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Stimulation of Insulin Secretion by Denatonium, One of the Most Bitter-Tasting Substances Known

From the Department of Molecular Medicine, College of Veterinary Medicine, Cornell University, Ithaca, New York

	ABSTRACT

TOP ABSTRACT RESEARCH DESIGN AND METHODS RESULTS DISCUSSION REFERENCES

 
Denatonium, one of the most bitter-tasting substances known, stimulated insulin secretion in clonal HIT-T15 ß-cells and rat pancreatic islets. Stimulation of release began promptly after exposure of the ß-cells to denatonium, reached peak rates after 4–5 min, and then declined to near basal values after 20–30 min. In islets, no effect was observed at 2.8 mmol/;l glucose, whereas a marked stimulation was observed at 8.3 mmol/;l glucose. No stimulation occurred in the absence of extracellular Ca²⁺ or in the presence of the Ca²⁺-channel blocker nitrendipine. Stimulated release was inhibited by ₂-adrenergic agonists. Denatonium had no direct effect on voltage-gated calcium channels or on cyclic AMP levels. There was no evidence for the activation of gustducin or transducin in the ß-cell. The results indicate that denatonium stimulates insulin secretion by decreasing KATP channel activity, depolarizing the ß-cell, and increasing Ca²⁺ influx. Denatonium did not displace glybenclamide from its binding sites on the sulfonylurea receptor (SUR). Strikingly, it increased glybenclamide binding by decreasing the K_d. It is concluded that denatonium, which interacts with K⁺ channels in taste cells, most likely binds to and blocks Kir6.2. A consequence of this is a conformational change in SUR to increase the SUR/;glybenclamide binding affinity.

Stimulation of taste cell membranes with denatonium activates not only gustducin but also transducin (9), with which it shares high sequence homology (21). Because transducin mRNA is reported to be present in pancreatic islets at approximately one-fifth the level of that in the retina (22), we were interested to determine the effects of denatonium on insulin secretion in ß-cells as a tool to study the potential involvement of gustducin or transducin in stimulus-secretion coupling. The experiments described here were performed on clonal HIT-T15 cells and rat pancreatic islets.

	RESEARCH DESIGN AND METHODS

TOP ABSTRACT RESEARCH DESIGN AND METHODS RESULTS DISCUSSION REFERENCES

 
Cell culture.
Clonal HIT-T15 cells were cultured in RPMI 1640 medium supplemented with 10% fetal bovine serum, 100 µg/ml streptomycin, and 100 units/ml penicillin in an atmosphere of 95% air and 5% CO₂. Cells of passage numbers 72–85 were used for the experiments.

Islet isolation.
Pancreatic islets from Sprague Dawley rats were isolated using the collagenase digestion technique of Lacy and Kostianovsky (23).

Insulin secretion.
Static incubation measurements of insulin release by HIT-T15 cells were performed in Krebs-Ringer-HEPES bicarbonate buffer (KRBH) of the following composition (in mmol/l): 129 NaCl, 5 NaHCO₃, 4.8 KCl, 1.2 KH₂PO₄, 1.2 MgSO₄, 10 HEPES, and 1 CaCl₂ at pH 7.4, with 0.1% BSA and the indicated glucose concentrations. Preincubation and test incubation periods were 30 min each. For dynamic measurements under perifusion conditions, 2 x 10⁶ HIT-T15 cells on glass coverslips were placed in 700-µl chambers and perifused with KRBH buffer at a flow rate of 1 ml/min. A preperifusion for 50 min was carried out to equilibrate the secretion rates before exposure of the cells to control and test conditions. Samples were collected at 1- or 2-min intervals and kept at -20°C before radioimmunoassay with a charcoal separation method. For the measurement of insulin release by rat islets under static incubation conditions, the KRBH contained 2.5 mmol/l CaCl₂. The islets were subjected to a 60-min preincubation before being exposed to fresh buffer solution containing the test agents for an additional 60 min. Samples of the incubation medium were collected and stored at -20°C before radioimmunoassay.

