Departments of 1 Medicine and 4 Physiology and Biophysics, and 2 Rammelkamp Center for Education and Research, Case Western Reserve University School of Medicine, Cleveland 44109; and 3 Louis B. Stokes Veterans Administration Medical Center, Cleveland, Ohio 44106
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ABSTRACT |
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Renal tubular epithelial cell (RTC) apoptosis causes tubular atrophy, a hallmark of renal disease progression. Apoptosis is generally characterized by reduced cell volume and cytosolic pH, but epithelial cells are relatively resistant to shrinkage due to regulatory volume increase, which is mediated by Na+/H+ exchanger (NHE) 1. We investigated whether RTC apoptosis requires caspase cleavage of NHE1. Staurosporine- and hypertonic NaCl-induced RTC apoptosis was associated with cell shrinkage and diminished cytosolic pH, and apoptosis was potentiated by amiloride analogs, suggesting NHE1 activity opposes apoptosis. NHE1-deficient fibroblasts demonstrated increased susceptibility to apoptosis, which was reversed by NHE1 reconstitution. NHE1 expression was markedly decreased in apoptotic RTC due to degradation, and preincubation with peptide caspase antagonists restored NHE1 expression, indicating that NHE1 is degraded by caspases. Recombinant caspase-3 cleaved the in vitro-translated NHE1 cytoplasmic domain into five distinct peptides, identical in molecular weight to NHE1 degradation products derived from staurosporine-stimulated RTC lysates. In vivo, NHE1 loss-of-function C57BL/6.SJL-swe/swe mice with adriamycin-induced nephropathy demonstrated increased RTC apoptosis compared with adriamycin-treated wild-type controls, thereby implicating NHE1 inactivation as a potential mechanism of tubular atrophy. We conclude that NHE1 activity is critical for RTC survival after injury and that caspase cleavage of RTC NHE1 may promote apoptosis and tubular atrophy by preventing compensatory intracellular volume and pH regulation.
cell death; nephropathy; regulatory volume increase; renal disease; tubular atrophy; Na+/H+ exchanger 1
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INTRODUCTION |
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TUBULAR ATROPHY IS A HALLMARK of chronic renal diseases and is superior to glomerular pathology as a histological predictor of clinical outcomes (41). We have previously shown that renal tubular epithelial cell (RTC) apoptosis is a mechanism of tubular atrophy (20, 43). In the original descriptions of apoptosis morphology, Wyllie et al. (51) termed the process "shrinkage necrosis" due to reductions in cell volume observed during apoptosis. Apoptotic cell shrinkage is achieved by net loss of intracellular osmoles and H2O, as well as by caspase-dependent proteolysis of housekeeping and structural proteins, which mediates cell disassembly. Many studies have also demonstrated that apoptosis is associated with a decrease in cytosolic pH (16, 27, 28, 30, 36, 46), which is required for activation of the caspase cascade (30, 45).
In contrast to neuronal cells and lymphocytes, epithelium-derived cells
are relatively resistant to apoptosis following exposure to
hypertonic extracellular conditions (6, 34), due to an enhanced capacity to rapidly expand intracellular volume through regulatory volume increase (RVI) pathways (17, 26, 31,
33). RVI is achieved by activation of the
Na+/H+ exchanger isoform NHE1 and, depending on
the cell type, the anion exchanger (AE) 2 isoform of the
Cl/HCO
symporter (26,
31, 33). The net effect is ion and H2O influx, which
leads to intracellular volume re-expansion. If RVI-dependent transporters are robustly activated after initiation of an
apoptotic stimulus, restoration of intracellular volume may preempt
apoptosis (26, 33). Alternatively, for a cell to
undergo apoptosis, RVI must be overcome or inhibited (26,
33). Neither AE2 nor the types 1 or 2 bumetanide-sensitive
cotransporter (BSC-1 or BSC-2, respectively) isoforms of the
Na+/K+/2Cl
symporter are
expressed in proximal tubule (1, 15, 18), the nephron
segment that demonstrates the most abundant apoptosis in animal
models of progressive renal disease (20, 43). However, NHE1 is ubiquitously expressed, including within the proximal tubule,
suggesting that NHE1 may be critical to RTC survival by promoting
resistance to apoptotic cell shrinkage.
