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J. Biol. Chem., Vol. 278, Issue 47, 46210-46218, November 21, 2003

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Cytosolic Multiple Inositol Polyphosphate Phosphatase in the Regulation of Cytoplasmic Free Ca²⁺ Concentration^*

Jia Yu, Barbara Leibiger, Shao-Nian Yang, James J. Caffery¶, Stephen B. Shears¶, Ingo B. Leibiger, Christopher J. Barker||, and Per-Olof Berggren

From the Department of Molecular Medicine, The Rolf Luft Center for Diabetes Research, L3, Karolinska Institutet, Karolinska Hospital, Stockholm SE-171 76, Sweden and ^¶Inositide Signaling Section, NIEHS, National Institutes of Health, Research Triangle Park, North Carolina 27709

Received for publication, April 10, 2003 , and in revised form, September 5, 2003.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Multiple inositol polyphosphate phosphatase (MIPP) is an enzyme that, in vitro, has the interesting property of degrading higher inositol polyphosphates to the Ca²⁺ second messenger, inositol 1,4,5-trisphosphate (Ins(1,4,5)P₃), independently of inositol lipid breakdown. We hypothesized that a truncated cytosolic form of the largely endoplasmic reticulum-confined MIPP (cyt-MIPP) could representanimportantnewtoolintheinvestigationofIns(1,4,5)P₃-dependent intracellular Ca²⁺ homeostasis. To optimize our ability to judge the impact of cyt-MIPP on intracellular Ca²⁺ concentration ([Ca²⁺]_i) we chose a poorly responsive -cell line (HIT M2.2.2) with an abnormally low [Ca²⁺]_i. Our results show for the first time in an intact mammalian cell that cyt-MIPP expression leads to a significant enhancement of Ins(1,4,5)P₃ concentration. This is achieved without a significant interference from other cyt-MIPP-derived inositol phosphates. Furthermore, the low basal [Ca²⁺]_i of these cells was raised to normal levels (35 to 115 nM) when they expressed cyt-MIPP. Noteworthy is that the normal feeble glucose-induced Ca²⁺ response of HIT M2.2.2 cells was enhanced dramatically by mechanisms related to this increase in basal [Ca²⁺]_i. These data support the use of cyt-MIPP as an important tool in investigating Ins(1,4,5)P₃-dependent Ca²⁺ homeostasis and suggest a close link between Ins(1,4,5)P₃ concentration and basal [Ca²⁺]_i, the latter being an important modulator of Ca²⁺ signaling in the pancreatic -cell.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Inositol polyphosphates have established roles in signal transduction (1–5). The first inositol polyphosphate recognized to be a second messenger was inositol 1,4,5-trisphosphate (Ins(1,4,5)P₃)¹ (1). Ins(1,4,5)P₃ releases Ca²⁺ from intracellular stores (6) and thus plays a central role in intracellular Ca²⁺ homeostasis. The normal source of Ins(1,4,5)P₃ in mammalian cells is the breakdown of phosphatidylinositol 4,5-bisphosphate (PtdIns(4,5)P₂/PIP₂) following the activation of G protein- or tyrosine kinase-coupled membrane receptors (1). Interestingly, in non-mammalian cells, namely Dictyostelium, evidence has been presented that the abundant higher inositol polyphosphates, particularly Ins(1,3,4,5,6)P₅ can also be degraded to Ins(1,4,5)P₃ (7, 8), thus separating Ins(1,4,5)P₃ production from PIP₂ breakdown and the diacylglycerol-mediated activation of protein kinase C. However, these studies did not report whether the Ins(1,4,5)P₃ so produced could impact intracellular Ca²⁺ homeostasis. In intact mammalian cells no such pathway has been reported; however, in vitro multiple inositol polyphosphate phosphatase (MIPP) has been shown to possess the enzymatic activity to convert Ins(1,3,4,5,6)P₅ to Ins(1,4,5)P₃ via inositol tetrakisphosphate intermediates (7). The role of this pathway in an intact cell is rather problematic because of the fact that the majority of the MIPP is confined in the ER (9, 10) and thus removed from Ins(1,3,4,5,6)P₅, which exists largely in the cytosol (11). There are, however, exceptions to this generalization as membrane-associated MIPP activity in human erythrocytes is exposed to the cytosol (12). However, in general, the limited access of MIPP to its substrates restricts its use as a tool to study the impact of higher inositol polyphosphate-generated Ins(1,4,5)P₃ on [Ca²⁺]_i. In contrast, we reasoned that a cytosolic version of this enzyme (cyt-MIPP) may be a such a tool, allowing us to separate the production of Ins(1,4,5)P₃ from the concomitant breakdown of lipids. To best judge the impact of cyt-MIPP expression we used a -cell line with an abnormally low basal intracellular Ca²⁺ concentration ([Ca²⁺]_i), reasoning that this system would represent the most sensitive assay for recording a possible impact of cyt-MIPP on [Ca²⁺]_i homeostasis.