Cyclic AMP determination.
After a 30-min preincubation period in KRBH buffer, HIT-T15 cells were exposed to the test agents for 5 min and the incubation was stopped by the addition of 0.25 ml of 6% trichloroacetic acid. The cells were scraped free, transferred to Eppendorf tubes, and centrifuged at 2,500g for 10 min at 4°C. The cell pellet was solubilized in 0.1 mmol/l NaOH, and protein content was assessed using the Bradford assay. The supernatant was used for determination of the cyclic AMP content by radioimmunoassay after ether extraction of the trichloroacetic acid.

Electrophysiology.
Cell membrane potentials and currents were measured under current clamp or voltage clamp conditions using either whole-cell or perforated-patch configurations (24,25). All recordings were made at room temperature. HIT-T15 cells plated on 18-mm circular glass coverslips were placed in a recording chamber mounted on an inverted microscope. Whole-cell recordings were performed within 2–4 days after the cells were plated. Patch-clamp recordings were done on cells that were not in contact with other cells to avoid possible cell-cell coupling artifacts. Patch-clamp electrodes were made from borosilicate glass capillaries (Garner Glass Company) using a two-stage puller (Narishighe). Electrodes were adjusted by firepolishing to obtain a tip resistance of 2–4 megohms. Amphotericin B was added to the intracellular solution from a stock solution to obtain a final concentration of 200 µg/ml. Recording electrodes were tip-filled with clean intracellular solution and then back-filled with intracellular solution containing 200 µg/ml Amphotericin B. The electrode tip resistance was monitored by applying a 1-mV square pulse of 10 ms at 10 Hz. Once a high resistance seal (2–10 gigohms) was formed between the recording pipette and the cell, the access resistance was continuously monitored until it reached values <30 megohms. Capacitance was monitored before, during, and after each experiment. All voltage clamp protocols were generated and currents were recorded using an EPC-7 amplifier (List Electronic) and pClamp 5.51 acquisition system (Axon Instruments). The extracellular solution was exchanged continuously during recordings to add/washout drugs. Bath perfusion was performed by exchanging the content of the 18-mm recording chamber with extracellular solutions at a rate of 2 ml/min using a gravity flow system. Drugs were added in the extracellular solution.

Solutions for the electrophysiological studies.
For recording currents through voltage-gated calcium channels, the solutions were (in mmol/l) as follows: external: 20 CaCl₂, 110 NaCl, 5 CsCl, and 10 HEPES (pH 7.4 with CsOH); and internal: 10 CsCl, 90 CsSO₄, 10 EGTA, 1 CaCl₂, and 10 HEPES (pH 7.2 with CsOH). Currents were recorded from cells stepped for 200 ms from a holding potential of -60 mV to test potentials between -50 and 40 mV at 10-mV increments.

For recording KATP currents, the solutions were (in mmol/l) as follows: external: 140 KCl, 2 CaCl₂, 1.1 MgCl₂, and 10 HEPES (pH 7.4 with NaOH) and internal: 70 K₂SO₄, 10 KCl, 10 NaCl, 2 MgCl₂, and 10 HEPES (pH 7.2). Currents were evoked by stepping from a -60 mV holding potential to membrane potentials ranging from -130 mV to -50 mV in 10-mV increments for 200 ms.

Western blot analysis.
Membrane and cytosol fractions were prepared from HIT-T15 cells. Briefly, cells were harvested and homogenized with a Dounce-glass homogenizer in a buffer containing 10 mmol/l Tris (pH 7.5), 10% (vol/vol) glycerol, 1 mmol/l EDTA, 1 mmol/l dithiothreitol, 10 µg/µl aprotinin, 10 µg/µl leupeptin, 10 µg/µl pepstatin A, and 1 mmol/l phenylmethylsulfonyl fluoride (PMSF). Unbroken cells were removed by centrifugation at 1,500g for 10 min. After a 30-min centrifugation at 96,000g at 4°C, the supernatant was collected as the cytosol fraction and the pellet was solubilized in the above buffer supplemented with 1% Triton X-100 and used as the membrane fraction. Cytosol and membrane samples were mixed with 5x Laemmli sample buffer, boiled for 5 min, and resolved on a 14% gel by SDS-PAGE. After transfer to a polyvinylidene fluoride membrane and blocking in a 5% BSA solution, Western blot analysis was performed using a gustducin or transducin antibody and visualizing the immunoreactive bands by enhanced chemiluminescence (Amersham).

