Am J Physiol Renal Physiol 295: F1303–F1312, 2008.
First published July 23, 2008; doi:10.1152/ajprenal.90343.2008.
Uncoupling of ER-mitochondrial calcium communication by transforming
growth factor-␤
Pál Pacher,1 Kumar Sharma,2,3 György Csordás,1 Yanqing Zhu,2 and György Hajnóczky1
1
Department of Pathology, Anatomy, and Cell Biology and 2Center for Novel Therapies for Kidney Disease, Department
of Medicine, Thomas Jefferson University, Philadelphia, Pennsylvania; and 3Translational Research in Kidney Disease,
University of California San Diego/VA San Diego Healthcare System, La Jolla, California
Submitted 3 June 2008; accepted in final form 21 July 2008
IP3 receptor; mitochondria; vascular smooth muscle cells; angiotensin II
TRANSFORMING GROWTH FACTOR-␤ (TGF-␤) 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-␤ 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-␤ (34). The IP3-linked cytoplas-
Address for reprint requests and other correspondence: K. Sharma, Translational Research in Kidney Disease, 9500 Gilman Drive, Stein Bldg., Rm 406
Univ. of California at San Diego, La Jolla, CA 92093-0711 (e-mail:
KumarSharma@ucsd.edu) or G. Hajnóczky, Dept. of Pathology and Cell
Biology, Rm 261, JAH Thomas Jefferson Univ., Philadelphia, PA 19107
(e-mail: Gyorgy.Hajnoczky@jefferson.edu).
http://www.ajprenal.org
mic 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-␤
impairs IP3R-mediated calcium release, we investigated
whether TGF-␤ 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-␤treated cells, we systematically evaluated the possible underlying mechanisms.
EXPERIMENTAL PROCEDURES
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-␮m 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 ␮g/ml;
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Pacher P, Sharma K, Csordás G, Zhu Y, Hajnóczky G. Uncoupling of ER-mitochondrial calcium communication by transforming
growth factor-␤. Am J Physiol Renal Physiol 295: F1303–F1312, 2008.
First published July 23, 2008; doi:10.1152/ajprenal.90343.2008.—
Transforming growth factor-␤ (TGF-␤) has been implicated as a key
factor in mediating many cellular processes germane to disease
pathogenesis, including diabetic vascular complications. TGF-␤ 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-␤ 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-␤ 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-␤ 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-␤. Also, TGF-␤ had no effect on mitochondrial
distribution and on the ER-mitochondrial contacts assessed by twophoton NAD(P)H imaging and electron microscopy. Downregulation
of both IP3R1 and IP3R3 was found in TGF-␤-treated PGASMC.
Thus, TGF-␤ causes uncoupling of mitochondria from the ER Ca2⫹
release. The sole source of this would be suppression of the IP3Rmediated 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-␤ production.
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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.
RESULTS
Effect of TGF-␤ on [Ca2⫹]c in PGASMCs. We first tested
the effect of TGF-␤ 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-␤-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-␤-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-␤-pretreated
cells; however, in TGF-␤-pretreated cells this response was
also delayed (peak rise in control at 294 s, TGF-␤ at 324 s; see
also traces in Fig. 1, A and B). TGF-␤ 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-␤-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-␤-pretreated cells (Table 1).
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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-␤1 (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 䡠 HCl (pH 7.2), 150 mM NaCl, 1% (wt/vol) Triton X-100, 1 mM
EDTA, 1 mM PMSF, and 5 ␮g/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% SDSPAGE 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
␤-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 ␮M fura2/AM for 20 min in the
presence of 100 ␮M sulfinpyrazone and 0.3% pluronic acid at room
temperature. For measurements of [Ca2⫹]m, cells were loaded with 4
␮M 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 ␮M EGTA/Tris at pH 7.4 and then permeabilized with
15–20 ␮g/ml digitonin for 4 min in intracellular medium (ICM)
composed of 120 mM KCl, 10 mM NaCl, 1 mM KH2PO4, 20 mM
Tris 䡠 HEPES, 2 mM MgATP at pH 7.2, supplemented with 1 ␮g/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 ␮M) 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.
