Mechanism of Cyclosporin-induced Inhibition of Intracellular Collagen Degradation*

Pamela D. Arora, Livia Silvestri, Bernhard Ganss, Jaro Sodek, and Christopher A. G. McCullochDagger

From the Canadian Institutes of Health Research Group in Periodontal Physiology, Faculty of Dentistry, University of Toronto, Toronto, Ontario M5S 3E2, Canada

Received for publication, November 13, 2000, and in revised form, January 12, 2001




    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The immunosuppressant cyclosporin A (CsA) markedly inhibits collagen degradation by an intracellular phagocytic pathway in fibroblasts, an effect that can lead to massive gingival overgrowth. We used a collagen bead model of collagen phagocytosis to determine whether CsA inhibits internalization by blocking efflux of calcium from endoplasmic reticulum (ER) and mitochondrial calcium stores. CsA caused dose-dependent inhibition of phagocytosis of collagen-coated (but not bovine serum albumin-coated) beads. Chelation of intracellular Ca2+ with BAPTA/AM or inhibition of Ca2+-ATPase of ER stores with thapsigargin reduced collagen bead phagocytosis. Measurement of intracellular calcium by ratio fluorometry showed increases in response to collagen-coated beads. Preincubation with CsA or thapsigargin caused a >3-fold decrease in intracellular calcium elevations in response to stimulation with collagen beads. Direct measurements of Ca2+ in mitochondrial and ER stores showed that CsA only slightly inhibited collagen bead-induced discharge of calcium from mitochondria, but almost completely blocked discharge from ER stores. We reduced the numbers of mitochondria with chronic ethidium bromide treatment to test for the importance of ER/mitochondrial interactions. In these cells, CsA delayed collagen bead-induced calcium discharge from mitochondria. Collectively, these data indicate that CsA inhibits collagen phagocytosis by blocking calcium release from ER stores and may perturb functional interactions between the ER and mitochondria that regulate calcium stores.




    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Cyclosporin A (CsA)1 is a cyclic endecapeptide immunosuppressant that is widely used to prevent organ rejection after transplantation. Despite the effectiveness of CsA in preventing rejection, its clinical application is limited by various side effects (1), including cardiotoxicity, nephrotoxicity, and gingival overgrowth (2). Previous studies have suggested that perturbation of intracellular Ca2+ signaling (3) is a common mechanism for CsA-induced cardiotoxicity (4) and many other side effects. In cardiac muscle cells, CsA inhibits contractility by altering the function of Ca2+ release channels and deregulating mitochondrial ion homeostasis (5). CsA induces mitochondrial dysfunction by binding to cyclophilin D (6), a mitochondrial protein that regulates conductance of the mitochondrial membrane permeability transition pore (PTP), a nonspecific, high conductance ion-permeable channel in mitochondrial membranes. CsA induces cyclophilin to detach from the PTP (7) and thus facilitates closure. However, the mechanism(s) by which CsA causes gingival overgrowth is unknown (2).

Under physiological conditions, gingival connective tissues exhibit a remarkably rapid rate of collagen turnover (8). The abundant synthesis of collagen in these tissues is balanced by an equally rapid rate of intracellular degradation (9), a process that is mediated by the phagocytic pathway in fibroblasts (10). In CsA-induced gingival overgrowth, there is a net increase in collagen (11), but CsA does not increase collagen expression (12) and does not substantially alter collagenase levels (13). Instead, the net increase in collagen is apparently due to reduced phagocytosis by fibroblasts (11). Indeed, although CsA-induced inhibition of collagen phagocytosis has been demonstrated in several in vivo studies (14, 15), the mechanisms and the intracellular locus of this inhibition are not defined.

In "professional" phagocytes such as neutrophils, efficient phagocytosis is dependent on calcium release from intracellular stores (16), which, in turn, is reliant on inositol 1,4,5-trisphosphate receptor function to regulate calcium levels within endoplasmic reticulum (ER) stores (17). These receptors are translocated to periphagosomal sites during particle internalization (18), suggesting that phagocytosis may be affected by calcium levels in the intracellular stores. Notably, alpha 2beta 1 integrin-mediated collagen phagocytosis in gingival fibroblasts is regulated by intracellular calcium (19), in which a calcium-dependent feedback loop may alter integrin affinity of matrix molecules and thereby enhance integrin-mediated cell adhesion (20). Since CsA binding to cyclophilin perturbs the function of the PTP (6) and can independently inhibit calcium release from ER stores (5, 21), we considered that CsA may inhibit collagen phagocytosis by deregulating calcium homeostasis in mitochondrial and/or ER stores.

To obtain an improved understanding of the role of calcium signaling in phagocytosis and CsA-induced gingival overgrowth, we have used a well characterized in vitro model of alpha 2beta 1 integrin-mediated collagen phagocytosis in gingival and Rat2 fibroblasts (19, 22-24). Cells were stimulated with collagen beads, and the effect of CsA on intracellular calcium signaling in cytosolic, mitochondrial, and ER stores was examined. The data show that CsA inhibits the binding step of collagen phagocytosis through a calcium-regulated pathway involving ER and mitochondrial stores.


    EXPERIMENTAL PROCEDURES
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Reagents-- Carbonyl cyanide m-chlorophenylhydrazone (CCCP) and cyclosporin A were obtained from Sigma. Mouse anti-human cytochrome oxidase subunit I monoclonal antibodies, BAPTA/AM, fura-2/AM, JC-1 (5,5',6,6'-tetrachloro-1,1'3,3'-tetraethylbenzimidazolecarbocyanine iodide), mag-fura-2/AM, MitoTracker Green®, Pluronic® F-127, and rhod-2/AM were obtained from Molecular Probes, Inc. (Eugene, OR). Ionomycin and thapsigargin were obtained from Calbiochem. Fluorescent and nonfluorescent latex beads were obtained from Polysciences (Warrington, PA).

Cell Culture-- Human gingival fibroblasts (HGFs; passages 8-15) were derived from primary explant cultures as described earlier (25). Cells from passages 8-15 were grown in alpha -minimal essential medium, 15% (v/v) fetal bovine serum (Flow Laboratories, McLean, VA), and antibiotics. The cells were maintained at 37 °C in a humidified incubator containing 5% CO2 and were passaged with 0.01% trypsin (Life Technologies, Inc., Burlington, Ontario, Canada). Rat2 cells (ATCC CRL 1764, American Type Culture Collection, Manassas, VA) were incubated in Dulbecco's modified Eagle's medium (high glucose) containing 10% fetal bovine serum and antibiotics.

