Uncoupling of ER-mitochondrial calcium communication by transforming growth factor-β

Pál Pacher, Kumar Sharma, György Csordás, Yanqing Zhu, György Hajnóczky


Transforming growth factor-β (TGF-β) has been implicated as a key factor in mediating many cellular processes germane to disease pathogenesis, including diabetic vascular complications. TGF-β alters cytosolic [Ca2+] ([Ca2+]c) signals, which in some cases may result from the downregulation of the IP3 receptor Ca2+ channels (IP3R). Ca2+ released by IP3Rs is effectively transferred from endoplasmic reticulum (ER) to the mitochondria to stimulate ATP production and to allow feedback control of the Ca2+ mobilization. To assess the effect of TGF-β on the ER-mitochondrial Ca2+ transfer, we first studied the [Ca2+]c and mitochondrial matrix Ca2+ ([Ca2+]m) signals in single preglomerular afferent arteriolar smooth muscle cells (PGASMC). TGF-β pretreatment (24 h) decreased both the [Ca2+]c and [Ca2+]m responses evoked by angiotensin II or endothelin. Strikingly, the [Ca2+]m signal was more depressed than the [Ca2+]c signal and was delayed. In permeabilized cells, TGF-β pretreatment attenuated the rate but not the magnitude of the IP3-induced [Ca2+]c rise, yet caused massive depression of the [Ca2+]m responses. ER Ca2+ storage and mitochondrial uptake of added Ca2+ were not affected by TGF-β. Also, TGF-β had no effect on mitochondrial distribution and on the ER-mitochondrial contacts assessed by two-photon NAD(P)H imaging and electron microscopy. Downregulation of both IP3R1 and IP3R3 was found in TGF-β-treated PGASMC. Thus, TGF-β causes uncoupling of mitochondria from the ER Ca2+ release. The sole source of this would be suppression of the IP3R-mediated Ca2+ efflux, indicating that the ER-mitochondrial Ca2+ transfer depends on the maximal rate of Ca2+ release. The impaired ER-mitochondrial coupling may contribute to the vascular pathophysiology associated with TGF-β production.

  • IP3 receptor
  • mitochondria
  • vascular smooth muscle cells
  • angiotensin II

transforming growth factor-β (TGF-β) has been closely linked to vascular smooth muscle cell growth and dysfunction (3, 10, 40). Several studies demonstrated anatomic and functional defects of arteriolar smooth muscle cells in diabetic tissues, including the kidney. A potential role for TGF-β in this context is the impairment of the normal mobilization of intracellular Ca2+ stores (1, 41). Prior studies demonstrated that the intracellular Ca2+ release channel, IP3R1, is reduced in diabetic aortic and preglomerular smooth muscle cells and the reduction is mediated by TGF-β (34). The IP3-linked cytoplasmic Ca2+ signaling is impaired in diabetic vascular smooth muscle cells and may thus contribute to vascular cell dysfunction.

Recently, it has been demonstrated that a key aspect of endoplasmic reticulum (ER) Ca2+ release is the Ca2+ coupling with the mitochondria (29, 31). In many cell types, Ca2+ mobilized through the IP3R and ryanodine receptors is effectively transferred to the mitochondria and stimulates in the mitochondrial matrix the Ca2+-sensitive steps of ATP production (11, 29). Furthermore, mitochondrial Ca2+ uptake exerts positive and negative feedback effects on the IP3R-mediated ER Ca2+ mobilization and affects the SERCA pump-mediated ER Ca2+ reuptake. In addition to these effects, Ca2+ uptake drives the mitochondrial phase of cell death (both apoptotic and necrotic) under Ca2+ overload or multistress conditions (2, 8, 12, 23). Thus, the mitochondrial Ca2+ uptake activated by Ca2+ released from the ER is essential for many aspects of cell function. In most cell types, mitochondria are closely associated with the ER and respond to the local [Ca2+] dynamics rather than to the global [Ca2+]c signal (6, 30). As TGF-β impairs IP3R-mediated calcium release, we investigated whether TGF-β may also affect the ER-mitochondrial communication pathway in preglomerular afferent arteriolar smooth muscle cells (PGASMC). Since severe suppression of the ER-mitochondrial Ca2+ transfer was found in the TGF-β-treated cells, we systematically evaluated the possible underlying mechanisms.


Cell isolation.