We now report for the first time that cyt-MIPP expression can generate Ins(1,4,5)P₃ in an intact cell. Furthermore, it significantly impacts on cellular Ca²⁺ homeostasis by both raising the low basal [Ca²⁺]_i in our model cell line to more normal levels and, as a likely consequence, considerably enhancing a previously poor Ca²⁺ response to glucose. This enhanced response was mediated by several interlinked factors. These included the sensitizing of the ryanodine receptor (RYR) to Ca²⁺-induced Ca²⁺ release (CICR) and the subsequent enhancement of Ca²⁺ entry, predominantly through store-operated Ca²⁺ channels. These data suggest that cyt-MIPP will be a useful tool in the investigation of Ins(1,4,5)P₃ in intracellular Ca²⁺ homeostasis, as evidenced by 1) data suggesting a link between basal Ins(1,4,5)P₃ concentration and basal [Ca²⁺]_i and 2) the importance of this basal [Ca²⁺]_i in determining the responsiveness of the RYR. Moreover, it should be noted that the full-length form of MIPP has unrestricted access to the cytosol in some mammalian cells (12). Hence, the pathway discussed here may have an impact on cellular Ca²⁺ handling in general.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Cell Culture, Transfection, and Labeling Protocols—HIT M2.2.2. cells were routinely maintained in RPMI 1640 medium with 10% fetal bovine serum, glutamine, and penicillin/streptomycin (Invitrogen) as described previously (13). For experiments a modified RPMI 1640, 1640-M, was used. This modified medium contained additional MgSO₄ (to give a final concentration of 0.8 mM) and 50 µM inositol (to make it more representative of the physiological milieu and to enable more efficient labeling) and 5.5 mM glucose. The 10% serum in this medium had been dialyzed (1000 molecular weight cut-off; Spectro Por) to remove inositol.

HIT M2.2.2 cells were plated at about 10% of their confluent density into 35- or 92-mm dishes and allowed to grow in the above RPMI 1640-M medium for 24 h. The cells were then changed into the transfection medium, Dulbecco's modified Eagle's medium, with the same inositol concentration as the experimental RPMI 1640-M. Cyt-MIPP expression plasmid (pCI.FLAG-MIPP) or vector alone were transfected into the cells using a Ca²⁺ phosphate precipitation technique (13). The next day the cells were washed and reintroduced into the RPMI 1640-M medium and maintained for a further 24 h. For some experiments measuring inositol polyphosphates and [Ca²⁺]_i, cells were washed twice in a Krebs buffer and preincubated for 45 min in the RPMI 1640-M medium, with a reduced glucose concentration (0.1 mM). To determine cell number and approximate volume, cells were trypsinized from the dish and counted, and the volume was estimated by measuring the diameter of spherical cells with a microscope and calibrated graticule. For the measurement of inositol phosphates cells were labeled for the entire growth period of 72 h with 6 µCi/ml myo-[³H]inositol (Amersham Biosciences) in the above medium in either 35- or 92-mm-diameter dishes.

Inositol Phosphate Analysis—Inositol phosphates were extracted using previously published methodologies (14–16). Briefly, medium was removed from the cells, and the cells were washed rapidly with Krebs-Ringer before addition of ice-cold 5% (w/v) trichloroacetic acid. The cell debris was pelleted, and the supernatant was removed and neutralized by ether washing and the addition of 0.1 M EDTA (pH 7.0) to bring the final pH to 6.5–7.0. To determine the mass of Ins(1,4,5)P₃ a commercial binding protein assay was used (Amersham Biosciences). The assay was carried out exactly as described in the supplied manual using a perchloric acid extraction protocol. Samples were then stored at –20 °C until required for HPLC.

HPLC—We performed HPLC on a 25-cm Whatman Partisphere-SAX column (Laserchrom, United Kingdom). For InsP separations HPLC was carried out as described previously (16). Initial identification of InsP species was carried out by the inclusion of [³H]inositol phosphate standards: inositol 1-phosphate, inositol 1,4-bisphosphate, inositol 1,3,4-trisphosphate, Ins(1,4,5)P₃, Ins(1,3,4,5)P₄, and InsP₆ in samples split into two. The two halves were run with and without standards. Standards were obtained from PerkinElmer Life Sciences, with the exception of inositol 1,3,4 trisphosphate, which was prepared from the Ins(1,3,4,5)P₄ by dephosphorylation using human erythrocyte membranes. Other inositol phosphates were identified by their relative elution positions and comparison of a previous study (16) on -cells using [¹⁴C]-labeled internal standards and identical elution conditions. Actual concentrations of InsPs were estimated based on the known volume of the cells and the known specific activity of the inositol used to label the majority of the cellular material. Radioactivity was determined by the addition of Packard Ultima Flo AP scintillant and counting on a Packard CA 2000 scintillation counter (both from CIAB, Stockholm, Sweden).

Measurement of [Ca²⁺]_i—Cells were cultured and transfected as above. They were then preincubated in basal glucose (0.1 mM) for 45 min. At the beginning of this preincubation period cells were loaded with Fura-2/AM to give a final concentration of 2 µM. Transfected cells were identified by DsRed (Molecular Probes, Leiden, Netherlands) fluorescence following coexpression with pRcCMVi.DsRed. DsRed was used rather than GFP because of the substantial interference of GFP in Fura-2 measurements, whereas DsRed has little impact. Fluorescence imaging was performed with a cool charged-coupled device camera (CH250 with KAF 1400; Photometrics, Tucson, AZ) connected to an imaging system (Inovision, Durham, NC) placed on a Zeiss Axiovert 135TV microscope (Carl Zeiss AGT, Göttingen, Germany) with a 515/40 emission filter using the Ratiotool software (Inovision). A SPEX fluorolog-2 CMlT11I spectrofluorimeter (SPEX industries, Edison, NJ) was used for fluorescence excitation at 340 and 380 nm. [Ca²⁺]_i was expressed as the ratio of fluorescence at 340 and 380 nm. To estimate basal [Ca²⁺]_i in the MIPP-versus mock-transfected cells, the experiments were calibrated as described previously (17). To verify that the pharmacological agents were working, we did parallel experiments on normal mouse -cells. Thus we alternated measurements of mouse -cell [Ca²⁺]_i changes with those of the HIT M2.2.2 cells. Normal mouse -cells have previously been extensively characterized and thus served as positive controls to all procedures and reagents during [Ca²⁺]_i measurements. The responses of the normal mouse -cells in the present study were similar to those reported previously in the literature (data not shown).

