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*
Sheffield Kidney Institute, Northern General Hospital Trust, Sheffield,
United Kingdom.
Sheffield University Division of Clinical Sciences, Northern General
Hospital Trust, Sheffield, United Kingdom.
Department of Histopathology, Northern General Hospital Trust, Sheffield,
United Kingdom.
Correspondence to Dr. Tim Johnson, Sheffield Kidney Institute, Northern General Hospital, Herries Road, Sheffield S5 7AU, England, United Kingdom. Phone: 44-0-114-271-5322; Fax: 44-0-114-271-4410; E-mail: T.Johnson{at}Sheffield.ac.uk
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Although many different signals initiate apoptosis, the phenotype of
apoptosis (a series of distinct morphologic and biochemical changes) is
surprisingly similar even in different cell types, suggesting that the final
stages of apoptotic death are highly conserved
(7). A recently identified
class of cysteine proteases termed caspases coordinates the execution
of the death program. A family of 14 different caspases that play a role in
inflammation and apoptosis have been identified
(8). They all are produced as
inactive precursors (zymogens) that contain an N-terminal prodomain and a
large (approximately 20 kD) and a small (approximately 10 kD) subunit. The
N-terminal domain is highly variable between the different members and
contains important structural domains for the regulation of the activation.
Activation requires processing of the precursor into the large and small
subunits, and in some cases the prodomain is removed as well. The individual
subunits dimerize and associate to form a 
2ß2 heterotetramer with
two independent active sites
(9). Caspases are highly
specific with an absolute requirement for cleavage after aspartic acid,
whereas individual caspases recognize different tetrapeptide motifs, which
might explain their individual substrate specificity
(10). Caspases have diverse
functions; members of this family play essential roles in both initial
signaling events (caspase-8, caspase-9) and the downstream proteolytic
cleavages (caspase-3)
(8,10).
Protease inhibitors, including macromolecular and peptide-based inhibitors of
caspases, are highly effective in preventing apoptotic cell death in both
in vitro and in vivo models of apoptosis
(11,12).
Among identified caspases, the best functionally correlated with the phenotype of apoptosis is caspase-3 (CPP32, YAMA, apopain) (12), which is mainly activated via both the death receptor and mitochondrial routes (8). However, there are few studies on the regulators of apoptosis in CRF models, with little data on the contribution of the caspases, except in an acute ischemic renal injury model (13) and experimental cyclosporin A-induced nephrotoxicity (14). The investigation of caspase-3—deficient mice suggested that this enzyme, in particular might be an appropriate target for therapeutic intervention in diseases that result from inappropriate apoptosis (11,15). Such an approach, based on the inhibition of caspase-3, has already been shown to be protective in a rat model of neonatal hypoxic-ischemic brain injury and in a murine model of renal ischemia (16,17).
In this study using the NTN model of experimental progressive glomerulonephritis in Wistar Kyoto rats, we measured caspase-3 at the mRNA, protein, and activity levels in both the early inflammatory and the late fibrotic stages of disease. Furthermore, we addressed the association between the observed changes of caspase-3 and apoptosis with the morphologic acute and chronic changes during the course of NTN.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Nephrotoxic serum nephritis was induced in 28 rats by a single intravenous injection of 0.5 ml of rabbit anti-rat glomerular basement membrane (GBM) serum (nephrotoxic serum) into the tail vein. Sixteen rats were used as controls and were injected with the same amount of normal rabbit serum (Vector Laboratories, Peterborough, UK) (18). Rats were killed in groups (n = 4 to 7) at days 7, 15, 30, and 45 after serum injection. Removed kidney tissues were fixed in 10% (wt/vol) neutral buffered formalin and paraffin embedded for histologic and immunohistochemical examination. For electron microscopy, tissues were fixed in 2.5% (vol/vol) glutaraldehyde solution in phosphate buffer (pH 7.4). Tissues were snap-frozen and stored in liquid nitrogen for mRNA, protein, and activity analyses. Serum creatinine (SCr) concentration (standard autoanalyzer techniques) and 24-h urinary protein excretion (Biuret method) were determined in each group at all time points.
Estimation of Renal Scarring
The extent of renal scarring after NTN was determined by observers who were
blinded to the experimental code according to a previously published arbitrary
scale
(19,20,21).
At x200 magnification, Masson's trichrome—stained sections were
examined and scored by two authors (B.Y. and G.L.T.). Glomerulosclerosis (GS):
a normal glomerulus scored 0, mild segmental GS affecting up to 25% of the
glomerular tuft scored 1, moderate GS affecting between 25 and 50% of the tuft
scored 2, and severe GS affecting in excess of 50% of the tuft scored 3.