Trypsin digestion assay for activated transducin.
The method was similar to those described previously (11,26). Membranes of HIT-T15 cells (prepared as above) were resuspended in DEAE buffer (in mmol/l: 150 NaCl, 25 Tris, 1 MgCl₂, 5 DTT, and 1 CHAPS at pH 7.5 with 100 µmol/l GDP and 1 µmol/l GTPS) and then incubated with the heterotrimeric form of rod transducin GGDP*ß (a gift of Drs. Richard Cerione and Jon Erikson) plus denatonium and other taste substances for 1 h at 30°C. AlF₄^- used as a positive control was prepared using 30 µmol/l AlCl₃ and 10 mmol/l NaF. Subsequently, trypsin digestions (1:25 wt/wt trypsin to total protein) were performed for different periods of time (0, 30, 60, and 90 min) at room temperature and stopped by the addition of soybean trypsin inhibitor at a 10-fold excess over trypsin. Samples were boiled for 5 min in SDS sample buffer and subjected to SDS-PAGE and immunoblotting as described.

[³H]Glybenclamide binding assay.
HIT-T15 cells were collected from confluent 175-cm² flasks and washed twice with ice-cold PBS. The cells were scraped off into 10 ml of the PBS. The cells were pelleted and resuspended in 20 volumes of 5 mmol/l Tris-HCl (pH 8.0) at 4°C containing 0.1 mmol/l PMSF. They were incubated on ice for 40 min and then homogenized with 15–20 strokes in a Dounce glass homogenizer. The homogenate was centrifuged for 10 min at 900g, and the supernatant was collected and centrifuged at 96,000g for 30 min. The supernatant was discarded, and the pellets from the second centrifugation were immediately frozen in liquid nitrogen and stored at -80°C until used.

The method was similar to that used by Hu et al. (27). HIT-T15 cell membranes were suspended in 20 mmol/l MOPS (pH 7.4) containing 0.1 mmol/l PMSF and rehomogenized. The protein concentration of the membrane preparation was determined using the Biorad DC protein assay with BSA as the standard. The "competition" assay was performed with 1.0 nmol/l [³H]glybenclamide, denatonium over the concentration range of 0, 0.01–3 mmol/l, and membranes at a protein concentration of 100–200 µg/ml at room temperature for 2 h. Nonspecific binding was determined in the presence of 2 µmol/l unlabeled glybenclamide. The assay was performed in 50 mmol/l MOPS (pH 7.4), 0.1 mmol/l CaCl₂ (buffer A), in a total volume of 1 ml/tube. All assays were performed in duplicate. Binding was terminated by rapid filtration through Whatman GF/F 25-mm-diameter glass microfiber filters (presoaked in buffer A) followed by five washes with 5 ml of cold 0.1 mol/l NaCl. The filters were placed in 10 ml of Formula 989 scintillation fluid (Packard, Meriden, CT), and radioactivity was determined after overnight incubation by liquid scintillation spectrometry. Saturation binding assays were performed under the same conditions with [³H]glybenclamide at concentrations from 0.1 to 10 nmol/l in the presence or the absence of 1 mmol/l denatonium. Nonspecific binding was determined in the presence of 2 µmol/l unlabeled glybenclamide. K_d and Bmax values were determined by nonlinear curve fitting of the saturation data using Kaleidagraph (Synergy Software).

Reagents.
Denatonium benzoate, amphotericin B, glybenclamide, DIDS, and clonidine were obtained from Sigma (St. Louis, MO). ¹²⁵I-insulin, ³H-glybenclamide, and the ¹²⁵I-cyclic AMP radioimmunoassay kit were obtained from DuPont/NEN (Boston, MA). The rabbit polyclonal antigustducin and antitransducin antibodies were from Santa Cruz Biotech (Santa Cruz, CA).

Statistical analysis.
Results are presented as mean ± SE. Statistical analysis was by Student’s t test for paired and unpaired data as appropriate, except for the results in Fig. 9A, which was by one-way ANOVA with pairwise comparison by Dunnett’s method.