In addition to regulating cell volume via RVI, NHE1 mediates other housekeeping functions, such as intracellular pH (pHi) regulation through electroneutral Na+ influx and H+ efflux. NHE1-dependent Na+/H+ exchange has been linked to essential cell functions, such as proliferation (4, 32), whereas diminished NHE1 activity has been associated with lymphocyte apoptosis (27, 40). Moreover, NHE1 has recently been recognized to function as a scaffold for binding with ezrin, radixin, and moesin (ERM) (14), adaptor molecules that link cytoskeleton to plasma membrane proteins, suggesting that NHE1 may facilitate apoptosis resistance by preserving cytoarchitecture and maintaining cell volume independent of Na+/H+ antiporter functions.
Because predicted sequelae of NHE1 inhibition-cell shrinkage, intracellular acidification, and cytoskeleton collapse mimic the apoptotic phenotype, we investigated whether RTC apoptosis is regulated by NHE1 caspase cleavage. We find that RTC apoptosis is associated with caspase-dependent NHE1 degradation. Furthermore, cell culture and whole animal data demonstrate that NHE1 loss-of-function mutations render RTC susceptible to apoptosis. The data are consistent with a mechanism whereby NHE1 degradation causes RTC apoptosis and tubular atrophy, which prevents RVI and promotes intracellular acidosis, an optimum condition for caspase activity.
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MATERIALS AND METHODS |
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Reagents. We used the following: amiloride, 4',6-diamidino-2-phenylindole (DAPI), ethyl-N-isopropylamiloride (EIPA), hexamethyleneamiloride (HMA), staurosporine (STS), adriamycin hydrochloride (Sigma, St. Louis, MO); z-VAD-fmk, z-DEVD-fmk, Ac-DEVD-CHO (Calbiochem, La Jolla, CA); 2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein (BCECF)-AM (Molecular Probes, Eugene, OR); anti-poly(ADP-ribose)polymerase (PARP) IgG, phycoerythrin (PE)-conjugated annexin V (Pharmingen, San Diego, CA); annexin V, anti-hemagglutinin (HA) IgG (Roche, Indianapolis, IN); horseradish peroxidase (HRP)-conjugated IgG (Santa Cruz Biotechnology, Santa Cruz, CA); green fluorescence protein (GFP) cDNA (Clontech, Palo Alto, CA); Red X-conjugated anti-mouse IgG, FITC-conjugated anti-mouse IgG (Vector Laboratories, Burlingame, CA); [35S]methionine (ICN, Irvine, CA); C57BL/6.SJL +/+, C57BL/6.SJL swe/+, and C57BL/6.SJL swe/swe mice (Jackson Laboratories, Bar Harbor, ME); COOH-terminal, HA epitope-tagged rat NHE1 cDNA (a gift from Dr. J. Orlowski, McGill University); and KR/A and E266I NHE1 mutant cDNAs (gifts from Dr. D. Barber, University of California at San Francisco). Rabbit polyclonal anti-NHE1 IgG was generated against an NHE1 cytoplasmic domain peptide as previously described (23) and affinity purified.
Cell lines. The human renal proximal tubule (HRPT) cell RTC line (gift from Dr. L. Racusen, Johns Hopkins University) has been extensively characterized (20, 21, 39, 43). HRPT and HEK 293 (ATCC, Manassas, VA) cells were maintained in DMEM-F12 (Gibco-BRL, Rockville, MD) plus 10% fetal bovine serum (Hyclone, Logan, UT) and 1% penicillin-streptomycin-fungizone solution (Sigma). PS120 cells are genetically deficient for NHE1 expression and were derived from control CCL39 fibroblasts (gifts from Drs. D. Grall and J. Pouysségur, University of Nice).
Flow cytometry. Cells were incubated with STS, lifted with trypsin-EDTA (10 min, 37°C), and washed twice in incubation buffer (10 mM HEPES/NaOH, pH 7.4, 140 mM NaCl, and 5 mM CaCl2) at 4°C. Washed cells were incubated with BCECF-AM (1.6 µM, 30 min, room temperature), PE-conjugated annexin V (15 min, room temperature), and DAPI (2 µg/ml, 15 min, room temperature). Cytosolic pH was measured by BCECF-AM fluorescence, and relative cell volumes were determined from forward vs. side light scatter characteristics with a Becton Dickinson FACS Vantage flow cytometer according to previously described methods (21, 50). Apoptosis was measured by annexin V binding (see below).
