|
Published in Volume
119, Issue 5 (May 1,2009)
J. Clin. Invest.
119(5):
1275-1285 (2009).
doi:10.1172/JCI37829.
Copyright © 2009, American Society for Clinical
Investigation
Research Article
Regulation of mitochondrial dynamics in acute kidney injury in cell
culture and rodent models
Craig Brooks,
Qingqing Wei,
Sung-Gyu Cho and
Zheng Dong
Department of Cellular Biology and Anatomy, Medical College of Georgia,
and Charlie Norwood VA Medical Center, Augusta, Georgia, USA.
Address correspondence to: Zheng Dong, Department of Cellular Biology and
Anatomy, Medical College of Georgia, 1459 Laney Walker Blvd., Augusta, Georgia 30912,
USA. Phone: (706) 721-2825; Fax: (706) 721-6120; E-mail:
zdong@mail.mcg.edu. First published April 6, 2009
Received for publication October 21,
2008, and
accepted in revised form February 18,
2009.
The mechanism of mitochondrial damage, a key contributor to renal tubular cell death
during acute kidney injury, remains largely unknown. Here, we have demonstrated a
striking morphological change of mitochondria in experimental models of renal
ischemia/reperfusion and cisplatin-induced nephrotoxicity. This change contributed to
mitochondrial outer membrane permeabilization, release of apoptogenic factors, and
consequent apoptosis. Following either ATP depletion or cisplatin treatment of rat
renal tubular cells, mitochondrial fragmentation was observed prior to cytochrome
c release and apoptosis. This mitochondrial fragmentation was
inhibited by Bcl2 but not by caspase inhibitors. Dynamin-related protein 1 (Drp1), a
critical mitochondrial fission protein, translocated to mitochondria early during
tubular cell injury, and both siRNA knockdown of Drp1 and expression of a
dominant-negative Drp1 attenuated mitochondrial fragmentation, cytochrome
c release, caspase activation, and apoptosis. Further in vivo
analysis revealed that mitochondrial fragmentation also occurred in proximal tubular
cells in mice during renal ischemia/reperfusion and cisplatin-induced nephrotoxicity.
Notably, both tubular cell apoptosis and acute kidney injury were attenuated by
mdivi-1, a newly identified pharmacological inhibitor of Drp1. This study
demonstrates a rapid regulation of mitochondrial dynamics during acute kidney injury
and identifies mitochondrial fragmentation as what we believe to be a novel mechanism
contributing to mitochondrial damage and apoptosis in vivo in mouse models of
disease.
Introduction
Sublethal and lethal injury to renal tubular cells is a major intrinsic cause of acute
renal failure, a disease associated with high mortality and increasing prevalence (1–9). In this condition, mitochondrial damage has been recognized as a crucial
contributor to tubular cell death in both necrosis and apoptosis (1, 3, 10–14). Tubular
cell necrosis may involve disruption of respiration complexes, loss of mitochondrial
membrane potential, and mitochondrial permeability transition, while apoptosis is
precipitated by mitochondrial outer membrane permeabilization and consequent release of
apoptogenic factors such as cytochrome c. Recent in vivo studies have
shown that ablation of proapoptotic Bcl2 family genes, such as Bid and
Bax, leads to preservation of mitochondrial integrity, suppression
of tubular cell apoptosis, and amelioration of ischemic and cisplatin nephrotoxic renal
failure (15, 16). Notably, in humans, mitochondrial membrane permeabilization and release of
cytochrome c seem to be key to tubular cell apoptosis in ischemically
injured kidneys (17). Despite these findings, the
mechanism underlying mitochondrial damage during tubular cell apoptosis remains elusive.
A new development in the understanding of mitochondrial regulation in apoptosis is the
discovery of a drastic morphological change of the organelles (18, 19). Mitochondria are
dynamic organelles that maintain their shape or morphology via 2 opposing processes,
fission and fusion (20–22). Mitochondrial fission involves the constriction
and cleavage of mitochondria by fission proteins, such as dynamin-related protein 1
(Drp1) and Fission 1 (Fis1). Mitochondrial fusion, on the other hand, is the lengthening
of mitochondria by tethering and joining together 2 adjacent mitochondria. Mitofusin-1
and -2 are mainly responsible for outer membrane fusion, while Opa1 is thought to
mediate inner membrane fusion (20–22). Under
physiological conditions, mitochondria are elongated and filamentous. Upon stress or
apoptotic stimulation, mitochondria become fragmented and, importantly, the
fragmentation may contribute to mitochondrial outer membrane permeabilization and the
release of apoptogenic factors from the mitochondrial intermembrane space. Thus far,
mitochondrial fragmentation has been demonstrated in a variety of mammalian cells and
also during programmed cell death in Caenorhabditis elegans and
Drosophila (18, 19). A role for mitochondrial fragmentation in
apoptosis has been suggested by notable studies but has also been seriously challenged
recently by others (23–25). Critically, evidence for the occurrence of
mitochondrial fragmentation and its involvement in relevant pathological or disease
conditions is scarce (26).
In the current study, we have demonstrated compelling evidence for mitochondrial
fragmentation in acute kidney injury using in vitro and in vivo experimental models. The
fragmentation involves the activation of mitochondrial fission via Drp1. Importantly, in
both cell cultures and whole animals, suppression of Drp1 and mitochondrial
fragmentation abrogates mitochondrial damage, cytochrome c release,
apoptosis, and renal injury. Regulation of mitochondrial dynamics may offer a novel
strategy for the prevention and treatment of acute renal failure.
ResultsMitochondrial fragmentation occurs in response to apoptotic stress in rat
proximal tubular cells.
Renal ischemia and nephrotoxicity are the major causes of acute kidney injury. To
examine mitochondrial morphological changes under this condition, we transfected
MitoRed into cultured rat proximal tubular cells (RPTCs) to label mitochondria with
red fluorescence. The cells were subsequently subjected to azide-induced ATP
depletion to model in vivo ischemia or cisplatin treatment for nephrotoxicity. As
shown in Figure 1A, mitochondria in control
cells were filamentous with a tubular or thread-like appearance and were often
interconnected to form a network. During azide treatment, the mitochondrial network
broke down and the mitochondria were fragmented into short rods or spheres.
