the size of RRP ({Sigma}{Delta}Cm1st-2ndpulse) in Munc13-1+/– ß-cells (7.18 ± 0.51 fF/pF) to a similar extent as Munc13-1+/+ ß-cells (8.20 ± 0.72 fF/pF) (Fig. 6A–C). Refilling of a releasable pool of vesicles ({Sigma}{Delta}Cm3rd-8thpulse) remained mildly but not significantly impaired (~12%) in Munc13-1+/– ß-cells (14.52 ± 1.08 vs. 16.57 ± 1.18 fF/pF in Munc13-1+/+ ß-cells; Fig. 6A–C). These results suggest that PKA activation alone has a tendency to overcome most of the exocytotic defects caused by Munc13-1 deficiency. Nonetheless, maximal PKA-potentiated exocytosis by N6-Bnz-cAMP was less than maximal cAMP potentiation (Fig. 6C), consistent with previous reports (28). When these results are further compared with 8-pCPT-2'-O-Me-cAMP (from Fig. 4), Epac signaling accounts for some of the cAMP rescue of the exocytotic defects in Munc13-1+/– ß-cells but seems to contribute somewhat less to the cAMP rescue than PKA signaling (Fig. 6D). Nonetheless, it also appears that synergistic actions of both Epac and PKA signaling are required for full cAMP rescue.


Figure 6
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  		FIG. 6. PKA-dependent potentiation of insulin exocytosis in single islet ß-cells of Munc13-1+/+ and Munc13-1+/– mice. Single ß-cells were subjected to the same depolarization protocol as in Fig. 2, and {Delta}Cm was determined. A: Summaries of changes in {Delta}Cm (fF/pF) of ß-cells when evoked in presence of dialyzed 100 µmol/l N6-Bnz-cAMP (N6Bnz; n = 7–8 cells). Representative Cm recordings and whole-cell current traces are shown in Supplemental Fig. 1E. B: Mean {Delta}Cmn – {Delta}Cmn-1 evoked by individual pulses in A. C and D: The size of RRP of insulin vesicles ({Sigma}{Delta}Cm1st-2ndpulse) and rate of refilling of a releasable pool of vesicles ({Sigma}{Delta}Cm3rd-8thpulse) (n = 7–8 cells), comparing the effects of N6-Bnz with cAMP (C) and with 8-pCPT-2'-O-Me-cAMP (8CPT) (D). Data are means ± SE. *P < 0.05 for Munc13-1+/– vs. Munc13-1+/+.

 
Blockade of Munc13-1–Rim interaction severely impairs cAMP rescue of exocytic defects in Munc13-1+/– islet ß-cells.
If PKA activation tends to account for much of the cAMP-mediated rescue of exocytotic defects in Munc13-1+/– ß-cells, then blockade of PKA signaling should block these potentiating actions of maximal cAMP stimulation. Here, we have used the potent and relatively selective PKA inhibitor H89 (3 µmol/l) plus 100 µmol/l cAMP, which would leave Epac2 signaling intact. In presence of H89, cAMP failed to fully amplify insulin exocytosis in both groups of ß-cells (Fig. 7A and B). Cm changes were much lower in Munc13-1+/– ß-cells in a majority of depolarization pulses (Fig. 7A and B). RRP ({Sigma}{Delta}Cm1st-2ndpulse) was reduced by 43% in Munc13-1+/– ß-cells (3.21 ± 0.70 vs. 5.66 ± 1.27 fF/pF in Munc13-1+/+ ß-cells; Fig. 7C). Refilling of a releasable pool of vesicles ({Sigma}{Delta}Cm3rd-8thpulse) was similarly impaired by 42% in Munc13-1+/– ß-cells (10.40 ± 0.81 vs. 17.99 ± 2.14 fF/pF in Munc13-1+/+ ß-cells; Fig. 7C). These results confirm that PKA signaling is a major contributor to cAMP potentiation and rescue of the exocytotic defects in Munc13-1+/– ß-cells.