Apoptosis assays. Cells were plated on glass coverslips at 0.25 × 106 cells/ml density, grown to 80% confluence, and then maintained in serum-free medium combined with apoptotic stimuli. In some experiments, cells were preincubated with NHE1 inhibitors for 2 h or caspase inhibitors for 1 h before apoptosis induction. Apoptotic cells were identified by simultaneous fluorescent labeling of chromatin with DAPI and externalized phosphatidylserine with annexin V as previously described (20, 21). Random fields were viewed at ×40 magnification with a Nikon epifluorescence microscope (Tokyo, Japan), and the percentage of apoptotic cells was separately determined by two blinded observers from a total of 100-200 cells per experimental condition. Representative fields were photographed with a Spot Digital System camera and analyzed using Image Pro software (Diagnostic Instruments, Sterling Heights, MI).
Plasmid transfections.
Plasmids were transformed into DH-5-competent bacterial
cells according to manufacturer's protocol (Gibco-BRL), extracted using a Maxiprep kit (Qiagen, Valencia, CA), and amplified by culture
in Luria-Bertani-ampicillin. GFP, HA-NHE1, KR/A (inhibits ERM binding),
and E226I (inhibits Na+/H+ exchange) mutant
NHE1 cDNAs were transiently transfected into cells according to
previously described methods (21). Briefly, cells were
plated in six-well dishes (0.25 × 106 cells/well) and
cultured overnight in DMEM-F12 plus 10% fetal bovine serum to achieve
80% confluence. Cells were then washed and incubated with 100 µl of
serum-free DMEM (Gibco-BRL) containing 6 µl of Fugene 6 transfection
reagent (Roche) and 2.0-3.0 µg of plasmid DNA for 20 min at room
temperature. Transfected cells were then cultured in complete media
containing DMEM-F12 and 10% fetal bovine serum for an additional
24 h.
Immunoblot analysis.
Methods have previously been described in detail (42).
Whole cell lysates were prepared in boiling 2× SDS sample buffer (125 mM Tris, pH 6.8, 2% SDS, 5% glycerol, 1% -mercaptoethanol, and
0.003% bromphenol blue). Samples were assayed for protein content
using protein assay reagents (Bio-Rad, Hercules, CA). Proteins were
denatured by boiling for 5 min, and samples (60 µg/lane) were
resolved by 8 or 14% SDS-PAGE (Novex, San Diego, CA). Proteins were
transferred to polyvinylidene difluoride membranes, blocked with 5%
nonfat milk, and incubated with either anti-PARP (1:2,000, 1 h,
room temperature) or anti-HA (1:5,000, 1 h, room temperature)
antibodies, followed by HRP-conjugated secondary antibody (1:5,000,
1 h, room temperature). Band intensity was detected by enhanced
chemiluminescence methods (Amersham Pharmacia Biotech, Arlington
Heights, IL) and exposure to Kodak Biomax ML film. In some
experiments individual bands were digitized by phosphorimager (Molecular Dynamics, Sunnyvale, CA), quantified with Image Quant 5 software (Molecular Dynamics), and normalized to control values.
Protein degradation by [35S]-labeled pulse chase. Cells were cultured to subconfluence, washed with PBS, and incubated with [35S]methionine in methionine-free DMEM (0.1 mCi/ml, 2 h, 37°C). Cells were washed with PBS and cultured in complete media (0-6 h, 37°C) with or without STS and peptide caspase inhibitors. Protein lysates (200 µg per sample) were immunoprecipitated with anti-HA (1 µg) or anti-NHE1 IgG (1 µg) and resolved by SDS-PAGE according to previously described methods (42). Autoradiograms were developed from dried gels. In some experiments, individual bands were digitized by phosphorimager (Molecular Dynamics), quantified with Image Quant 5 software (Molecular Dynamics), and normalized to control values.