Mitochondrial Depletion-- Rat2 cells were grown in Dulbecco's modified Eagle's medium (high glucose) containing 10% fetal bovine serum and antibiotics as described above. Cells were grown in the presence of 100 ng/ml ethidium bromide for 30 passages before the dose was increased gradually to 1 µg/ml by passage 40. The ethidium bromide-treated Rat2 cells (Rat2EtBr) were always cultured at 1 µg/ml ethidium bromide for at least 4 weeks before using for experiments to maintain mitochondrial depletion as described (26).

[Ca2+]i Measurement-- For measurement of [Ca2+]i, cells on glass coverslips were incubated at 37 °C with 3 µM fura-2/AM in bicarbonate-free medium (Life Technologies, Inc.) for 25 min. The attached cells were washed twice with bicarbonate-free calcium buffer (145 mM NaCl, 5 mM KCl, 5 mM MgCl2, 10 mM D-glucose, 10 mM HEPES, and 1 mM CaCl2, pH 7.4; osmolarity = 291 mosM) and transferred to a microscope mounting stage. CaCl2 was omitted from the buffer solution where indicated. Measurements of [Ca2+]i were made on single cells using a Nikon Diaphot II inverted microscope optically interfaced to an epifluorescence spectrofluorometer and analysis system (Photon Technology International Inc., London, Ontario) operating on a 386SX personal computer. The dual-excitation fluorochrome fura-2 was excited at alternating (~100 Hz) wavelengths of 346 and 380 nm from dual monochromators with slit widths set at 2 nm. Emitted fluorescence was collected by a 40× quartz 1.30-NA oil-immersion Nikon Fluor objective, passed through a 520/30-nm barrier filter (Omega Optical Inc., Brattleboro, VT), and detected by a photomultiplier tube. A variable-aperture intrabeam mask was used to restrict measurements to single cells. Estimates of [Ca2+]i independent of the precise intracellular concentration of fura-2 were calculated from dual-excitation emitted fluorescence according to the equation of Grynkiewicz et al. (27) (i.e. [Ca2+]i = Kd × Sf2/Sb2 × (R - Rmin)/(Rmax - R).

For analysis of calcium in ER compartments, cells were incubated with 4.5 µM mag-fura-2/AM for 140 min at 37 °C as described (28). Mag-fura-2 measurements were made with the Photon Technology International instrument and single cell photometry. For analysis of mitochondrial calcium, Rat2 fibroblasts were loaded with 4.5 µM rhod-2/AM in 0.005% (v/v) Pluronic F-127 gel for 30-140 min at 37 °C. The cells were washed twice and incubated in calcium buffer for imaging. The magnitude of fluorescence of rhod-2-stained samples was observed in a Nikon inverted microscope equipped with a CCD camera (Pentamax, Princeton Research Instruments Inc., Princeton, NJ) and analyzed using the Winview software program (Princeton Research Instruments Inc.). As ratio imaging cannot be used to overcome problems of dye leakage and photobleaching in time course experiments with rhod-2, single excitation/single emission imaging analyses were conducted with a number of corrective procedures as described earlier (29). First, the fluorescence intensity of rhod-2 was measured in small sampling grids (~4 µm2) in the lamellipodia of well spread cells to avoid the inclusion of nucleoli, which stain brightly with rhod-2. Second, background fluorescence and adjustments for time-dependent photobleaching (29) were made for each measurement by calculating the bleach-induced rate constant (i.e. time-dependent loss of fluorescence) in untreated cell samples that were not incubated with collagen-coated beads. Third, for verification of the appropriate spatial localization of the rhod-2 staining and to ensure that rhod-2 was not sequestered to lysosomes (30), in some experiments, Rat2 fibroblasts were co-loaded with 100 nM MitoTracker Green and 4.5 µM rhod-2/AM in 0.005% (v/v) Pluronic F-127 for 30 min. The cells were washed twice and left in calcium buffer before imaging.

Flow Cytometric Analysis of Phagocytosis-- Green fluorescent microbeads (2 µm in diameter) were incubated with 1 ml of a 2.9 mg/ml acidic bovine collagen solution (Vitrogen, Celtrix Laboratory, Palo Alto, CA) neutralized with 1 N NaOH to produce collagen-coated beads (22). In some experiments, beads coated with either bovine serum albumin or poly-L-lysine were used as controls (19). Binding of beads was assessed by flow cytometry as described (22, 23). Cell samples were analyzed with a FACStar Plus flow cytometer (Becton Dickinson FACS Systems, Mountain View, CA). Photomultiplier tube voltage settings were determined for each experiment on the basis of thresholds established from negative and positive control samples for each sample that was analyzed. To reduce the likelihood of measuring cells with loosely attached, nonspecifically bound beads, the cells were washed with calcium- and magnesium-free phosphate-buffered saline and trypsinized prior to analysis by flow cytometry.

Analysis of Mitochondrial Membrane Potential (Psi m)-- Changes in Psi m were estimated using JC-1. This cyanine dye accumulates in the mitochondrial matrix under the influence of Psi m and forms J-shaped aggregates that have characteristic absorption and emission spectra (31). Untreated cells and cells treated with CsA were incubated in 3 ml of phosphate-buffered saline supplemented with 10% serum containing 0.5 µM JC-1 for 1 h. As a control for cells in which Psi m was dissipated, a group of cells were treated with the uncoupling agent CCCP before labeling with JC-1 (32). Cell suspensions were prepared for flow cytometry (33), and the 488-nm line of an argon ion laser was used for excitation. Orange and green emitted fluorescence was collected through 585/42-nm (FL2) and 530/30-nm (FL1) band-pass filters. Flow cytometry was performed on a FACStar Plus flow cytometer and analyzed using FACStation software (Becton Dickinson FACS Systems). After gating out noncellular debris, 10,000 cells were analyzed for each sample. Bivariate plots of FL2 versus FL1 and the computation of FL2:FL1 ratios were used to estimate Psi m as described (32, 34).