Mitochondrial fragmentation was azide-treatment time dependent, concurring with
apoptosis (Figure 1B). Cisplatin also induced
mitochondrial fragmentation in RPTCs in a time-dependent manner (Figure 1C). Notably, mitochondrial fragmentation during
cisplatin treatment clearly preceded apoptosis and, as a result, fragmentation was
observed at 4 hours, whereas apoptosis was not detected until 12 hours (Figure 1C). A time-lapse study further recorded
mitochondrial fragmentation during azide-induced cell injury. A very rapid/sudden
fragmentation of all mitochondria in individual cells was shown after about 2 hours
of azide treatment (Supplemental Video 1; supplemental material available online with
this article; doi:10.1172/JCI37829DS1). The results demonstrate an early and striking
morphological change of mitochondria during tubular cell injury.
Mitochondrial fragmentation can be inhibited by Bcl2 but not by caspase
inhibitors.
While mitochondrial fragmentation could play a causative role in mitochondrial injury
and apoptosis, it may also be a result of cell death. We therefore tested the effects
of carbobenzoxy-valyl-alanyl-aspartyl-(O-methyl)-fluoromethylketone (VAD), a
pan-caspase inhibitor that blocks apoptosis during ATP depletion and cisplatin
treatment of RPTCs. Although caspase activation and apoptosis were inhibited (not
shown), VAD did not suppress mitochondrial fragmentation during either azide or
cisplatin treatments (Figure 2, A and B),
suggesting that mitochondrial fragmentation is not secondary to caspase activation.
We further tested the effects of Bcl2 on mitochondrial fragmentation during RPTC
apoptosis. As shown in Figure 2, mitochondrial
fragmentation induced by azide and cisplatin was suppressed in Bcl2-transfected
cells. For example, azide treatment induced mitochondrial fragmentation in 70% of
RPTCs, which was reduced to 20% in Bcl2-overexpressing RPTCs (Figure 2A). Interestingly, Bcl2 but not VAD attenuated
azide-induced mitochondrial permeabilization, reflected by lower cytochrome
c release into cytosol (Figure 2C). The results suggest that Bcl2 may protect against mitochondria injury
and apoptosis in part by suppressing mitochondrial fragmentation.
Drp1 is activated and translocates to mitochondria early during azide-induced ATP
depletion.
The morphology of mitochondria is determined by 2 opposing processes, fission and
fusion (20–22). Accordingly, mitochondrial fragmentation observed in our
study could be a result of activated fission, suppressed fusion, or both. We
demonstrated the activation of Drp1, a fission protein, during azide treatment of
RPTCs. As shown in Figure 3A, compared with
untreated cells, 1–3 hours of azide treatment induced an accumulation of
Drp1 in the mitochondrial fraction. Azide-induced Drp1 translocation was also
confirmed by dual immunofluorescence staining of Drp1 and Fis1, an integral
mitochondrial protein involved in fission (20–22). As shown in
Figure 3B, colocalization in Drp1 and Fis1 was
shown in some azide-treated cells. Of note, consistent with previous reports (27, 28),
Drp1 appeared to be restricted to specific sites along the mitochondrial membrane,
where fission may have been occurring.
Dominant-negative Drp1 inhibits mitochondrial fragmentation during RPTC injury.
To determine whether Drp1 is indeed involved in mitochondrial fragmentation during
tubular cell apoptosis, we tested the effects of dominant-negative Drp1 (DN-Drp1)
that had a K38A point mutation (28). RPTCs
were transfected with DN-Drp1 or empty vector and then treated with azide or
cisplatin. As shown in Figure 4A, DN-Drp1
reduced mitochondrial fragmentation from 64% to 28% during azide treatment.
Similarly, DN-Drp1 significantly inhibited mitochondrial fragmentation during
cisplatin incubation (Figure 4B). Representative
images are shown in Figure 4C. Untreated cells
had typical filamentous mitochondria; after azide treatment, the mitochondria became
fragmented and punctate, but the cells transfected with DN-Drp1 retained their
filamentous mitochondria. These data suggest a role for Drp1 and associated fission
in mitochondrial fragmentation during tubular cell apoptosis.
Inhibition of cytochrome c release during RPTC injury by DN-Drp1.
Next, we determined whether blocking mitochondrial fragmentation with DN-Drp1 would
inhibit cytochrome c release. RPTCs were cotransfected with MitoRed
and DN-Drp1 or empty vector and then subjected to azide-induced ATP depletion.
Cytochrome c release was examined by immunofluorescence. Of note,
the transfection efficacy in RPTCs was 20%–30%. Thus, our examination was
focused on the transfected (MitoRed labeled) cells to determine the effects of
DN-Drp1. Representative images are shown in Figure 5A. In control cells, cytochrome c was sequestered in
mitochondria and overlapped with the MitoRed signal. After azide treatment,
mitochondria became fragmented and cytochrome c was released from
mitochondria and distributed throughout the cytosol. However,
DN-Drp1–transfected cells retained filamentous mitochondria and
maintained cytochrome c in the organelles. Cell counting showed that
DN-Drp1 significantly inhibited cytochrome c release during azide
treatment, reducing cytochrome c–released cells from 61%
to 26% (Figure 5B). In contrast, DN-Drp1 did not
suppress azide-induced Bax accumulation in mitochondria (Figure 5C). The results suggest that Drp1-mediated mitochondrial
fragmentation contributes to mitochondrial damage and cytochrome c
release, although it does not affect initial Bax activation or translocation.
Inhibition of RPTC apoptosis by DN-Drp1.
Once released, cytochrome c activates caspases to result in
apoptosis (29). By blocking mitochondrial
fragmentation and cytochrome c release (Figures 4 and 5), DN-Drp1 was
expected to inhibit caspase activation and apoptosis. To test this, we cotransfected
RPTCs with GFP and wild-type Drp1, DN-Drp1, or empty vector. The cells were then
subjected to azide treatment followed by recovery in normal medium to analyze
apoptosis and caspase activation. By TUNEL assay, azide induced 51% apoptosis in
vector-transfected cells, which was decreased to 27% in
DN-Drp1–transfected cells (Figure 6A). Consistently, DN-Drp1 suppressed the development of apoptotic morphology
in transfected (GFP-labeled) cells, including cellular shrinkage and nuclear
fragmentation (Figure 6B). DN-Drp1 also
suppressed caspase activation (not shown). In addition, DN-Drp1 reduced
cisplatin-induced apoptosis from 55% to 32% (Figure 6C).
siRNA knockdown of Drp1 blocks mitochondrial fragmentation, cytochrome c release,
and apoptosis.