Immunocytochemistry and fluorescence microscopy. Methods have previously been described in detail (20, 21, 43). Cells were maintained on sterile glass coverslips within six-well plates, fixed in paraformaldehyde (4%, 10 min, room temperature), and blocked with 5% low-IgG BSA and 0.2% Triton X-100 (Sigma) for 30 min at room temperature. Cells were incubated with anti-HA IgG (1:200, 2 h, room temperature), followed by either red X-conjugated or FITC-conjugated anti-mouse IgG (1:200, 2 h, 4°C). Negative controls were cells incubated with isotype-identical IgG, which was immunoreactive with an irrelevant epitope. Coverslips were mounted in antifade, aqueous media containing DAPI (Vectashield; Vector Laboratories) on standard microscope slides. Random fields were viewed by two observers blinded to experimental condition, using a Nikon epifluorescence microscope with appropriate fluorescence filters. Representative fields were photographed with a Spot Digital System camera and analyzed using Image Pro software.
Assay for caspase-3 cleavage of in vitro-translated NHE1. The DNA template for in vitro translation was created by PCR amplification of the NHE1 cytosolic domain (cNHE1) from rat cDNA with upstream primer 5'-CTACCGCTC- GAGCCACCATGCCCAAGGACCAGTTCATCATTGCC-3' that contains Xho I restriction endonuclease, Kozak and ATG start sites, and downstream primer 5'-TGCTCTAGACTAGCCCTGCCCTTTGGGGATGAAAGG-3' containing an Xba I restriction site and stop codon. The cNHE1 construct included the open reading frame encoding amino acids 447-820, which corresponds to the 58 COOH-terminal amino acids within the transmembrane domain and the entire cytosolic domain. The resulting DNA was digested with Xho I and Xba I (Gibco-BRL) and 1.1 kb product was cloned into pTNT vector (Promega, Madison, WI). PCR-generated cNHE1 nucleotide sequence was verified by automated sequencing (Cleveland Genomics, Cleveland, OH). [35S]Met-labeled cytoplasmic NHE1 substrate was generated using the reticulocyte lysate-based TNT Quick T7-coupled transcription/translation system (Promega) according to manufacturer's instructions. Briefly, cNHE1 plasmid template (1 µg) was labeled with [35S]Met (50 µCi, 90 min, 37°C). Autoradiograms from dried gels yielded a single 45-kDa band within 2- to 3-h film exposure (not shown). [35S]Met-labeled, in vitro-translated cNHE1 (3 µl) was incubated with or without Ac-DEVD-CHO (100 µM, 2 h, 37°C), followed by 1-2 µl of purified caspase-3 (6 h, 30°C; Pharmingen) in 5 µl of caspase buffer (100 mM HEPES, pH 7.5, 10% sucrose, and 0.1% 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate) according to published protocols (10, 22). Peptide products were resolved by 14% SDS-PAGE and examined for cleavage by autoradiography.
Animal models.
C57BL/6.SJL swe/swe mice harbor an NHE1 A1639T point
mutation, which introduces a premature stop codon, resulting in
truncation between the 11th and 12th NHE1 transmembrane domains and
loss of NHE1-dependent Na+/H+ activity
(13). Four-week-old swe/swe homozygotes have a
brain phenotype that includes ataxia and seizures, but a gross renal phenotype was not observed, as determined by kidney histology and serum
Na+, K+, Cl,
HCO
Statistics. Data are representative of three to five experiments per condition. Graphical results are expressed as means ± SE unless otherwise indicated. Comparisons between paired samples were made by the Student's t-test. Comparisons between groups containing more than two samples were made by one-way analysis of variance with the Bonferroni, Student-Newman-Keuls, or Kruskal-Wallis tests for multiple comparisons. Statistical significance is defined as P < 0.05.
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RESULTS |
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Apoptotic RTC are shrunken and acidic.
To determine whether apoptotic RTC develop reduced cell volume and
cytosolic pH, as has been described in leukocytes, we stimulated cultured RTC to undergo apoptosis with STS. Cell size and pH
were determined by flow cytometry. As shown in Fig.
1, a greater proportion of RTC incubated
with STS displayed smaller cell volumes and lower cytosolic pH, similar
to STS- and Fas-induced Jurkat T cell apoptosis (30). These data demonstrate that the apoptotic RTC
phenotype includes shrinkage and acidification, consistent with a role
for NHE1 inhibition.
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Hypertonicity stimulates RTC apoptosis.