Electrophysiology—HIT M2.2.2 cells were transfected with either pCI.FLAG-MIPP (cyt-MIPP) in combination with pRcCMVi.EGFP or pRcCMVi.EGFP alone as mock transfection. In this case DsRed was not needed as no Fura-2 Ca²⁺ measurements were carried out. The transfected cells were cultured from 2 to 4 days on coverslips. The cells expressing EGFP were selected for single channel recordings. Pipettes were pulled from borosilicate glass capillaries (Hilgenberg, Malsfeld, Germany) on a horizontal programmable puller (DMZ Universal Puller; Zeitz-Instrumente, Ausburg, Germany). Typical electrode resistance was 2–4 megohms. Cell-attached single channel recordings were made with Ba²⁺ as the charge carrier (in mM): 110 BaCl₂, 10 tetraethyl ammonium chloride, 5 HEPES-Ba(OH)₂, pH 7.4. A depolarizing external recording solution, containing (in mM) 125 KCl, 30 KOH, 10 EGTA, 2 CaCl₂, 1 MgCl₂, 5 HEPES-KOH, pH 7.15, was used to bring the intracellular potential to approx 0 mV. Recordings were made with an Axopatch 200 amplifier (Axon Instruments, Foster City, CA). Voltage pulses (200 ms) were applied at a frequency of 0.5 Hz to depolarize cells from a holding potential of –70 mV to a membrane potential of 0 mV. Resulting currents were filtered at 1 kHz, digitized at 5 kHz, and analyzed with the software program pCLAMP 6 (Axon Instruments, Foster City, CA).

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
The construct used in this study was based on the N-terminal truncated rat MIPP sequence first reported by Craxton et al. (10). This sequence lacks the signal peptide necessary for translocation into the ER and was used to degrade cytosolic inositol polyphosphates. Although our cyt-MIPP had no ER signal peptide, recent reports have suggested that other internal peptide sequences can confer ER targeting (18). Furthermore, the only other published study using a truncated MIPP (19) had both the signal peptide and the ER retention sequence removed, whereas in the current study the construct still had the ER retention sequence. To verify that the cyt-MIPP was indeed cytosolic and not ER-localized, a cyt-MIPP bearing a GFP tag was expressed in HIT M2.2.2 cells. Examination with confocal microscopy discerned that GFP-cyt-MIPP was present throughout the cytosol. There was no co-localization with the fluorescent ER marker Brefeldin A, BODYiPY 558/568 conjugate, Molecular Probes (data not shown). Our experience (20), along with the experience of many others (21), is that a GFP tag itself does not present a restriction for the entry of peptides into the ER.

Changes in Inositol Polyphosphate Profile following cyt-MIPP Expression—In our initial experiments we established that cyt-MIPP expression decreased the concentration of its principal substrates, Ins(1,3,4,5,6)P₅ and InsP₆. Note, however, that only 30% of the cells were transfected (data not shown), so changes in concentration of inositol polyphosphates in a single cyt-MIPP-transfected cell were considerably underestimated because of the dilution with non-transfected cells. Fig. 1A indicates a significant 25% reduction in the concentrations of both Ins(1,3,4,5,6)P₅ and InsP₆ after cyt-MIPP expression.

View larger version (20K): [in this window] [in a new window]   FIG. 1. Effect of cyt-MIPP on InsP6 and Ins(1,3,4,5,6)P5 concentration and their downstream products including Ins(1,4,5)P3. HIT M2.2.2 cells were grown in either 35-mm (A) or 92-mm dishes (B), transfected and labeled with [3H]inositol (6 µCi/ml). Cells were then either immediately acid-killed (A) or preincubated for 45 min in RPMI 1640-M (see "Experimental Procedures") with basal glucose (0.1 mM) and then acid-killed (B). The InsPs were extracted and then separated using inositol phosphate HPLC protocol, and the radioactivity was determined (see "Experimental Procedures"). A, the concentrations of Ins(1,3,4,5,6)P5 and InsP6 are plotted from cyt-MIPP-transfected cells relative to mock-transfected cells with appropriate adjustments for differences in cell volume and number. The data are the mean ± S.E. of five separate experiments each carried out in triplicate. *, p < 0.05 (unpaired t test). B, D-Ins(1,4,5)P3 was determined in mock-versus cyt-MIPP-transfected cells using a commercial kit. cyt-MIPP-transfected data are presented as percent of mock-transfected cells with adjustments made for differences in cell volume and number. The data shows the mean ± S.E. from three separate experiments each carried out in triplicate. Significance was carried out using an unpaired Student's t test; **, p < 0.01.

Fig. 2 presents a scheme depicting the putative products of cyt-MIPP-catalyzed metabolism of Ins(1,3,4,5,6)P₅ and InsP₆. There were significant increases in Ins(1,2,3)P₃ (20%) and in peaks with the elution characteristics of D/L-Ins(1,4,6)P₃ (56%) and Ins(1,3,4,5)P₄ (79%) in cyt-MIPP-compared with mock-transfected cells, (data not shown). However, it is important to note that the concentrations of these inositol polyphosphates were initially so low (0.2 µM) that even an increased production is unlikely to have any cellular impact. These inositol polyphosphates could all be derived directly or indirectly from cyt-MIPP degradation of Ins(1,3,4,5,6)P₅ and InsP₆ (see Fig. 2). The shape of the mock- but not the cyt-MIPP-transfected Ins(1,4,5)P₃ peak was skewed (data not shown), indicating that more than one InsP₃ isomer was present in the peak. The fact that only the mock-transfected Ins(1,4,5)P₃ peak was skewed suggests that this contaminant did not go up after cyt-MIPP transfection. Noteworthy is that the contribution of the contaminant to the Ins(1,4,5)P₃ signal in the mock-transfected cells reduced our ability to detect an increase in genuine Ins(1,4,5)P₃ against a high non-Ins(1,4,5)P₃ background. In a previous study in normal mouse -cells, the peak of Ins(1,4,5)P₃ (14) also indicated contamination from an unknown InsP₃, which masked bona fide Ins(1,4,5)P₃. Thus, we used a commercial mass assay, specific for D-myo-Ins(1,4,5)P₃, to determine the true increase in Ins(1,4,5)P₃. We estimated that approximately half of the HPLC-resolved Ins(1,4,5)P₃ was another InsP₃. Fig. 1B illustrates a significant 36% increase in Ins(1,4,5)P₃ concentration between cyt-MIPP- and mock-transfected cells, when only genuine Ins(1,4,5)P₃ was measured. It is important to note that because of the 30% transfection rate (see above) the actual rise in concentration of Ins(1,4,5)P₃ in a transfected cell would be considerably greater, about double the concentration in the mock-transfected cells. Thus cyt-MIPP expression significantly enhanced basal Ins(1,4,5)P₃ concentration in intact cells and not just in vitro as has been reported previously (7).

