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Role of the Cytoskeleton in Calcium Signaling in NIH 3T3 Cells

1997; Elsevier BV; Volume: 272; Issue: 42 Linguagem: Inglês

10.1074/jbc.272.42.26555

1083-351X

Carla M. P. Ribeiro, Jeffrey M. Reece, James W. Putney,

Neuroscience and Neuropharmacology Research

Treatment of NIH 3T3 cells with cytochalasin D (10 μm, 1 h at 37 °C) disrupted the actin cytoskeleton and changed the cells from a planar, extended morphology, to a rounded shape. Calcium mobilization by ATP or by platelet-derived growth factor was abolished, while the ability of thapsigargin (2 μm) to empty calcium stores and activate calcium influx was unaffected. Similar experiments with nocodazole to depolymerize the tubulin network yielded identical results. Platelet-derived growth factor induced an increase in inositol phosphates, and this increase was undiminished in the presence of cytochalasin D. Therefore, the blockade of agonist responses by this drug does not result from decreased phospholipase C. Injection of inositol 1,4,5-trisphosphate (IP3) released calcium to the same extent in control and cytochalasin D-treated cells. Confocal microscopic studies revealed a significant rearrangement of the endoplasmic reticulum after cytochalasin D treatment. Thus, disruption of the cytoskeleton blocks agonist-elicited [Ca2+] i mobilization, but this effect does not result from a lower calcium storage capacity, impaired function of the IP3 receptor, or diminished phospholipase C activity. We suggest that cytoskeletal disruption alters the spatial relationship between phospholipase C and IP3 receptors, impairing phospholipase C-dependent calcium signaling. Capacitative calcium entry was not altered under these conditions, indicating that the coupling between depletion of intracellular calcium stores and calcium entry does not depend on a precise structural relationship between intracellular stores and plasma membrane calcium channels. Treatment of NIH 3T3 cells with cytochalasin D (10 μm, 1 h at 37 °C) disrupted the actin cytoskeleton and changed the cells from a planar, extended morphology, to a rounded shape. Calcium mobilization by ATP or by platelet-derived growth factor was abolished, while the ability of thapsigargin (2 μm) to empty calcium stores and activate calcium influx was unaffected. Similar experiments with nocodazole to depolymerize the tubulin network yielded identical results. Platelet-derived growth factor induced an increase in inositol phosphates, and this increase was undiminished in the presence of cytochalasin D. Therefore, the blockade of agonist responses by this drug does not result from decreased phospholipase C. Injection of inositol 1,4,5-trisphosphate (IP3) released calcium to the same extent in control and cytochalasin D-treated cells. Confocal microscopic studies revealed a significant rearrangement of the endoplasmic reticulum after cytochalasin D treatment. Thus, disruption of the cytoskeleton blocks agonist-elicited [Ca2+] i mobilization, but this effect does not result from a lower calcium storage capacity, impaired function of the IP3 receptor, or diminished phospholipase C activity. We suggest that cytoskeletal disruption alters the spatial relationship between phospholipase C and IP3 receptors, impairing phospholipase C-dependent calcium signaling. Capacitative calcium entry was not altered under these conditions, indicating that the coupling between depletion of intracellular calcium stores and calcium entry does not depend on a precise structural relationship between intracellular stores and plasma membrane calcium channels. A variety of hormone and neurotransmitter agonists activate phospholipase C, promoting the breakdown of phosphatidylinositol 4,5-bisphosphate into two intracellular messengers: inositol 1,4,5-trisphosphate (IP3) 1The abbreviations used are: IP3, inositol 1,4,5-trisphosphate; PDGF, platelet-derived growth factor; DMEM, Dulbecco's modified Eagle's medium; PMA, phorbol 12-myristate 13-acetate; PBS, phosphate-buffered saline; DIOC6(3), dihexaoxacarbocyanine; DIC, differential interference contrast; (2,4,5)IP3, inositol 2,4,5-trisphosphate; PIP2, phosphatidylinositol 4,5-bisphosphate. and diacylglycerol (1Berridge M.J. Nature. 