Figure 7
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  		FIG. 7. Assessment of blockade of PKA-dependent potentiation of insulin exocytosis in single islet ß-cells of Munc13-1+/+ and Munc13-1+/– mice. Single ß-cells were subjected to the same depolarization protocol as in Fig. 2, and {Delta}Cm was determined. A: Summaries of changes in {Delta}Cm (fF/pF) ß-cells when evoked in presence of dialyzed 100 µmol/l cAMP plus 3 µmol/l H89 (n = 8–10 cells). Representative Cm recordings and whole-cell current traces are shown in Supplemental Fig. 1F. B: Mean {Delta}Cmn – {Delta}Cmn-1 evoked by individual pulses in A. C: The size of RRP of insulin vesicles ({Sigma}{Delta}Cm1st-2ndpulse) and rate of refilling of a releasable pool of vesicles ({Sigma}{Delta}Cm3rd-8thpulse) (n = 8–10 cells). Data are means ± SE. *P < 0.05 for Munc13-1+/– vs. Munc13-1+/+.

 
Rim2 has been suggested to mediate PKA-dependent potentiation of insulin secretion (18). Rim isoform requires PKA-dependent phosphorylation for activation (21) and is believed to interact with Munc13-1 to enhance the exocytic action of Munc13-1 (24), but the precise mechanism remains unclear. We hypothesized that functional interaction of Munc13-1 and Rim was critical for PKA signaling to overcome the Munc13-1 deficiency. We blocked endogenous Rim2–Munc13-1 interaction in ß-cells by dialysis of 1 µmol/l GST-Munc13-1–Rim-binding domain (RBD) peptide (35). Here, cAMP rescue of RRP size and refilling of a releasable pool of vesicles in Munc13-1+/– ß-cells were greatly reduced (Fig. 8A and B), with RRP ({Sigma}{Delta}Cm1st-2ndpulse) reduced by 54% in Munc13-1+/– ß-cells (4.46 ± 0.74 vs. 9.65 ± 1.63 fF/pF in Munc13-1+/+ ß-cells; Fig. 8C) and refilling ({Sigma}{Delta}Cm3rd-8thpulse) impaired by 37% in Munc13-1+/– ß-cells (12.56 ± 0.57 vs. 19.95 ± 1.93 fF/pF in Munc13-1+/+ ß-cells; Fig. 8C). These results strongly support a putative role of Munc13-1–Rim interaction in cAMP-mediated rescue of Munc13-1 exocytotic defects.


Figure 8
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  		FIG. 8. Assessment of blockade of Munc13-1–Rim interaction in cAMP-mediated rescue of insulin exocytosis in single Munc13-1+/– islet ß-cells. Single ß-cells were subjected to the same depolarization protocol as in Fig. 2, and {Delta}Cm was determined. A: Summaries of changes in {Delta}Cm (fF/pF) ß-cells when evoked in presence of dialyzed 100 µmol/l cAMP plus 1 µmol/l GST-Munc13-1–RBD peptide (n = 7–9 cells). Representative Cm recordings and whole-cell current traces are shown in Supplemental Fig. 1G. B: Mean {Delta}Cmn – {Delta}Cmn-1 evoked by individual pulses in A. C: The size of RRP of insulin vesicles ({Sigma}{Delta}Cm1st-2ndpulse) and rate of refilling of a releasable pool of vesicles ({Sigma}{Delta}Cm3rd-8thpulse) (n = 7–9 cells). Data are means ± SE. *P < 0.05 for Munc13-1+/– vs. Munc13-1+/+.