To isolate PGASMC from the renal resistance vessels, we used a technique previously described (38) for the rat kidney. Three normal Sprague-Dawley male rats (4 wk of age) were anesthetized with pentobarbital sodium (60 mg/kg ip), and the abdominal aorta was cannulated below the renal arteries. The kidneys were perfused with ice-cold PBS, followed by 5 ml of a magnetized iron oxide suspension (1% Fe3O4 in PBS), excised, and placed in fresh cold PBS, passed through needles of decreasing size (22- and 23-gauge), and filtered through a 120-μm sieve. The microvessels were recovered from the retentate and purified by magnetic separation. The final preparation was digested with collagenase (8 mg/10 ml type 1A; Worthington Biochemical, Lakewood, NJ) for 30 min with constant shaking at 37°C to disperse the cells and iron oxide. Cells of the digested microvessels were collected by brief centrifugation, washed once with PBS, and plated in RPMI-1640 medium supplemented with 10% heat-inactivated FCS, 5.5 mM d-glucose, 10 ml/l l-glutamine, penicillin (100 U/ml), and streptomycin (100 μg/ml; Cellgro). In all experiments, cells that were in passage 3–5 were used. Before biochemical and imaging analysis, cells were rested in 1% serum overnight and then treated with vehicle or TGF-β1 (50 nM for 24 h; R&D Systems). For imaging experiments, cells were plated onto poly-d-lysine-coated glass coverslips.

Western analysis.

Immunoblotting of PGASMC was performed by obtaining a protein homogenate with lysis buffer containing 50 mM Tris·HCl (pH 7.2), 150 mM NaCl, 1% (wt/vol) Triton X-100, 1 mM EDTA, 1 mM PMSF, and 5 μg/ml each of aprotonin and leupeptin. Protein concentration of samples was quantitated (Bio-Rad DC, Hercules, CA), and equal amounts of protein were run on a 7% SDS-PAGE gel, transferred to nitrocellulose, and immunoblotted with an antibody raised to the COOH terminus of the IP3R1 from brain, as previously described (35) or a murine monoclonal anti-NH2-terminal IP3R3 antibody (Transduction Laboratories). For standardization, the blots were stripped and immunoblotted with a monclonal antibody to β-actin (Sigma).

Loading of cells with fluorescent indicators.

Loading of PAGSMC with fluorescent dyes was performed in an extracellular buffer composed of 121 mM NaCl, 5 mM NaHCO3, 4.7 mM KCl, 1.2 mM KH2PO4, 1.2 mM MgSO4, 2 mM CaCl2, 10 mM glucose, and 10 mM HEPES/NaOH, pH 7.4, supplemented with 2% BSA. To monitor [Ca2+]c, cells were loaded with 5 μM fura2/AM for 20 min in the presence of 100 μM sulfinpyrazone and 0.3% pluronic acid at room temperature. For measurements of [Ca2+]m, cells were loaded with 4 μM rhod2/AM in the presence of 0.003% (wt/vol) pluronic acid at 37°C for 50 min. To simultaneously measure [Ca2+]c and [Ca2+]m, cells were first loaded with rhod2/AM and after washout of rhod2, incubation with fura2/AM was performed as described above. Intact cell measurements were performed in the extracellular buffer used for labeling with dyes except BSA was 0.25%. To visualize the mitochondria, cells were loaded with 50 nM Mitotracker Green (Molecular Probes) for 20 min after the rhod2 loading was completed. Loading of the cells with the dyes was regularly followed by 15 min after incubation to facilitate complete enzymatic processing of the accumulated probes.

For measurements of [Ca2+]m in permeabilized PGASMC, the rhod2-loaded cells were washed with Ca2+-free extracellular buffer composed of 120 mM NaCl, 20 mM Na-HEPES, 5 mM KCl, 1 mM KH2PO4, 100 μM EGTA/Tris at pH 7.4 and then permeabilized with 15–20 μg/ml digitonin for 4 min in intracellular medium (ICM) composed of 120 mM KCl, 10 mM NaCl, 1 mM KH2PO4, 20 mM Tris·HEPES, 2 mM MgATP at pH 7.2, supplemented with 1 μg/ml each of antipain, leupeptin, and pepstatin. All the measurements in permeabilized cells were performed in the presence of 2 mM succinate, 2 mM MgATP, and an ATP regenerating system composed of 5 mM phosphocreatine, 5 U/ml creatine kinase. After permeabilization, the cells were washed into fresh buffer without digitonin and incubated in the imaging chamber at 35°C. For monitoring of [Ca2+]c fluo3/FA (5 μM) was included in the buffer.

Microscopic imaging studies.

