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J. Biol. Chem., Vol. 283, Issue 15, 10184-10197, April 11, 2008

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Investigation of Transport Mechanisms and Regulation of Intracellular Zn²⁺ in Pancreatic -Cells^*

Armen V. Gyulkhandanyan, Hongfang Lu, Simon C. Lee, Alpana Bhattacharjee, Nadeeja Wijesekara¹, Jocelyn E. Manning Fox, Patrick E. MacDonald, Fabrice Chimienti^¶, Feihan F. Dai, and Michael B. Wheeler²

From the Department of Physiology, University of Toronto, Toronto, Ontario M5S 1A8, Canada, the Department of Pharmacology, University of Alberta, Edmonton, Alberta T6G 2E1, Canada, and ^¶Mellitech, DRMFC/SC1B, CEA Grenoble, 38054 Grenoble, France

Received for publication, August 21, 2007 , and in revised form, January 31, 2008.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
During insulin secretion, pancreatic -cells are exposed to Zn²⁺ released from insulin-containing secretory granules. Although maintenance of Zn²⁺ homeostasis is critical for cell survival and glucagon secretion, very little is known about Zn²⁺-transporting pathways and the regulation of Zn²⁺ in -cells. To examine the effect of Zn²⁺ on glucagon secretion and possible mechanisms controlling the intracellular Zn²⁺ level ([Zn²⁺]_i), we employed a glucagon-producing cell line (-TC6) and mouse islets where non-β-cells were identified using islets expressing green fluorescent protein exclusively in β-cells. In this study, we first confirmed that Zn²⁺ treatment resulted in the inhibition of glucagon secretion in -TC6 cells and mouse islets in vitro. The inhibition of secretion was not likely via activation of K_ATP channels by Zn²⁺. We then determined that Zn²⁺ was transported into -cells and was able to accumulate under both low and high glucose conditions, as well as upon depolarization of cells with KCl. The nonselective Ca²⁺ channel blocker Gd³⁺ partially inhibited Zn²⁺ influx in -TC cells, whereas the L-type voltage-gated Ca²⁺ channel inhibitor nitrendipine failed to block Zn²⁺ accumulation. To investigate Zn²⁺ transport further, we profiled -cells for Zn²⁺ transporter transcripts from the two families that work in opposite directions, SLC39 (ZIP, Zrt/Irt-like protein) and SLC30 (ZnT, Zn²⁺ transporter). We observed that Zip1, Zip10, and Zip14 were the most abundantly expressed Zips and ZnT4, ZnT5, and ZnT8 the dominant ZnTs. Because the redox state of cells is also a major regulator of [Zn²⁺]_i, we examined the effects of oxidizing agents on Zn²⁺ mobilization within -cells. 2,2'-Dithiodipyridine (-SH group oxidant), menadione (superoxide generator), and SIN-1 (3-morpholinosydnonimine) (peroxynitrite generator) all increased [Zn²⁺]_i in -cells. Together these results demonstrate that Zn²⁺ inhibits glucagon secretion, and it is transported into -cells in part through Ca²⁺ channels. Zn²⁺ transporters and the redox state also modulate [Zn²⁺]_i.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Blood glucose homeostasis is maintained by appropriate secretion of insulin and glucagon from pancreatic islet β- and -cells, respectively (1, 2). It is quite clear that when ambient glucose concentrations rise, glucagon secretion is inhibited and insulin secretion is stimulated. The mechanisms that control the suppression of glucagon secretion under high glucose conditions have not been completely agreed upon, but it is likely that -cells sense and respond to changes in blood glucose through direct and indirect mechanisms (3-6). To date the most likely candidates are glucose itself, paracrine and endocrine factors, and neuronal modulation. Some of the paracrine/endocrine factors that could facilitate the suppression of glucagon secretion include insulin and other factors released upon β-cell exocytosis, including -aminobutyric acid and Zn²⁺ (3, 7, 8).

In pancreatic β-cells a fraction of the intracellular Zn²⁺ pool is stored with insulin in intracellular vesicles as a complex of zinc-insulin (9). The concentration of Zn²⁺ in these vesicles is about 20 mM (10, 11). Following exocytosis of the intracellular vesicles, it is likely that the Zn²⁺ released into the extracellular islet space would be transported back into the host cell or into neighboring cells. The effects of β-cell secretory products, Zn²⁺ and insulin, on glucagon secretion are still controversial and may be species-specific. It was shown that Zn²⁺ had inhibitory actions on glucagon secretion from rat islets (8, 12, 13). Recently it was found that switching off pancreatic artery infusions of Zn²⁺ stimulated glucagon secretion in rats (14). Contrary to these experiments, a lack of Zn²⁺ effect on glucagon secretion was observed in mouse islets (15).

In mammalian cells the uptake of Zn²⁺ and its outward transport are regulated by two families of membrane proteins that work in opposite directions. The influx of Zn²⁺ is facilitated by ZIP³ (SLC39) proteins (16-20) and by different types of Ca²⁺ channels: dihydropyridine-sensitive Ca²⁺ channels in heart cells (21) and the N-methyl-D-aspartic acid and -amino-3-hydroxy-5-methyl-4-isoxazole propionate/kainate-activated Ca²⁺-permeable channels in neurons (22, 23). The efflux of Zn²⁺ or intracellular Zn²⁺ sequestration is promoted by CDF/ZnT (SLC30) family of transporters, which work in opposition to the ZIP transporters (16-20). In addition, a Na⁺/Zn²⁺ exchange mechanism that is different from both ZnT1 and Na⁺/Ca²⁺ exchanger utilizes the transmembrane electro-chemical Na⁺ gradient and mediates extrusion of Zn²⁺ in some cells (24). A Zn²⁺/H⁺ antiport mechanism, associated with intracellular ZnTs, could be responsible for Zn²⁺ accumulation in intracellular organelles (25). It is likely that ZnT proteins serve as secondary active transporters, using the gradient of other ions to drive the Zn²⁺ transport (20).