Epithelial cells are relatively resistant to apoptosis
induction by anisotonic conditions due to robust RVI (6,
34), which is mediated by multiple transporters (26, 31,
33). To determine whether cell stress imposed by hypertonicity
causes apoptosis, we exposed RTC to increasing extracellular
concentrations of the impermeant sugars mannitol and sucrose or NaCl
and then simultaneously assayed them for apoptosis by annexin V
labeling of externalized phosphatidylserine and chromatin condensation. Figure 2 shows that hypertonicity caused
RTC apoptosis in response to all three stimuli in a
concentration-dependent fashion, indicating that RVI was surmounted and
RTC were susceptible to hypertonic stress-induced apoptosis.
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Hypertonicity stimulates RTC apoptosis by an NHE1-dependent
mechanism.
To determine the role of NHE1 in apoptosis induced by
hypertonic conditions, we examined the effect of NHE1 inhibitors on RTC
apoptosis, assayed by DAPI labeling of condensed chromatin (Fig. 3A) and annexin V
labeling of externalized phosphatidylserine (Fig. 3B). These
studies show that the Na+/H+ inhibitor
amiloride caused modest apoptosis but significantly potentiated
RTC apoptosis from hypertonic stress. Because amiloride, particularly at high concentrations, may inhibit multiple sodium transporters, including other NHE isoforms expressed in proximal tubule, NaCl-induced apoptosis was assayed following
preincubation with NHE1-specific amiloride analogs EIPA and HMA
(35, 47). Figure 3, C and D,
demonstrates that, like amiloride, EIPA and HMA caused modest
apoptosis and both inhibitors enhanced hypertonic NaCl-dependent apoptosis (HMA > EIPA), suggesting that
NHE1 inactivation potentiates RTC apoptosis induced by
stresses, such as hypertonicity.
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Hypertonicity-induced RTC apoptosis is mediated by
caspase-3 activation and decreased NHE1 expression.
To determine whether NHE1 is linked to caspase activation in
apoptosis due to hypertonic stimuli, we incubated RTC with
hypertonic NaCl and amiloride and then probed for activation of
caspase-3, which is the final downstream executioner caspase in many
apoptosis signaling cascades. Figure
4A shows that amiloride
accentuated hypertonicity-induced cleavage of the caspase-3 substrate
PARP, suggesting that NHE1 activity opposes apoptosis, perhaps
by inhibiting decreases in cytosolic pH, which enhance caspase-3
activity (30, 45). Figure 4, B and
C, demonstrates that hypertonic NaCl induction of caspase-3
activity was associated with diminished NHE1 expression, indicating
that NHE1 may represent a caspase-3 target, which is consistent with
the possibility that NHE1 dysfunction could contribute to the acidic
and shrunken cell phenotype, as shown in Fig. 1.
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RTC apoptosis under isotonic conditions is associated with
decreased NHE1 expression.
Because the data indicated that cell shrinkage-induced RTC
apoptosis is mediated by NHE1 inhibition and caspase-3
activation, we queried whether NHE1 could be a caspase-3 substrate. To
explore this possibility, we transiently cotransfected HEK 293 cells
with an NHE1 cDNA construct containing a carboxy-terminal HA tag and GFP cDNA (to mark transfected cells), followed by STS incubation to
stimulate caspase-3-dependent apoptosis. Nuclear morphology and
NHE1 expression patterns were determined by standard, fluorescence microscopy. Figure 5, A
and B, demonstrates representative, transfected, apoptotic, and nonapoptotic cells. Approximately 20% of all
cells underwent apoptosis, but only a small percentage of
transfected, apoptotic cells expressed NHE1 on the cell surface
(Fig. 5, C and E). Conversely, almost all
transfected, nonapoptotic cells expressed NHE1 in a plasma membrane
distribution (Fig. 5, C and E). Similar results
were observed in RTC (data not shown). The results from these
experiments suggest that NHE1 is cleaved during apoptosis and
that loss of NHE1 expression renders cells susceptible to
apoptosis.
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NHE1 reconstitution promotes resistance to apoptosis.
To further investigate the role of NHE1 in apoptosis, we
compared NHE1-deficient PS120 cells, which were derived from Chinese hamster ovary fibroblasts (37), and NHE1-expressing
control fibroblasts (CCL39 cells) for susceptibility to STS-induced
apoptosis. As shown by DAPI and annexin V assays in Fig.