View larger version (20K): [in this window] [in a new window]   FIG. 9. The effects of denatonium on the binding of [3H]glybenclamide to HIT cell membranes. A: Effects of denatonium at concentrations from 0.01 to 3 mmol/l on the binding of 1 nmol/l [3H]glybenclamide. B: Amounts of [3H]glybenclamide bound (at concentrations from 0.3 to 11 nmol/l) in the absence and presence of 1 mmol/l denatonium. A Scatchard analysis of the data is shown in the inset.

	RESULTS

TOP ABSTRACT RESEARCH DESIGN AND METHODS RESULTS DISCUSSION REFERENCES

View larger version (35K): [in this window] [in a new window]   FIG. 1. Concentration-response characteristics for the effect of denatonium to stimulate insulin secretion in HIT-T15 cells. Denatonium was tested at concentrations from 0.03 to 3 mmol/l under conditions of static incubation (n = 4). *P < 0.01 vs. 4 mmol/l glucose.

View larger version (20K): [in this window] [in a new window]   FIG. 2. A: The effect of 1 mmol/l denatonium (solid circles) and lack of effect of 1 mmol/l sodium benzoate () in the presence of 4 mmol/l glucose on HIT-T15 cells under perifusion conditions. B: The effect of 10 µmol/l clonidine () on insulin secretion stimulated by 4 mmol/l glucose () and the effect of clonidine () on the insulin secretion stimulated by glucose plus 1 mmol/l denatonium (•). n = 4.

View larger version (32K): [in this window] [in a new window]   FIG. 3. Concentration-response characteristics for the effect of denatonium to stimulate insulin secretion by isolated rat pancreatic islets. Denatonium was tested at concentrations from 0.1 to 3 mmol/l under conditions of static incubation (n = 5). *P < 0.03; **P < 0.002; #P < 0.05 (all vs. 8.3 mmol/l glucose alone).

View larger version (23K): [in this window] [in a new window]   FIG. 4. The lack of effect of denatonium to stimulate insulin release in the absence of extracellular Ca2+ or in the presence of 10 µmol/l nitrendipine. The experiments were performed on HIT-T15 cells in the presence of 4 mmol/l glucose. Both glucose- and denatonium-stimulated release were inhibited by the absence of extracellular Ca2+ and by nitrendipine in the presence of extracellular Ca2+ (n = 4). *P < 0.02 vs. 4 mmol/l glucose; **P < 0.001 vs. 4 mmol/l glucose + denatonium.

Effect of denatonium on membrane potential.
Denatonium induced a glucose-dependent depolarization and increase in membrane potential fluctuations. Initial experiments examined changes in HIT-T15 cell membrane potential and electrical activity after exposure to denatonium at two different glucose concentrations. The resting potential of cells, measured after a 15- to 30-min exposure to 0.2 mmol/l glucose, was -62.2 ± 6 mV (n = 6). Application of 500 µmol/l denatonium caused a slight depolarization (-55.7 ± 5 mV, n = 5), which was rapidly reversible after washout of the denatonium. Generally, no action potentials developed after denatonium application to cells maintained in 0.2 mmol/l glucose. However, application of 500 µmol/l denatonium to cells maintained in 4 mmol/l glucose resulted in a substantial depolarization (13.5 ± 7 mV, n = 10) and an increase in membrane activity that was often accompanied by generation of multiple action potentials. The denatonium-stimulated depolarization and increase in electrical activity was quickly reversible after washing of the cell. Figure 5 shows membrane potential changes from a representative single cell exposed to denatonium at glucose concentrations of 0.2 and 4 mmol/l. Subsequent experiments were performed to investigate the underlying ionic mechanisms that contribute to the denatonium-induced glucose-dependent depolarization and increase in membrane activity in HIT-T15 cells.

View larger version (13K): [in this window] [in a new window]   FIG. 5. The effect of denatonium on the membrane potential of an isolated HIT-T15 cell using the whole-cell perforated-patch technique. The cell was equilibrated in 0.2 mmol/l glucose for 15 min, and 500 µmol/l denatonium was added to the bath for the period indicated by the rectangle. The cell was then exposed to 4 mmol/l glucose as indicated by the upper line and to denatonium for the period indicated by the rectangle.