View larger version (11K): [in this window] [in a new window]   FIG. 2. Diagram illustrating possible metabolic interactions resulting from cyt-MIPP expression. This figure is a synthesis of known downstream products of MIPP activity toward Ins(1,3,4,5,6)P5 and conjectured downstream products of InsP6 based on the current data shown in Fig. 1B and InsP6 metabolism experiments carried out in mammalian cell homogenates (19). Thus the actual intermediates between InsP6 and Ins(1,2,3)P3 are undefined, and a number of species are bracketed. Dashed lines reflect minor pathways.

HIT M2.2.2 cells transfected with cyt-MIPP or mock-transfected were loaded with the Ca²⁺-sensitive dye Fura-2/AM, and the [Ca²⁺]_i in individual DsRed-positive cells were recorded using digital fluorescence imaging. Fig. 3 represents a quantification of basal [Ca²⁺]_i in mock- and cyt-MIPP-transfected cells, determinations made in 45 separate cells from more than three separate preparations. Basal [Ca²⁺]_i in cyt-MIPP-transfected cells was significantly higher (3-fold) than in mock-transfected cells (115.1 ± 4.1 versus 35.5 ± 2.5 nM ± S.E.). Untransfected cells had a [Ca²⁺]_i similar to mock-transfected cells (data not shown). Because the parent HIT T15 cell line has a basal [Ca²⁺]_i between 100 and 150 nM, the untransfected HIT M2.2.2 cells clearly have an abnormally low basal [Ca²⁺]_i. Thus cyt-MIPP expression is able to raise the basal [Ca²⁺]_i in HIT M2.2.2 cells to a more normal level.

View larger version (10K): [in this window] [in a new window]   FIG. 3. Effect of MIPP expression on basal Ca2+. HIT M2.2.2 cells, either cyt-MIPP- or mock-transfected, were prepared in experimental medium (see "Experimental Procedures"), preincubated for 45 min in RPMI 1640-M with basal (0.1 mM) glucose, and simultaneously loaded with Fura-2/AM (2 µM). Fluorescence in single transfected cells was measured by digital fluorescence imaging. [Ca2+]i is expressed in nM. The data are the mean ± S.D. of 45 separate determinations; ****, p < 0.0001.

View larger version (18K): [in this window] [in a new window]   FIG. 4. Effect of cyt-MIPP expression on glucose-stimulated Ca2+ increase. Cells were prepared identically to Fig. 5. Traces corresponding to cells mock- or cyt-MIPP-transfected are denoted in the figures. A, cells were stimulated with 16.7 mM glucose. B,10 µM nimodipine was added simultaneously with the low glucose medium. Following preincubation in low glucose, cells were then stimulated with 16.7 mM glucose at the time indicated. The figure shows a typical trace, with a reduced response in the presence of nimodipine, the average reduction produced over seven experiments was 18%. In the case of the glucose-stimulated cyt-MIPP-transfected cells 80% of the traces showed the repetitive spiking pattern, and the periodicity varies between 20 and 40 s dependent on the individual preparation of cells; however, in an individual cell the spikes are remarkably regular. The mock-transfected cells gave only a single small response, often less clear than depicted here. C, mock- or cyt-MIPP-expressing cells were preincubated in a medium with no added Ca2+ and then stimulated with 16.7 mM glucose for the time indicated. Fig. 6, A–C shows typical data from seven experiments.

Role of Voltage-dependent L-type Ca²⁺ Channels—To address the role of voltage-dependent L-type Ca²⁺ channels in the improved Ca²⁺ response following cyt-MIPP expression in more detail, single channel activity was recorded, using the cell-attached mode of the patch clamp technique. Both mock- and cyt-MIPP-transfected cells were examined for L-type Ca²⁺ current. Transfected cells were identified by co-expression of GFP. Fig. 5A illustrates examples of Ba²⁺ currents flowing through single Ca²⁺ channels recorded from patches attached to cells subjected to mock transfection (left panel) and cyt-MIPP transfection (right panel), respectively. The cell transfected with cyt-MIPP displayed more frequent openings of the Ca²⁺ channel than the control cell. Open time (Fig. 5B) and current amplitude (Fig. 5C) distributions from the same recordings were fit by a single exponential function and exhibit Gaussian distributions with a clear separation between closed and open states, respectively. This indicates that only one channel was present in the patches recorded. Compiled data demonstrated that cyt-MIPP expression significantly increased open probability (Fig. 5E) and availability (Fig. 5F) of single L-type Ca²⁺ channels without influencing mean open time (Fig. 5D). Thus L-type Ca²⁺ channels from cyt-MIPP-transfected cells have changed characteristics that may favor increased Ca²⁺ flux into the cell.