6, A and B,
respectively, apoptosis was observed in a significantly greater
percentage of PS120 cells compared with the CCL39 cells, consistent
with a recent report in these two cell lines (2). NHE1
function was subsequently addressed by add-back experiments, in which
PS120 cells were transiently transfected with increasing concentrations
of NHE1 then stimulated with STS and assayed for apoptosis.
Figure 6C demonstrates that NHE1 reconstitution in PS120
cells conferred resistance to apoptosis. At the highest NHE1
expression levels, apoptosis was equivalent to control CCL39
cells (Fig. 6, A and B).
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RTC apoptosis leads to diminished NHE1 expression by
protein degradation.
Because NHE1 has a long (~24 h) half-life (9, 12) and is
not significantly regulated by membrane cycling (38),
rapid NHE1 disappearance with STS incubation suggests an NHE1
degradation mechanism, rather than suppressed synthesis. To determine
more definitively whether STS-induced decreases in cell surface NHE1 were due to protein degradation, we conducted 35S-labeled
pulse-chase experiments in HEK 293 cells transfected with HA-NHE1 cDNA
and then induced them to undergo apoptosis with STS.
Immunoprecipitation with anti-HA IgG revealed diminished 35S-labeled NHE1 beginning 2.5 h after STS incubation,
which was markedly more pronounced at 5-6 h (Fig.
7), indicating that STS stimulates NHE1
degradation. 35S-labeled NHE1 levels did not appreciably
change in unstimulated cells from 0 to 4 h, consistent with the
long NHE1 half-life (not shown). More importantly,
35S-labeled NHE1 levels were significantly greater in
unstimulated compared with STS-stimulated cells at 4 h (not
shown), further indicating that NHE1 is degraded with
apoptosis. Because STS is a known caspase-3 activator, the data
also suggest a caspase-3 mechanism of NHE1 degradation.
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NHE1 is degraded by caspase-3.
To determine whether apoptosis-associated NHE1 degradation is
due to caspase cleavage, we transiently transfected RTC with carboxy-terminal HA, epitope-tagged NHE1, and stimulated them with STS
to undergo apoptosis in the presence or absence of the cell-permeable, broad-spectrum peptide caspase inhibitor z-VAD-fmk or
the peptide caspase-3 inhibitor z-DEVD-fmk. Whole cell lysates were
probed for NHE1 expression and PARP cleavage by immunoblot analysis.
Figure 8A demonstrates that
STS induced concomitant caspase-3 activity and NHE1 degradation in HEK
293 cells, and both caspase-3 activity and NHE1 degradation were
partially inhibited by z-DEVD-fmk or z-VAD-fmk preincubation. Similar
results were observed in RTC (Fig. 8B). These data strongly
suggest that NHE1 is cleaved by caspase-3 during apoptosis.
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NHE1 regulates RTC apoptosis in vivo.
To test the role of NHE1 inactivation as a mechanism of RTC deletion in
an in vivo model of progressive renal disease, wild-type (C57BL/6.SJL
+/+), NHE1 loss-of-function mutant (C57BL/6.SJL
swe/swe), and heterozygote (C57BL/6.SJL
swe/+) mice underwent adriamycin infusion to induce
nephropathy (49). Kidneys removed 10 days postinfusion
revealed little histopathological damage in any of the genotypes (data
not shown). RTC apoptosis was rarely observed in wild-type
controls (Fig. 10A),
consistent with a failure of adriamycin to cause nephropathy in mice
with a C57BL/6 genetic background (49). However,
significant increases in RTC apoptosis were observed in C57BL/6
swe/swe and C57BL/6 swe/+ compared with C57BL/6
+/+ mice (Fig. 10, B and C). Because
neither C57BL/6 mice infused with adriamycin nor NHE1 loss-of-function
mutant C57BL/6 swe/swe mice demonstrate histological or
functional kidney abnormalities (13, 49), the data suggest
that NHE1 inhibition unmasked a renal phenotype in adriamycin-treated
C57BL/6 mice. In accordance with our previous reports demonstrating
that RTC apoptosis precedes and contributes to tubular atrophy
(20), these studies indicate that NHE1 confers
cytoprotection, whereas loss of RTC NHE1 function is associated with
apoptosis and tubular atrophy.