The fluorescence of the Ca2+-sensitive dyes was measured as described previously (25, 26, 37). Fluorescence images were acquired using an Olympus IX70 inverted microscope fitted with a ×40 (UApo, numerical aperture 1.35) oil immersion objective and a cooled CCD camera (PXL, Photometrics) under computer control. The computer also controlled a scanning monochromator (DeltaRam, PTI) to select the excitation wavelength. Excitation at 340 and 380 nm was used with a broad band emission filter passing 460–600 nm for measurements of [Ca2+]c with fura2. Fluorescence images of rhod2 were acquired using 545-nm excitation and 590-nm emission. Fluorescence images of fluo3 were acquired using 485-nm excitation and 520-nm emission. Dual dichroic/emission filter cubes were used to perform simultaneous measurements of two dyes. For confocal microscopy, we used standard fluorescein and rhodamine filter sets. Image pairs or triplets were acquired in every 2 or 3 s.

Confocal imaging of [Ca2+]c and [Ca2+]m and mitochondria were carried out using a BioRad MRC1024/2P imaging system equipped with a Kr/Ar-ion laser source (488- and 568-nm excitation) fitted to an Olympus IX70 inverted microscope (see Fig. 2B: ×40, UApo340, numerical aperture 1.35; Figs. 2A, 3, 4: ×60, PlanApo, numerical aperture 1.4 oil-immersion objectives). Rhod2 was excited at 568 nm, and fluo3 and Mitotracker Green at 488 nm. Two-photon (2P) imaging of NAD(P)H fluorescence was carried out using a pulsed femtosecond laser system (Millennia V/Tsunami, tuned to 720 nm, ≈80-fs pulses) and nondescanned detectors for recording the fluorescent signal. To calculate [Ca2+]c, fura2 fluorescence ratios were obtained in 50–100 cells and monitored in each experiment. The lag time was calculated as the time in seconds to attain half-maximal [Ca2+]c peak following agonist stimulation. Experiments were carried out with three different cell cultures, at least three parallel experiments on each occasion. The data are shown as means ± SE. Significance of differences from the relevant controls was calculated by Student's t-test.

Transmission electron microscopy.

For embedding a standard protocol was used (24). Briefly, cells were fixed using 2% glutaraldehyde, stained with 1% OsO4 and 0.5% uranyl acetate (UA; omitted for tomography samples), pelleted in 2% agarose (Sigma, Type IX ultra low gelling temperature), dehydrated in an acetone/water dilution series, and finally embedded in Spurr's resin (Araldite 6005 or 506 Epon 812-Polybed+DDSA 1:1:2.7 volumes catalyzed with DMP30 0.1 ml/5 ml resin, from Electron Microscopy Sciences). Ultrathin sections for transmission electron microscopy were poststained with UA and sodium bismuth (24). The sections were examined with a Hitachi 7000 scanning transmission electron microscope.

Micrographs were collected in all areas of the sections that showed mitochondria and ER. Before measurements, the shape and distribution of organelles were inspected in every micrograph and the organelles visible in more than one section were marked to avoid double counting of ER-mitochondrial interfaces. The minimum distance between mitochondrial outer membrane and the nearest ER membrane and the lengths of the ER-mitochondrial interface was measured using Image J for all mitochondria included in images.


Effect of TGF-β on [Ca2+]c in PGASMCs.

We first tested the effect of TGF-β pretreatment (50 nM for 24 h) on angiotensin II (AII)-induced [Ca2+]c responses in cultured PGASMC (Fig. 1, Table 1). Figure 1A shows AII-induced [Ca2+]c signals in naive and Fig. 1B in TGF-β-pretreated PGASMC measured with fura2. Top row of images and corresponding traces from four cells (marked with numbers) show that most of the naive cells responded to 2 nM AII by a rapid and large rise of [Ca2+]c and gradual decay (green-red shift shows [Ca2+]c rise), while in most of the TGF-β-pretreated cells this response was absent or largely delayed and attenuated (bottom row of images and corresponding traces from 4 cells). Cumulative addition of a supramaximal dose of AII (100 nM) evoked a [Ca2+]c rise both in naive and TGF-β-pretreated cells; however, in TGF-β-pretreated cells this response was also delayed (peak rise in control at 294 s, TGF-β at 324 s; see also traces in Fig. 1, A and B). TGF-β pretreatment also caused attenuation of the 100 nM AII-induced [Ca2+]c rise but this effect was more apparent when the cells were not preexposed to 2 nM AII (see Fig. 3B). In the combined experiments, 93.3% of naive PGASMC responded to 2 nM AII with a [Ca2+]c elevation, in contrast only 51% of the TGF-β-pretreated cells responded to 2 nM AII (Table 1). One hundred nanomolar AII evoked a [Ca2+]c response in 100% of naive and 92.8% of TGF-β-pretreated cells (Table 1).