View larger version (33K): [in this window] [in a new window]   FIG. 5. Effect of cyt-MIPP expression on single L-type Ca2+ channel activity. A, examples of single Ca2+ channel currents recorded from cell-attached patches on a control cell (mock transfection, left) and a cell expressing cyt-MIPP (right). Both patches contain one L-type Ca2+ channel. The channel in the patch on the cell expressing cyt-MIPP opens more frequently than that in the patch on the control cell subjected to mock transfection. B, open time distributions from the same patches were fit by a single exponential function, indicating only one channel in the patch. C, current amplitude distributions from the same patches (left, mock transfection; right, cyt-MIPP-expression) as in A. Current amplitudes in both patches exhibit Gaussian distributions with a clear separation between closed (C) and open (O) states, indicating only one conductance level in the patch. D, summary graph of the percentage change in mean open time shows that there was no significant difference between control cells (n = 9) and cells overexpressing cyt-MIPP (n = 9). E, summary graph of the change in open probability shows that cells expressing cyt-MIPP (n = 9) display significantly higher open probability than control cells (n = 9; *, p < 0.05). F, summary graph of the change in availability shows that cells expressing cyt-MIPP (n = 9) display significantly higher availability than control cells (n = 9; *, p < 0.05). Data are presented as means ± S.E. Statistical significance was evaluated by unpaired Student's t test.

Fig. 6A shows cells preincubated with the Ins(1,4,5)P₃ receptor blocker 2-aminoethoxydiphenyl-borate (2-APB) and then stimulated with glucose. 2-APB completely blocked the glucose-induced rise in [Ca²⁺]_i in both mock- and cyt-MIPP-transfected cells (cf. Fig 4A). During the blockade of the Ins(1,4,5)P₃-sensitive stores it should still be possible to measure Ca²⁺ entry through L-type Ca²⁺ channels by depolarizing the cells with KCl. Addition of KCl, together with glucose or on its own, resulted in a distinct but small change in [Ca²⁺]_i. This confirms the nimodipine data that suggested that the L-type channel only made a small contribution to the overall [Ca²⁺]_i increase and raises the notion that this -cell variant has a low number of functional L-type Ca²⁺ channels. Recent data have indicated that 2-APB has additional effects on capacitative Ca²⁺ entry (reviewed in Ref. 25). It has been reported to inhibit store-operated Ca²⁺ entry at a lower concentration than required to block the Ins(1,4,5)P₃-receptor (25). Thus a block by 2-APB of the Ins(1,4,5)P₃ receptor will always be accompanied by an effective inhibition of capacitative Ca²⁺ entry. Therefore, we checked our conclusions regarding the cyt-MIPP-expressing cells by the use of an alternative cell-permeant Ins(1,4,5)P₃ receptor blocker, Xestospongin C. Xestospongin C was also able to block the glucose-mediated increase in [Ca²⁺]_i (Fig. 6B versus Fig. 4A), though the effect was not as complete as with 2-APB (Fig. 6, B versus A), which suggests that capacitative Ca²⁺ entry may be involved in the enhanced glucose-induced Ca²⁺ response.

View larger version (20K): [in this window] [in a new window]   FIG. 6. Role of Ins(1,4,5)P3 and ryanodine-sensitive Ca2+ stores. Cells were prepared as described in Fig. 4. A, cells were preincubated with 100 µM 2-APB and then stimulated with 16.7 mM glucose or KCl at the times indicated. B, cells were preincubated with 20 µM Xestospongin C and then stimulated with glucose as indicated. C, cells were preincubated with a mixture of 5 mM caffeine and 100 µM ryanodine, to block ryanodine sensitive Ca2+ stores. The traces are representative of 10 other experiments.

The above data suggest that cyt-MIPP enables the coupling between glucose metabolism and the release of Ca²⁺ from intracellular stores. This mechanism was apparently defective in the parent HIT M2.2.2 cell. One possible explanation for the difference between mock- and cyt-MIPP-transfected cells could be the Ca²⁺ content of the stores themselves. Therefore we assessed the content of the stores by adding the store Ca²⁺-ATPase inhibitor thapsigargin (Tg) (Fig. 7). Fig. 7 indicates that application of Tg releases Ca²⁺ from intracellular stores of both mock- and MIPP-transfected cells; however, the cyt-MIPP-transfected cells had a lower amount of Ca²⁺ in the stores. Fig. 7 also illustrates that if the cells were stimulated with 16.7 mM glucose, subsequent to the addition of Tg and thereby increasing basal [Ca²⁺]_i in mock-transfected cells to a more normal level, the deficient [Ca²⁺]_i response was partially restored.

View larger version (18K): [in this window] [in a new window]   FIG. 7. Effect of cyt-MIPP expression on handling of intracellular Ca2+ pools. Cells were prepared as described under "Experimental Procedures" and in Fig. 5. 1 µM Tg or 16.7 mM glucose were added at the times indicated to mock- or cyt-MIPP-transfected cells. These are representative traces of 10.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Cyt-MIPP Degrades Higher Inositol Polyphosphates to Ins(1,4,5)P₃ and Other Downstream Products—The expected primary consequence of cyt-MIPP-transfection would be the depletion of its principal substrates, in this case Ins(1,3,4,5,6)P₅ and InsP₆. In this respect, the 25% reduction in both Ins(1,3,4,5,6)P₅ and InsP₆ in cyt-MIPP-expressing cells is therefore not surprising. Because about one-third of the cells were transfected the impact in an individual cell would be considerably greater than this. These data are also consistent with the one other published report (19) documenting the expression of a cytosolic version of MIPP. In this latter case, an even greater reduction in Ins(1,3,4,5,6)P₅ and InsP₆ was observed (57 and 40%, respectively; see Ref. 19). This difference is likely to be a reflection of the different levels of expression of cyt-MIPP in the two cell systems.