Recently a genome-wide association study identified novel risk loci for type 2 diabetes mellitus (26-28). These loci include a nonsynonymous polymorphism in the zinc transporter SLC30A8, which is expressed in insulin-producing β-cells (29). Based upon these recent linkage studies, the mechanisms of Zn²⁺ transport as well as the modulation of intracellular Zn²⁺ in islet cells takes on an important clinical significance. Despite the importance of Zn²⁺ in islet of Langerhans function, a limited number of articles are dedicated to Zn²⁺ transport (30-33). It was demonstrated that Zn²⁺ accumulation into insulin-secreting β-cells occurred by the following two pathways: through membrane Zn²⁺ transporters under low glucose conditions, and through L-type voltage-gated Ca²⁺ channels (VGCC) under high glucose (32). To date, Zn²⁺ transport mechanisms have not been studied in other islet cell types, including -cells.

Taking into account the Zn²⁺-rich environment surrounding -cells, it is expected that they should have an effective mechanism for maintaining Zn²⁺ homeostasis. Dysfunction of the transporters responsible for Zn²⁺ export can lead to dramatic changes in cell viability, because an excess of Zn²⁺ can induce oxidative damage, mitochondrial depolarization, and opening of mitochondrial permeability transition pores (34-36). In addition, as stated above, Zn²⁺ released from β-cells may be a required component of the "off switch" for glucagon secretion. Thus the inhibition of glucagon secretion may require Zn²⁺ transport and the modulation of the cytoplasmic Zn²⁺ level ([Zn²⁺]_i). -Cells are electrically active and equipped with different types of channels, including K_ATP channels and four types of VGCC, which could be involved in regulating Zn²⁺ as in β-cells (3, 37). In addition, -cells may also modulate [Zn²⁺]_i through ZnTs and Zips. Finally, it is well known that free Zn²⁺ in cells is modulated by redox state and the level of reactive oxygen species in cells (38, 39), which in turn are regulated by glucose and fatty acid metabolism and β-oxidation. Thus the redox state of the -cell may also regulate -cell [Zn²⁺]_i.

In this study we showed an inhibitory effect of Zn²⁺ on glucagon secretion in a glucagon-producing -cell line (-TC). Using the Zn²⁺-selective dye FluoZin-3 AM (32, 40) we studied, for the first time, Zn²⁺ transport in intact mouse pancreatic islet -cells, as well as in the -TC cell line, and showed the possibility of Zn²⁺ transport through both Ca²⁺ channels and Zn²⁺ influx transporter(s). We demonstrated the expression of ZIP and ZnT family of Zn²⁺ transporters in -TC cells, as well as in isolated mouse islets. In addition, we investigated the regulation of [Zn²⁺]_i by oxidizing agents in -cells. Our results suggest that cytoplasmic Zn²⁺ homeostasis in -cells is maintained by a concerted action of Zn²⁺ transporters, Ca²⁺ channels, and the redox state of thiol groups.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Reagents—Fluorescent dyes FluoZin-3 AM, bis-(1,3-dibutylbarbituric acid) trimethine oxonol (DiBAC₄(3)), rhodamine 123 (Rh123), MitoTracker Red CMXRos, Hoechst 33342, 3-morpholinosydnonimine (SIN-1) were obtained from Molecular Probes (Eugene, OR). Dispase II (neutral protease) was from Roche Diagnostics. Carbonyl cyanide p-(trifluorometoxy)phenylhydrazone (FCCP), 2,2'-dithiodipyridine (DTDP), 2-mercaptopyridine N-oxide (pyrithione), N,N,N',N',-tetrakis-(2-pyridylmethyl)-ethylenediamine (TPEN), nitrendipine, tolbutamide, tetraethylammonium, N-ethylmaleimide (NEM), iodoacetamide, dithioerythritol (DTE), azide, menadione, poly-L-lysine, annexin V-FITC, and propidium iodide were from Sigma. 4-Hydroxynonenal (HNE) was from Cayman Chemical Co. ZnSO₄ was used in all experiments.

-TC6 Cells—-TC6 (-TC) glucagon-producing cell line was kindly provided by Dr. Y. Moriyama, Okayama University, Japan. Cells were cultured in Dulbecco's modified Eagle's medium with 25 mM glucose, L-glutamine, supplemented with 10% bovine serum, 100 units/ml penicillin, 100 µg/ml streptomycin, at 37 °C and 5% CO₂, 95% air. Culture medium was changed every 48 h, and cells were grown in monolayer to 80-90% confluence. The cells were plated onto glass coverslips for fluorescence measurements. Experiments with cells were performed typically 2-3 days after plating.

Dispersed Islet Cells from Mouse Insulin Promoter-Green Fluorescent Protein (MIP-GFP) Mice—To identify non-β-cells we used MIP-GFP mice (CD1 background), in which GFP is specifically expressed in the islet β-cells. MIP-GFP mice were a gift from Dr. M. Hara, University of Chicago, Chicago, IL (41). Islets were isolated as described previously (32). To obtain dispersed cells, isolated islets were incubated for 10 min in Ca²⁺-free phosphate-buffered solution supplemented with 2 mM EGTA, 3 mM glucose, 100 units/ml penicillin, and 100 µg/ml streptomycin. Islets then were centrifuged and incubated with dispase II, followed by addition of RPMI 1640 medium with 11.1 mM glucose, 10% bovine serum, 100 units/ml penicillin, 100 µg/ml streptomycin, and 10 mM HEPES, pH 7.4. The suspension was centrifuged, and the pellet was resuspended in the same medium. The cells were plated on 22-mm glass coverslips coated with poly-L-lysine and maintained for 1-3 days at 37 °C and 5%CO₂, 95% air. We selected GFP-negative (non-green) cells for experiments.

Glucagon Secretion Assay—-TC cells were plated onto 24-well plates and cultured for 48 h in Dulbecco's modified Eagle's medium with L-glutamine supplemented with serum, streptomycin, and penicillin at 37 °C. Culture medium was aspirated, and cells were preincubated for 15 min with buffer containing (mM) 130 NaCl, 5 KCl, 2 CaCl₂, 1 MgCl₂, 5 NaHCO₃, 10 HEPES, pH 7.4, supplemented with 20 mM glucose. Buffer was then aspirated, and cells were incubated in 0.5 ml of buffer with 1 or 20 mM glucose with or without different concentrations of Zn²⁺ for 1 h at 37 °C. After incubation, 450 µl of medium was collected and centrifuged at 2000 rpm, and supernatant was assayed for glucagon using glucagon radioimmuno-assay kit (Linco Research) according to the manufacturer's instructions. The DNA amount in samples was determined spectrophotometrically for normalization. The procedure for mouse (FVB background) islet glucagon secretion assay was similar.