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DISCUSSION |
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NHE1 is ubiquitously expressed, and activation has been linked to vital housekeeping functions such as cell volume regulation (31, 33) and growth factor-dependent proliferation (4, 32, 44). Decreased NHE1 activity has been associated with lymphocyte apoptosis (27, 40), but a specific mechanism of NHE1 inhibition in apoptosis has not previously been described. A major finding in our study is that RTC NHE1 is inhibited by caspase cleavage. The results have potentially broad implications to disease pathogenesis, inasmuch as the data were generated in both epithelial and mesenchymal cells and in response to apoptosis induction by hypertonic stress or STS.
Many epithelium-derived cell lines were previously considered to be
resistant to apoptotic cell volume reduction due to robust expression and function of RVI pathway components (6).
However, our data demonstrate that RTC RVI can be overcome, permitting apoptosis to proceed. Of the transporters that mediate RVI, we focused on NHE1 partly because it is responsible for intracellular volume regulation, and an anticipated consequence of NHE1 inhibition, cell shrinkage, is a characteristic apoptotic feature. Indeed, we
found that RTC apoptosis is associated with diminished cell volume, consistent with NHE1 inhibition. Our data do not exclude roles
for regulatory volume decrease pathways, which can be activated in
apoptosis (33). In addition, other transporters
(7, 29, 52) may participate in RTC RVI triggered by
apoptotic stimuli. For example, inhibition of NHE3, which is
expressed on the apical proximal RTC membrane, could conceivably
contribute to RTC shrinkage and acidosis. However, because hypertonic
cell shrinkage suppresses NHE3 activity (19), we reasoned
that apoptotic cleavage of NHE3 was unlikely to result in
significant further suppression of transporter activity. Inhibition of
other transporters that have been implicated in RVI, such as the AE2
Cl/HCO
cotransporter, could also be involved in RTC apoptosis, by
impeding intracellular volume expansion. However, prominent functions
for these proteins in proximal RTC apoptosis are unlikely,
since none are abundantly expressed in the proximal tubule (1,
15, 18).
In addition to cell shrinkage, RTC NHE1 degradation was also associated
with intracellular acidification (16). The data are in
agreement with studies in leukocyte cell lines, which demonstrate that
apoptosis is preceded by decreased pHi (16,
27, 36). Importantly, NHE1 stimulation prevented intracellular
acidification and abrogated apoptosis (16, 36),
indicating NHE1 activation opposes apoptosis. Rich et al.
(40) demonstrated that human leukemia cells exhibit a
significantly higher pHi compared with normal leukocyte
lineage cells. Moreover, decreased pHi was associated with
increased apoptosis, which was exacerbated by exposure to the
NHE1 inhibitor HMA. Barrière et al. (2) recently
reported that apoptosis was also associated with intracellular
acidification in Chinese hamster lung fibroblasts and that NHE1-induced
increases in pHi were sufficient to prevent
apoptosis. All of these findings are consistent with
observations that the optimum pH for endonuclease and caspase
activation is 6.3-6.8 and support the notion that intracellular
acidification is critical for apoptosis execution (16,
30). Grinstein's group (5, 25) has shown that
expression of 566 NHE1 mutants (cytoplasmic domain deletion membrane
distal to Met566) resulted in impaired osmoregulation, as well as a
constitutively decreased resting pHi, suggesting that
apoptotic NHE1 cleavage upstream from this domain would yield a
similar phenotype. Together, our data in RTC are consistent with NHE1
inhibition contributing to both apoptotic acidification and
intracellular volume dysregulation.