Western Blot Analysis-- Whole cell extracts were prepared by rinsing trypsinized cells with calcium- and magnesium-free phosphate-buffered saline containing protease inhibitors (0.5 µg/ml leupeptin, 0.5 µg/ml pepstatin, and 1 mM phenylmethylsulfonyl fluoride). Cells (5 × 106 cells/ml) were pelleted, solubilized for 30 min in calcium- and magnesium-free phosphate-buffered saline containing the above protease inhibitors plus 1.5% lauryl maltoside at 4 °C, and centrifuged for 20 min at 16,600 × g, and the supernatants were saved for biochemical analysis. Equivalent amounts of protein (by Bio-Rad assay) were separated on polyacrylamide gels and transferred electrophoretically to 0.2-µm nitrocellulose membranes. The blots were developed with chemiluminescent reagents and exposed to x-ray films, which were developed and analyzed.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Effect of CsA on Collagen Bead Phagocytosis-- For study of CsA regulation of collagen phagocytosis, we utilized an in vitro model in which collagen bead binding to gingival fibroblasts facilitates analysis of regulatory processes involved in collagen internalization (19, 22-24). To overcome the problems of phenotypic variability associated with primary isolates of gingival fibroblasts (35), we first rationalized the use of Rat2 fibroblasts, a stable and readily propagated cell line previously used for studies of collagen phagocytosis and that also exhibits many of the characteristic features of periodontal fibroblasts (24). We compared collagen bead-induced phagocytosis in HGFs and Rat2 fibroblasts by flow cytometry, a protocol that enables quantitative and unbiased evaluations of the bead binding step of phagocytosis (36). This integrin-dependent step of collagen phagocytosis is rate-limiting for subsequent steps involving intracellular collagen degradation (19, 22, 23). In HGFs, incubation with collagen-coated beads for 1 h produced a bead number-dependent relationship with phagocytosis (eight beads/cell, 70.2 ± 0.7% phagocytic cells; four beads/cell, 45.4 ± 1.2% phagocytic cells; and two beads/cell, 21.4 ± 0.5% phagocytic cells). At eight beads/cell, binding of BSA-coated beads was >4-fold less compared with collagen-coated beads (17.2 ± 1.4%; p < 0.001). In Rat2 cells, incubation with collagen-coated beads at two beads/cell for 1 h produced a result similar to that obtained with HGFs (19.0 ± 1.5%). In CsA-treated HGFs, addition of very low doses of CsA (10 nM) induced a large reduction (>2-fold; p < 0.05) in the mean percent of phagocytic cells and the number of beads/cell (p < 0.05) (Fig. 1A), which was also seen in Rat2 fibroblasts (19.0% in controls compared with 6.6 ± 0.7% after CsA; p < 0.001). When Rat2 cells were treated with increasing doses of CsA for 30 min prior to 1-h incubations with collagen-coated beads, we found that CsA doses resembling tissue levels obtained therapeutically in humans (37) produced a dose-dependent reduction of phagocytosis (Fig. 1B). These reductions of collagen bead binding were not due to direct interference of CsA in the binding of collagen to integrins because preincubation of collagen beads with equivalent concentrations of CsA followed by washing and subsequent incubations with cells and no CsA caused no significant inhibition of short-term (i.e. 5 min) bead binding.



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Fig. 1.   Effect of CsA on collagen bead binding. A, flow cytometric analysis of the binding step of collagen phagocytosis in human gingival fibroblasts after incubation with 10 nM CsA for 30 min. Note the marked reduction of collagen bead fluorescence (FL1-height) in the CsA-treated cells. n = 10,000 cells per analysis. B, quantification of means ± S.E. of percent collagen bead binding in Rat2 cells after 30-min preincubations with CsA at the indicated doses. CsA caused dose-dependent reduction of bead binding (p < 0.01; n = five separate cell samples/group).

Phagocytic Response and Calcium Signaling-- Integrin activation by extracellular matrix ligands stimulates calcium rises (38, 39) that may be important in regulating integrin affinity and the bead binding step of phagocytosis. Accordingly, we assessed the role of intracellular calcium signaling in the regulation of collagen bead-induced phagocytosis. The treatment of cells with thapsigargin (1 µM, 10 min) to inhibit store uptake and to prevent subsequent store release dramatically reduced the phagocytic capacity of Rat2 fibroblasts when incubated with collagen beads for 1 h (10.2-fold reduction, from 19.0 to 1.86 ± 0.45% of phagocytic cells; p < 0.01; a reduction that was comparable in size to the 8-fold decrease seen when Rat2 fibroblasts were pretreated with 10 µM CsA (p < 0.02); n = five samples/group). A similarly sized reduction of phagocytosis was obtained after pretreatment with BAPTA/AM, which chelates intracellular calcium (from 19.0 to 2.2 ± 0.3%; p < 0.01; n = five samples/group). These data indicate that intracellular calcium signaling may have an important regulatory function in collagen bead-induced phagocytosis.

We compared calcium signaling in HGFs and Rat2 cells after collagen bead stimulation to rationalize the use of Rat2 cells as surrogates for HGFs in studies of calcium signaling in phagocytosis. For all collagen, BSA, or poly-L-lysine bead incubations, a ratio of eight beads to one cell was used. In fura-2/AM-loaded cells, the response of [Ca2+]i to collagen bead-induced phagocytosis was similar (the mean [Ca2+]i rise above the base-line level in HGFs was 124 ± 31 nM, and that in Rat2 fibroblasts was 130 ± 9 nM; p > 0.2; n = 10 cells/group). The rise to maximum [Ca2+]i after collagen bead stimulation also proceeded over a similar time course (time to peak: HGFs, 105 s; and Rat2 cells, 110 s). In both types of cells, incubation with BSA- or poly-L-lysine-coated beads produced no significant rise in [Ca2+]i above the base-line levels (<5 nM; n = five cells). These data indicate that analogous to the phagocytosis data, Rat2 cells are a sensitive and ligand-specific model for study of collagen bead-induced calcium signaling in gingival fibroblasts.