Fig. 2 illustrates the possible impact of cyt-MIPP expression on inositol phosphate biochemistry. In contrast to the current study, the previous report documenting the impact of cyt-MIPP on Ins(1,3,4,5,6)P₅ and InsP₆ (19) did not characterize the downstream products in detail. The principal detected increases in concentration resulting from cyt-MIPP expression are found in the InsP₃ and inositol tetrakisphosphate regions of the chromatogram, namely, Ins(1,2,3)P_3, D/L-Ins(1,4,6)P₃, Ins(1,4,5)P₃, and Ins(1,3,4,5)P₄. Ins(1,2,3)P₃ is a downstream product of InsP₆ dephosphorylation in mammalian cells (16). D/L-Ins(1,4,6)P₃ is a putative inositol 1,3,4,6-tetratrisphosphate metabolite (see Fig. 2 and Ref. 27). Both inositol 1,3,4,6-tetratrisphosphate and Ins(1,3,4,5)P₄ can be derived by the action of MIPP on Ins(1,3,4,5,6)P₅ (see Fig. 2 and Ref. 7). However, Ins(1,3,4,5)P₄ can also be produced by the action of the Ca²⁺-activated Ins(1,4,5)P₃ 3'-kinase on Ins(1,4,5)P₃ (Fig. 2). It is worth reinforcing at this time that the relatively small increase in Ins(1,4,5)P₃ (35%) will be an underestimate because of the 30% transfection rate. Thus in a single transfected cell the actual Ins(1,4,5)P₃ concentration would be doubled by cyt-MIPP. This is important to note when marrying the Ins(1,4,5)P₃ changes with the [Ca²⁺]_i changes discussed below.

The usefulness of cyt-MIPP as a tool is dependent on whether other products of MIPP activation, besides Ins(1,4,5)P₃, have an impact on cell Ca²⁺ homeostasis or on general cellular physiology. However, at the outset it is important to note that these other inositol polyphosphates exist at low concentrations, and their increase upon MIPP expression is relatively small. The inositol polyphosphates with detectable increases that also may impact on Ca²⁺ handling are D/L-Ins(1,4,6)P₃ (28, 29) and Ins(1,3,4,5)P₄ (30, 31). D/L-Ins(1,4,6)P₃ is a partial agonist of the Ins(1,4,5)P₃ receptor with the D-isoform being the most potent and thus relevant enantiomer (29). We estimate the increase in this mixed peak in a single transfected cells to be maximally 0.17 µM over basal concentrations, in contrast to an increase in Ins(1,4,5)P₃ concentration of 0.7 µM over basal concentrations. The combination of the 4-fold higher concentration of Ins(1,4,5)P₃ allied with the 3–80-fold lower potency of D-Ins(1,4,6)P₃ (28, 29), dependent on Ins(1,4,5)P₃ receptor subtype (29), suggests that the contribution of D-Ins(1,4,6)P₃ will be negligible, especially in cells in which the type 2 or type 3 receptors dominate (29), which includes -cells. Ins(1,3,4,5)P₄ also has a role in Ca²⁺ homeostasis, aiding Ins(1,4,5)P₃ -induced capacitative Ca²⁺ entry (30, 31). However, Ins(1,3,4,5)P₄ is only a minor product of MIPP action (see Ref. 7 and Fig. 2), and the most likely source of Ins(1,3,4,5)P₄ in the current experiments is the Ins(1,4,5)P₃ derived directly from cyt-MIPP activity and not from higher inositol polyphosphates (see above). Nonetheless, we estimate the maximal concentration of Ins(1,3,4,5)P₄ to be 0.4 µM in contrast to 2 µM, the minimally effective concentration of Ins(1,3,4,5)P₄ required to enhance capacitative Ca²⁺ entry (30). The only other inositol phosphate to increase in concentration is the Ins(1,2,3)P₃, which is ineffective as an agonist to the Ins(1,4,5)P₃ receptor (28). Ins(1,2,3)P₃ is an iron chelator and proposed iron shuttle (16). Could the very small increase in Ins(1,2,3)P₃ impact on intracellular iron metabolism? This is not a likely scenario, because Ins(1,2,3)P₃ has related properties to the iron chelator desferrioxamine but is less potent. In this context desferrioxamine is required at a 500-fold higher concentration than the actual Ins(1,2,3)P₃ concentration obtained subsequent to MIPP expression to elicit any impact on cellular iron metabolism (32). Thus cyt-MIPP is able to generate Ins(4,5)P₃ independently of PtdIns(1,4,5)P₂ breakdown without serious interference from other inositol phosphate products. It is interesting to note that raising Ins(1,4,5)P₃ by an alternative strategy, namely the knock-down of the Ins(1,4,5)P₃/Ins(1,3,4,5)P₄ 5-phosphatase produced results similar to those described in the current work (33–35).

These data indicate for the first time that Ins(1,4,5)P₃ can be generated in the intact cell following cyt-MIPP expression. The fact that Ins(1,4,5)P₃ can be derived from higher inositol polyphosphates, principally Ins(1,3,4,5,6)P₅, has previously only been shown in a non-mammalian cell, Dictyostelium (7, 8), although the identity of the enzyme responsible remains unknown. Full-length MIPP is confined in the ER in a number of cell types (9, 19) and thus separated from its principal substrates. However, studies on human erythrocytes have suggested that MIPP activity is associated with the inner leaflet of plasma membranes and thus with easy access to its principal substrates (12). This suggests that, in at least some mammalian cells, a pathway exists that makes the generation of Ins(1,4,5)P₃ independent of lipid breakdown a real physiological possibility.