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To determine whether the inhibition by TGF-␤ was upstream to IP3 formation, we evaluated the effect of TGF-␤
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-␤-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-␤, the effect of TGF-␤ pretreatment
on the [Ca2⫹]c elevation evoked by thapsigargin (Tg), an
inhibitor of the SERCA Ca2⫹ pumps, was studied. Tg (2 ␮M)
elicited a gradual [Ca2⫹]c rise in 100% of both control and
TGF-␤-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-␤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-␤-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-␤-pretreated intact cells. These results indicate that TGF-␤
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-␤ failed to affect the Tg- or ionomycin-induced [Ca2⫹]c rise,
the effect of TGF-␤ 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 mitochondrionspecific probe, Mitotracker Green (Fig. 2). Mitotracker Green
Table 1. Effect of TGF-␤ on IP3-linked calcium signal
Cells Showing (% of Total) 关Ca2⫹兴m Response
Cells Showing (% of Total) 关Ca2⫹兴c Response
Stimulus
Naive
TGF-␤-Pretreated
Naive
TGF-␤-Pretreated
All 2 nM # cells
All 100 nM # cells
93.3% (323/345, n ⫽ 23)
100% (178/178, n ⫽ 14)
51.0% (149/292, n ⫽ 21)
92.8% (141/152, n ⫽ 12)
80.8% (143/177)
97.5% (78/80)
23.1% (36/156)
76.2% (64/84)
Effect of transforming growth factor (TGF)-␤ pretreatment on the % of cells showing 关Ca2⫹兴c (measured with fura2 or rhod2) and 关Ca2⫹兴m responses
(measured with rhod2).
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Fig. 1. Effect of transforming growth factor (TGF)-␤ 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 ␮M) in naive (red) and TGF-␤-pretreated (green) PGASMC. The data are representative of experiments repeated at
least 5 times.
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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).
Effect of TGF-␤ 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
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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).
Consistent with the results of the fura2 measurements,
TGF-␤ 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-␤pretreated cells (compare Fig. 3, B to A, images and red traces).
While 51% of the TGF-␤-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-␤-pretreated: 5.2 s vs. naive: 1.6 s for 100 nM AII; Table
2). Notably, TGF-␤ pretreatment did not affect the rhod2
distribution in the cells; after cell permeabilization, similar
compartmentalized fluorescence intensities were retained in
both naive and TGF-␤-pretreated cells. TGF-␤ pretreatment
also did not change the response of the compartmentalized
rhod2 to Ca2⫹ as evidenced by the fluorescence increase
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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.
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evoked by elevation of the bulk cytosolic [Ca2⫹] in permeabilized cells.
Collectively, these data provide evidence that TGF-␤ 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-␤
engages another mechanism to inhibit the delivery of the Ca2⫹
release to the mitochondria.
Effect of TGF-␤ 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 perme-
abilization, 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 ␮M) and subsequently were exposed to an
increase in bulk [Ca2⫹]c attained by the addition of 30 ␮M
CaCl2. IP3 evoked a rapid [Ca2⫹]c rise in both naive and
TGF-␤-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-␤-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-␤-
Table 2. Effect of TGF-␤ 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
Lag Time of Response 关Ca2⫹兴c, s
Lag Time of Response 关Ca2⫹兴m, s
Coupling Time, s
Stimulus
Naive
TGF-␤-Pretreated
Naive
TGF-␤-Pretreated
Naive
TGF-␤-Pretreated
All 2 nM
All 100 nM
11.5⫾1.2
1.9⫾0.2
64.5⫾3.1
30.4⫾1.5
14.9⫾1
3.5⫾0.3
79.2⫾5.7
33.26⫾1.6
5.33⫾0.5
1.6⫾0.1
12.9⫾2.6
5.2⫾0.3
Values represent means ⫾ SE responses from 36 –166 cells.
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Fig. 3. Effect of TGF-␤ 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-␤-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-␤-pretreated cells.
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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 ␮M CaCl2 was similar in both naive and
TGF-␤-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 ␮M CaCl2), while in
TGF-␤-pretreated cells the [Ca2⫹]m was increased only by
5.78 ⫾ 3.2% (n ⫽ 13, 171 cells). Thus, TGF-␤ 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.