We further confirmed the role of Drp1 in mitochondrial regulation and apoptosis by
using an RNA interference approach. Stable Drp1-siRNA cell lines, including R3 and
R24, were generated by transfection of RPTCs with a specific short hairpin siRNA of
Drp1 (30). Drp1 knockdown in R3 and R24 cells
was verified by immunoblot analysis (Figure 7A).
In response to azide treatment, both R3 and R24 cells showed significantly lower
mitochondrial fragmentation than parental RPTCs (Figure 7B). Moreover, these cells demonstrated less apoptosis and cytochrome
c release (Figure 7, C and
D). Cisplatin-induced mitochondrial fragmentation, cytochrome c
release, and apoptosis were also suppressed in these cells (data not shown).
Mitochondrial fragmentation and apoptosis in primary proximal tubular cells are
inhibited by DN-Drp1.
To further substantiate the role of mitochondrial fragmentation in renal injury, we
examined mitochondrial fragmentation during cisplatin treatment of primary proximal
tubular cells. Primary cultures of proximal tubular cells had highly filamentous
mitochondria (Figure 8A). Upon cisplatin
treatment, however, the mitochondria became fragmented and punctate (Figure 8A). The fragmentation was inhibited by DN-Drp1
(Figure 8A). Cell counting indicated that
DN-Drp1 reduced mitochondrial fragmentation from 55% to 31% (Figure 8B). DN-Drp1 also suppressed cisplatin-induced
apoptosis in the primary cells (Figure 8C). To
determine whether Drp1 suppression affects upstream signaling during cisplatin
treatment, we examined cisplatin-induced p53 phosphorylation and p53-upregulated
modulator of apoptosis α (PUMA-α) induction (31) in RPTCs stably transfected with Drp1 siRNA.
Downregulation of Drp1 in the siRNA cells was verified above, as shown in Figure
7A. As shown in Supplemental Figure 1, Drp1
siRNA cells showed p53 and PUMA-α induction similar to that of the
parental wild-type RPTCs, suggesting that Drp1 suppression does not affect upstream
signaling during cisplatin treatment. Together, these data strongly support a role
for Drp1-mediated mitochondrial fragmentation in tubular cell apoptosis.
Mitochondrial fragmentation occurs in vivo in proximal tubular cells following
renal ischemia.
To examine mitochondrial fragmentation in vivo, C57BL/6 mice were subjected to 30
minutes of bilateral clamping to induce renal ischemia followed by brief (15 minutes)
reperfusion. Kidneys were collected for EM. EM micrographs of proximal tubular cells
from both cortex and outer stripe of outer medulla were selected for evaluation. Due
to the orientation of mitochondria, proximal tubular cells from control animals
usually displayed 10%–20% long (≥ 2 μm)
filamentous mitochondria at the basal lateral side, whereas the perinuclear
mitochondria were cross-sectioned and thus appeared
“fragmented” (Figure 9A). After ischemia/reperfusion, however, many proximal tubular cells
completely fragmented their mitochondria into small, punctate suborganelles (Figure
9A). For quantification, we determined the
percentage of cells that had completely lost their long mitochondria. As shown in
Figure 9B, sham control had approximately 7%
proximal tubular cells with fragmented mitochondria, which was increased to 42%
during renal ischemia/reperfusion. To confirm mitochondrial fragmentation in ischemically injured tubular cells, we
performed 3D reconstruction of mitochondria from 100 serial section EM micrographs.
The control tubular cell showed many filamentous mitochondria at the basolateral side
in 2D EM (not shown); in the reconstruction, we purposely selected a perinuclear area
where mitochondria appeared “fragmented” (Figure 10A). The 3D image showed clearly that these
mitochondria were actually long and filamentous (Figure 10B). In sharp contrast, mitochondria in the ischemic cells were
completely fragmented (Figure 10, C and D).
These data indicate that mitochondrial fragmentation indeed occurs in vivo in renal
tubular cells during ischemic injury.
Amelioration of ischemic and cisplatin nephrotoxic renal injury and tubular
apoptosis by mdivi-1, a pharmacological inhibitor of Drp1.
Cassidy-Stone and colleagues have recently screened several chemical libraries and
identified mdivi-1 as an efficacious inhibitor of mitochondrial division that
operates by selectively inhibiting Drp1 (32).
To determine the role of Drp1 and mitochondrial fragmentation in vivo, we examined
the effects of mdivi-1 in the mouse model of ischemia/reperfusion (Figure 11). Ischemia/reperfusion induced a rapid loss of
renal function, as indicated by increases in serum creatinine and blood urea nitrogen
(BUN) (Figure 11, A and B), which was partially
but significantly reduced in animals pretreated with mdivi-1 (Figure 11, A and B). Consistently, mdivi-1 ameliorated
tubular damage in renal cortical and outer medulla tissues, as determined by
histological examination (Figure 11, C and D).
We further analyzed renal apoptosis by TUNEL assay. Cell counting showed that
ischemia/reperfusion induced 24 apoptotic cells/mm2 renal tissue in mice
that were pretreated with vehicle solution but only 11 in
mdivi-1–pretreated mice (Figure 11E). Using EM, we confirmed that mdivi-1 partially suppressed
ischemia-induced mitochondrial fragmentation in proximal tubular cells, from 43% to
32%. We further analyzed mitochondrial fragmentation in a mouse of cisplatin
nephrotoxicity (15, 33). It was shown that cisplatin (30 mg/kg) treatment for 3 and 4
days led to mitochondrial fragmentation in 33% and 53% of proximal tubular cells,
respectively. We then determined the effects of mdivi-1 on cisplatin-induced renal
injury and nephrotoxicity. As shown in Supplemental Figure 2, both BUN and serum
creatinine increases during cisplatin treatment were reduced by daily mdivi-1
injections. Consistently, mdivi-1 ameliorated renal tissue damage, especially in
renal tubules. Cisplatin-induced apoptosis was also decreased by mdivi-1
(Supplemental Figure 2). Collectively, the results support an important role for
Drp1-mediated mitochondrial fragmentation in the pathogenesis of both ischemic and
nephrotoxic acute kidney injury.
Discussion
Tubular cell apoptosis via the mitochondrial pathway contributes to ischemic as well as
nephrotoxic acute kidney injury (1, 3, 4, 6, 10, 11, 13, 14). The inference has been supported not only by in
vitro cell culture and in vivo animal studies but also by biopsy of ischemically injured
human kidneys (17). In this condition, Bax and
Bak are activated and form oligomers on mitochondria, leading to permeabilization of the
outer membrane and release of apoptogenic factors such as cytochrome c.