Fig. 1.

Effect of transforming growth factor (TGF)-β pretreatment (50 nM for 24 h) on angiotensin II (AII)- and thapsigargin (Tg)-induced cytosolic Ca2+ signals in intact rat preglomerular afferent arteriolar smooth muscle cells (PGASMCs). A and B: [Ca2+]c increase is shown by a green to red shift in the overlay of the green (excited at 380 nm) and red (excited at 340 nm) fura2 fluorescence images. Cells were sequentially stimulated with 2 and 100 nM AII. Time courses of the fluorescence ratio of 340- and 380-nm excitations (R340/380) calculated for the single cells marked by the numbers are shown in the graphs. C: mean time courses of [Ca2+]c signal evoked by Tg (2 μM) in naive (red) and TGF-β-pretreated (green) PGASMC. The data are representative of experiments repeated at least 5 times.

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Table 1.

Effect of TGF-β on IP3-linked calcium signal

To determine whether the inhibition by TGF-β was upstream to IP3 formation, we evaluated the effect of TGF-β pretreatment on the [Ca2+]c rise evoked by endothelin (ET), another IP3-linked agonist. Forty-one of 42 control cells, but only 16 of 41 TGF-β-pretreated cells gave a [Ca2+]c signal in response to ET (25 nM; data not shown).

To test the possibility of whether the IP3-sensitive ER Ca2+ store was depleted by TGF-β, the effect of TGF-β pretreatment on the [Ca2+]c elevation evoked by thapsigargin (Tg), an inhibitor of the SERCA Ca2+ pumps, was studied. Tg (2 μM) elicited a gradual [Ca2+]c rise in 100% of both control and TGF-β-pretreated cells (n = 10 and 8 experiments, 503 and 382 cells, respectively). No difference in the kinetics of the Tg-induced [Ca2+]c rise appeared between control and TGF-β-pretreated cells (Fig. 1C). Ionomycin, a Ca2+ ionophore that also discharges the ER Ca2+ store, evoked comparable [Ca2+]c rises in both control and TGF-β-pretreated cells (n = 3, not shown).

The experiments described to this point demonstrated that both the AII- and ET-induced [Ca2+]c signaling was suppressed in TGF-β-pretreated intact cells. These results indicate that TGF-β likely targets a step in the IP3-linked [Ca2+]c signaling and this step is shared by both AII- and ET-activated pathways. Since TGF-β failed to affect the Tg- or ionomycin-induced [Ca2+]c rise, the effect of TGF-β does not seem to affect the total Ca2+ storage in the ER and in nonacidic Ca2+ pools.

Simultaneous measurements of [Ca2+]c and [Ca2+]m in PGASMC cells using rhod2.

To conduct simultaneous confocal imaging measurements of the [Ca2+]c and [Ca2+]m signals evoked by AII and ET in intact single PGASMC, the cells were loaded with rhod2/AM using a protocol that favors the compartmentalization of rhod2 in the mitochondria (see experimental procedures). The mitochondrial localization of rhod2 distribution was tested via coloading of the cells with a mitochondrion-specific probe, Mitotracker Green (Fig. 2). Mitotracker Green labeled both globular and tubular structures that were present at highest density in the perinuclear area (Fig. 2i). In unstimulated cells, the rhod2 fluorescence was hardly visible (Fig. 2ii) but the distribution of the weak signal was similar to the distribution of Mitotracker Green. In response to AII, a large increase in rhod2 fluorescence occurred (Fig. 2 bottom ii), which was colocalized with the Mitotracker Green fluorescence (overlay of Mitotracker Green and rhod2 images in Fig. 2 bottom iii), providing evidence that rhod2 was in the mitochondria and its fluorescence showed the [Ca2+]m. For the evaluation of [Ca2+]m, rhod2 fluorescence was taken from regions showing characteristic mitochondrial structure. A minor fraction of the rhod2 fluorescence was detected in the nuclear matrix (<10%). The rhod2 fluorescence in the nucleus provided information on the nuclear matrix [Ca2+] that closely follows the IP3-linked [Ca2+]c signal. In mitochondrial uncoupler pretreated cells, the agonist-induced rhod2 response was abolished in the mitochondrial region. Only a small transient in the nucleus was maintained, providing further evidence for the rhod2 compartmentalization (not shown).