View larger version (19K): [in this window] [in a new window]   FIG. 6. Voltage-gated calcium currents are not directly influenced by denatonium. A: Current-voltage relationship for peak currents from a representative HIT-T15 cell held at -60 mV and stepped from -50 mV to 50 mV at 10-mV increments. B: Time course showing activation of voltage-gated calcium current in a representative HIT-T15 cell held at -60 mV and stepped to 10 mV in control solution and after the application of 500 µmol/l denatonium. Solutions contained 4 mmol/l glucose.

View larger version (38K): [in this window] [in a new window]   FIG. 7. Denatonium reduces current flow through KATP channels in HIT-T15 cells. A: Effect of denatonium on KATP currents in the presence of 0.2 and 4 mmol/l glucose. B: Summary of the effect of denatonium on KATP currents in solutions containing 0.2 mmol/l (n = 5 cells), 4 mmol/l (n = 6 cells), or 10 mmol/l (n = 5 cells) glucose. Each point represents the peak current ± SD from cells held at -60 mV and stepped to -90 mV. *P = 0.03, **P < 0.01. C: Summary of the effect of denatonium on KATP currents in cells pretreated with 1 µmol/l glibenclamide. Solutions contained 0.2 mmol/l glucose (n = 5 cells). Each point represents the peak current from cells held at -60 mV and stepped to -90 mV ± SE. **P < 0.01.

View larger version (31K): [in this window] [in a new window]   FIG. 8. Denatonium treatment of HIT-T15 cell membranes fails to activate transducin (Gt) in the trypsin proteolysis assay. With the exception of lanes 1 and 2, the lanes represent incubations of HIT-T15 cell membranes with Gt for 1 h followed by a 60-min digestion with trypsin. Lane 1 is the marker lane with Gt alone (39 kDa). Lane 2 is the 0-min control (Gt reaction mixture for 1 h followed by the simultaneous addition of trypsin and soybean trypsin inhibitor). Lanes 3, 6, and 9 are 60-min control lanes without denatonium in the Gt reaction mixture). Lanes 4 and 7 tested the effect of 1 mmol/l denatonium, and lanes 5 and 8 tested the effect of 3 mmol/l denatonium. Lane 10 was the positive control with AlF4-. The proteolysis of the 39-kDa Gt (shown in lanes 1 and 2) to a 23-kDa band was similar for all of the conditions shown in lanes 3–9, i.e., both control and denatonium-treated conditions. In lane 10, in contrast, activation of Gt by AlF4- resulted in the almost complete proteolysis of Gt to the 32-kDa fragment.

The increased affinity of glybenclamide binding induced by denatonium demonstrates that denatonium does not interact with the glybenclamide binding site on the SUR. Possible mechanisms include that denatonium binds to a separate site on the SUR or that it interacts directly with the KATP channels in a similar manner to its interaction with K⁺ channels in taste cells. Furthermore, this interaction with Kir6.2 induces a conformational change in the SUR that is responsible for the increased affinity.

	DISCUSSION

TOP ABSTRACT RESEARCH DESIGN AND METHODS RESULTS DISCUSSION REFERENCES

 
The results of these studies demonstrate a concentration- and glucose-dependent stimulation of insulin secretion in response to denatonium that is due to depolarization of the ß-cell and increased Ca²⁺ entry. The action of denatonium to reduce KATP channel activity would be the primary cause of the depolarization. Denatonium does not seem to have any major effect on K⁺ channels other than the KATP channel or any direct effect on the L-type Ca²⁺ channel. However, part of the strong depolarizing effect of denatonium in the presence of glucose will be explained by an indirect action on L-type Ca²⁺ channels. Certainly, in the presence of denatonium, voltage-gated Ca²⁺ channels are activated as a result of the membrane depolarization caused by the inhibition of KATP channels. Therefore, the additional KATP channel-related depolarization observed in the presence of both denatonium and glucose would trigger the opening of more voltage-activated Ca²⁺ channels purely as a result of increased membrane depolarization. The inhibition of the KATP channel with no apparent effect on other K⁺ channels and the lack of a direct effect on voltage-gated Ca²⁺ channels in the ß-cell illustrate the selectivity and specificity of the effects of denatonium.