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 ␮M
CaCl2 addition in naive and TGF-␤-pretreated permeabilized
cells. At the conclusions of these experiments, we also added
30 ␮M 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 ␮M CaCl2) in naive and 30.6 ⫾ 6.7%/s in
TGF-␤-pretreated (24 h) permeabilized PGASMCs. Thus, the
mitochondrial uptake of added Ca2⫹ is not affected by TGF-␤.
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These findings narrowed down the potential sites of an
inhibitory effect of TGF-␤ 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-␤-pretreated cells.
Effect of TGF-␤ on IP3R levels in PGASMCs. Western
blotting of cell lysates showed a marked reduction of IP3R1
and IP3R3 protein in TGF-␤-treated (50 nM for 24 h) isolated
PGASMC (Fig. 5). Complementing these results, real-time
Fig. 5. TGF-␤-induced downregulation of type 1 and 3 IP3Rs in PGASMC.
Cells were treated with TGF-␤1 (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.
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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-␤-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-␤-pretreated permeabilized cells.
TGF-␤ AND MITOCHONDRIAL CALCIUM SIGNALING
To evaluate whether the spatial relationship between ER and
mitochondria at the sites of close associations between the
organelles was altered in TGF-␤-pretreated cells, transmission
electron micrographs of both naive and TGF-␤-pretreated cells
were analyzed (Fig. 7). In the TGF-␤-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-␤ 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-␤-pretreated cells (data not shown).
DISCUSSION
This work reveals that TGF-␤ 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
Fig. 6. Two-photon imaging of the NAD(P)H in permeabilized naive and TGF-␤-pretreated (50 nM for 24 h) PGASMC. The gray images show the NAD(P)H
fluorescence in intact naive (A) and TGF-␤-pretreated PGASMC (B) before and after the treatment (5 min) with an uncoupler, FCCP (5 ␮g/ml, right). The
uncoupler was used to stimulate mitochondrial oxidation.
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PCR data showed an approximate 40% decrease of the mRNA
for both IP3R1 and IP3R3 in TGF-␤-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-␤ 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.
Effect of TGF-␤ on the mitochondrial morphology and
ER-mitochondrial associations. To elucidate whether TGF-␤
pretreatment affects mitochondrial morphology, we first
compared the pattern of the NAD(P)H fluorescence in naive
and TGF-␤-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-␤pretreated cells (Fig. 6, A and B, high-magnification images on
the right).
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Fig. 7. ER-mitochondrial ultrastructure in naive and TGF-␤-treated (50 nM for 24 h)
PGASMC. A: micrograph of naive and TGF␤-pretreated PGASMC. B: dimensions of the
ER-mitochondrial interface. The average ERmitochondrial distance (rough and smooth ER
to OMM, surface to surface; top) and interface
length (with ⱕ100-nm gap distance) determined from the electron micrographs of
PGASMC (150 associations for each condition).
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-␤-related loss of
healthy vessel structure and function (10, 21).
Previous studies demonstrated changes in global cytoplasmic calcium signaling in TGF-␤-treated cells (1, 20, 34, 35,
41). However, the effect of TGF-␤ on local Ca2⫹ transfer between ER and mitochondria and the ensuing changes in mitochondrial calcium signaling have not been previously studied.
AJP-Renal Physiol • VOL
Ca2⫹-dependent control of both the mitochondrial metabolism
and the apoptotic permeabilization is of particular significance in
many target tissues of TGF-␤ (7, 12). The present calcium
imaging studies revealed that mitochondria fail to respond to the
attenuated Ca2⫹ mobilization mediated by the IP3Rs in TGF-␤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-␤ on
mitochondrial structure and function. An effect of TGF-␤ on both
ER and plasma membrane Ca2⫹ fluxes has been reported (20, 33).
The present studies confirmed that TGF-␤ induced downregulation of IP3Rs in PGASMC. However, neither the amount of ER
Ca2⫹ storage nor spontaneous Ca2⫹ leak was affected. Thus,
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ACKNOWLEDGMENTS
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.
AJP-Renal Physiol • VOL
GRANTS
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.
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