Despite these findings, it is not entirely clear how the outer membrane of mitochondria
is permeabilized. In this study, we have revealed a striking morphological change of
mitochondria in both in vitro and in vivo experimental models of acute kidney injury.
Mitochondria become fragmented early during renal cell injury. Importantly, prevention
of mitochondrial fragmentation by use of genetic and pharmacological approaches
abrogates mitochondrial damage, tubular cell apoptosis, and renal injury.
A role for mitochondrial fragmentation in apoptosis has been suggested and supported by
notable studies (18); however, it has also been
seriously challenged by others (24, 34). For example, the latest work by Sheridan et al.
showed that Bcl-xL and Mcl1, 2 antiapoptotic Bcl2 family proteins, can antagonize
cytochrome c release without blocking mitochondrial fragmentation in
HeLa cells, suggesting a separation of the morphological change from mitochondrial
damage (25). Breckenridge et al. further
demonstrated that regulation of mitochondrial morphological dynamics does not have a
significant role in the regulation of programmed cell death in C.
elegans (23), disputing the previous
finding by Jagasia et al. (35). The contrasting
results may certainly be caused by differences in the experimental conditions or models.
While the controversy may not be solved easily, a critical question is whether
mitochondrial fragmentation occurs in vivo under disease conditions and how much it
contributes to pathological apoptosis. Unfortunately, examination of mitochondrial
fragmentation in vivo in mammalian tissues is not a trivial task. Thus far, the only
reported examination is from Barsoum et al., who showed that mitochondria were
fragmented in neurons in ischemically injured brain (26). However, whether the fragmentation is important to neuronal apoptosis in
the brain was not determined. Our current study has demonstrated convincing evidence for
mitochondrial fragmentation in kidneys following ischemia/reperfusion by using 2D and 3D
EM. Importantly, we have further shown that pharmacological blockade of mitochondrial
fission can attenuate tubular cell apoptosis, tissue damage, and renal injury,
demonstrating what we believe is the first in vivo evidence for a role of the regulation
of mitochondrial morphological dynamics in pathological apoptosis in disease models.
Of note, in many tissue types, mitochondria are randomly oriented and, as a result,
tissue sectioning for EM results in cross sections of most mitochondria within a cell.
These mitochondria would then appear to be small and round, difficult to distinguish
from fragmented mitochondria in 2D EM. In this regard, tubular cells in the kidneys make
an excellent model for studying mitochondrial fragmentation in vivo because in these
cells a significant portion of mitochondria line up perpendicular to the
basement membrane and, as a result, cross sections of tubules would normally reveal
10%–20% longitudinally sectioned long mitochondria in a given cell. In
contrast, when mitochondria are fragmented, they appear as short rods or spherical
fragments, no matter how they are sectioned. The unique feature of mitochondrial
organization in renal tubular cells not only makes it easier to count and quantify
mitochondria-fragmented cells but also ensures the reliability of the results. In this
study, we have further verified the 2D EM results by 3D reconstruction of serial section
EM images.
The morphology of mitochondria is determined by a balance between fission and fusion
(20–22). Thus, fragmentation of the organelles can be a result of
increased fission, suppressed fusion, or a combination of both. In this study, during
renal tubular cell injury, we detected an early activation and mitochondrial
translocation of Drp1, a critical fission protein. Moreover, suppression of Drp1 by
dominant-negative mutants, siRNA, or mdivi-1 can block mitochondrial fragmentation.
These results, though not excluding a role for mitochondrial fusion regulation, suggest
that mitochondrial fission is activated under experimental conditions and contributes to
the observed mitochondrial fragmentation. It is unclear how Drp1 is activated.
Posttranslational modification of Drp1 including phosphorylation and sumoylation has
been documented and may play a role in Drp1 activation upon apoptotic stress (36, 37).
Further investigations should examine these possibilities during renal cell injury. In
addition, Drp1 is also involved in the fission of peroxisomes (38, 39). Whether and to what
extent Drp1-related changes in peroxisomes contribute to mitochondrial damage and
apoptosis has yet to be determined.
It is currently unclear how mitochondrial fragmentation, a seemingly morphological
change, can have an impact on mitochondrial membrane permeabilization (18, 19). It
is clear from previous studies that mitochondrial fragmentation alone is not effective
or sufficient to trigger porous defects in the outer membrane (18, 19). In support, our
recent work showed that although all apoptotic cells have fragmented mitochondria, not
all mitochondria-fragmented cells have cytochrome c release or
apoptosis (40). We further showed that fragmented
mitochondria can re-fuse if the injurious stress is removed before permanent damage
occurs to trigger apoptosis (our unpublished observations). However, when the injury is
prolonged, mitochondria cannot re-fuse and become irreversibly damaged. These
observations suggest that mitochondrial fragmentation is not the “point of
no return”; rather, it is a facilitating event for irreversible damage. It
is likely that fragmentation sensitizes mitochondria to additional injurious events,
which precipitate in the membrane leakage. One such event, as proposed in our recent
work (40), is initiated and mediated by Bax and
Bak. Mitochondrial fragmentation is virtually a process of membrane bending, scission,
and remelding. The associated biochemical and biophysical changes of the membrane are
expected to significantly affect the interaction, activation, and function of the
molecules residing in or accumulating near the membrane. Mitochondrial outer membrane
permeabilization involves the activation of Bax and Bak and significant conformational
changes of the proteins (41–45). Thus, fragmentation may predispose the
mitochondria to Bax/Bak-induced development of porous defects. This scenario, also
called the “2-hit” hypothesis (46), emphasizes 2 critical events of outer membrane permeabilization:
mitochondrial fragmentation and Bax/Bak-mediated pore formation. It is noteworthy that
these 2 events are not separately regulated, as both may be subjected to regulation by
Bcl2 family proteins (18, 46).