1993; 361: 315-325Crossref PubMed Scopus (6258) Google Scholar). IP3 binds to channel receptors located in the endoplasmic reticulum, promoting channel opening and increasing cytoplasmic calcium ([Ca2+] i ) as a consequence of depletion of Ca2+ sequestered within the lumen of the endoplasmic reticulum; diacylglycerol activates protein kinase C, an enzyme involved in the modulation of diverse cellular functions. Depletion of intracellular Ca2+ stores activates a pathway for Ca2+ influx (presumably mediated by a channel) across the plasma membrane, which has been termed capacitative Ca2+ entry (2Putney Jr., J.W. Cell Calcium. 1986; 7: 1-12Crossref PubMed Scopus (2165) Google Scholar). Numerous studies utilizing reagents that inhibit the Ca2+-ATPase responsible for Ca2+storage within the endoplasmic reticulum (e.g. thapsigargin) have demonstrated that Ca2+ store depletion provides a full and sufficient signal for activation of capacitative Ca2+entry (3Putney Jr., J.W. Bird G.S.J. Endocr. Rev. 1993; 14: 610-631Crossref PubMed Scopus (490) Google Scholar, 4Putney Jr., J.W. Capacitative Calcium Entry. Landes Biomedical Publishing, Austin, TX1997Crossref Google Scholar). However, the nature of the signal linking pool depletion to opening of the capacitative Ca2+ influx pathway remains unknown. Two general mechanisms have been proposed to explain how information is transferred from the endoplasmic reticulum to the plasma membrane (reviewed in Putney and Bird (3Putney Jr., J.W. Bird G.S.J. Endocr. Rev. 1993; 14: 610-631Crossref PubMed Scopus (490) Google Scholar) and Berridge (5Berridge M.J. Biochem. J. 1995; 312: 1-11Crossref PubMed Scopus (1051) Google Scholar)). One model describes a direct protein-protein association between the IP3 receptor in the endoplasmic reticulum and the capacitative entry channel: after IP3 binding and release of stored Ca2+, the IP3 receptor would undergo a conformational change, which allows for its interaction with the capacitative Ca2+ channel, resulting in its opening. In the other proposal, a diffusible messenger, released and/or formed as a consequence of Ca2+ pool depletion, would stimulate the Ca2+ influx pathway to open. The first model implies a close physical interaction between the endoplasmic reticulum and the plasma membrane; on the other hand, such a direct interaction would not be required for a diffusible factor to act. In a previous report, we provided indirect evidence for a physical interaction between IP3 receptors and the plasma membrane that involved the actin cytoskeleton, and we speculated that this interaction might be important in signaling capacitative calcium entry (6Rossier M.F. Bird G.S.J. Putney Jr., J.W. Biochem. J. 1991; 274: 643-650Crossref PubMed Scopus (123) Google Scholar). Therefore, the present studies were conducted to assess agonist- and thapsigargin-induced [Ca2+] i mobilization in NIH 3T3 cells under cytoskeletal-disrupting conditions. Whereas cytoskeletal disruption blocked agonist-induced Ca2+signaling, release of sequestered Ca2+ by thapsigargin and the resulting activation of capacitative Ca2+ entry were not affected, despite profound alterations in cellular structure and endoplasmic reticulum morphology. Thus, these data indicate that the coupling between Ca2+ pool depletion and capacitative Ca2+ entry does not depend on the cytoskeleton or on cell shape and suggest that signaling can occur without maintenance of a close proximity between intracellular stores and plasma membrane Ca2+ channels. NIH 3T3 cells were maintained at 37 °C under 5% CO2 in Dulbecco's modified Eagle's medium (DMEM) containing 10% fetal bovine serum, 5 mmglutamine, 50 units/ml penicillin, and 50 units/ml streptomycin. After 3 days in culture, subconfluent cells were passed at a dilution of 1:10. For measurements of [Ca2+] i, the cells were plated to subconfluence on glass coverslips 2 days before use. The cells were then incubated with fura-2/AM as described below. The attached cells were mounted in a Teflon chamber (Bionique, Saranac Lake, NY) and incubated in DMEM containing 1 μm fura-2/AM (Molecular Probes, Eugene, OR) for 15 min at 37 °C and 5% CO2. The cells were then washed and bathed in a HEPES-buffered physiological saline solution (HPSS, in mm: NaCl, 116; KCl, 