Fig. 2.

Propagation of AII-induced [Ca2+]c signals to the mitochondria in intact PGASMCs. Cells were loaded with both Mito Tracker Green (i images, shown in green) and rhod2 (ii images, shown in red). Following stimulation with 100 nM AII, there is a large increase in rhod2 fluorescence (bottom ii image, shown in red), which is colocalized with MitoTracker Green (overlaid image iii, shown in yellow). Notably, the images shown at the bottom were taken 1–2 min after the stimulation. By then, the [Ca2+]c signal had already decayed, thus above the nuclear region no visible increase of rhod2 fluorescence appears.

Effect of TGF-β on propagation of the AII-evoked [Ca2+]c rise to the mitochondria in intact PGASMCs.

The spatial and temporal distribution of the AII-induced rhod2 signal is shown in Fig. 3 as difference images (increase in rhod2 fluorescence appears in purple) and the corresponding time course traces. In control cells, the agonist stimulation induced a steep rise in [Ca2+]c (green trace) that was closely followed by a rise in [Ca2+]m (red trace, Fig. 3A). While the rising phase of the [Ca2+]m was fast and synchronized to the rising phase of the [Ca2+]c signal, the decay of the [Ca2+]m was much slower as it was shown previously in other cells (14). Based on the summary of all experiments, the vast majority of the cells that exhibited a [Ca2+]c rise also showed a [Ca2+]m signal (Table 1) and the delay of the [Ca2+]m rise relative to the [Ca2+]c signal was 1.6 s for 100 nM AII, indicating a very effective Ca2+ delivery to the mitochondria (Table 2).

Fig. 3.

Effect of TGF-β pretreatment (50 nM for 24 h) on 2 and 100 nM AII-induced cytosolic and mitochondrial calcium signals. The cytosolic and mitochondrial calcium signals were measured in rhod2-loaded cells. Difference images (increase visualized in purple) and corresponding graphs show that in naive cells (A) stimulation evokes rapid cytosolic (green traces) and closely coupled mitochondrial (red traces) [Ca2+] signals, while in TGF-β-pretreated cells (B) there is a considerable delay and decrease both of cytosolic and mitochondrial [Ca2+] signals. [Ca2+]c traces were taken from the nuclear regions, and [Ca2+]m from regions showing mitochondrial structure after the stimulation. Note the difference in the time scale of the images of control and TGF-β-pretreated cells.

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Table 2.

Effect of TGF-β pretreatment on the lag time of [Ca2+]c and [Ca2+]m responses (measured at half height of the signal) and on the propagation time (coupling time) of the cytosolic signal to the mitochondria

Consistent with the results of the fura2 measurements, TGF-β pretreatment (50 nM for 24 h) substantially delayed and suppressed the [Ca2+]c responses evoked by AII (compare Fig. 3, B to A, images and green traces). However, the [Ca2+]m signal was even more depressed and delayed in the TGF-β-pretreated cells (compare Fig. 3, B to A, images and red traces). While 51% of the TGF-β-pretreated cells showed a [Ca2+]c signal, only 23.1% displayed a [Ca2+]m elevation (Table 1). The propagation time to the mitochondria was also longer (TGF-β-pretreated: 5.2 s vs. naive: 1.6 s for 100 nM AII; Table 2). Notably, TGF-β pretreatment did not affect the rhod2 distribution in the cells; after cell permeabilization, similar compartmentalized fluorescence intensities were retained in both naive and TGF-β-pretreated cells. TGF-β pretreatment also did not change the response of the compartmentalized rhod2 to Ca2+ as evidenced by the fluorescence increase evoked by elevation of the bulk cytosolic [Ca2+] in permeabilized cells.

Collectively, these data provide evidence that TGF-β pretreatment inhibits the calcium signal propagation to the mitochondria. Suppression of the [Ca2+]m signal may be a consequence of the attenuation of the [Ca2+]c rise but the depression of the mitochondrial response is larger than that in the cytosol. Thus, either a supralinear relationship exists between Ca2+ mobilization and mitochondrial Ca2+ uptake in the PGASMC or parallel to the control of IP3-induced Ca2+ release, TGF-β engages another mechanism to inhibit the delivery of the Ca2+ release to the mitochondria.

Effect of TGF-β on the IP3- and Ca2+-induced [Ca2+]c and [Ca2+]m rise in permeabilized PGASMCs.