Our present results show that temporary blockade of fission can prevent mitochondrial
fragmentation and tubular cell apoptosis during acute kidney injury. However, it is
important to note that permanent blockade of mitochondrial fission may adversely affect
mitochondrial, cellular, and renal function or physiology. In this regard, Chan and
colleagues showed that mitochondrial fragmentation due to loss of fusion proteins leads
to defects in mitochondrial respiration and oxygen consumption accompanied by loss of
mitochondrial membrane potential and cell growth inhibition (47). Interestingly, Rossignol and colleagues demonstrated that
forced fusion of mitochondria by Drp1 knockdown also results in a reduction in
respiration and oxygen consumption (48). These
results, seemingly contradictory, suggest that mitochondrial morphological dynamics is a
key to mitochondrial physiology and perturbation of the dynamics to either fission or
fusion direction would reduce mitochondrial function.
In conclusion, this study has demonstrated mitochondrial fragmentation in both ischemic
and nephrotoxic models of acute renal failure. Mitochondrial fragmentation occurs early
and contributes to subsequent development of mitochondrial membrane permeabilization,
release of apoptogenic factors, and tubular cell apoptosis. Importantly, inhibition of
mitochondrial fragmentation protects against tubular cell apoptosis and renal injury,
suggesting what we believe is a novel strategy for the prevention and treatment of acute
renal failure.
MethodsAntibodies, plasmids, and other reagents.
Antibodies and sources are as follows: monoclonal anti–cytochrome
c (7H8.2C12 and 6H2.B4), anti-active caspase-3, and anti-Drp1
from BD Biosciences — Pharmingen; polyclonal anti-Fis1 from ALEXIS
Biochemicals; polyclonal anti–phospho-p53 from Cell Signaling Technology;
monoclonal anti-Bax from NeoMarkers; polyclonal anti-PUMA from Jian Yu (University of
Pittsburgh, Pittsburgh, Pennsylvania, USA); and all secondary antibodies from Jackson
ImmunoResearch Laboratories Inc. Plasmids containing Drp1 or its dominant-negative
mutant were kindly provided by Alexander van der Bliek (UCLA School of Medicine, Los
Angeles, California, USA) (28) and were
further subcloned into pcDNA3.1 (Invitrogen) for this study. The short hairpin Drp1
siRNA plasmid was a gift from Ansgar Santel (Silence Therapeutics, Berlin, Germany)
(30). pDsRed2-Mito (MitoRed), an expression
vector encoding red fluorescent protein with a mitochondrial targeting sequence, was
purchased from Clontech. All other reagents were purchased from Sigma-Aldrich unless
otherwise noted.
RPTC lines.
RPTCs were originally provided by Ulrich Hopfer (Case Western Reserve University,
Cleveland, Ohio, USA). RPTC lines stably transfected with Bcl2 were generated in
previous work (49). RPTC lines with Drp1
knockdown were generated by transfection with the short hairpin Drp1 siRNA plasmid
(30), followed by hygromycin selection. In
brief, RPTCs were cotransfected with the Drp1 siRNA plasmid and a hygromycin
resistance vector. 24 hours after transfection, cells were treated with 400
μg/ml hygromycin (Sigma-Aldrich). Hygromycin treatment was continued
until the original monolayer of cells was reduced to individual cells. The hygromycin
concentration was then reduced to 200 μg/ml to allow the individual cells
to proliferate into colonies. Individual colonies were extracted and cultured, then
tested for Drp1 expression. All RPTCs were maintained in Ham’s F-12/DMEM
supplemented with 10% FBS, 5 μg/ml transferrin, 5 μg/ml
insulin, 1 ng/ml EGF, 4 μg/ml dexamethasone, and 1% antibiotics.
Primary renal proximal tubular cells.
C57BL/6 mice (8 to 12 weeks, male) were purchased from The Jackson Laboratory. Renal
cortical tissues were harvested from animals to isolate proximal tubular cells for
culture as previously described (15, 33). In brief, cortical tissues were minced,
digested with collagenase, and centrifuged in 32% Percoll medium to purify proximal
tubular cells. Cells were then plated in collagen-coated dishes and maintained in
DMEM/F-12 medium supplemented with 5 μg/ml transferrin, 5
μg/ml insulin, 0.05 μM hydrocortisone, and 50 μM
vitamin C.
Apoptotic treatment.
For ATP depletion, cells were treated with 10 mM azide in glucose-free Krebs-Ringer
bicarbonate solution for 3 hours or time as indicated (50). After the incubation, cells were fractionated for analysis
of cytochrome c release, fixed for immunostaining and microscopic
examination, or returned to glucose-containing culture medium for 2 hours to observe
apoptosis (51, 52). For cisplatin treatment, RPTC lines were incubated with 20
μM cisplatin in culture medium for 4 to 16 hours (53), whereas primary mouse kidney tubular cells were incubated
with 50 μM cisplatin for 20 hours (15).
Transfection.
RPTCs were plated at approximately 50% confluence and transfected with 1.0
μg plasmid DNA using Lipofectamine PLUS reagent (Invitrogen). Primary
cultures of isolated kidney tubular cells were plated at approximately 70% confluence
for transfection with 1.0 μg plasmid DNA using Lipofectamine 2000
according to the manufacturer’s instructions (Invitrogen). To visualize
mitochondria, cells were transfected with 0.1 μg pDsRed2-Mito to label
mitochondria with the red fluorescent MitoRed protein.
Mitochondrial and cell morphology.
Mitochondrial morphology was examined by fluorescence microscopy in
pDsRed2-Mito–transfected cells as described in our recent study (40). pDsRed2-Mito transfection leads to the
expression in mitochondria of the red fluorescent protein MitoRed, a fusion protein
containing a red fluorescent sequence and a mitochondrial targeting domain of
cytochrome c oxidase, subunit VIII. MitoRed signal is not sensitive
to changes of mitochondrial conditions including membrane potential and redox status.
In brief, cells were transfected with pDsRed2-Mito. After various treatments, the
cells were examined by fluorescence microscopy to evaluate mitochondrial morphology
in individual cells. Mitochondria of untreated cells were filamentous and showed a
threadlike tubular structure, while mitochondria in stressed cells were fragmented
and appeared shortened and punctate. Consistent with earlier studies (15, 54,
55), the mitochondria within a cell were
often either filamentous or fragmented. In uncommon cases of mixed mitochondrial
morphology, we classified the cells based on the majority (>70%) of
mitochondria. Cytochrome c release and apoptotic morphology were
often evaluated in the same cells that were assessed for mitochondrial morphology.
Cytochrome c release was indicated by the loss of mitochondrial
cytochrome c staining and the appearance of cytochrome
c in cytosol. Typical apoptotic morphology was indicated by cellular
condensation, formation of apoptotic bodies, and condensation and fragmentation of
the nucleus. For each sample, several random fields of cells (≥100 cells
per dish) were evaluated for mitochondrial morphology, apoptosis, and cytochrome
c release.