Prior to measuring the impact of CsA on collagen bead-induced calcium signaling, we examined the effect of CsA alone on Ca2+ mobilization. In Rat2 fibroblasts, CsA induced rapid increases in [Ca2+]i, which returned to base-line levels within 100 s (Fig. 2a). The rapid removal of Ca2+ from the extracellular buffer (5 mM EGTA immediately before addition of CsA) completely abolished the CsA-induced cytosolic calcium transients, indicating the importance of extracellular calcium influx for the CsA mechanism of action (Fig. 2b). To determine whether these cytosolic calcium changes involved release of calcium from thapsigargin-sensitive stores, we pretreated cells with thapsigargin (1 µM) to deplete internal Ca2+ stores (Fig. 2c) and then added CsA (Fig. 2d). The increases in [Ca2+]i were reduced in comparison with control cells, in which thapsigargin stores were not depleted; there was a very slow subsequent reduction of Ca2+ after peak levels were attained (Fig. 2d), an expected result due to the reduced uptake of calcium into the ER. Mag-fura-2-loaded cells treated with CsA also showed no discharge of Ca2+ from the ER (Fig. 2e). These results are consistent with earlier data on CsA-induced calcium fluxes in LLC-PK1 cells (40) and indicate that influx of extracellular calcium is likely important for CsA-mediated effects on collagen phagocytosis.



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Fig. 2.   CsA regulation of calcium signaling in Rat2 cells. a, in fura-2-loaded cells, CsA (10 nM) induced a steep and rapid increase in intracellular calcium, which returned to base-line levels within 50 s in normal calcium-containing medium. b, cells switched to a medium containing 5 mM EGTA showed no calcium increase after CsA treatment. c, cells induced with thapsigargin (Tg; 1 µM) in normal calcium-containing medium showed rapid calcium flux, indicating emptying of ER stores. d, cells in normal medium preincubated with thapsigargin (1 µM) and then treated with CsA showed a large increase in intracellular calcium, but the calcium did not return to base-line levels. e, in cells loaded with mag-fura-2 to measure calcium in ER stores, CsA caused no change in calcium concentration when cells were in normal medium. These data are representative of five separate experiments for each treatment. Tx, pretreated.

Following incubation with collagen beads, there was a steep rise in [Ca2+]i, which dissipated within 100 s (Fig. 3, A and B, panel a). In experiments using CsA pretreatment, there was a large decrease in [Ca2+]i responses to collagen bead-induced phagocytosis (3.4-fold; p < 0.01; cells pretreated with 10 nM CsA for 1 h) (Fig. 3A). This effect may be due to CsA-induced inhibition of intracellular calcium store release. Accordingly, the Ca2+i levels were investigated after intracellular stores of calcium were depleted with 1 µM thapsigargin pretreatment or cytoplasmic calcium was chelated with 3 µM BAPTA/AM. For thapsigargin, the response to collagen bead-induced phagocytosis was reduced by >3-fold (Fig. 3A). Following calcium chelation with BAPTA/AM, the amplitude of the [Ca2+]i transients induced by collagen beads was reduced by >5-fold (Fig. 3). In comparison with vehicle controls, pretreatment with CsA reduced the amplitude of the collagen bead-induced calcium responses an additional 20 nM when cells were treated with thapsigargin, but there was no additional reduction conferred by CsA when cells were pretreated with BAPTA/AM. These data indicate that CsA inhibits the calcium responses induced by collagen bead binding to integrins.



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Fig. 3.   Collagen bead-induced calcium signaling in Rat2 cells. A, shown are histograms (means ± S.E.) of peak calcium amplitudes induced by collagen beads in cells treated with CsA (10 nM) or vehicle (untreated). In controls, CsA reduced the collagen bead-induced calcium peak by >3-fold (p < 0.01). In cells pretreated with thapsigargin (Tg), collagen bead-induced calcium signals were also reduced by >3-fold (p < 0.01), and CsA reduced the signal by an additional 20 nM. With BAPTA/AM pretreatment, the collagen bead-induced calcium signal was reduced by >6-fold (p < 0.01), and no additional reduction was produced by CsA. Data are means ± S.E. (n = five cells/group). B, Rat2 cells were loaded with fura-2/AM and subsequently incubated with collagen-coated beads (ccb). Panel a, in medium containing 1 mM Ca2+; panel b, bead-induced calcium signal in medium with reduced calcium (0.1 mM) suggesting a requirement of calcium influx for collagen bead-induced calcium changes. These individual traces are representative of five independent experiments.

We next studied the role of collagen-induced influx of extracellular calcium using culture medium in which nominal amounts of calcium (100 µM) were present. This approach was employed since the collagen receptor most abundantly expressed in these cells (alpha 2beta 1 integrin) (22) requires cations for ligand binding (e.g. Ca2+ and Mg2+) to maintain their active conformation (41). In low calcium medium, there was a small rise in [Ca2+]i after incubation with collagen-coated beads (Fig. 3B, panel b). These data indicate that both intracellular and extracellular calcium sources are important in the [Ca2+]i responses during the bead binding step of collagen phagocytosis and that CsA may affect both pathways.

To obtain an improved understanding of cellular Ca2+ homeostasis and the role of the ER calcium stores in response to collagen bead stimulation, we loaded fibroblasts with mag-fura-2/AM for 140 min according to published methods (42-44). Briefly, cells were incubated with mag-fura-2/AM for 140 min at 37 °C. In contrast with cells loaded for 30 min, which showed diffuse fluorescence (Fig. 4A, panel a), cells loaded for 140 min exhibited preferential accumulation in discrete organelles (panel b), and virtually no dye was released by subsequent digitonin permeabilization of the plasmalemma (panel c) (45). We treated mag-fura-2-loaded cells with ATP to determine whether the presumptive ER store-loaded mag-fura-2 was reporting changes in an inositol 1,4,5-trisphosphate-sensitive internal store (Fig. 4A, panel c). Mag-fura-2-loaded cells showed rapid reductions of ER calcium following ATP treatment, which were followed by a rapid store refilling back to base-line calcium levels. When cells were incubated with collagen-coated beads, there were rapid reductions of ER calcium, which preceded increases in cytosolic calcium measured by conventional fura-2 (Fig. 3B, panel a). Following incubation with BSA-coated beads, there was no significant change over 1000 s (Fig. 4B, panel b), indicating that the collagen bead response was indeed specific. Pretreatment with CsA reduced the base-line levels of ER Ca2+ and also inhibited the bead-induced reduction of ER [Ca2+] by >5-fold (Fig. 4B, panel c).