Fluorescent Measurements—During fluorescent measurements we used the incubation and perfusion buffer containing (mM) 130 NaCl, 5 KCl, 2 CaCl₂, 1 MgCl₂, 5 NaHCO₃, 10 HEPES, pH 7.4, or the same medium without Ca²⁺. In selected experiments, to depolarize the cells, 30 or 50 mM NaCl was replaced with 30 or 50 mM KCl. Fluorescent experiments were carried out using an Olympus BX51W1 fluorescent microscope fitted with a 20x/0.95 water immersion objective and cooled CCD camera equipped with a magnification changer (U-TVCAC, Olympus). For excitation, a xenon lamp-based DeltaRam high speed monochromator from Photon Technology International (PTI, Lawrenceville, NJ) was used. For control of the monochromator and video camera, as well as for analysis of fluorescent imaging, ImageMaster 3 software (PTI) was used as we have described previously (32). The cells were transferred to an open chamber on the microscope stage for imaging and perfused at 1 ml/min. All time-dependent experiments were performed at 36-37 °C using TC-324B Heater Controller (Warner Instruments, Hamden, CT). In static experiments a Delta T Culture Dish Controller (Bioptechs, Butler, PA) was used for heating. One to 30 randomly chosen cells were analyzed per coverslip.

Dynamic Measurements of Zn²⁺, Ca²⁺, Plasma Membrane (_P), and Mitochondrial Membrane Potential (_m)—For Zn²⁺ measurements, coverslips with -TC cells, or dispersed islet cells, were loaded with 2 µM FluoZin-3 AM for 50 min in incubation buffer in the presence of 2 mM glucose at 37 °C and 5% CO₂, 95% air. Then coverslips with -TC cells or islet cells were subsequently incubated for 10 min in the same buffer without dye. FluoZin-3 AM was excited at 480 nm, and emission was measured with a 525-nm band pass filter using 505 nm beam splitter. Changes in intracellular Ca²⁺ concentrations were assessed using Fura-2 AM. Coverslips with cells were loaded with 2.5 µM Fura-2 AM for 45 min in incubation buffer under the same conditions as with FluoZin-3 AM. Cells were excited by dual excitation at 340/380 nm, and emission was detected by a 510-nm band pass filter using 415-nm beam splitter. Cell _P was measured by negatively charged oxonol dye DiBAC₄(3). Coverslips with -TC cells or islet cells were loaded with 100 nM dye for 20 min in incubation buffer. The perfusion solution contained the same concentration of oxonol dye. The excitation and emission wavelengths were similar to FluoZin-3 AM. Mitochondrial _m was measured by positively charged dye Rh123 with excitation and emission wavelengths similar to FluoZin-3 AM. Cells were loaded with 5 µM Rh123 for 45 min. To minimize possible photodamaging effects, the shutter on the monochromator was closed for 3 s between each acquisition.

Measurements of Cells Simultaneously Stained with FluoZin-3 AM, MitoTracker Red CMXRos, and Hoechst 33342—-TC cells were loaded with 2 µM FluoZin-3 AM for 30 min in incubation buffer in the presence of 2 mM glucose at 37 °C and 5% CO₂, 95% air. Then 1 µM of mitochondria-staining dye MitoTracker Red and 2 µM of DNA-specific dye Hoechst 33342 were added, and cells were incubated for 20 min. Cells were washed in incubation buffer, transferred to the chamber on the microscope stage, and maintained at 36-37 °C. Excitation wavelengths for MitoTracker Red and Hoechst 33342 were 540 and 350 nm, respectively, and emission was measured using 660-nm band pass filter and 550-nm beam splitter for Mito-Tracker Red and 465-nm band pass filter and 400-nm beam splitter for Hoechst 33342.

Confocal Fluorescent Measurements—Confocal imaging was performed using a Zeiss LSM510 laser scanning microscope. Coverslips with -TC cells co-loaded with FluoZin-3 AM (2 µM) and MitoTracker Red (500 nM) or FluoZin-3 AM and endoplasmic reticulum (ER)-staining dye FM 4-64 (2 µM) were transferred into the chamber on the stage of the microscope fitted with a 40 x 0.75 water immersion objective. For FluoZin-3 AM and MitoTracker Red, as well as for FluoZin-3 AM and FM 4-64 excitations, the 488- and 514-nm argon laser lines were used, and emissions were acquired using FITC and TRITC set of filters, respectively. The images were analyzed using Zeiss LSM Image software.

Measurements of NAD(P)H Fluorescence—Changes in NAD(P)H redox state of -TC were observed at perfusion of cells with incubation buffer in the presence of 1 mM glucose at 37 °C. The excitation wavelength for NAD(P)H autofluorescence was 360 nm, and emission was measured using 465-nm band pass filter and 400-nm beam splitter. To assess the maximal signal (maximal NAD(P)H reduction) in cells, the mitochondrial electron transport chain was inhibited by azide.

Assessment of -TC Cells Apoptosis and Necrosis—For apoptosis and necrosis measurements, cells plated on glass coverslips were incubated at 37 °C during 1 h with different concentrations of Zn²⁺ in incubation buffer containing 1 mM glucose. Then coverslips were simultaneously incubated with annexin V-FITC (0.4 µg/ml) and propidium iodide (PI, 1 µM) for 20 min at room temperature in darkness in incubation buffer with 1 mM glucose. Coverslips were washed in the same buffer, transferred to an open chamber on the microscope stage, and maintained at 36-37 °C. The fluorescence of annexin V-FITC was excited at 480 nm and emission measured with 525-nm band pass filter using a 505-nm beam splitter. The PI fluorescence was excited at 540 nm and emission measured with a 660-nm band pass filter using a 550-nm beam splitter. Percent of apoptosis and necrosis was calculated as the ratio of annexin V or PI-positive cells to all cells in the field of view. For these experiments 400-500 cells in the field of view were analyzed per coverslip.