Immunofluorescence and TUNEL staining.
Cells were grown on collagen-coated glass coverslips and subjected to various
treatments. The cells were then fixed with a modified Zamboni’s fixative
containing 4% paraformaldehyde and picric acid and permeabilized with 0.1% SDS. Cells
were blocked with 5% normal goat serum and 2% BSA. After blocking, cells were
incubated with primary antibodies (monoclonal mouse anti–cytochrome
c, anti-Drp1, anti-Fis1, or anti-active caspase-3) and secondary
antibodies (Cy-3–labeled or FITC-labeled goat anti-mouse IgG). TUNEL
staining was performed using the In Situ Cell Death Detection Kit
from Roche Applied Science according to the manufacturer’s instructions.
In brief, cells were grown on collagen-coated glass coverslips for experiment, then
fixed with 4% paraformaldehyde and permeabilized with 0.1% Triton X-100 in 0.1%
sodium citrate. Cells were then incubated with the TUNEL reaction mixture for 1 hour
at 37°C. Immunofluorescence and TUNEL staining were examined by
fluorescence and confocal microscopy, using a LSM 510 Zeiss microscope.
Cellular fractionation.
To study cytochrome c release, cells were incubated with 0.05%
digitonin in an isotonic sucrose buffer (250 mM sucrose, 10 mM HEPES-NaOH, 10 mM KCl,
1.5 mM MgCl2, 1 mM EDTA, 1 mM EGTA, and 0.5 mM phenylmethylsulfonyl
fluoride; pH 7.2) at room temperature as described previously (40, 49). At this
concentration, digitonin selectively permeabilizes the plasma membrane but not the
mitochondrial membrane. The digitonin extract was collected as the cytosolic fraction
for immunoblot analysis. To examine Drp1 translocation to mitochondria during
apoptosis, cells were collected in ice-cold isotonic sucrose buffer by gentle
scraping. The cells were then homogenized with a Wheaton homogenizer. The homogenate
was centrifuged at 1000 g to remove debris and nuclei to collect the
supernatant for further centrifugation at 10,000 g to yield the
mitochondrial fraction. Homogenization and centrifugation procedures were conducted
at 4°C.
Immunoblot analysis.
Standard procedure was performed for immunoblotting using NuPAGE (Invitrogen) or
Bio-Rad Gel Systems. In brief, 25 μg protein from each sample was
resolved by SDS-PAGE electrophoresis under reducing conditions. Proteins were then
transferred to PVDF membranes and blocked using 5% milk. The membranes were then
incubated with primary antibody overnight at 4°C and with HRP-conjugated
secondary antibodies for 1 hour at room temperature. The antigen-specific signal was
then detected through incubation with enhanced chemoluminescence substrate (Pierce
Biotechnology).
Renal ischemia/reperfusion in C57BL/6 mice.
C57BL/6 mice purchased from The Jackson Laboratory were maintained in the animal
facility of Charlie Norwood VA Medical Center at Augusta under a 12-hour
light/12-hour dark cycle with free access to food and water. Renal ischemia was
induced in male mice of 8 to 10 weeks as described previously (15, 16, 33). In brief, the animals were anesthetized with
pentobarbital (i.p., 50 mg/kg) and were kept on a homeothermic table. Flank incisions
were made to expose the renal pedicles for bilateral clamping to induce 30 minutes of
renal ischemia. The clamps were then released for reperfusion for indicated times.
Control animals were subjected to sham operation without renal pedicle clamping. To
test the effects of mdivi-1, the animals were injected (i.p.) with 50 mg/kg body
weight mdivi-1 1 hour prior to renal ischemia/reperfusion. All experiments involving
animals were conducted according to a protocol approved by the Institutional Animal
Care and Use Committees of Charlie Norwood VA Medical Center.
Renal function, histology, and TUNEL assay.
Serum creatinine and BUN were determined to monitor renal function as previously
reported (15, 16, 33). For histology, kidneys
were fixed with 4% paraformaldehyde, paraffin embedded, and stained with
H&E. Histopathological changes evaluated in this study included loss of
brush border, tubular dilation, cast formation, and cell lysis. Tissue damage was
examined in a blind manner and scored according to the percentage of damaged tubules:
0, no damage; 1, less than 25% damage; 2, 25%–50% damage; 3,
50%–75% damage; and 4, more than 75% damage. TUNEL assay was performed to
evaluate apoptosis in renal tissues using the In Situ Cell Death
Detection Kit from Roche Applied Science as described previously (15, 16,
33). In brief, renal tissues were fixed
with 4% paraformaldehyde and paraffin embedded. Tissue sections of 4 μm
were exposed to a TUNEL reaction mixture containing terminal deoxynucleotidyl
transferase and nucleotides, including tetramethylrhodamine–labeled
(TMR-labeled) dUTP. The slides were examined by fluorescence microscopy.
Examination of mitochondrial fragmentation in renal tissues by 2D EM.
Sham-operated control and ischemia/reperfused animals were perfused with 10 ml (at 10
units/ml) heparin, followed by 50 ml fixative containing 100 mM sodium cacodylate, 2
mM CaCl2, 4 mM MgSO4, 4% paraformaldehyde, and 2.5%
glutaraldehyde. Kidneys were then harvested and postfixed in the same fixative. A
tissue block of approximately 1 mm3 was collected from each kidney,
including a portion of renal cortex and outer medulla for standard processing for EM.
The tissue block was examined initially at low magnification (×3,000) to
identify representative proximal tubules. Cells in these tubules were then examined
at high magnification (×15,000) to reveal mitochondria. To determine
mitochondrial fragmentation, digital images with scale bars were collected in EM. The
lengths of individual mitochondria in a cell were measured by tracing using NIH
ImageJ software (http://rsbweb.nih.gov/ij/). For each cell, approximately 100
mitochondria in a representative area were measured to determine the percentage
distribution of mitochondria with various lengths (0–1 μm,
1–2 μm, >2 μm). We showed that the
mitochondrial diameter of tubular cells is approximately 0.3 to 0.5 μm
and that a mitochondrion with length greater than 2 μm is clearly
filamentous (Figure 9). In 2D EM, proximal
tubular cells in control tissues consistently showed 10%–20% long
filamentous mitochondria. If a cell had undergone mitochondrial fragmentation, then
very few or no filamentous mitochondria would appear. Thus, we determined the
percentage of cells that had less than 1% long filamentous mitochondria to indicate
the degree of mitochondrial fragmentation.