Impact of cyt-MIPP Expression on [Ca²⁺]_i Homeostasis in HIT M2.2.2 Cells—Given that Ins(1,4,5)P₃ (1) was generated in the intact cell we next wanted to know whether production of Ins(1,4,5)P₃ in this manner could affect [Ca²⁺]_i. In choosing the HIT M2.2.2 cell with its abnormally low [Ca²⁺]_i, we hoped to maximize our ability to detect the impact of cyt-MIPP expression on basal [Ca²⁺]_i. We studied [Ca²⁺]_i by loading mock- and cyt-MIPP-transfected cells with the Ca²⁺-sensitive dye Fura-2. We identified transfected cells by co-transfection with DsRed, and thus the [Ca²⁺]_i measurements were restricted to cells expressing cyt-MIPP or mock-transfected controls. Therefore, the magnitude of the response was not diluted by untransfected cells, unlike the Ins(1,4,5)P₃ and other inositol polyphosphate data. There were two critical changes rendered by cyt-MIPP expression on [Ca²⁺]_i. The first was a rise in [Ca²⁺]_i under basal conditions. The second was the enhancement of a glucose-induced Ca²⁺ response.

Cyt-MIPP Expression Raises Basal [Ca²⁺]_i—It might seem self-evident that the increased basal concentration of the Ca²⁺ mobilizing second messenger Ins(1,4,5)P₃ would lead to a concomitant rise in [Ca²⁺]_i. However, the literature is more circumspect on this issue (36, 37). This is because a number of contributing factors, in addition to Ins(1,4,5)P₃ concentration, can play a role in setting basal [Ca²⁺]_i, in particular plasma membrane and ER Ca²⁺-ATPases and various plasma membrane Ca²⁺ channels (36). Also, the mechanism of Ins(1,4,5)P₃ action is normally associated with an acute rise and fall in [Ca²⁺]_i, characteristic of its second messenger function (1). Nonetheless, it is interesting to note that this is the third study that we are aware of that has correlated Ins(1,4,5)P₃ concentration or Ins(1,4,5)P₃-sensitive Ca²⁺ stores with the steady state basal [Ca²⁺]_i. The first approach to chronically raising Ins(1,4,5)P₃ concentration was achieved by an antisense knockdown of the principal degradatory enzyme of Ins(1,4,5)P₃, the Ins(1,4,5)P₃/Ins(1,3,4,5)P₄ 5-phosphatase (33–35). The authors recorded both chronically raised Ins(1,4,5)P₃ and a concomitantly increased basal [Ca²⁺]_i (33). The elevated level of [Ca²⁺]_i found in the cells with reduced Ins(1,4,5)P₃/Ins(1,3,4,5)P₄ 5-phosphatase activity was still largely maintained in the absence of extracellular Ca²⁺ (33). This suggests that the increased [Ca²⁺]_i was being maintained by the components of the intracellular Ca²⁺ handling apparatus. The second study relates to an examination of kidney glomerula cells from mice with the Ins(1,4,5)P₃ type 1 receptor knocked out and with no compensatory change in other types of Ins(1,4,5)P₃ receptors (37). In this study the basal [Ca²⁺]_i was significantly lower in the knock-out mouse cells than that of the controls, again implying that Ins(1,4,5)P₃ gated stores influence basal [Ca²⁺]_i. In all three cases and particularly the knock-out mouse, what we are observing is the result of chronic changes that could have been implemented by some longer term adaptive responses in [Ca²⁺]_i homeostasis. However, we believe the simplest explanation is that basal Ins(1,4,5)P₃ can set basal [Ca²⁺]_i. Evident from all three studies is that the relationship between basal Ins(1,4,5)P₃ and basal [Ca²⁺]_i must be given a greater weight in the future modeling of basal [Ca²⁺]_i homeostasis in mammalian cells in general.

Cyt-MIPP Expression Enhances Stimulus-induced Increases in [Ca²⁺]_i—The second area of impact of cyt-MIPP expression was to considerably enhance the glucose-stimulated [Ca²⁺]_i rise in the previously rather unresponsive HIT M2.2.2 cells. In normal -cells the glucose-stimulated increase in [Ca²⁺]_i is mediated by Ca²⁺ entry through the voltage activated L-type Ca²⁺ channel and Ca²⁺ mobilization from intracellular stores (23), supported by Ca²⁺-induced Ca²⁺ release mechanisms (38). Both these principal signal transduction mechanisms could be positively enhanced by the increase in basal [Ca²⁺]_i we observed following cyt-MIPP expression.

Increased basal Ca²⁺ can enhance L-type Ca²⁺ channel activity via the increased stimulation of Ca²⁺-calmodulin-sensitive adenylate cyclases present in -cells (39) and the subsequent enhancement of the protein kinase A phosphorylation of the channel that results in its increased activity (40). Fig. 5 indicates that there are significant increases in open probability and channel availability following cyt-MIPP expression. These particular changes in channel characteristics specifically relate to increased phosphorylation of this channel by protein kinase A (40) (see above). Thus there is a clear mechanism linking the raised basal [Ca²⁺]_i created by cyt-MIPP expression and the observed enhancement of L-type Ca²⁺ channel activity. However, the improved response did not come primarily from this enhancement of the L-type Ca²⁺ channel activity (23), because the specific L-type Ca²⁺ channel blocker nimodipine only reduced the enhanced response by about 20% (see Fig. 4B). Therefore, the Ca²⁺ sensitivity of the intracellular Ca²⁺ stores is likely to be the more important player in the restored glucose-induced Ca²⁺ response.