 

   DISCUSSION
TOP
 ABSTRACT
 RESEARCH DESIGN AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Biphasic insulin secretion is attributed to exocytosis of distinct pools of insulin vesicles, which become severely perturbed in type 2 diabetes (3). GLP-1 mimetics, acting on cAMP- and PKA-signaling pathways, have emerged as promising therapeutic agents in treating type 2 diabetes (4), by normalizing insulin exocytotic defects. However, the molecular substrates and corresponding steps in exocytosis (vesicle priming and refilling) acted on by GLP-1 remain unclear. In this work, we have gained much insight into the functional interactions of Munc13-1, Epac, and PKA (including Rim), which compose the priming machinery for vesicle priming and refilling steps of exocytosis. We have used Munc13-1–haplodeficient mouse islets, which exhibit defective biphasic insulin secretion mimicking type 2 diabetes and whose ß-cells exhibit major defects in priming and refilling steps in exocytosis. We examined how these defects caused by Munc13-1 deficiency are selectively rescued by agents acting on distinct cAMP and PKA signaling pathways. Below, we discuss our major findings.

First, we characterized the kinetics of insulin secretion and exocytosis of adult haplodeficient Munc13-1+/– islets. Islet perifusion studies showed that Munc13-1+/– islets had severe reduction in F-PIS and S-PIS. Consistently, Cm measurements revealed severe impairment in the size of RRP and rate of refilling of a releasable pool of vesicles. Using homozygous Munc13-1–/– knockout mouse islets, Kang et al. (12) showed nearly abolished S-PIS and severely reduced F-PIS. Their Cm measurements showed Munc13-1–/– ß-cells to exhibit severe reduction in rate of refilling of a releasable pool of vesicles, but surprisingly, the size of RRP was normal. Of note, in the study by Kang et al., ß-cells examined were from perinatal mice because homozygous Munc13-1–/– mice die shortly after birth (7). The size of RRP of insulin vesicles in Munc13-1–/– mice could be larger initially and may then undergo normal progressive reduction during development. Alternatively, progressive reduction of RRP and F-PIS could have been induced by the diabetic state because adult Munc13+/– mice are glucose intolerant (11). A more prolonged period of Munc13-1 deficiency in adult Munc13+/– mice could lead to reduced tightness of coupling between Ca2+ channels and insulin vesicles, resulting in less efficient excitosome complex to effect exocytosis (36).

Second, GLP-1 stimulation normalized biphasic insulin secretory defects caused by Munc13-1 deficiency. Consistently, examination of kinetics of exocytosis showed that maximal cAMP stimulation completely restored the size of RRP and almost completely recovered the rate of refilling of a releasable pool of vesicles. The restoration was not due to alteration in Ca2+ entry through voltage-gated Ca2+ channels, which we previously showed to be normal (11), but was in part due to rescue of impaired sensitivity to a Ca2+ trigger as demonstrated in the Ca2+ infusion experiments. The rescue of refilling of a releasable pool of vesicles was somewhat less efficient than the complete rescue of S-PIS because serial depolarization is "supraphysiological" compared with physiological GSIS. Nonetheless, such an acute supraphysiological stimulation may simulate the prolonged and high demand on pancreatic islets imposed by diabetes. Islet levels cAMP receptors Epac and PKA substrate Rim2, as well as cAMP production, were normal in Munc13+/– mice, indicating that GLP-1 could activate these cAMP receptors to bypass Munc13-1 deficiency or interact with residual Munc13-1 proteins to promote insulin exocytosis. Below, we discussed the independent contributions of Epac and PKA signaling.

Third, Epac activation partially normalized the defects in priming and refilling of a releasable pool of vesicles caused by Munc13-1 deficiency. Epac2 and Rim2 interaction is essential for GLP-1–enhanced insulin secretion (18). Epac2/Rim2 complex was postulated to serve as a scaffold to bind and orchestrate additional exocytic proteins, including not only Munc13-1 (24,27) and SNARE proteins but also other proteins involved in exocytosis, including Piccolo (Ca2+ sensor of this complex), L-type Ca2+ channel {alpha}-subunit, and nucleotide-binding fold of sulfonylurea receptor 1 (SUR1) (18). Epac activation through SUR1 inhibits KATP channels (30) and enhances Cl– influx from insulin vesicles (37), leading to enhanced insulin exocytosis, and was abolished in SUR1-null mice (29). Ca2+-induced Ca2+ release could be amplified by Epac activation (17). Epac2 therefore integrates ATP, cAMP, and Ca2+ signals in pancreatic ß-cells to facilitate stimulus-exocytosis coupling. Our results showed that Epac signaling contributes to but is insufficient to account for the full cAMP rescue of exocytic defects in Munc13-1+/– ß-cells.