5.4; MgSO4, 0.8; HEPES, 20, and glucose, 10) at room temperature for at least 20 min before Ca2+ measurements were made. The fluorescence of the fura-2-loaded cells was monitored with a photomultiplier-based system, mounted on a Nikon Diaphot microscope equipped with a Nikon 40× (1.3 N.A.) Neofluor objective. The fluorescence light source was provided by a Deltascan D101 (Photon Technology International Ltd., Monmouth Junction, NJ), equipped with a light path chopper and dual excitation monochromators. The light path chopper enabled rapid interchange between two excitation wavelengths (340 nm and 380 nm), and a photomultiplier tube monitored the emission fluorescence at 510 nm, selected by a barrier filter (Omega Optical, Brattleboro, VT). All experiments were carried out at 25 °C on a field of four to six cells. Calculation of [Ca2+] i was carried out as described previously (7Grynkiewicz G. Poenie M. Tsien R.Y. J. Biol. Chem. 1986; 260: 3440-3450Abstract Full Text PDF Google Scholar). NIH 3T3 cells (partially loaded with fura-2/AM to allow the measurement of baseline [Ca2+] i ) were microinjected with a solution containing 2 mm fura-2 and 1 mm or 1 μm (2,4,5)IP3 via a glass micropipette attached to a WPI PV830 Picopump (World Precision Instruments, New Haven, CT) as described previously (8Ribeiro C.M.P. Putney Jr., J.W. J. Biol. Chem. 1996; 271: 21522-21528Abstract Full Text Full Text PDF PubMed Scopus (57) Google Scholar). [Ca2+] i was measured immediately following microinjection as described above. Cultures were plated directly into culture medium containing 50 μCi/ml myo-[3H]inositol and grown to subconfluence in six-well plates. After 3 days, the medium was aspirated and cells were kept for 4 h in serum-free culture medium containing equivalent myo-[3H]inositol. During the final hour, cells were treated with vehicle or 10 μmcytochalasin D as described above. 100 ng/ml platelet-derived growth factor (PDGF) was added to the incubation medium during the last 5 min. The medium was aspirated, cells were quickly washed in Puck's A saline, and inositol polyphosphates were extracted into 1 ml of 6% perchloric acid + 5 mm EDTA + 250 μg/ml phytate on ice for 20 min. For separation by Dowex anion exchange columns, acid extracts were neutralized with 0.5 m KOH plus 9 mm sodium borax, salt was precipitated overnight and pelleted, and the sample supernatant was loaded onto 1-ml Dowex columns. The columns were first washed extensively with water to remove free [3H]inositol, and then [3H]inositol phosphates were eluted sequentially with ammonium formate/formic acid as described previously (9Sugiya H. Tennes K.A. Putney Jr., J.W. Biochem. J. 1987; 244: 647-653Crossref PubMed Scopus (66) Google Scholar) and subjected to liquid scintillation counting. Data were normalized to total cell [3H]inositol lipids as determined by a 0.1% Triton X-100, 0.1 mm NaOH extraction of the acid-precipitated cell monolayer. Subconfluent cells, plated on coverslips, were treated with vehicle (Me2SO), 10 μm cytochalasin D (60 min in DMEM at 37 °C and 5% CO2), or 10 μm nocodazole (30 min at 25 °C in HPSS buffer containing 1.8 mmCaCl2). For [Ca2+] i determination, following the cytochalasin D treatment cells were loaded with fura-2/AM as described above; for assessing the effect of disruption of the tubulin network on [Ca2+] i mobilization, cells were first loaded with fura-2/AM, followed by treatment with nocodazole as described. Cells, grown to subconfluence on coverslips, were mounted on Teflon chambers in the same manner as for [Ca2+] i determination and treated with Me2SO, 10 μm cytochalasin D, or 10 μm nocodazole, as described above, or 1.6 μm phorbol 12-myristate 13-acetate (PMA) for 60 min at 37 °C in DMEM and 5% CO2 as reported previously (8Ribeiro C.M.P. Putney Jr., J.W. J. Biol. Chem. 1996; 271: 21522-21528Abstract Full Text Full Text PDF PubMed Scopus (57) Google Scholar). Cells were subsequently washed with phosphate-buffered saline (PBS: 120 mm NaCl, 2.6 mm KCl, 8.1 mmNa2HPO4, 1.5 mmKH2PO4, pH 7.4) and fixed in PBS containing 2% formaldehyde and 0.1% glutaraldehyde for 15 min at 25 °C (all incubations and rinses were performed with PBS). Cells were rinsed three times, permeabilized with 1% Triton X-100 for 10 min at 25 °C, rinsed three times, blocked with 3% bovine serum albumin for 30 min at 37 °C, and washed three times. For staining the tubulin cytoskeleton, cells were incubated with a mouse monoclonal anti-β-tubulin antibody (Amersham Corp.) at a 1:250 dilution for 60 min at 37 °C, followed by three more washes. Cells were then incubated with a fluorescein-conjugated affinity-purified goat anti-mouse IgG (Kirkegaard & Perry Laboratories Inc.), diluted 1:10, for 30 min at 25 °C. For labeling the actin cytoskeleton, the primary antibody step was omitted, and cells were incubated with BODIPY-phalloidin (Molecular Probes) at a 1:25 dilution for 30 min at 25 °C. Cells were then rinsed three times, kept in PBS, and their actin and tubulin cytoskeletons were visualized as described below. After vehicle or cytochalasin D treatment as described above, cells were fixed in DMEM containing 0.625% glutaraldehyde for 10 min at room temperature. Cells were rinsed three times in HPSS buffer containing 1.8 mm CaCl2, and the endoplasmic reticulum was subsequently stained with the fluorescent dihexaoxacarbocyanine dye DIOC6(3) (250 ng/ml, 1 min at room temperature in HPSS buffer-containing 1.8 mmCaCl2) according to Terasaki (10Terasaki M. Methods Cell Biol. 1989; 29: 125-135Crossref PubMed Scopus (91) Google Scholar). A laser scanning confocal microscope (LSM 410, Carl Zeiss, Inc.) was used to study cell morphology, the actin and tubulin cytoskeletons, and the endoplasmic reticulum network. The differential interference contrast (DIC) transmission images were collected using the 488-nm line of a krypton-argon mixed gas laser (Omnichrome, Inc.). The objective lens used was a C Apo 40 × 1.25 numeric aperture water immersion lens. For the DIC/DIOC6(3) images, DIC was collected as just described while simultaneously exciting DIOC6(3) and collecting its emission with a band pass filter of 515–565 nm. At a pinhole setting of 10, a zresolution of 1 μm was obtained. For the actin and tubulin double-labeling images, the fluorescein (representing tubulin) and BODIPY (representing actin) were excited simultaneously with the 488- and 568-nm lines (respectively) of the laser while collecting fluorescein fluorescence with a 515–540-nm band pass filter and BODIPY fluorescence with a 610-nm long pass filter. For these experiments, a Plan Apo 100 × 1.4 numeric aperture objective lens was used with different pinhole settings for the two channels, resulting in az resolution of 0.8 μm for both images. All cells were imaged across a plane as close as possible to where they were attached to the coverslip. Representative cells are shown in the figures. All individually shown experiments (fluorescent stainings, Ca2+ tracings, and electron micrographs are representative of at least three separate experiments; the inositol phospholipid data depict the average of 10 experiments, performed in duplicate and subjected to analysis of variance. Results were considered statistically significant with p < 0.05. NIH 3T3 cells treated with vehicle, 10 μmcytochalasin D to depolymerize the actin microfilaments, or 10 μm nocodazole to disrupt the tubulin microtubules, were visualized by differential interference contrast microscopy (Fig.1). Control cells were characterized as being flat, extended, and elongated, expressing a typical fibroblast phenotype (Fig. 1 A); on the other hand, both cytochalasin D and nocodazole induced significant alterations in cellular morphology, causing the cells to assume a spherical shape (Fig. 1, B andC, respectively). To confirm that cytochalasin D and nocodazole induced the predicted cytoskeletal disruptions, a double-labeling fluorescence protocol was used to stain the actin and the tubulin cytoskeletons (Fig. 2). The filamentous actin, as evidenced by the presence of stress fibers (red staining), as well as the microtubules (green staining) were found to be intact and distributed throughout the cytoplasm in control cells (Fig. 2 A); the tubulin stain, however, was more prominent in the cell periphery. Pretreatment with cytochalasin D (Fig.2 B) or nocodazole (Fig. 2 C) specifically depolymerized either the stress fibers or the microtubules, respectively. In the cytochalasin D-treated cells, the actin bundles were replaced by a punctate