In normal insulin-secreting -cells, the L-type Ca²⁺ channel has a partner, the RY-sensitive intracellular Ca²⁺ store (22, 38). It is now better appreciated that, like in cardiac cells, in -cells the Ca²⁺ entering the cell via the L-type Ca²⁺ channel triggers CICR from the RYR-operated intracellular Ca²⁺ stores (22, 38). The RYR is a good candidate for linking the elevated basal [Ca²⁺]_i to the increased glucose-induced Ca²⁺ response, because the proper operation of this receptor is sensitive to prevailing [Ca²⁺]_i (38). Thus, the lowered [Ca²⁺]_i in wild type HIT M2.2.2 cells is likely to be inhibitory to RYR function whereas the restored basal [Ca²⁺]_i in cyt-MIPP-expressing cells may permit RYR activation. Indirect evidence that the raised basal [Ca²⁺]_i in the cyt-MIPP-expressing cells might activate the RYR comes from the pattern of multiple peaks of [Ca²⁺]_i observed upon glucose stimulation. These multiple peaks are a common observation in glucose-stimulated -cells (38), but their magnitude is often small. However, sensitization of the RYR reveals a strikingly regular repetitive Ca²⁺ spiking pattern of greater amplitude thought to be associated with CICR (22, 38, 41). More directly, our data clearly show that blockade of the RYR prevents the enhanced Ca²⁺ response to glucose in cyt-MIPP cells, confirming this idea. Our discussions above indicate that in the case of the HIT M2.2.2 cells the majority of the Ca²⁺ trigger for CICR is not coming from the L-type channel, which suggests other intracellular Ca²⁺ stores, possibly those sensitive to Ins(1,4,5)P₃, are responsible for the initiation of the CICR. Glucose can stimulate increased levels of Ins(1,4,5)P₃ in -cells leading to the mobilization of intracellular Ca²⁺ (42). Our data support the idea that this mechanism contributes to the enhanced glucose response as the blockade of the Ins(1,4,5)P₃ receptor can completely or partially block the response, dependent on the pharmacological agent used, 2-APB or Xestospongin C, respectively. Because 2-APB is a better blocker of store-operated capacitative Ca²⁺ entry than it is of the Ins(1,4,5)P₃ receptor (25), the partial block afforded by the Xestospongin is probably more relevant. Thus Ca²⁺ released from Ins(1,4,5)P₃-sensitive stores is an important contributor to the proposed Ca²⁺ activation of the RYR. Glucose can also stimulate RYR operation in other ways. One is the direct priming by the glycolytic intermediate fructose 1,6-bisphosphate (43) and another is the phosphorylation of the channel by protein kinase A (44) whose activity, we have already argued, is enhanced by raised [Ca²⁺]_i (see above).

One other important final element needs to be integrated into this model. The fact that both the omission of extracellular Ca²⁺ (Fig. 4C) and the application of 2-APB (an inhibitor of capacitative Ca²⁺ entry (Fig. 6A) (see above)) can completely block the response, suggests that store-operated Ca²⁺ entry plays an important role (24). Hence, the apparent depletion of the intracellular stores in cyt-MIPP-expressing cells (Fig. 7) may act to enhance capacitative Ca²⁺ entry as the stores will require less Ca²⁺ released from them to reach the threshold needed for store-operated Ca²⁺ influx to be activated. In short, we hypothesize that the primary consequence of the raised basal [Ca²⁺]_i is to prime the RYR to Ca²⁺. The RYR can then be activated by glucose metabolism and the glucose-induced increase in [Ca²⁺]_i coming largely from Ins(1,4,5)P₃-sensitive stores.

We have shown that expression of cyt-MIPP can raise Ins(1,4,5)P₃ concentration in an insulin-secreting cell, without a significant contribution of other cyt-MIPP side products. The raised Ins(1,4,5)P₃ has an impact on basal [Ca²⁺]_i, raising a previously low [Ca²⁺]_i to a more physiological basal concentration. This increase in basal [Ca²⁺]_i is mechanistically linked to the dramatically enhanced glucose-stimulated [Ca²⁺]_i seen following cyt-MIPP expression. These data support the use of cyt-MIPP as an important tool in investigating Ins(1,4,5)P₃-dependent Ca²⁺ homeostasis and the idea of a close link between basal Ins(1,4,5)P₃ concentration and basal [Ca²⁺]_i, the latter being an important modulator of Ca²⁺ signaling in the pancreatic -cell.

	FOOTNOTES

 
^* This work was supported by grants from Karolinska Institutet, Novo Nordisk Foundation, the National Institutes of Health (DK-58508), the Swedish Research Council, the Swedish Diabetes Association, Juvenile Diabetes Research Foundation International, Åke Wibergs Foundation, and Berth von Kantzow's Foundation. 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.

Contributed equally to this work

^|| To whom correspondence should be addressed. Tel.: 46-8-517-79454; Fax: 46-8-517-79450; E-mail: chris.barker{at}molmed.ki.se.

¹ abbreviations used are: Ins(1,4,5)P₃, inositol 1,4,5-trisphosphate; PtdIns(4,5)P₂/PIP₂, phosphatidylinositol 4,5-bisphosphate; Ins (1,3,4,5,6)P₅, inositol 1,3,4,5,6-pentaphosphate; MIPP, multiple inositol polyphosphate phosphatase; ER, endoplasmic reticulum; RYR, ryanodine receptor; CICR, Ca²⁺-induced Ca²⁺ release; HPLC, high performance liquid chromatography; Ins(1,3,4,5)P₄, inositol 1,3,4,5-tetraphosphate; InsP, inositol phosphate; GFP, green fluorescent protein; EGFP, enhanced GFP; Ins(1,2,3)P₃, inositol 1,2,3-trisphosphate; D/L-Ins (1,4,6)P₃, D/L-inositol 1,4,6-trisphosphate; InsP₆, inositol hexakisphosphate; InsP₃, inositol triphosphate; RY, ryanodine; 2-APB, 2-aminoethoxydiphenyl-borate; Tg, thapsigargin.

	ACKNOWLEDGMENTS

 
We thank Dr. T. Moede for help with Ca²⁺ imaging and Drs. M. S. Islam and S. R. James for useful discussions.

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