Examination of mitochondria in renal tissues by 3D EM.
Renal tissues were fixed and processed for EM as described above. The tissues were
initially examined to identify representative proximal tubules for serial section.
One hundred serial sections (45 nm/section) were collected from representative
tubules or cells. EM images of the serial sections were obtained and then aligned to
reconstruct a 3D image of mitochondria using Reconstruct Software, version 1.1.0.0
(http://synapses.clm.utexas.edu/tools/reconstruct/reconstruct.stm).
Statistics.
Quantitative data were expressed as mean ± SD from at least 3 separate
experiments. Statistical differences between the means were determined using ANOVA
followed by Tukey’s post test. P < 0.05 was
considered significant. Qualitative data including cell images and immunoblots are
representative of at least 3 separate experiments.
Supplemental dataView Supplemental data View Supplemental video
AcknowledgmentsWe thank Alexander van der Bliek (University of California School of Medicine, Los
Angeles, California, USA) and Ansgar Santel (Silence Therapeutics, Berlin, Germany) for
providing plasmids. We also thank Robert Smith and Libby Perry at the Electron
Microscopy Core Laboratory of the Medical College of Georgia for their assistance with
electron microscopy in this study. Craig Brooks was supported in part by the
Multidisciplinary Predoctoral Training Program in Integrative Cardiovascular Biology
from the NIH. Zheng Dong is a Research Career Scientist at the United States Department
of Veterans Affairs (VA). The study was supported by grants from the NIH and the VA.
Footnotes
Authorship note: Craig Brooks and Qingqing Wei contributed equally to
this work. Conflict of interest: The authors have declared that no conflict of
interest exists. Nonstandard abbreviations used: BUN, blood urea nitrogen; DN-Drp1,
dominant-negative Drp1; Drp1, dynamin-related protein 1; Fis1, Fission 1;
PUMA-α, p53-upregulated modulator of apoptosis α; RPTC, rat
proximal tubular cell; VAD,
carbobenzoxy-valyl-alanyl-aspartyl-(O-methyl)-fluoromethylketone. Citation for this article:J. Clin. Invest.119:1275–1285 (2009). doi:10.1172/JCI37829 Craig Brooks’s present address is: Renal Division, Department of
Medicine, Brigham and Women’s Hospital, Harvard Medical School, Boston,
Massachusetts, USA.
References-
Bonventre, J.V., Weinberg, J.M. 2003. Recent advances in the pathophysiology of ischemic acute renal
failure. J. Am. Soc. Nephrol. 14:2199-2210.
-
Lieberthal, W., Nigam, S.K. 1998. Acute renal failure. I. Relative importance of proximal vs. distal
tubular injury. Am. J. Physiol. 275:F623-F631.
-
Devarajan, P. 2006. Update on mechanisms of ischemic acute kidney injury. J. Am. Soc. Nephrol. 17:1503-1520.
-
Molitoris, B.A. 2003. Transitioning to therapy in ischemic acute renal failure. J. Am. Soc. Nephrol. 14:265-267.
-
Lieberthal, W., Nigam, S.K. 2000. Acute renal failure. II. Experimental models of acute renal failure:
imperfect but indispensable. Am. J. Physiol. Renal Physiol. 278:F1-F12.
-
Pabla, N., Dong, Z. 2008. Cisplatin nephrotoxicity: Mechanisms and renoprotective strategies. Kidney Int. 73:994-1007.
-
Schrier, R.W., Wang, W., Poole, B., Mitra, A. 2004. Acute renal failure: definitions, diagnosis, pathogenesis, and
therapy. J. Clin. Invest. 114:5-14.
-
Waikar, S.S., Curhan, G.C., Wald, R., McCarthy, E.P., Chertow, G.M. 2006. Declining mortality in patients with acute renal failure, 1988 to
2002. J. Am. Soc. Nephrol. 17:1143-1150.
-
Xue, J.L., et al. 2006. Incidence and mortality of acute renal failure in Medicare
beneficiaries, 1992 to 2001. J. Am. Soc. Nephrol. 17:1135-1142.
-
Kaushal, G.P., Basnakian, A.G., Shah, S.V. 2004. Apoptotic pathways in ischemic acute renal failure. Kidney Int. 66:500-506.
-
Padanilam, B.J. 2003. Cell death induced by acute renal injury: a perspective on the
contributions of apoptosis and necrosis. Am. J. Physiol. Renal Physiol. 284:F608-F627.
-
Weinberg, J.M. 1991. The cell biology of ischemic renal injury. Kidney Int. 39:476-500.
-
Sanz, A.B., Santamaria, B., Ruiz-Ortega, M., Egido, J., Ortiz, A. 2008. Mechanisms of renal apoptosis in health and disease. J. Am. Soc. Nephrol. 19:1634-1642.
-
Servais, H., et al. 2008. Renal cell apoptosis induced by nephrotoxic drugs: cellular and
molecular mechanisms and potential approaches to modulation. Apoptosis. 13:11-32.
-
Wei, Q., Dong, G., Franklin, J., Dong, Z. 2007. The pathological role of Bax in cisplatin nephrotoxicity. Kidney Int. 72:53-62.
-
Wei, Q., Yin, X.M., Wang, M.H., Dong, Z. 2006. Bid deficiency ameliorates ischemic renal failure and delays animal
death in C57BL/6 mice. Am. J. Physiol. Renal Physiol. 290:F35-F42.
-
Castaneda, M.P., et al. 2003. Activation of mitochondrial apoptotic pathways in human renal
allografts after ischemiareperfusion injury. Transplantation. 76:50-54.
-
Suen, D.F., Norris, K.L., Youle, R.J. 2008. Mitochondrial dynamics and apoptosis. Genes Dev. 22:1577-1590.
-
Herzig, S., Martinou, J.C. 2008. Mitochondrial dynamics: to be in good shape to survive. Curr. Mol. Med. 8:131-137.
-
Chan, D.C. 2006. Mitochondrial fusion and fission in mammals. Annu. Rev. Cell Dev. Biol. 22:79-99.
-
Okamoto, K., Shaw, J.M. 2005. Mitochondrial morphology and dynamics in yeast and multicellular
eukaryotes. Annu. Rev. Genet. 39:503-536.