The detailed mechanism by which denatonium selectively affects the KATP channels remains to be understood. It is possible that denatonium interacts with the sulfonylurea receptor that is an essential component of the functional KATP channel (29,30). Denatonium has some structural similarity to nateglinide, meglitinide, and other recently introduced blood glucose–lowering compounds that also act on the KATP channel and that displace glybenclamide from its binding site on SUR (27,31–34). However, denatonium did not displace glybenclamide from its binding site on the SUR. Instead, it actually increased [³H]glybenclamide binding by an increase in affinity. Thus, the structural similarity of denatonium to nateglinide and meglitinide is not such that it can bind to the same site. Two possible explanations come to mind: (1) denatonium is binding to a site on the SUR that is distinct from the glybenclamide binding site but that still induces channel closure; and (2) denatonium acts directly on the channel, closes it, and induces a conformational change in SUR. Evidence in favor of this is that denatonium is known to act directly on K⁺ channels in taste cells, including delayed rectifier and Ca²⁺-activated K⁺ channels (17). In view of this, an action of denatonium on the Kir6.2 channel that affects the conformation of SUR would suffice to produce the observed effects. The nature of the observed glucose dependence for the action of denatonium to stimulate insulin secretion is not known but is similar to that seen with the sulfonylureas at low glucose concentrations, e.g., 2.8 mmol/l glucose for rat islets. The presence or absence of 5 mmol/l ATP in the binding assays did not influence the effect of denatonium to increase the K_d for glybenclamide (data not shown), indicating a lack of ATP dependence to the binding and to the conformational change required to affect the SUR. Also, denatonium inhibits the KATP current in the presence of low glucose concentrations. Thus, the glucose dependence lies at some point in stimulus-secretion coupling distal to membrane depolarization.

A question that arises from these observations is their relationship to the responses of taste receptor cells to denatonium and whether the signal transduction mechanisms are similar. An obvious similarity is that a K⁺ channel is involved, although denatonium has not been shown previously to affect a KATP channel in taste cells. This may be because the effects of bitter substances on KATP channels in taste cells have not been investigated yet, or more likely because this channel is not expressed in taste cells.

In summary, the ß-cell effects of denatonium are not mediated by T2R/TRB receptors and activation of either gustducin or transducin. The known effects of agonist-T2R/TRB interactions are gustducin -subunit activation of cyclic NPDE and ß-subunit activation of PLCß2, neither of which has been associated with inhibition of K⁺ channels. In this connection, there were no changes in intracellular cyclic AMP levels that might reflect increased cyclic NPDE activity or of any mobilization of intracellular Ca²⁺ to reflect PLC activation. Finally, using the trypsin digestion assay, we found no evidence that transducin was activated by denatonium that would have implicated an interaction with T2R/TRB receptors. In view of all of the available data, we conclude that denatonium most likely stimulates insulin secretion by an action on the Kir6.2 channel and depolarization of the ß-cell. This raises the possibility that structure-function studies on denatonium could give rise to other compounds with greater potency and specificity for the KATP channel that could be used orally as insulin secretagogues.

	ACKNOWLEDGMENTS

 
This work was supported by a Mentor-Based Postdoctoral Fellowship from the American Diabetes Association and grants DK-42063, DK-54243, and DK-59813 from the National Institutes of Health (to G.W.G.S.).

We are grateful to Dr. Rick Cerione and Dr. Jon Ericson for advice on gustducin/transducin signaling pathways and the trypsin-digestion assay; Dr. Leslie Satin for advice on the electrophysiological studies; and Dr. Troitza Bratanova-Tochkova for tissue culture of the clonal HIT-T15 cells.

	FOOTNOTES

 
Address correspondence and reprint requests to Dr. Geoffrey W.G. Sharp, Department of Molecular Medicine, College of Veterinary Medicine, Cornell University, Ithaca, NY 14853-6401. E-mail: gws2{at}cornell.edu.

Received for publication 8 April 2002 and accepted in revised form 17 October 2002.

PMSF, phenylmethylsulfonyl fluoride; SUR, sulfonylurea receptor.

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