ZIP and ZnT mRNA Expression Analysis in -TC6 Cells—Total RNA was isolated from -TC cells, mouse islets (FVB background), and whole mouse brain using TRIzol Reagent (Invitrogen) according to the manufacturer's instruction. The extracted total RNA was treated with rDNase I (Ambion, Houston, TX). One µg of the isolated RNA was reverse-transcribed using Moloney murine leukemia virus reverse transcriptase according to the manufacturer's instructions (Invitrogen). The resulting cDNA was used for amplification in quantitative real time PCR (qPCR). qPCR was performed as described previously (32). Primers were designed using Primer Express version 2.0 software (Applied Biosystems, Foster City, CA). Primer sequences are indicated in supplemental Table 1. 10 ng of -TC6 cDNA per well was used as the template for amplification. The real time PCR protocol employed was as follows: heat activation of polymerase at 95 °C for 3 min, followed by 40 cycles of 95 °C for 10 s, 65 °C for 15 s, and 72 °C for 20 s. Readings were carried out on an ABI Prism® 7900HT Sequence Detection System (Applied Biosystems) and compared against a standard curve created from mouse genomic DNA by serial dilutions. Data were normalized to mouse β-actin mRNA.

View larger version (26K): [in this window] [in a new window]   FIGURE 1. Effect of Zn2+ on glucagon secretion in -TC cells and mouse islets. A, -TC cells were incubated for 1 h with 1 or 20 mM glucose with or without 5, 10, or 20 µM Zn2+, and glucagon secretion was measured. B, islets were incubated for 1 h with 1 mM glucose with or without 20 µM Zn2+, and glucagon secretion was measured. Data were averaged from triplicates in three independent experiments. *, p < 0.05; **, p < 0.01; ***, p < 0.005; error bars represent S.E.

Electrophysiology—K_ATP current was measured in the whole-cell patch clamp configuration during 200-ms voltage excursions of ±10 mV from a holding potential of -70mV. Mean outward current at each potential was measured during the last 50 ms of depolarization. The bath solution contained (mM) NaCl 135, KCl 5.4, CaCl₂ 1, MgCl₂ 1.2, glucose 5, HEPES 10, pH 7.3, with NaOH, with 10 µM ZnSO₄ added as indicated. The pipette solution contained (mM) KCl 140, MgCl₂ 1, HEPES 10, MgATP 0.3, pH 7.3, with KOH. Data were acquired using an EPC 10 amplifier (HEKA Instruments, Southboro, MA) and analyzed using Pulsefit software (HEKA Instruments, Southboro, MA).

Statistics—All experiments were repeated three or more times in -TC cells or dispersed islet cells, and typical results are presented. The raw data were processed using PSI-PLOT. Student's t test was used for determination of statistical significance. p values less than 0.05 were considered statistically significant.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Effect of Zn²⁺ on Glucagon Secretion—Pancreatic -cells are exposed to a large amount of Zn²⁺ when insulin is secreted from β-cells. To test the effect of Zn²⁺ on glucagon secretion in -TC cells, we incubated -TC cells with 1 or 20 mM glucose in the presence or absence of 5, 10, and 20 µM Zn²⁺ (see "Materials and Methods"). As seen in Fig. 1A, at low glucose, glucagon secretion was decreased by 26.2% (p < 0.05, n = 9), 50.3% (p < 0.005, n = 9), and by 29.4% (p < 0.01, n = 9) with 5, 10, and 20 µM Zn²⁺, respectively. At high glucose conditions, glucagon secretion was reduced by 45.5% (p < 0.01, n = 9), by 58.3% (p < 0.005, n = 9), and by 36.6% (p < 0.05, n = 9) with 5, 10, and 20 µM Zn²⁺, respectively. Glucagon secretion at 1 mM glucose was 32% higher than at 20 mM glucose (p < 0.01, n = 9) (Fig. 1A). Similar changes were observed in isolated mouse islets (Fig. 1B). This result is in agreement with the role of glucose as one of the physiological inhibitors of glucagon secretion. As seen in Fig. 1 in -TC cells, the inhibitory effect of 10 µM Zn²⁺ on glucagon secretion is greater than that of 20 µM Zn²⁺. Experiments using an expanded concentration range showed that a further increase in Zn²⁺ concentration led to a further decline of the inhibition (supplemental Fig. 1A). To investigate the possible effect of Zn²⁺ treatment on apoptosis or necrosis, we incubated -TC cells for 1 h with buffer supplemented with 1 mM glucose in the presence of various Zn²⁺ concentrations and measured cell viability (supplemental Fig. 1B). The analysis of fluorescence intensity of annexin V and PI showed that compared with control, treatment with 10 µM Zn²⁺ had no effect on the necrotic processes, whereas at 20, 50, and 100 µM Zn²⁺ necrosis was significantly increased (supplemental Fig. 1B). None of the Zn²⁺ concentrations used had any significant effect on the degree of apoptosis (not shown). Based on these results, it is likely that the mechanism underlying the -TC secretory response to high concentrations of Zn²⁺ involves necrosis. We hypothesize that the increased glucagon secretion (compared with 10 µM Zn²⁺) in the presence of 20, 50, or 100 µM Zn²⁺ could be explained by an interplay between a direct inhibitory effect of Zn²⁺ and injury/partial destruction of cells resulting in the release of intracellular glucagon.

Fluorescent Visualization of Zn²⁺ Accumulation in -TC Cells—Our previous results showed the partial co-localization of FluoZin-3 and a mitochondrial marker in β-cells (32). In this study -TC cells stained simultaneously with FluoZin-3 AM and the mitochondrial marker dye MitoTracker Red (Fig. 2A, upper panel) or FluoZin-3 AM and the ER indicator FM 4-64 (Fig. 2A, bottom panel) were imaged using confocal microscopy. Experiments showed nonhomogeneous fluorescence of FluoZin-3 in the cell interior with insignificant nuclear fluorescence and prominent location of mitochondria or ER in the perinuclear region. Partial co-localization of FluoZin-3 and MitoTracker Red or FluoZin-3 and FM 4-64 was observed. Punctate Zn²⁺ staining and significant co-localization suggest that the distinct amount of extractable Zn²⁺ is present not only in the cytoplasm but also in other cellular compartments like mitochondria and ER.