To directly access the IP3Rs, we also established simultaneous single cell confocal measurements of the IP3-induced [Ca2+]c and [Ca2+]m signals in permeabilized PGASMC (Fig. 4). After cell permeabilization, the rhod2/AM-loaded cells retained only the compartmentalized rhod2 that was used for monitoring of [Ca2+]m and the incubation medium was supplemented with fluo3 to measure the [Ca2+]c signal. In these experiments, permeabilized PGASMCs were first stimulated with a supramaximal dose of IP3 (7.5 μM) and subsequently were exposed to an increase in bulk [Ca2+]c attained by the addition of 30 μM CaCl2. IP3 evoked a rapid [Ca2+]c rise in both naive and TGF-β-pretreated permeabilized PGASMC. In both conditions, the [Ca2+]c elevation appeared as a fairly homogeneous increase in fluo3 fluorescence to similar maximal intensities; however, the rate of rise was smaller in TGF-β-pretreated cells (Fig. 4, A and B, see the lower rows of difference images and corresponding graphs with the fluo3 fluorescence increase visualized in green). The IP3-induced [Ca2+]c rise in naive cells was closely followed by a large [Ca2+]m signal, whereas a barely detectable [Ca2+]m increase occurred in TGF-β-pretreated cells (Fig. 4, A vs. B, see top rows of difference images with increases in rhod2 fluorescence visualized in purple and corresponding graphs). The fluorescence increase evoked by 30 μM CaCl2 was similar in both naive and TGF-β-pretreated cells and was used for the normalization of the IP3-induced [Ca2+]c and [Ca2+]m responses when the mean responses were calculated. On average, in naive permeabilized PGASMCs, IP3 evoked 42.5 ± 8% (n = 9, 127 cells) increase in [Ca2+]m (normalized to the effect of 30 μM CaCl2), while in TGF-β-pretreated cells the [Ca2+]m was increased only by 5.78 ± 3.2% (n = 13, 171 cells). Thus, TGF-β treatment caused slower kinetic but unchanged volume of the IP3-induced Ca2+ mobilization and exerted substantial depression of the [Ca2+]m responses in permeabilized PGASMC.

Fig. 4.

Propagation of the IP3-induced [Ca2+]c signals to the mitochondria in permeabilized PGASMC. Bottom: difference images and corresponding graphs (green traces) show IP3-induced rapid [Ca2+]c rise (increase visualized in green) both in naive (A) and TGF-β-pretreated (50 nM for 24 h; B) permeabilized PGASMC. Top: difference images (increase visualized in purple) and corresponding graphs (red traces) show large mitochondrial [Ca2+] signals closely coupled to the IP3-induced [Ca2+]c rise in naive and only a small increase of [Ca2+]m in TGF-β-pretreated permeabilized cells.

To check whether an impairment of the mitochondrial Ca2+ uptake would be important for the decreased IP3R-mitochondrial coupling, in a separate set of experiments we measured and compared the mitochondrial uptake rates following 10 μM CaCl2 addition in naive and TGF-β-pretreated permeabilized cells. At the conclusions of these experiments, we also added 30 μM CaCl2 to achieve maximal [Ca2+]m responses. The rate of [Ca2+]m rise was 28.6 ± 3.1%/s (normalized to the increase evoked by 30 μM CaCl2) in naive and 30.6 ± 6.7%/s in TGF-β-pretreated (24 h) permeabilized PGASMCs. Thus, the mitochondrial uptake of added Ca2+ is not affected by TGF-β.

These findings narrowed down the potential sites of an inhibitory effect of TGF-β on the mitochondrial calcium signal to the processes of IP3-induced Ca2+ release and to the transfer of Ca2+ to the mitochondrial Ca2+ uptake sites. The mitochondrial Ca2+ uptake by itself seems to be preserved in the TGF-β-pretreated cells.

Effect of TGF-β on IP3R levels in PGASMCs.

Western blotting of cell lysates showed a marked reduction of IP3R1 and IP3R3 protein in TGF-β-treated (50 nM for 24 h) isolated PGASMC (Fig. 5). Complementing these results, real-time PCR data showed an approximate 40% decrease of the mRNA for both IP3R1 and IP3R3 in TGF-β-treated cells (L. Deelman and K. Sharma, unpublished data) similar to what was previously found in aortic smooth muscle cells and mesangial cells (33, 34). As both IP3R1 and IP3R3 have been demonstrated to preferentially transmit [Ca2+]c signals to the mitochondria (15, 22), the downregulation of the IP3Rs by TGF-β may provide the mechanism for the decrease in the IP3-induced [Ca2+]c signal. However, it remained possible that the suppression of the [Ca2+]m calcium signal involves additional factors. A change in the subcellular mitochondrial distribution or in their positioning relative to the ER could also contribute to suppression of the local Ca2+ transfer from IP3R to the mitochondrial Ca2+ uptake sites.