Fourth, PKA activation is a major contributor to cAMP rescue of exocytic defects caused by Munc13-1 deficiency and is in part mediated via Rim2–Munc13-1 interaction. We show that PKA activation fully amplified the size of RRP of insulin vesicles to the same extent as maximal cAMP stimulation and partially accelerated refilling of a releasable pool of vesicles. PKA-catalyzed protein phosphorylation is known to potentiate insulin secretion, but the precise molecular substrates remain largely unknown (18). Rim proteins are major phosphorylation targets of PKA and can directly bind Munc13-1 (7,21). We here demonstrated that Rim2–Munc13-1 interaction was critical for the rescue by PKA. Rim binds Munc13-1 even when it is not PKA-phosphorylated, and disruption of Rim-Munc13-1 binding reduces fusion-competent synaptic vesicles mimicking Munc13-1 knockout neurons (25). Although the NH2-terminal Rim-binding L-region of Munc13-1 does not exert a priming reaction per se (25,38), this binding is thought to influence indirectly the priming activity of Munc13-1 by promoting recruitment of Munc13-1 or causing conformational changes in Munc13-1 that affect its priming activity (27). These observations suggest that decrease in number of primed insulin vesicles in Munc13-1 ß-cells is contributed by reduction in Rim-Munc13-1 binding, thus indirectly impairing the priming activity of Munc13-1. Following this reasoning, improvement in priming and refilling of a releasable pool of vesicles in Munc13-1+/– ß-cells by PKA activation is likely due to more effective binding of PKA-phosphorylated Rim to Munc13-1, which would enhance the priming reaction of Munc13-1 (21). Conversely, increase in Rim-Munc13-1 binding could be facilitated by PKA-phosphorylated Munc13-1. In fact, it was recently shown that surfaces of Munc13-1 responsible for heterodimerization with Rim contain several putative phosphorylation sites (39). Although Rim-Munc13-1 binding is important for maintenance of normal RRP, it may not be the case for recovery of vesicles (26), implying that upregulation of Rim-Munc13-1 interactions in Munc13-1+/– ß-cells may not be sufficient to fully account for the rescue of the exocytotic defects by PKA activation. PKA potentiation of exocytosis is contributed by phosphorylation of other exocytotic proteins, including SNAP25, {alpha}1.2-subunit of voltage-dependent Ca2+ channels, KATP channel (18), {alpha}-SNAP (40), synapsin I (41), snapin (42), cysteine string protein (43), tomosyn (44), and syntaphilin (45), the majority of which are also involved in insulin secretion (46–48). Although few studies have attempted to explain the precise functional consequences of PKA-catalyzed phosphorylation of these substrates on insulin and neurotransmitter release (49,50), much is still unknown and remains to be further studied, including the possibility that they could functionally interact with Munc13-1.


   ACKNOWLEDGMENTS
 
E.P.K. has received graduate studentship awards from the Canada Graduate Scholarships Doctoral Awards from the Canadian Institutes of Health Research (CIHR) and support from the University of Toronto Banting and Best Diabetes Center (BBDC) and Unilever-Lipton Fellowship in Neuroscience. H.Y.G. has received grants from the CIHR (grant MOP-64465 and equipment grant MMA-66939) and the University of Toronto BBDC.


   FOOTNOTES
 
Published ahead of print at http://diabetes.diabetesjournals.org on 16 July 2007. DOI: 10.2337/db06-1207.

Additional information for this article can be found in an online appendix at http://dx.doi.org/10.2337/db06-1207.

E.P.K. and L.X. contributed equally to this work.

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.

Received for publication August 30, 2006 and accepted in revised form July 10, 2007


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