View larger version (55K): [in this window] [in a new window]   FIGURE 2. Fluorescent Zn2+ visualization in -TC cells. A, detection of Zn2+ in -TC cell mitochondria (panels i) and ER (panels ii) using confocal microscopy. Coverslips with cells were simultaneously loaded with 2 µM FluoZin-3 AM and 500 nM MitoTracker Red (panels i) or with 2 µM FluoZin-3 AM and 2 µM FM4-64 (panels ii) (see "Materials and Methods") and then washed twice without dyes and transferred to the open chamber with incubation medium with 1 mM glucose at room temperature. B and C, Zn2+ detection in -TC cells at low (B) and high (C) glucose conditions using conventional microscopy. Coverslips with cells were simultaneously loaded with 3 µM FluoZin-3 AM, 1 µM MitoTracker Red, and 2 µM Hoechst 33342, then washed twice without dyes, and transferred to the open chamber with incubation medium at 37 °C. B, corresponding DIC, fluorescent and merged images at 1 mM glucose concentration before and after 15 min of exposure to 10 µM Zn2+. C, corresponding DIC, fluorescent and merged images at 20 mM glucose concentrations before and after 15 min of exposure to 10 µM Zn2+. The corresponding line profiles of FluoZin-3 fluorescence intensity along the line drawn in DIC images at 1 mM (B) and 20 mM (C) glucose conditions before and after 10 µM Zn2+ addition are shown in the right panels. Fluorescence was normalized to background (100%).

Time-dependent Kinetics of Zn²⁺ Transport in -TC and Dispersed Islet MIP-GFP-negative Cells at Low and High Glucose Conditions and under KCl Depolarization—To examine the dynamics of Zn²⁺ influx, a perfusion system was employed using 5-10 µM Zn²⁺. Fig. 3A shows the kinetics of FluoZin-3 fluorescence was almost linear (at least during the 25 min), reflecting the linearity of increase in [Zn²⁺]_i. Treatment with the artificial Zn²⁺ ionophore pyrithione sharply increased the rate of Zn²⁺ entry (Fig. 3A). The subsequent application of the cell-permeable highly selective Zn²⁺ chelator TPEN, which has much higher affinity for Zn²⁺ than for Ca²⁺ or Mg²⁺ (49, 50), resulted in the decline of fluorescence (decrease of[Zn²⁺]_i) to initial levels.

In insulin-secreting β-cells, plasma membrane potential is tightly regulated by K_ATP channel activity, and an increased glucose concentration results in closure of the K_ATP channel and depolarization of cells. -Cells are also electrically active, but unlike β-cells an increase in glucose concentrations leads to a more hyperpolarized state of -cell plasma membrane (51-53). To check the effect of glucose on the rate of Zn²⁺ accumulation, we perfused -TC cells with low followed by high glucose concentrations (Fig. 3B). As seen in Fig. 3B, perfusion with Zn²⁺ at 1 mM glucose resulted in an increase in FluoZin-3 fluorescence. No significant change in the rate of Zn²⁺ accumulation was observed when cells were challenged with 20 mM glucose after low glucose perfusion. Thus in -cells the change in glucose concentration from low to high did not have any significant effect on the rate of Zn²⁺ accumulation. Switching from high to low glucose (Fig. 3C) also had little effect on the rate of Zn²⁺ influx.

Depolarization of -TC cells with KCl resulted in an increase of the Zn²⁺ accumulation rate. Fig. 3D and pie chart i demonstrated the effect of KCl on the rate of Zn²⁺ influx following low glucose treatment. In -TC cells (75 cells in four independent experiments), the increase in the rate of Zn²⁺ accumulation because of perfusion with KCl was observed in 57 cells (76%). In 14 cells (18.7%) KCl did not change the rate of Zn²⁺ accumulation, and in 4 cells (5.3%) treatment led to a decrease. As seen in Fig. 3B, treatment with KCl following high glucose also resulted in increased Zn²⁺ influx in -TC cells. In 50 investigated cells (four independent experiments), we observed an increased rate of Zn²⁺ accumulation in 32 cells (64%), no effect in 14 cells (28%), and a decrease in 4 cells (8%). Fig. 3B, pie chart ii, illustrates these data.

To determine whether the conditions used above resulted in changes of _P in -cell, we used the potential-sensitive anionic dye DiBAC₄(3). The initial level of DiBAC₄(3) fluorescence was monitored at low glucose conditions, and stimulation with high glucose led to some decrease in DiBAC₄(3) fluorescence (hyperpolarization) (Fig. 3E). Further perfusion with 30 mM KCl resulted in a fluorescence increase, reflecting depolarization of these cells (Fig. 3E). It is of note that fast increases in intracellular Zn²⁺ because of treatment with Zn²⁺/pyrithione also caused depolarization of both the plasma (Fig. 3F) and mitochondrial membrane (Fig. 3G). Perfusion with FCCP caused more complete depolarization of mitochondria (Fig. 3G).

Additionally, we performed a similar study in primary -cells to look at the Zn²⁺ transporting characteristics. It is known that pancreatic islets contain four types of cells (-, β-, -, and PP-cells), and the dominant are β-, -, and -cells with following distribution in rodents: β-cells, 75-80%; -cells, 10-18%, and -cells, 5-10% (54-58). We used dispersed islet cells isolated from MIP-GFP mice, where β-cells are GFP-positive. In our study the smallest GFP-negative dispersed islet cells in each field of view were chosen for fluorescent measurements, and the responses of these cells were evaluated. We took into account that two types (- and -cells) of GFP-negative cells are possible and that the mean diameter of -cells (10.6 µm) is smaller than -cells (11.8 µm) (59, 60). Fig. 4 shows representative experiments reflecting the rate of Zn²⁺ accumulation in primary -cells both at low and high glucose conditions, as well as at treatment with KCl. A depolarizing concentration of KCl changed the rate of Zn²⁺ accumulation in most cells. In 17 investigated GFP-negative cells (five independent experiments) treated with KCl following low glucose incubation, we observed an increased rate of Zn²⁺ accumulation in 11 cells (64.7%), no effect in 5 cells (29.4%), and decreased Zn²⁺ influx in 1 cell (5.9%) (Fig. 4A). In 12 investigated GFP-negative cells (three independent experiments) treated with KCl following high glucose, increased rate of Zn²⁺ accumulation was observed in 10 cells (83.3%), no effect in 1 cell (8.3%), and decreased Zn²⁺ influx in 1 cell (8.3%) (Fig. 4B). Pie charts in Fig. 4 indicate the percentage of cells responding to KCl. Similar to our observations in -TC cells, switching from high to low glucose did not change the rate of Zn²⁺ accumulation (Fig. 4C). Because the mean diameter of - and -cells is distinguished only by a 10% difference, one is never completely sure that all traces presented in Fig. 4 belong to -cells. However, taking into account that the average abundance of -cells (10-18%) is significantly higher than that of -cells (5-10%), we can assume that the majority of traces presented in Fig. 4 reflect the -cell responses.