Fig. 5.

TGF-β-induced downregulation of type 1 and 3 IP3Rs in PGASMC. Cells were treated with TGF-β1 (50 nM) or vehicle for indicated periods before harvesting. Protein was resolved on a 10% SDS-PAGE and immunoblotted with antibody to IP3R1, IP3R3, and b-actin.

Effect of TGF-β on the mitochondrial morphology and ER-mitochondrial associations.

To elucidate whether TGF-β pretreatment affects mitochondrial morphology, we first compared the pattern of the NAD(P)H fluorescence in naive and TGF-β-pretreated PGASMC by 2P imaging (Fig. 6). NAD(P)H fluorescence appeared as globular and tubular structures and waned in the presence of a mitochondrial uncoupler (Fig. 6, right). The spatial distribution of the fluorescence did not show any clear difference between naive and TGF-β-pretreated cells (Fig. 6, A and B, high-magnification images on the right).

Fig. 6.

Two-photon imaging of the NAD(P)H in permeabilized naive and TGF-β-pretreated (50 nM for 24 h) PGASMC. The gray images show the NAD(P)H fluorescence in intact naive (A) and TGF-β-pretreated PGASMC (B) before and after the treatment (5 min) with an uncoupler, FCCP (5 μg/ml, right). The uncoupler was used to stimulate mitochondrial oxidation.

To evaluate whether the spatial relationship between ER and mitochondria at the sites of close associations between the organelles was altered in TGF-β-pretreated cells, transmission electron micrographs of both naive and TGF-β-pretreated cells were analyzed (Fig. 7). In the TGF-β-pretreated cells, the appearance of the mitochondria and ER and the distance between the organelles at the ER/mitochondrial contacts were similar to the controls (Fig. 7). Collectively, these experiments indicate that TGF-β does not target the mechanisms that determine the mitochondrial or ER distribution to uncouple mitochondria from the cytoplasmic calcium signaling. The electron micrographs also provided information on the intramitochondrial structure and did not indicate a change in the amount or size of the cristae in the TGF-β-pretreated cells (data not shown).

Fig. 7.

ER-mitochondrial ultrastructure in naive and TGF-β-treated (50 nM for 24 h) PGASMC. A: micrograph of naive and TGF-β-pretreated PGASMC. B: dimensions of the ER-mitochondrial interface. The average ER-mitochondrial distance (rough and smooth ER to OMM, surface to surface; top) and interface length (with ≤100-nm gap distance) determined from the electron micrographs of PGASMC (150 associations for each condition).


This work reveals that TGF-β causes a marked depression of the ER-mitochondrial calcium signaling in PGASMC. The effect is not due to altered Ca2+ sequestration by ER or mitochondria nor a change in the morphology or spatial relationship of ER and mitochondria. Rather, the only source of the ER-mitochondrial calcium uncoupling appears to be the downregulation of the IP3Rs, which inhibits the ER-mitochondrial Ca2+ transfer more efficiently than the Ca2+ mobilization from the ER. Thus, the results demonstrate a complex, nonlinear relationship between the cytoplasmic and mitochondrial calcium signal evoked by IP3-linked hormones in PGASMC. Since PGASMCs represent a cell type in which the contractile function is primarily dependent on calcium signaling and on a matching ATP supply, dysregulation of the recruitment of the mitochondria to the Ca2+ mobilization may serve as an important mechanism for the well-documented TGF-β-related loss of healthy vessel structure and function (10, 21).