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Fig. 4.   Response of calcium in ER stores to collagen beads. A, in cells loaded with mag-fura-2 for 30 min, fluorescence was diffusely distributed throughout the cytoplasm (panel a). In contrast, when cells were loaded for 140 min, discrete fluorescence labeling identified the putative ER stores (panel b), which was confirmed by digitonin permeabilization (Dig) (panel c). Treatment with ATP caused rapid loss of calcium and subsequent refilling. B, in cells loaded with mag-fura-2 as described for panel b in A, addition of collagen-coated beads (CCB) caused a rapid calcium efflux from ER stores, followed by a slow refilling as measured by ratio fluorometry (panel a). Addition of BSA beads caused no significant change in the mag-fura-2 ratio (panel b). Pretreatment of cells with CsA (10 nM) reduced the base-line mag-fura-2 ratio and blunted the collagen bead-induced efflux of calcium (panel c). Treatment with CCCP (10 µm) alone caused no change in the mag-fura-2 ratio (panel d), and pretreatment with CCCP blocked the collagen bead-induced calcium efflux (panel e). The traces are representative of five independent experiments. Tx, pretreated.

Mitochondrial Function-- A role for mitochondria in the regulation of collagen bead-induced calcium signaling was suggested by experiments in which the protonophore CCCP (10 µM) completely blocked collagen bead-induced efflux of calcium from ER stores (Fig. 4, B, panels d and e). As CsA has also been shown to affect mitochondrial function (32, 34), which, in turn, could regulate phagocytic processes, we assessed if dissipation of the mitochondrial membrane potential with CCCP (10 µM) or inhibition of ATP formation with sodium azide (0.1%) would affect the collagen binding step in phagocytosis. After short-term (10 min) treatments with either CCCP or sodium azide followed by 1-h collagen bead incubations, both agents significantly reduced collagen phagocytosis to a level similar to that seen with 100 nM CsA (CCCP, >3-fold reduction, 5.5 ± 0.7%; sodium azide, >4-fold reduction, 3.5 ± 0.8%; and controls, 19.0%; p < 0.01). These data suggest that CsA may exert its effects on phagocytosis by regulating mitochondrial ion homeostasis. Accordingly, we investigated if the mitochondrial PTP was a candidate locus of action for CsA in Rat2 cells. To determine whether CsA was indeed affecting mitochondrial function, we assessed the ability of CsA to block the dissipation of Psi m, an effect that is stimulated by suspension culture of fibroblasts (34). Psi m was estimated in JC-1-loaded cells by ratio flow cytometry (32). Treatment of Rat2 fibroblasts in suspension with increasing concentrations of CsA was accompanied by an elevated ratio of JC-1 aggregates to monomers (untreated suspended cells, 1.3 ± 0.12; CCCP-treated controls, 0.013 ± 0.001; 10 nM CsA, 1.5 ± 0.08; 100 nM CsA, 2.9 ± 1.07; 1 µM CsA, 3.0 ± 0.08; and 10 µM CsA, 3.5 ± 0.06), indicating that CsA preserved Psi m as expected (32) and that CsA was affecting the function of the PTP.

Mitochondrial calcium changes were monitored with compartmentalized rhod-2. Double labeling with the vital mitochondrial dye rhod-2 (29) (Fig. 5A, panel a) and MitoTracker Green (panel b) demonstrated that rhod-2 fluorescence and MitoTracker Green were completely coincident in Rat2 cells (panel c), indicating that rhod-2 loading reported calcium changes associated with mitochondria and not with lysosomes (30). Comparison of rhod-2-loaded cells before (Fig. 5, A, panel d) and after (panel e) treatment with CCCP showed that dissipation of Psi m by CCCP caused loss of mitochondrion-specific fluorescence and the spread of diffuse fluorescence throughout the cytoplasm. When rhod-2-loaded cells were stimulated with collagen-coated beads alone, there were progressive and linear reductions of mitochondrial calcium that occurred over a 20-min interval, thereby exhibiting much slower kinetics than the discharge of calcium from the ER (Fig. 5B). The collagen bead-induced reduction of fluorescence was not due to photobleaching or dye loss, as these factors were already compensated for during the data acquisition step using previously described methods (29). Incubation with poly-L-lysine- or BSA-coated beads caused no substantial reduction of fluorescence over time (data not shown). Notably, pretreatment with CsA (10 or 100 nM) caused only small reductions (p > 0.2, not statistically significant) in the overall slope (i.e. kinetics) of calcium discharge from the mitochondria (Fig. 5B), although there was a statistically significant reduction (p < 0.05) at 10 min. This result may be due to a short-term effect of CsA on calcium discharge rates. Collectively, these data show that binding of collagen beads can induce the specific discharge of mitochondrial Ca2+, but that CsA, at doses that strongly inhibit phagocytosis, does not substantially alter the overall calcium discharge rates from mitochondria.



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Fig. 5.   Mitochondrial calcium. A, shown is the colocalization of the calcium indicator rhod-2 (panel a) and MitoTracker Green (panel b) in Rat2 fibroblasts. Panel c shows colocalization. Rat2 cells loaded with the mitochondrion-specific dye JC-1 showed discrete staining in mitochondria (panel d). After dissipation of Psi m with CCCP, JC-1 fluorescence was diffusely distributed throughout the cytoplasm (panel e). B, mitochondrial calcium (means ± S.E.) was estimated by fluorescence after adjustment for photobleaching and dye leakage as described under "Experimental Procedures." Note that following incubation with collagen-coated beads at time 0, there was a slow and progressive reduction of mitochondrial calcium. There was no significant difference in the rate of reduced fluorescence in CsA-treated cells and vehicle-treated controls (p > 0.2; n = 10 cells/group; slope for controls, -1.9 × 104 fluorescence units/min; slope for CsA-treated cells, -1.7 × 104 fluorescence units/min). Note that there was a statistically significant individual difference at 10 min between controls and CsA-treated cells (p < 0.05).