View larger version (27K): [in this window] [in a new window]   FIGURE 3. Time-dependent kinetics of Zn2+ accumulation, plasma membrane (p), and mitochondrial membrane (m) potential changes in -TC cells. A, kinetics of Zn2+ uptake in -TC cells. The averaged trace ± S.E. of FluoZin-3 AM loaded cells from randomly chosen separately delineated cells (n = 17) of the same experiment is presented. Inset shows all traces. As a control Zn2+ ionophore pyrithione and a membrane-permeable Zn2+ chelator TPEN were added. As a base line (100%) here and in subsequent figures, the mean level of fluorescence during the initial 100 s was taken. In right panels the corresponding DIC and fluorescent images of cells at different times of the experiment are shown. B, kinetics of Zn2+ transport in -TC cells after switching from low (1 mM) to high (20 mM) glucose. C, kinetics of Zn2+ transport in -TC cells after switching from high (20 mM) to low (1 mM) glucose. D, kinetics of Zn2+ transport in -TC cells after depolarization with KCl followed by low glucose perfusion. Dashed lines show the slopes of FluoZin-3 fluorescence changes, which reflect the relative rate of Zn2+ accumulation in cells. Pie charts show the percent of -TC cells that responded to KCl treatment followed by low (panel i) or high (panel ii) glucose perfusion (see text). E, changes of p in -TC cells at low and high glucose conditions. At the end of the experiment KCl was added for complete plasma membrane depolarization. F, effect of pyrithione-induced fast Zn2+ influx into -TC cells on p. G, effect of pyrithione-induced fast Zn2+ influx into -TC cells on m. At the end of experiment FCCP was added for mitochondrial depolarization. The averaged traces ± S.E. of n = 16 (B), 12 (C), 21 (D), 12 (E), 19 (F), and 15 (G) cells are shown. In all cases the traces are representative of at least three independent experiments. Bars indicated perfusion with corresponding reagents.

View larger version (26K): [in this window] [in a new window]   FIGURE 4. Time-dependent kinetics of Zn2+ accumulation in MIP GFP-negative dispersed mouse islet cells at low and high glucose conditions. Upper panels show DIC and fluorescent images of dispersed MIP GFP islet cells loaded with 3 µM FluoZin-3 AM at time 0. The arrows in DIC images show GFP-negative (non-green) cells. A, Zn2+ uptake by dispersed islet cells during the perfusion with 1 mM glucose followed by 30 mM KCl. B, Zn2+ uptake by dispersed islet cells during the perfusion with 10 mM glucose followed by 30 mM KCl. Pie charts show the corresponding percent responses of MIP GFP-negative cells to KCl perfusion following low or high glucose. C, Zn2+ uptake by dispersed islet cells during the perfusion 10 mM glucose followed by 1 mM glucose. In all cases the traces are representative of 3-5 independent experiments.

Effect of Zn²⁺ on K_ATP Currents in -Cells—To determine whether Zn²⁺ was inhibiting glucagon secretion via an effect on the K_ATP channel, we examined whether extracellularly applied Zn²⁺ had any effect on K_ATP channel current density in mouse islet -cells. Because capacitance of cells is proportional to the size of cells, -cells were identified by their small size (capacitance <4 pF) and the presence of Na⁺ current when depolarized from a holding potential of -70 mV (6). K_ATP current measured in these cells was very small under the recording conditions applied, and 10 µM Zn²⁺ was found to have no significant effect on the current density (Fig. 7). As was mentioned above, in these experiments we anticipate that the majority of cells recorded would be -cells.

View larger version (32K): [in this window] [in a new window]   FIGURE 5. Effects of nonselective Ca2+ channel inhibitor Gd3+ and L-VGCC channel inhibitor nitrendipine on Zn2+ accumulation in -TC cells. Effect of -TC cell treatment with Gd3+ on Zn2+ uptake during low (A) and high glucose conditions (B). Effect of -TC cell treatment with nitrendipine (C) and Gd3+ (D) on KCl-induced Zn2+ uptake. The averaged traces ± S.E. of n = 17 (A and B), 18 (C), and 16 (D) cells are shown. The slopes reflect the relative rate of Zn2+ accumulation. In all cases the traces are representative at least three independent experiments.

The redox status of the cells is known to play a vital role in the regulation of different cellular processes. We investigated the effect of oxidative modification of -SH groups on the change of the intracellular Zn²⁺ profile. A small amount (10 µM) of -SH group oxidant DTDP led to a significant increase of the FluoZin-3 fluorescence level, indicating elevation of free intracellular Zn²⁺ in -TC cells (Fig. 8A) or in GFP-negative dispersed mouse islet cells (supplemental Fig. 2A). Other oxidizing compounds menadione (generator of superoxide) (Fig. 8B) or SIN-1 (generator of peroxynitrite by simultaneous production of nitric oxide and superoxide) (Fig. 8C) also elevated FluoZin-3 fluorescence, further supporting mobilization of intracellular Zn²⁺ by redox state. Products of lipid peroxidation can trigger multiple signaling cascades. HNE, a reactive aldehydic product of lipid peroxidation, which itself increases reactive oxygen species production, can form adducts with proteins containing histidine, cysteine, or lysine residues (65, 66). As seen in Fig. 8D, treatment of -cells with HNE also led to mobilization of intracellular Zn²⁺. The cells treated with the above oxidizing reagents were loaded with cell-impermeable dye PI, and no fluorescence increase was detected (not shown), which indicated the maintenance of cell membrane integrity during the time of experiments.