accumulation of actin in the cytoplasm; on the other hand, in the nocodazole-treated cells the depolymerized tubulin redistributed toward the cell periphery and also accumulated within surface blebs. Although disruption of either the actin or the tubulin cytoskeletons caused the cells to round up, a specific feature of the cytochalasin D treatment was the formation of cellular processes radiating from the cytoplasm, whereas the nocodazole-treated cells expressed cell surface blebs. This laboratory has recently reported that activation of protein kinase C by PMA promotes cell shape alterations as well as changes in Ca2+ storage capacity in NIH 3T3 fibroblasts (8Ribeiro C.M.P. Putney Jr., J.W. J. Biol. Chem. 1996; 271: 21522-21528Abstract Full Text Full Text PDF PubMed Scopus (57) Google Scholar). Since protein kinase C activation can lead to actin depolymerization in some cells, including fibroblasts (11Rifkin D.B. Crowe R.M. Cell. 1979; 18: 361-368Abstract Full Text PDF PubMed Scopus (97) Google Scholar, 12Schliwa M. Nakamura T. Porter K.R. Euteneuer U. J. Cell Biol. 1984; 99: 1045-1059Crossref PubMed Scopus (243) Google Scholar, 13Hedberg K.K. Birrell G.B. Griffith O.H. Cell Regul. 1991; 2: 1067-1079Crossref PubMed Scopus (12) Google Scholar), we also examined the actin and tubulin cytoskeletons in PMA-treated NIH 3T3 cells. Fig. 2 D confirms that activation of protein kinase C in these cells leads to disruption of actin fibers. Unlike cytochalasin D, PMA causes accumulation of depolymerized actin in the cell periphery, toward membrane ruffles. The integrity of the microtubules was not affected by PMA treatment as previously reported (12Schliwa M. Nakamura T. Porter K.R. Euteneuer U. J. Cell Biol. 1984; 99: 1045-1059Crossref PubMed Scopus (243) Google Scholar).Figure 2Cytochalasin D or nocodazole treatment specifically disrupts the actin or the tubulin cytoskeleton, respectively, in NIH 3T3 cells. Cells were treated with vehicle (Me2SO, control; A), cytochalasin D (B), nocodazole (C), or PMA (D) and double stained for actin (red) or tubulin (green) as described under "Materials and Methods." A, Control fibroblasts express intact actin stress fibers and tubulin microtubules throughout the cytoplasm. B, cytochalasin D treatment promotes disruption of the stress fibers, with the depolymerized actin shown as a punctate cytoplasmic accumulation, but the microtubules are left intact.C, nocodazole disrupts the microtubules, resulting in accumulation of the depolymerized tubulin on the cell periphery and in surface blebs, without altering the stress fibers. D, in PMA-treated cells, the stress fibers are disrupted but the tubulin cytoskeleton is spared. However, contrary to the effect of cytochalasin D, the PMA-depolymerized actin accumulates in the cell periphery toward membrane ruffles.View Large Image Figure ViewerDownload Hi-res image Download (PPT) We next examined the effect of treatment with cytoskeletal-disrupting agents on [Ca2+] i mobilization. Only cells that expressed the spherical phenotype described above were selected for comparison with control fibroblasts. Fig.3 A depicts the Ca2+-mobilizing action of 1 mm adenosine triphosphate, ATP, an agonist that acts through purinergic receptors to couple to phospholipase Cβ activation. In control cells (Fig.3 A, solid line), ATP induced a rapid and transient increase in [Ca2+] i . This action was completely abolished in cells pretreated with 10 μmcytochalasin D (Fig. 3 A, broken line). Activation of phospholipase Cγ by 50 ng/ml PDGF led, after a lag period of few hundred seconds, to a Ca2+ mobilization of longer duration compared with ATP in control cells (Fig. 3 B, solid line) as previously reported (8Ribeiro C.M.P. Putney Jr., J.W. J. Biol. Chem. 1996; 271: 21522-21528Abstract Full Text Full Text PDF PubMed Scopus (57) Google Scholar). Disruption of the actin cytoskeleton with cytochalasin D also abolished the Ca2+response to PDGF (Fig. 3 B, broken line). Disruption of the tubulin network with nocodazole yielded similar results, with complete blockade of ATP- or PDGF-induced Ca2+ mobilization (Fig. 3, C and D, control, solid lines; nocodazole, broken lines). The failure of cytoskeleton-disrupted cells to respond to phospholipase C-activating agonists was