Previous studies demonstrated changes in global cytoplasmic calcium signaling in TGF-β-treated cells (1, 20, 34, 35, 41). However, the effect of TGF-β on local Ca2+ transfer between ER and mitochondria and the ensuing changes in mitochondrial calcium signaling have not been previously studied. Ca2+-dependent control of both the mitochondrial metabolism and the apoptotic permeabilization is of particular significance in many target tissues of TGF-β (7, 12). The present calcium imaging studies revealed that mitochondria fail to respond to the attenuated Ca2+ mobilization mediated by the IP3Rs in TGF-β-pretreated cells. Since the [Ca2+]c signal was only delayed and partially attenuated, further clues were sought to the mechanism of the ER Ca2+ mobilization and to possible effects of TGF-β on mitochondrial structure and function. An effect of TGF-β on both ER and plasma membrane Ca2+ fluxes has been reported (20, 33). The present studies confirmed that TGF-β induced downregulation of IP3Rs in PGASMC. However, neither the amount of ER Ca2+ storage nor spontaneous Ca2+ leak was affected. Thus, the sole source of the attenuated [Ca2+]c signaling appears to be the decrease in the IP3R-mediated Ca2+ release. In addition to the lesser Ca2+ release, the severely depressed mitochondrial Ca2+ accumulation could also result from a TGF-β-induced change in mitochondrial metabolism (9, 39) or a change in the spatial relationship between ER and mitochondria (5, 30). However, mitochondrial accumulation of added Ca2+, the pyridine nucleotide redox, and the ER-mitochondrial morphology were not altered in TGF-β-treated PGASMC. Therefore, mitochondria seem to be competent to respond to ER Ca2+ release both in control and TGF-β-treated cells. Collectively, these results suggest that the decrease in the IP3R-mediated Ca2+ flux leads to lesser Ca2+ exposure of the mitochondria in TGF-β-treated PGASMC.

The data demonstrating the dependency of [Ca2+]m to intact [Ca2+]c rise parallel the observations that the IP3 dose-response relationship for the [Ca2+]m rise was rightward shifted compared with that for the simultaneously measured [Ca2+]c rise (6) and that enhanced intracellular IP3 buffering caused only moderate inhibition of the [Ca2+]c signal but effectively suppressed the [Ca2+]m signal (19). A decrease in the number of IP3Rs or in the activator molecules does not have to greatly dampen the Ca2+ release to interfere with Ca2+ transfer to the mitochondria. Presumably, even modest changes in the spatial and temporal organization of the Ca2+ release suppress the local [Ca2+]c signal sensed by the mitochondria. Indeed, we found that slightly slower mobilization of the entire IP3-sensitive Ca2+ store resulted in severe suppression of the Ca2+ transfer to the mitochondria. Notably, a recent study provided evidence that phosphorylation of the IP3R by Akt caused a modest decrease in the IP3R-mediated [Ca2+]c increase and essentially abolished the [Ca2+]m rise (36). Thus, posttranslational control of the IP3Rs may also affect the Ca2+ transfer to the mitochondria more so than the Ca2+ release from IP3Rs.

The greatly attenuated [Ca2+]m signal in TGF-β-treated cells may affect the Ca2+-dependent steps of energy metabolism in the mitochondrial matrix, which are controlled in the physiological range of the IP3-linked [Ca2+]m signal (17, 32). This pathway may be of great importance in smooth muscle where ATP production has to be continuously coordinated with the energy needs of contraction. Exposure to TGF-β may affect the delicate regulation of vascular tone and the response to autoregulatory challenges in afferent arteriolar smooth muscle cells (33) in progressive kidney disease, such as diabetic nephropathy. Failure of mitochondrial Ca2+ uptake would predispose mitochondria to fail to meet the ATP demand of the vascular cells. Furthermore, mitochondrial Ca2+ uptake in the vicinity of the IP3-induced Ca2+ release gives rise to important feedback effects on both IP3Rs and SERCA pumps (13, 16, 18). Since both positive and negative mitochondrial feedback effects on the IP3Rs have been described in various cell types, it is likely that suppression of the mitochondrial Ca2+ uptake may have marked effects on ER Ca2+ mobilization. The disturbance of these mechanisms can be particularly relevant in vascular smooth muscle, where regulation of the vascular tone depends on spatially and temporally coordinated local interactions between IP3Rs, mitochondria, and Ca2+-activated K+ channels (4, 27, 28).


This work was supported by a Juvenile Diabetes Research Foundation postdoctoral fellowship 3-2000-143 (P. Pacher) and by National Institutes of Health Grants R01-DK-053867 (K. Sharma) and DK-51526 (G. Hajnóczky).

Present address of P. Pacher: Section on Oxidative Stress and Tissue Injury, Laboratory of Physiological Studies, National Institutes of Health, National Institute of Alcohol Abuse and Alcoholism, Bethesda, MD 20892-9413.


We thank Dr. T. Taraschi and T. Schneider for help with the electron microscopy. We also thank Dr. S. K. Joseph for critical reading of the manuscript.


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