NAD(P)H autofluorescence was used to monitor the changes in cellular redox state following oxidant treatment. We found that perfusion of -TC cells with 20 µM DTDP led to a drop in NAD(P)H fluorescence, indicating the oxidation of pyridine nucleotides (not shown). NEM, a sulfhydryl alkylating agent that covalently modifies cysteine residues in proteins, also significantly increased intracellular Zn²⁺ mobilization in both -TC cells (Fig. 8E) and GFP-negative dispersed mouse islet cells (supplemental Fig. 2B). Similarly, iodoacetamide, another alkylating agent for cysteine and histidine residues in proteins, also mobilized intracellular Zn²⁺ (Fig. 8F).

The comparison of Fig. 8, B and C with E and F, shows the difference in the maximal values of FluoZin-3 fluorescence after treatment with various compounds. This diversity reflects the ability of compounds used to extract Zn²⁺ from intracellular Zn²⁺ stores, as well as the duration of treatment.

Our data suggest that these reagents extracted Zn²⁺ from Zn²⁺-containing metallothioneins or other possible Zn²⁺ stores. Because this extractable Zn²⁺ reflects a pool of total Zn²⁺ present in the cells, these results demonstrated a significant amount of bound Zn²⁺ in this cell type. Thus, bound Zn²⁺ reflects another significant pool that is highly modulated by redox state and potentially oxidative stress. To further examine this, we investigated whether the removal of oxidants or alkylating agents as well as a shift in redox state to more reduced conditions resulted in a decrease in [Zn²⁺]_i. As seen in Fig. 8G, the DTDP-stimulated increase in [Zn²⁺]_i was reversible. The fast increase in [Zn²⁺]_i was observed (Fig. 8G) if cells were perfused with 20 µM DTDP. The removal of the -SH group oxidant from the perfusion medium led to restoration of the initial FluoZin-3 fluorescence during 10 min (Fig. 8G). Similar effects were observed during the treatment of -TC cells with or SIN-1 (not shown). Removal of the -SH group modifier (NEM, 15 µM) resulted in a decrease in the rate of [Zn²⁺]_i accumulation but not to the restoration of the initial fluorescence level (not shown), possibly indicating the higher affinity of NEM to its target. We also used the thiol-reducing agent DTE to study the effect of -SH group reduction on [Zn²⁺]_i. Experiments showed that perfusion of -TC (Fig. 8H) with DTE (100 µM) following exposure to DTDP (20 µM) led to the dramatic decrease (during 2-3 min) in FluoZin-3 fluorescence indicating the decreased level of free Zn²⁺ in these cells.

View larger version (34K): [in this window] [in a new window]   FIGURE 6. Real time quantitative PCR analysis for Zip and ZnT transporter transcripts in-TC cells (A), mouse islets (B), mouse brain (C), and expression of ZnT8 protein in dispersed mouse islet cells (D). Mouse brain was used as a positive control for expression of Zip and ZnT transporters. KCNJ11 (KATP channel Kir6.2 subunit) expression was measured for comparison of relative levels of Zn2+ transporters. Expression of Zn2+ transporters and KCNJ11 was calculated using mouse β-actin as the normalizing gene. Results represent the average ± S.E. of 3-5 independent experiments. D, immunofluorescent detection of ZnT8 protein in insulin-(upper panel) and glucagon-expressing cells (lower panel).

View larger version (12K): [in this window] [in a new window]   FIGURE 7. Effect of Zn2+ on KATP current in mouse islet -cells. Extracellularly applied Zn2+ (10 µM) had no effect on KATP current density. Results represent the average ± S.E. of 9 cells.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

The effects of glucose and Zn²⁺ on glucagon secretion remain uncertain at the present time. In intact rat and mouse islets increased glucose decreases glucagon release (15, 67, 68). Contrary to this, glucose stimulates glucagon release in isolated rat pancreatic -cells (13). In glucagonoma cell lines InR1-G9 (69) and -TC1-9 (15), an increase in the glucose concentration led to a decrease in glucagon secretion. Our results in this study showed that glucose caused the inhibition of glucagon secretion (Fig. 1) in both an -cell line and isolated islets. However, Zn²⁺ reduced glucagon secretion both at low and high glucose conditions suggesting its effects are not entirely glucose-dependent (Fig. 1). Our experiments also demonstrated that Zn²⁺ accumulated in -cells both at low and high glucose conditions (Fig. 2, B and C, Fig. 3, B and C, and Fig. 4, A and B). Unlike β-cells (32), treatment with tolbutamide or nitrendipine did not have any visible effect on the rate of Zn²⁺ accumulation under the conditions studied. The nonselective Ca²⁺ channel blocker Gd³⁺ moderately attenuated Zn²⁺ accumulation both at low and high glucose conditions (Fig. 5, A and B), as well as during depolarization of cells with KCl (Fig. 5D). These findings indicate that Zn²⁺ is transported into -cells in part through dihydropyridine-insensitive Ca²⁺ channels. Four types of VGCC (L-, T-, N-, and R-type) and tetrodotoxin-sensitive voltage-gated Na⁺-channels have been observed in -cells and thus could be involved in Zn²⁺ transport (3, 37). Further studies are required to examine if other ion channels are involved.

As stated, the mechanism of the inhibition of glucagon release by Zn²⁺ is unclear. It was suggested that Zn²⁺ inhibits glucagon secretion in rat -cells because of activation of the K_ATP channel resulting in hyperpolarization of cells (12). However, the activation of the K_ATP channel by Zn²⁺ was not found in mouse -cells (70). In these studies both groups used different protocols for measuring K_ATP channel modulation. Our results are in agreement with previous findings in mouse; we see no effect of Zn²⁺ on -cell K_ATP current, suggesting that under the conditions studied this mechanism is not involved in Zn²⁺ modulation of glucagon secretion. The inhibition of secretion therefore appears to require transport as part of the mechanism (Fig. 7).

In addition to ion channels, another possible pathway of Zn²⁺ transport is through plasma membrane Zn²⁺ transporter(s). Although there are some data on Zn²⁺ influx transporters in the pancreas, information about Zip transporters in - and β-cells is limited. Zip1 transcripts were previously found in human (71), but not mouse, pancreas (72). The expression of Zip5 mRNA in the whole mouse pancreas was also observed (73, 74). Our previous work using quantitative real time PCR demonstrates a number of Zip transcripts in mouse insulinoma β-cells (MIN6), mouse islets, and mouse brain (32). In this work we have identified in glucagon-producing -TC cells the expression of a number of genes of the ZIP and ZnT families (Fig. 6A). The most abundantly expressed Zip transporter genes in -TC cells were Zip1, Zip10, and Zip14 (Fig. 6A). The expression profile of Zip transporters in mouse islets (Fig. 6B) and mouse brain (Fig. 6C) mainly confirms our previous results (32).