not due to a loss of Ca2+ stores, because 2 μm thapsigargin released Ca2+ in cytochalasin D- or nocodazole-treated cells to the same extent as in control (Fig. 4, A andB, respectively); moreover, in cytochalasin D- or nocodazole-treated cells, the kinetics of activation and the magnitude of capacitative Ca2+ entry after thapsigargin-induced pool depletion were indistinguishable from those in control cells (Fig. 4,A and B, control, solid lines; cytochalasin D or nocodazole, broken lines). These findings indicate that inhibition of agonist-promoted Ca2+mobilization by cytoskeletal disruption does not result from a diminished Ca2+ pool storage capacity or capacitative entry or from increased Ca2+ buffering. In addition, they demonstrate that the signaling mechanism for capacitative Ca2+ entry is not rigorously dependent on cell morphology or on the integrity of the cytoskeleton.Figure 4Disruption of the actin or tubulin cytoskeleton does not block thapsigargin-induced [Ca2+] i mobilization. NIH 3T3 cells grown on coverslips were treated with Me2SO (control), 10 μm cytochalasin D or 10 μm nocodazole and cells were loaded with fura-2/AM as described under "Materials and Methods." To assess the capacity of the Ca2+ pool and the extent of Ca2+ entry in control, cytochalasin D or nocodazole-treated cells, 2 μm thapsigargin was added to nominally Ca2+ free HPSS buffer, followed by restoration of the extracellular Ca2+ to 1.8 mm where indicated. Panel A, control, solid line; cytochalasin D, broken line. Panel B, control, solid line; nocodazole, broken line.View Large Image Figure ViewerDownload Hi-res image Download (PPT) The failure of agonists to induce Ca2+ mobilization in cytoskeleton-disrupted cells could be accounted for by the following possibilities: (a) one or more of the steps involved in coupling cell surface receptor activation and phospholipase C stimulation may be affected by cytoskeletal disruption, the result being a failure of agonists to increase IP3 levels; (b) the activation of the IP3 receptor in the endoplasmic reticulum may be affected by cytoskeletal disruption, impairing IP3-induced pool depletion; (c) cytoskeletal disruption-induced cell shape changes may alter the spatial-temporal relationship between phospholipase C activation and Ca2+ pool depletion, resulting in diminished Ca2+ mobilization. To address the first possibility, PDGF-induced phospholipase C activation was investigated in control and cytochalasin D-treated cells. As shown in Fig.5, either PDGF or cytochalasin D pretreatment alone increased the levels of IP3 compared with control but, more importantly, when cells were pretreated with cytochalasin D, the effect of PDGF on IP3 production was not blocked and was, if anything, augmented. 2The data shown in Fig. 5 represent the IP3 fraction from Dowex separations of inositol polyphosphates, and thus include both the 1,4,5 and 1,3,4 isomers of IP3. It is unlikely that the increases due to PDGF, cytochalasin D, or the combination of the two represent significant net increases in cellular content of IP3; we have previously shown that the increase of this isomer is too small in NIH 3T3 cells to be statistically detectable against the sizable background of basal IP3, whether assayed by high performance liquid chromatography-resolved radiolabeled inositol polyphosphates, or be mass assay of IP3 (25Lobaugh L.A. Eisfelder B. Gibson K. Johnson G.L. Putney Jr., J.W. Mol. Pharmacol. 1996; 50: 493-500PubMed Google Scholar). We note that in addition to increases in the IP3 fraction from the Dowex columns, the fractions containing isomers of inositol diphosphate and of inositol tetraphosphate, were also significantly increased by PDGF, and in each case this was not prevented by treatment with cytochalasin D. This suggested that the coupling between agonist receptor activation and phospholipase C stimulation was not affected by cytoskeletal disruption. These results are in agreement with studies in mast cells showing that actin or tubulin depolymerization did not inhibit agonist-increased IP3 levels (14Tasaka K. Mio M. Fujisawa K. Aoki I. Biochem. Pharmacol. 