Previous reports have emphasized the spatially localized nature of the interaction between ER and mitochondrial calcium stores (29, 46). As mitochondria can regulate the release kinetics of calcium from ER stores (46), we considered that CsA may indirectly affect calcium signaling by virtue of local interactions with the ER stores (29). Accordingly, we reduced the numbers of potential ER/mitochondrial interactions in Rat2 cells using chronic treatment with ethidium bromide according to well established methods (26). Long-term monitoring of cells showed that this protocol did not significantly affect cell viability or the ability of the cells to be passaged in culture. After staining with JC-1 (0.5 µM) and rhod-2 (4 µM) to localize mitochondria (Fig. 6A, panels a and c), the depleted Rat2EtBr cells showed fewer JC-1-stained mitochondria and fewer organelles stained with rhod-2 (panels b and d). Immunoblotting also showed >3-fold less of the mitochondrial-specific protein cytochrome oxidase I in comparison with normal cells (panels e and f). These data indicate that the numbers of mitochondria/cell were reduced and that, correspondingly, potential ER/mitochondrial interactions were likely decreased. Similar to Rat2 cells without EtBr treatment (Fig. 5B), after collagen bead incubation, there was a linear reduction of mitochondrial calcium over 25 min in EtBr-treated cells, indicating that the reduction of mitochondrial numbers per se did not substantially affect the regulation of mitochondrial calcium by collagen bead phagocytosis. However, in contrast to normal Rat2 cells, CsA pretreatment of these cells induced a 2-fold reduction of mitochondrial [Ca2+] at base-line levels, and the discharge rate of calcium from the mitochondrial stores following collagen bead incubation was ~55% slower than from the untreated Rat2EtBr cells. These data indicate that when ER/mitochondrial interactions are curtailed by simply reducing the numbers of mitochondria, regulation of mitochondrial calcium is impaired. Notably, collagen phagocytosis in Rat2EtBr cells was reduced by 2-fold (from 19.0% in controls to 9.8% in Rat2EtBr cells), an effect that could be due in part to CsA causing deregulation of mitochondrial and ER calcium signaling. Collectively, these data indicate that mitochondria play an important but perhaps not absolutely central role in the collagen-induced calcium signaling that is required for collagen phagocytosis.



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Fig. 6.   Mitochondrial depletion and calcium signaling. A, Rat2 cells were chronically treated over 30 passages with ethidium bromide to reduce numbers of mitochondria and ER/mitochondrial interactions. In control cells stained with JC-1, mitochondria were brightly stained and numerous (panel a), whereas ethidium bromide-treated cells exhibited more sparse staining (panel b). rhod-2 fluorescence identified calcium in brightly staining organelles in normal Rat2 cells (panel c); but following ethidium bromide treatment, there was a substantial reduction in the number (but not the brightness) of individual rhod-2-stained mitochondria (panel d). Immunoblotting for the mitochondrial protein cytochrome oxidase I in Rat2 cells (panel e) and in mitochondrion-depleted cells (panel f) showed >3-fold reduced blot density. B, following collagen bead incubation, rhod-2 fluorescence decreased linearly over 30 min of sampling (-1.5 × 104 fluorescence units/min), but CsA treatment diminished this by 60% (-0.8 × 104 fluorescence units/min; p < 0.05). Data are means ± S.E. (n = 10 cells/group).



    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Although some of the modes of action of CsA as an immunosuppressant are understood (37), the mechanism(s) by which CsA induces gingival overgrowth is unknown (2). The main findings in this study are that the CsA-induced inhibition of collagen phagocytosis observed in vivo (11, 14, 15) can be replicated in the collagen bead model and that CsA mediates phagocytic inhibition by deregulating intracellular calcium signaling. In this study, we have considered previous observations that CsA binding to cyclophilin inhibits the conductance of the mitochondrial PTP (6) and can independently act upon ER stores to inhibit calcium release (5, 21). Furthermore, several functional interactions between the regulation of calcium in mitochondrial and ER calcium stores have been characterized (46) that underline the importance of communication between these organelles in calcium homeostasis (48). Accordingly, we have shown here that calcium signaling is an important regulatory mechanism of collagen bead phagocytosis and that CsA markedly perturbs calcium responses to collagen bead incubations. Collagen phagocytosis was strongly inhibited by clamping [Ca2+]i to low levels with BAPTA, by depleting intracellular stores with thapsigargin, or by treatment with pharmacologically relevant doses of CsA. These data implicate intracellular calcium homeostasis and CsA perturbation of collagen bead-induced calcium signaling as important target processes in CsA inhibition of collagen phagocytosis.

Phagocytosis and [Ca2+]i Signals-- When [Ca2+]i was monitored during collagen bead-induced phagocytosis, Rat2 fibroblasts exhibited a calcium peak within 150 s, a similar but more rapid response than that seen in the spreading of endothelial cells on fibronectin substrata (38) or after incubation of kidney epithelial cells with RGD-coated beads (20). This calcium response was evidently important for the collagen bead binding step of phagocytosis because chelation of intracellular calcium with BAPTA/AM or inhibition of ER stores with thapsigargin strongly inhibited phagocytosis. These results are consistent with the previous demonstration of a requirement for calcium fluxes in cells forming productive adherent interactions with beads (20). However, the important requirement of calcium release from ER stores that we have shown here is in marked contrast to previous data showing that matrix protein-coated beads induce calcium rises that are largely dependent on extracellular sources (20). In this context, the importance of ER calcium stores in collagen bead-induced phagocytosis was shown by direct measurement of ER calcium stores with mag-fura-2 and by thapsigargin pretreatment, in which collagen beads induced only a small calcium rise; this effect was seen in both Rat2 fibroblasts and Rat2 fibroblasts pretreated with CsA. The chelation of Ca2+i with BAPTA also caused a significant decrease in the [Ca2+]i response compared with untreated cells, and the magnitude of this response was similar to that following thapsigargin treatment. Notably and consistent with previous findings (20, 38), we found that [Ca2+]i responses to collagen phagocytic stimuli also involve entry of extracellular calcium and thus are not due solely to release from intracellular stores (47). However, in this study, we have concentrated on the role of CsA in regulating intracellular calcium responses to collagen beads since previous data showed that CsA can down-regulate calcium release from both ER (5, 21) and mitochondrial (48, 49) stores and thus is likely to suggest a mechanism for how CsA inhibits collagen phagocytosis (11, 14, 15).

ER and Mitochondrial [Ca2+] Responses to Collagen Phagocytic Stimuli-- Preincubation of Rat2 fibroblasts with CsA caused a rapid and significant inhibition of the calcium efflux from the ER in response to collagen beads. Previous data (38) suggest that a large part of the increase in [Ca2+]i following cell spreading on matrix proteins may be due to calcium release from intracellular calcium stores, and our direct measurements of ER [Ca2+] with mag-fura-2 show that ER stores are indeed a locus for this inhibition. Notably, CsA inhibits inositol 1,4,5-trisphosphate binding to its cognate receptors and inhibits calcium release from ER stores in macrophages (21); CsA can also alter the characteristics of the calcium release channel in the ER of cardiac cells (5). In view of these data, we suggest that an important mechanism for CsA inhibition of collagen phagocytosis is blocking calcium efflux from ER stores; this would also account for the greatly reduced [Ca2+]i following CsA pretreatment and collagen bead stimulation. The CsA-induced attenuation of this calcium signal may be sufficient to inhibit collagen binding to collagen receptors, perhaps by regulating the affinity of the alpha 2beta 1 integrin (20, 41).