-
Westermann, B. 2008. Molecular machinery of mitochondrial fusion and fission. J. Biol. Chem. 283:13501-13505.
-
Breckenridge, D.G., et al. 2008. Caenorhabditis elegans drp-1 and fis-2 regulate distinct cell-death
execution pathways downstream of ced-3 and independent of ced-9. Mol. Cell. 31:586-597.
-
James, D.I., Martinou, J.C. 2008. Mitochondrial dynamics and apoptosis: a painful separation. Dev. Cell. 15:341-343.
-
Sheridan, C., Delivani, P., Cullen, S.P., Martin, S.J. 2008. Bax- or Bak-induced mitochondrial fission can be uncoupled from
cytochrome C release. Mol. Cell. 31:570-585.
-
Barsoum, M.J., et al. 2006. Nitric oxide-induced mitochondrial fission is regulated by
dynamin-related GTPases in neurons. EMBO J. 25:3900-3911.
-
Karbowski, M., et al. 2002. Spatial and temporal association of Bax with mitochondrial fission
sites, Drp1, and Mfn2 during apoptosis. J. Cell Biol. 159:931-938.
-
Smirnova, E., Griparic, L., Shurland, D.L., van der Bliek, A.M. 2001. Dynamin-related protein Drp1 is required for mitochondrial division in
mammalian cells. Mol. Biol. Cell. 12:2245-2256.
-
Wang, X. 2001. The expanding role of mitochondria in apoptosis. Genes Dev. 15:2922-2933.
-
Tondera, D., et al. 2005. The mitochondrial protein MTP18 contributes to mitochondrial fission
in mammalian cells. J. Cell Sci. 118:3049-3059.
-
Jiang, M., et al. 2006. Regulation of PUMA-alpha by p53 in cisplatin-induced renal cell
apoptosis. Oncogene. 25:4056-4066.
-
Cassidy-Stone, A., et al. 2008. Chemical inhibition of the mitochondrial division dynamin reveals its
role in Bax/Bak-dependent mitochondrial outer membrane permeabilization. Dev. Cell. 14:193-204.
-
Wei, Q., et al. 2007. Activation and involvement of p53 in cisplatin-induced nephrotoxicity. Am. J. Physiol. Renal Physiol. 293:F1282-F1291.
-
Perfettini, J.L., Roumier, T., Kroemer, G. 2005. Mitochondrial fusion and fission in the control of apoptosis. Trends Cell Biol. 15:179-183.
-
Jagasia, R., Grote, P., Westermann, B., Conradt, B. 2005. DRP-1-mediated mitochondrial fragmentation during EGL-1-induced cell
death in C. elegans. Nature. 433:754-760.
-
Cribbs, J.T., Strack, S. 2007. Reversible phosphorylation of Drp1 by cyclic AMP-dependent protein
kinase and calcineurin regulates mitochondrial fission and cell death. EMBO Rep. 8:939-944.
-
Wasiak, S., Zunino, R., McBride, H.M. 2007. Bax/Bak promote sumoylation of DRP1 and its stable association with
mitochondria during apoptotic cell death. J. Cell Biol. 177:439-450.
-
Waterham, H.R., et al. 2007. A lethal defect of mitochondrial and peroxisomal fission. N. Engl. J. Med. 356:1736-1741.
-
Schrader, M. 2006. Shared components of mitochondrial and peroxisomal division. Biochim. Biophys. Acta. 1763:531-541.
-
Brooks, C., et al. 2007. Bak regulates mitochondrial morphology and pathology during apoptosis
by interacting with mitofusins. Proc. Natl. Acad. Sci. U. S. A. 104:11649-11654.
-
Adams, J.M., Cory, S. 2001. Life-or-death decisions by the Bcl-2 protein family. Trends Biochem. Sci. 26:61-66.
-
Danial, N.N., Korsmeyer, S.J. 2004. Cell death: critical control points. Cell. 116:205-219.
-
Harris, M.H., Thompson, C.B. 2000. The role of the Bcl-2 family in the regulation of outer mitochondrial
membrane permeability. Cell Death Differ. 7:1182-1191.
-
Martinou, J.C., Green, D.R. 2001. Breaking the mitochondrial barrier. Nat. Rev. Mol. Cell Biol. 2:63-67.
-
Youle, R.J., Strasser, A. 2008. The BCL-2 protein family: opposing activities that mediate cell death. Nat. Rev. Mol. Cell Biol. 9:47-59.
-
Brooks, C., Dong, Z. 2007. Regulation of mitochondrial morphological dynamics during apoptosis by
Bcl-2 family proteins: a key in Bak? Cell Cycle. 6:3043-3047.
-
Chen, H., Chomyn, A., Chan, D.C. 2005. Disruption of fusion results in mitochondrial heterogeneity and
dysfunction. J. Biol. Chem. 280:26185-26192.
-
Benard, G., et al. 2007. Mitochondrial bioenergetics and structural network organization. J. Cell Sci. 120:838-848.
-
Saikumar, P., et al. 1998. Role of hypoxia-induced Bax translocation and cytochrome c release in
reoxygenation injury. Oncogene. 17:3401-3415.
-
Brooks, C., Wang, J., Yang, T., Dong, Z. 2007. Characterization of cell clones isolated from hypoxia-selected renal
proximal tubular cells. Am. J. Physiol. Renal Physiol. 292:F243-F252.
-
Dong, Z., Wang, J. 2004. Hypoxia selection of death-resistant cells: a role for Bcl-XL. J. Biol. Chem. 279:9215-9221.
-
Wang, J., et al. 2004. Minocycline up-regulates Bcl-2 and protects against cell death in
mitochondria. J. Biol. Chem. 279:19948-19954.
-
Pabla, N., Huang, S., Mi, Q.S., Daniel, R., Dong, Z. 2008. ATR-Chk2 signaling in p53 activation and DNA damage response during
cisplatin-induced apoptosis. J. Biol. Chem. 283:6572-6583.
-
Youle, R.J., Karbowski, M. 2005. Mitochondrial fission in apoptosis. Nat. Rev. Mol. Cell Biol. 6:657-663.
-
Lee, Y.J., Jeong, S.Y., Karbowski, M., Smith, C.L., Youle, R.J. 2004. Roles of the mammalian mitochondrial fission and fusion mediators
Fis1, Drp1, and Opa1 in apoptosis. Mol. Biol. Cell. 15:5001-5011.
|