1991; 41: 1031-1037Crossref PubMed Scopus (38) Google Scholar). The possibility that the function of the IP3 receptor could have been depressed by cytoskeletal disruption was addressed by experiments such as the one shown in Fig.6. Fig. 6 A (solid line) shows that microinjection of a single control NIH 3T3 cell incubated in nominally Ca2+-free buffer with a maximal Ca2+-mobilizing concentration (1 mm pipette concentration) of the nonhydrolyzable IP3 analogue, (2,4,5)IP3, led to a rapid and transient Ca2+mobilization as previously reported (8Ribeiro C.M.P. Putney Jr., J.W. J. Biol. Chem. 1996; 271: 21522-21528Abstract Full Text Full Text PDF PubMed Scopus (57) Google Scholar, 15Bird G.S.J. Rossier M.F. Hughes A.R. Shears S.B. Armstrong D.L. Putney Jr., J.W. Nature. 1991; 352: 162-165Crossref PubMed Scopus (138) Google Scholar); pretreatment with cytochalasin D did not affect this response (Fig. 6 A,broken line). Likewise, the Ca2+-mobilizing action of a submaximal concentration (1 μm pipette concentration) of (2,4,5)IP3 was the same in control and cytochalasin D-treated cells (Fig. 6 B, control, solid line; cytochalasin D, broken line). To confirm that the Ca2+ stores were only partially depleted by the lower concentration of (2,4,5)IP3, 2 μmthapsigargin was subsequently applied; again, there was no difference in the response to thapsigargin in cytochalasin D-treatedversus control cells (Fig. 6 B). Thus, the ability of the IP3 receptor to bind IP3 and release Ca2+ was not altered by cytoskeletal disruption. Since we found no evidence for alterations in the formation or action of IP3 under cytoskeletal-dismantled conditions, we tentatively concluded that the cell shape changes induced by cytochalasin D or nocodazole were altering the spatial relationship between phospholipase C and IP3 receptors, impairing phospholipase C-dependent Ca2+ signaling. Thus, we next investigated the effect of cytoskeletal depolymerization on the cellular distribution of the endoplasmic reticulum. To localize the endoplasmic reticulum in control and cytoskeleton-disrupted cells, this organelle was labeled in fixed cells with the fluorescent dihexaoxacarbocyanine dye, DIOC6(3), as previously reported (10Terasaki M. Methods Cell Biol. 1989; 29: 125-135Crossref PubMed Scopus (91) Google Scholar). By abolishing the mitochondrial membrane potential, cell fixation resulted in better labeling of the endoplasmic reticulum. Fig.7 illustrates the fluorescent staining of the endoplasmic reticulum (orange) superimposed with the differential interference contrast image (green) in control (left) or cytochalasin D-treated (right) cells (DIOC6(3) images were ∼ 1 μm thick). In control cells, the endoplasmic reticulum appeared to be spread throughout the cytoplasm, although in the interior regions of the cells some labeling of mitochondria undoubtedly occurs, confusing the interpretation of the images. However, in the flattened regions in the cell periphery, single tubule-like structures were observed and confirmed to correspond to ribosome-studded endoplasmic reticulum in electron micrographs (not shown). As illustrated in Fig. 1, disruption of the actin cytoskeleton caused the cells to round up, and thus the flattened cellular regions containing diffuse endoplasmic reticulum strands were retracted into the more compacted cells. Although it is not possible from such images to determine the precise relationship between endoplasmic reticulum and plasma membrane effectors, it is nonetheless clear that the shape of the endoplasmic reticulum is dramatically changed in cytochalasin D-treated cells, at the very least to accommodate the drastic changes in cell shape. Thus, it is possible that this severe rearrangement of endoplasmic reticulum membranes alters the spatial relationship between the site for phospholipase C generation of IP3 and IP3-sensitive sites on the endoplasmic reticulum. Previous work from this laboratory provided evidence that the IP3 receptors in the endoplasmic reticulum are linked to the plasma membrane through the actin cytoskeleton (6Rossier M.F. Bird G.S.J. Putney Jr., J.W. Biochem. J. 1991; 274: 643-650Crossref PubMed Scopus (123) Google Scholar). As a result, investigators from this laboratory (6Rossier M.F. Bird G.S.J. Putney Jr., J.W. Biochem. J. 1991; 274: 643-650Crossref PubMed Scopus (123) Google Scholar) and elsewhere (5Berridge M.J. Biochem. J. 1995