Unlike Zip transporters, the Zn²⁺ efflux transporters (ZnT) in pancreatic β-cells are better characterized. Among the ZnT family of transporters, the ZnT8 was recently detected in insulin-secreting INS-1 cells and human pancreatic islets and was co-localized with insulin in these cells, suggesting they are in vesicular membranes (29, 63). Our results identified the expression of ZnT1, ZnT4, ZnT5, ZnT6, ZnT7, and ZnT8 transporter genes in -TC cells (Fig. 6A) and mouse islets (Fig. 6B), with ZnT4, ZnT5, and ZnT8 showing the highest level of expression in -TC cells and ZnT5 and ZnT8 in islets. The high level of ZnT5 in -TC cells (Fig. 6A) and islets (Fig. 6B) is in agreement with a previous report, in which ZnT5 was found to be abundantly expressed in pancreas (75). Our data demonstrated that ZnT8 was not exclusively a β-cell Zn²⁺ transporter. It is noteworthy that others have shown that ZnT8 was also expressed in adipose tissue with comparable abundance as ZnT1, ZnT5, and ZnT7 (76). The role and localization of ZnT8 in -TC cells are not clear. As stated above, in β-cells ZnT8 is thought to be localized to the insulin secretory granule, and overexpression studies in β-cells suggests it functions to modulate [Zn²⁺]_i and regulate (enhance) insulin secretion (29, 63). It is important to reiterate that polymorphisms in the ZnT8 gene link ZnT8 transporter to T2D (26, 29), and this pathological association may involve effects on both - and β-cell function. It will be important to determine the compartmental distribution of -cell ZnTs in future studies and further assess their function in islet cells.

View larger version (28K): [in this window] [in a new window]   FIGURE 8. Effect of treatment with -SH group oxidant and blockers on the level of intracellular Zn2+. Increase of intracellular Zn2+ level in -TC cells after treatment with oxidants (A) DTDP, (B) menadione, (C) SIN-1, (D) HNE, -SH group blockers (E) NEM, and (F) iodoacetamide. G, reversible changes of intracellular Zn2+ after cyclic addition and removal of DTDP. H, effect of treatment with reducing agent DTE on intracellular Zn2+ level. The averaged traces ± S.E. of n = 13 (A), 17 (B), 12 (C), 14 (D), 18 (E), 17 (F), 16 (G), and 14 (H) cells are shown. In all cases the traces are representative of at least three independent experiments.

Recently, a vicious cycle of Zn²⁺-dependent injury was suggested in brain cells (77). This cycle includes oxidative stress-induced Zn²⁺ release from metallothioneins, followed by mitochondrial uptake of Zn²⁺ leading to organelle dysfunction and mitochondrial reactive oxygen species generation, which in turn caused further Zn²⁺ release from metallothioneins. The increase in [Zn²⁺]_i activates cytotoxic mitochondrial Zn²⁺ sequestration and apoptosis (77). A similar scenario could also take place in islet -cells in the presence of high glucose or free fatty acid levels, termed glucolipotoxicity, a condition characterized by increased reactive oxygen species production (78-80). Our experiments showed that a significant increase in [Zn²⁺]_i led to mitochondrial depolarization (Fig. 3G). One could speculate that processes leading to perturbations of Zn²⁺ homeostasis could be through a disturbance in the function of Zn²⁺ efflux transporter(s), including ion channels. In this case, a shift to the reduced state of cells, leading to Zn²⁺ binding and a decrease in [Zn²⁺]_i, could decrease the damage caused by excess of Zn²⁺ and may have a protective role. Our experiments with the thiol-reducing agent DTE (Fig. 8H), which led to a more reduced intracellular environment and decreased [Zn²⁺]_i, support such a conclusion.

In summary the data presented demonstrate that Zn²⁺ accumulates in glucagon-producing -cells under low and high glucose conditions through both dihydropyridine-insensitive Ca²⁺ channels and other Zn²⁺-transporting mechanisms. [Zn²⁺]_i in -cells is therefore likely regulated by Ca²⁺ channels and Zn²⁺ transporters as well as through the redox sensitivity of thiol groups.

	FOOTNOTES

 
^* This work was supported by Canadian Institutes of Health Research Operating Grant MOP-49521 (to M. B. W.). 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.

The on-line version of this article (available at http://www.jbc.org) contains supplemental Table 1 and Figs. 1 and 2.

¹ Supported by a studentship from Canadian Institute of Health Research.

² To whom correspondence should be addressed: Dept. of Physiology, University of Toronto, 1 King's College Circle, Rm. 3352, Toronto, Ontario M5S 1A8, Canada. Tel.: 416-978-6737; Fax: 416-978-4940; E-mail: michael.wheeler{at}utoronto.ca.

³ The abbreviations used are: ZIP, Zrt/Irt-like protein; GFP, green fluorescent protein; MIP-GFP, mouse insulin promoter-GFP; ZnT, Zn²⁺ transporters; VGCC, voltage-gated calcium channel; L-VGCC, L-type VGCC; DiBAC₄(3), bis-(1,3-dibutylbarbituric acid) trimethine oxonol; TPEN, N,N,N',N',-tetrakis-(2-pyridylmethyl)ethylenediamine; DTDP, 2,2'-dithiodipyridine; SIN-1, 3-morpholinosydnonimine; FCCP, carbonyl cyanide p-(trifluorometoxy)phenylhydrazone; NEM, N-ethylmaleimide; DTE, dithioerythritol; HNE, 4-hydroxynonenal; DIC, differential interference contrast; qPCR, quantitative real time PCR; FITC, fluorescein isothiocyanate; ER, endoplasmic reticulum; TRITC, tetramethylrhodamine isothiocyanate; PI, propidium iodide.

	ACKNOWLEDGMENTS

 
We thank Dr. E. Allister for review of the manuscript and helpful discussions.

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