Although the role of NHE1 as a Na+/H+ exchanger is applicable to apoptotic cell volume decrease, a number of structural proteins must also be cleaved to achieve a shrunken cell morphology. Denker et al. (14) recently demonstrated that NHE1 is tethered to the plasma membrane and cytoskeleton through direct interaction with actin-binding ERM proteins. On the basis of this discovery, NHE1 would appear to function as a cytoarchitecture scaffold and require disassembly during apoptosis, consistent with ERM protein dissociation from plasma membrane during apoptosis (24). We reasoned that NHE1 may therefore confer apoptosis resistance by anchoring the actin cytoskeleton to the plasma membrane via interactions with ERM proteins. However, expression of NHE1 constructs with point mutations (KR/A) that abolish ERM binding (14) resulted in partial rescue of RTC from apoptosis, whereas expression of E266I mutants with intact ERM binding domains had no effect on apoptosis. Results from these studies suggest that mechanisms in addition to NHE1-ERM interactions must be required for apoptosis resistance. Furthermore, our data do not exclude the possibility that interactions between other NHE1 domains and the cytoskeleton may be important for maintenance of cell volume and resistance to apoptosis. Because the NHE1 KR/A mutant has normal Na+/H+ exchange activity (14), we conclude that Na+ influx and/or H+ efflux are critical NHE1 functions for apoptosis resistance.
Evidence to support the hypothesis that NHE1 is cleaved by caspases includes apoptosis-dependent loss of NHE1 expression due to protein degradation, rescue of NHE1 expression by preincubation with cell-permeable peptide caspase inhibitors, and direct cleavage of in vitro-translated NHE1 by caspase-3. Although consensus caspase cleavage sites are identified in the carboxy-terminal human NHE1 cytosolic tail (e.g., 755-DEED-758), deletion mutation studies predict that cleavage at this site would not result in Na+/H+ exchange-dependent osmoregulatory dysfunction (5). Furthermore, cleavage at distal carboxy-terminal site(s) would result in generation of very small fragments and, therefore, would not account for the peptide band pattern observed in Fig. 9, indicating that cleavage at additional membrane-proximal, noncanonical sites is required. Further degradation of NHE1 and/or NHE1 caspase cleavage products by other protease pathways is also possible, although preliminary studies revealed that STS-induced changes in NHE1 expression are not altered by pretreatment with the proteosome inhibitors lactacystin, MG132, and PS-1 (Wu KL and Schelling JR, unpublished observations).
Although neither in vitro stimulus of apoptosis (STS, hypertonicity) is encountered by RTC in vivo, these agents were employed to mimic cell stresses that result in RTC apoptosis in vivo. We have previously shown that RTC apoptosis and tubular atrophy are caused by the in vivo stresses hypoxia and Fas activation in murine models of progressive renal disease (20, 43). In the current studies, the in vivo role of NHE1 was established by demonstration of increased RTC susceptibility to apoptosis in NHE1-deficient mice with adriamycin-induced nephropathy. To maximize the likelihood of detecting apoptotic RTC before obliteration of tubulointerstitial architecture by renal scarring, we killed mice after only 10 days. Because of the short observation interval, animals did not develop tubulointerstitial pathology, although we predict that the natural history of enhanced RTC apoptosis is tubular atrophy and interstitial fibrosis.
NHE1 is commonly referred to as a housekeeping protein, implying that it is pedestrian and unregulated. To the contrary, NHE1 has been shown to mediate vital cell functions, and the current studies establish a new role for NHE1 as a defender against RTC death. We speculate that in initial stages of RTC apoptosis in vivo, e.g., due to inflammation or uremia, NHE1 is likely to be activated in response to cell volume reduction cues. NHE1-dependent RVI may then be sufficient to prevent further cell volume shrinkage and perhaps even promote cell survival, provided the apoptotic stimulus is not too robust. However, once NHE1 is cleaved by caspases, the combined sequelae of NHE1 inhibition, cytosolic acidification, and cell shrinkage promote inexorable RTC apoptosis by optimizing pHi for further caspase activity and by bringing caspases in proximity to substrates via cell shrinkage.
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ACKNOWLEDGEMENTS |
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We are grateful to Drs. Barber, Grall, Orlowski, Pouysségur, and Racusen for donation of reagents and to Dr. Eleanor Lederer for thoughtful observations and comments.
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FOOTNOTES |
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This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-54178, DK-38558, and DK-57933. J. R. Schelling is an Established Investigator of the American Heart Association.
Address for reprint requests and other correspondence: J. R. Schelling, Case Western Reserve Univ., MetroHealth Medical Center Campus, Rammelkamp Center for Education and Research, 2500 MetroHealth Dr., G531, Cleveland, OH 44109-1998 (E-mail: jrs15{at}po.cwru.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
First published November 26, 2002;10.1152/ajprenal.00314.2002
Received 3 September 2002; accepted in final form 18 November 2002.
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