Collagen bead-induced phagocytosis provoked a time-dependent and equivalent decrease in mitochondrial [Ca2+] in untreated and CsA-pretreated Rat2 cells. Since CsA greatly attenuated [Ca2+]i responses to collagen beads, it seems unlikely that the efflux of calcium from mitochondrial stores contributed significantly to the increased [Ca2+]i after bead incubation. This result was not because CsA had no effect on mitochondrial function: indeed, CsA pretreatment preserved Psi m following suspension challenge (34), indicating that the PTP was inhibited, as reported earlier (32). A more likely explanation for the lack of effect of CsA on mitochondrial calcium is that calcium ions can exit via other channels or pumps that are not affected by CsA (48). In view of these data, does mitochondrial regulation of calcium impact on collagen bead binding? Conceivably, and as suggested by recent data on ER/mitochondrial store interactions (29, 46), mitochondria may sense high local concentrations of calcium within the cell and act to buffer [Ca2+] (44). We investigated this possibility using mitochondrial depletion methods previously described for studies of ER/mitochondrial calcium store interactions (26). In mitochondrion-depleted cells, collagen bead-induced calcium efflux from mitochondria was inhibited by CsA, indicating that ER/mitochondrial regulation of calcium signaling is dependent on functional interactions between the two sets of organelles. Conceivably, the effect of CsA on mitochondrial regulation of calcium may serve to inhibit the explosive release of calcium from ER stores (46); when these processes are dampened, the phagocytosis-induced calcium signal in the cytosol is inhibited.

We are aware of the potent inhibition of the serine/threonine phosphatase calcineurin by CsA (50) and the possibility that CsA may modulate integrin affinity for collagen through a calcineurin pathway. However, in the context of this study, we note that FK506, which also potently inhibits calcineurin, has no effect on collagen phagocytosis and does not cause gingival overgrowth (51). Consequently, we conclude that the effect of CsA perturbation on the whole cell is independent of a calcineurin-mediated decrease in collagen binding and phagocytic efficiency. In the CsA-treated gingival fibroblast, this inhibition leads to decreased collagen phagocytosis, a net increase in matrix proteins, and gingival overgrowth. Although the exact role of the mitochondrial calcium stores is not shown by these data, the importance of signaling interactions between discrete calcium stores is evident.


    ACKNOWLEDGEMENT

We thank Milton Charlton for invaluable advice.


    FOOTNOTES

* This work was supported by Canadian Institutes of Health Research operating and group grants (to C. A. G. M.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger To whom correspondence should be addressed: University of Toronto, Fitzgerald Bldg., Rm. 244, 150 College St., Toronto, Ontario M5S 3E2, Canada. Tel.: 416-978-1258; Fax: 416-978-5956; E-mail: christopher.mcculloch@utoronto.ca.

Published, JBC Papers in Press, January 22, 2001, DOI 10.1074/jbc.M010298200


    ABBREVIATIONS

The abbreviations used are: CsA, cyclosporin A; PTP, permeability transition pore; ER, endoplasmic reticulum; CCCP, carbonyl cyanide m-chlorophenylhydrazone; BAPTA/AM, 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid acetoxymethyl ester; HGFs, human gingival fibroblasts; BSA, bovine serum albumin.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES


1. Thiru, S. (1989) in Cyclosporin, Mode of Action and Clinical Application (Thomson, A. W., ed) , pp. 324-359, Kluwer Academic Publishers, Lancaster, United Kingdom
2. Carranza, F. A., Jr. (1996) in Cinical Periodontology (Carranza, F. A., Jr. , and Newman, M. G., eds), 8th Ed. , W. B. Saunders Co., Philadelphia
3. Fomina, A. F., Fanger, C. M., Kozak, J. A., and Cahalan, M. D. (2000) J. Cell Biol. 150, 1435-1444[Abstract/Free Full Text]
4. Olbrich, H. G., Dols, S., Ver Donck, L., Kober, G., Kaltenbach, M., and Mutschler, E. (2000) Int. J. Cardiol. 27, 319-325
5. Park, K. S., Kim, T. K., and Kim, D. H. (1999) Am. J. Physiol. 276, H865-H872[Medline] [Order article via Infotrieve]
6. Crompton, M., Ellinger, H., and Costi, A. (1988) Biochem. J. 255, 357-360[Medline] [Order article via Infotrieve]
7. Nicolli, A., Basso, E., Petronilli, V., Wenter, R. M., and Bernardi, P. (1996) J. Biol. Chem. 271, 2185-2192[Abstract/Free Full Text]
8. Sodek, J. (1978) Arch. Oral Biol. 23, 977-982[Medline] [Order article via Infotrieve]
9. Sodek, J., and Ferrier, J. (1988) Collagen Relat. Res. 1, 11-21
10. Everts, V., van der Zee, E., Creemers, L., and Beertsen, W. (1996) Histochem. J. 28, 229-245[Medline] [Order article via Infotrieve]
11. McGaw, W. T., and Porter, H. (1988) Oral Surg. Oral Med. Oral Pathol. 65, 186-190[Medline] [Order article via Infotrieve]
12. Redlich, M., Greenfeld, Z., Cooperman, H., Pisanty, S., and Shoshan, S. (1997) Arch. Oral Biol. 42, 277-282[CrossRef][Medline] [Order article via Infotrieve]
13. Tipton, D. A., Stricklin, G. P., and Dabbous, M. K. (1991) J. Cell. Biochem. 46, 152-165[Medline] [Order article via Infotrieve]
14. Ayanoglou, C. M., and Lesty, C. (1999) J. Periodontal Res. 34, 7-15[Medline] [Order article via Infotrieve]
15. Kataoka, M., Shimizu, Y., Kunikiyo, K., Asahara, Y., Yamashita, K., Ninomiya, M., Morisaki, I., Ohsaki, Y., Kido, J. I., and Nagata, T. (2000) J. Cell. Physiol. 182, 351-358[CrossRef][Medline] [Order article via Infotrieve]
16. Stendahl, O., Krause, K.-H., Krischer, J., Jerström, P., Theler, J.-M., Clark, R. A., Carpentier, J. L., and Lew, D. P. (1994) Science 265, 1439-1441[Medline] [Order article via Infotrieve]
17. Bultynck, G., De Smet, P., Weidema, A. F., Ve Heyen, M., Maes, K., Callewaert, G., Missiaen, L., Parys, J. B., and De Smedt, H. (2000) J. Physiol. (Lond.) 3, 681-693
18. Favre, C. J., Jerstrom, P., Foti, M., Stendahl, O., Huggler, E., Lew, D. P., and Krause, K.-H. (1996) Biochem. J. 316, 137-142[Medline] [Order article via Infotrieve]
19. Arora, P. D., Manolson, M., Downey, G. P., and McCulloch, C. A. G. (2000) J. Biol. Chem. 275, 35432-35441[Abstract/Free Full Text]
20. Sjaastad, M. D., Angres, B., Lewis, R. S., and Nelson, W. J. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 8214-8218[Abstract]
21. Misra, U. K., Gawdi, G., and Pizzo, S. V. (1998) J. Immunol. 161, 6122-6127[Abstract/Free Full Text]
22. Lee, W., Sodek, J., and McCulloch, C. A. G. (1996) J. Cell. Physiol. 168, 695-704[CrossRef][Medline] [Order article via Infotrieve]
23. Knowles, G., McKeown, M., Sodek, J., and McCulloch, C. A. G. (1991) J. Cell Sci. 98, 551-558[Abstract]
24. Hui, M.-Z., Tenenbaum, H. C., and McCulloch, C. A. G. (1997) J. Cell. Physiol. 172, 323-333[CrossRef][Medline] [Order article via Infotrieve]
25. Pender, N., and McCulloch, C. A. G. (1991) J. Cell Sci. 100, 187-193[Abstract]
26. Biswas, G., Adebanjo, O. A., Freedman, B. D., Anandatheerthavarada, H. K., Vijayasarathy, C., Zaidi, M., Kotlikoff, M., and Avdhani, G. (1999) EMBO J. 18, 522-533[Abstract/Free Full Text]
27. Grynkiewicz, G., Poenie, M., and Tsien, R. Y. (1985) J. Biol. Chem. 260, 3440-3450[Abstract]
28. Hofer, A. M., Landolfi, B., Debellis, L., Pozzan, T., and Curci, S. (1998) EMBO J. 17, 1986-1995[Free Full Text]
29. Babcock, D. F., Herrington, J., Goodwin, P. C., Park, Y. B., and Hille, B. (1997) J. Cell Biol. 136, 833-844[Abstract/Free Full Text]
30. Trollinger, D. R., Cascio, W. E., and Lemasters, J. J. (2000) Biophys. J. 79, 39-50[Abstract/Free Full Text]
31. Smiley, S. T., Reers, M., Mottola-Hartshorn, C., Lin, M., Chen, A., Smith, T. W., Steele, G. D., Jr., and Chen, L. B. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 3671-3675[Abstract]
32. Zamzami, N., Marchetti, P., Castedo, M., Decaudin, D., Mach, A., Hirsch, T., Susin, S. A., Petit, X. A., Mignotte, B., and Kroemer, G. (1995) J. Exp. Med. 182, 367-377[Abstract]
33. Kulkarni, G. V., and McCulloch, C. A. G. (1995) J. Cell. Physiol. 165, 119-133[Medline] [Order article via Infotrieve]
34. Kulkarni, G. V., Lee, W., Seth, A., and McCulloch, C. A. G. (1998) Exp. Cell Res. 244, 170-178[CrossRef]
35. Narayanan, A. S., and Page, R. C. (1983) Collagen Relat. Res. 3, 33-64
36. McKeown, M., Knowles, G., and McCulloch, C. A. G. (1990) Cell Tissue Res. 262, 523-530[Medline] [Order article via Infotrieve]
37. Kay, J. E. (1989) in Cyclosporin, Mode of Action and Clinical Application (Thomson, A. W., ed) , p. 4, Kluwer Academic Publishers, Lancaster, United Kingdom
38. Schwartz, M. A. (1993) J. Cell Biol. 120, 1003-1010[Abstract]
39. Sjaastad, M. D., Lewis, R. S., and Nelson, W. J. (1996) Mol. Biol. Cell 7, 1025-1041[Abstract]
40. Gordjani, N., Epting, T., Fischer-Riepe, P., Greger, R. F., Brandis, M., Leipziger, J., and Nitschke, R. (2000) Pfluegers Arch. Eur. J. Physiol. 439, 627-633[CrossRef][Medline] [Order article via Infotrieve]
41. Onley, D. J., Knight, C. G., Tuckwell, D. S., Barnes, M. J., and Farndale, R. W. (2000) J. Biol. Chem. 275, 24560-24564[Abstract/Free Full Text]
42. Hofer, A. M., and Machen, T. E. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 2598-2602[Abstract]
43. Hofer, A. M., Curci, S., Machen, T. E., and Schulz, I. (1996) FASEB J. 10, 302-308[Abstract/Free Full Text]
44. Rizzuto, R., Brini, M., Murgia, M., and Pozzan, T. (1993) Science 262, 744-747[Medline] [Order article via Infotrieve]
45. Golovina, V. A., and Blaustein, M. P. (1997) Science 275, 1643-1648[Abstract/Free Full Text]
46. Landolfi, B., Curci, S., Debellis, L., Pozzan, T., and Hofer, A. M. (1998) J. Cell Biol. 142, 1235-1243[Abstract/Free Full Text]
47. Bengtsson, T., Jaconi, M. E. E., Gustafson, M., Magnusson, K.-E., Theler, J.-M., Lew, D. P., and Stendahl, O. (1993) Eur. J. Cell Biol. 62, 49-58[Medline] [Order article via Infotrieve]
48. Bernardi, P. (1999) Physiol. Rev. 79, 1127-1155[Abstract/Free Full Text]
49. Kristian, T., Gertsch, J., Bates, T. E., and Siesjo, B. K. (2000) J. Neurochem. 74, 1999-2009[CrossRef][Medline] [Order article via Infotrieve]
50. Rusnak, F., and Mertz, P. (2000) Physiol. Rev. 80, 1483-1521[Abstract/Free Full Text]
51. Thorp, M., DeMattos, A., Bennett, W., Barry, J., and Norman, D. (2000) Transplantation 69, 1218-1220[Medline] [Order article via Infotrieve]


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