Tubulointerstitial scarring: normal tubules (tubule cell number approximately
1000/x200 magnification field) and interstitium scored 0, mild tubular
atrophy (TA; tubule cell number approximately 800) and interstitial edema or
fibrosis (IF) affecting up to 25% scored 1, moderate TA (tubule cell number
approximately 600) and IF affecting 25 to 50% scored 2, and severe TA (tubule
cell number approximately 400) and IF scored 3. To determine the level of TA,
tubule cells per x200 magnification field were counted. The data were
collected from a minimum series of 12 randomly selected fields in the cortex
or such number of fields until 30 glomeruli had been counted.
In Situ End Labeling for the Detection of Apoptotic Cells
Using 10% neutral buffered formalin-fixed, paraffin-embedded sections,
fragmented nuclear DNA associated with apoptosis was labeled in situ
with digoxigenin-deoxyuridine (dUTP) by terminal deoxynucleotidyl transferase
(TdT), using the ApopTag Plus peroxidase kit (Appligene Oncor, Illkirch,
France) according to the manufacturer's instructions
(22). For negative controls,
slides were incubated in TdT buffer without TdT. For biochemically induced
positive controls, slides were pretreated with 10 µg/ml DNAse I (Sigma,
Poole, UK) in DNA buffer.
For each experimental animal, more than 30 glomerular cross sections and 20 high-power (x400 magnification) fields of tubulointerstitium were examined by two authors who were blinded to the experimental code (B.Y. and G.L.T.). The number of in situ end labeling (ISEL) positive-staining nuclei per glomerulus (Gapo), per 100 tubule cells (%) (Tapo), or per interstitial field (Iapo) was determined respectively. ISEL of DNA while associated with apoptosis can also be seen in necrotic (nonspecific DNA degradation) and mitotic (transient DNA strand break) cells. To substantiate the specificity of our results, we confirmed apoptosis by light microscopic evaluation of the characteristic morphologic features; only strongly positive ISEL cells with observable morphologic features of apoptosis, such as shrunken cells with condensed nuclei surrounded by a narrow cytoplasmic halo, were counted (2,23,24). Fewer than 5% of ISEL-labeled cells in both NTN and control kidneys failed to demonstrate morphologic features of apoptosis.
Evaluation of Cellular Proliferation and Inflammation by
Proliferating Cell Nuclear Antigen and ED1 Immunostaining
Cellular proliferation and inflammation were evaluated by proliferating
cell nuclear antigen (PCNA) and ED1 (a specific monocyte/macrophage marker)
immunostaining. Localization of PCNA and ED1 was performed in 10% (wt/vol)
neutral buffer formalin-fixed, paraffin-embedded kidney tissues by
immunohistochemistry using a standard avidin-biotin peroxidase complex
technique as described previously
(19). Primary antibodies
(monoclonal mouse anti-human PCNA [clone PC10; Dako, Glostrup, Denmark];
monoclonal mouse anti-rat ED1 antibody [Serotec Ltd., Oxford, UK]) both were
diluted 1:50 and applied overnight at 4°C in a humid atmosphere.
Thereafter, antibody binding was revealed using the ABC Elite Kit (Vector
Laboratories). 3'-amino-9-ethylcarbazole (AEC) was used as the
substrate. Sections were counterstained with hematoxylin and mounted in
Glycergel (Dako). Negative control sections were incubated with normal mouse
IgG at the same protein concentration as the primary antibody. The
immunohistochemical staining pattern of PCNA and ED1 was semiquantitatively
assessed using the same counting system used with ISEL staining.
Double Staining for Apoptosis with ED1 and -Smooth Muscle
Actin
Double immunohistochemical staining was undertaken on paraffin sections.
ISEL was carried out as described above. Before application of the
antidigoxigenin antibody, sections were preincubated with blocking serum for
30 min, labeled with the anti-ED1 or -smooth muscle actin (-SMA;
monoclonal mouse anti-human -SMA, diluted 1:250, Dako) antibody at
4°C overnight. Sections were labeled with biotinylated secondary
anti-mouse IgG at 37°C for 30 min and with alkaline phosphatase
streptavidin for another 30 min and were developed with Fast Red TR/Naphthol
AS-MX solution to produce a bright pink color. Subsequently,
anti—digoxigenin-peroxidase antibody was applied on the sections and the
development was achieved by the solution of diaminobenzidine to provide
positive staining as yellow brown. Control sections were incubated with
nonimmune normal mouse IgG in place of primary antibody and with the omission
of TdT enzyme as ISEL controls.
Northern Blot Analysis of Caspase-3 mRNA
Northern blot analysis was carried on the snap-frozen kidney tissues. Total
RNA was extracted using the TRIzol reagent (Life Technologies BRL, Paisley,
UK) and quantified by scanning spectrophotometer at 260 nm. Fifteen µg of
total RNA were electrophoresed on a 1% (wt/vol)
agarose/3-(N-morpholino)propanesulfonic acid (MOPS)/formaldehyde gel. RNA was
then transferred to a nylon membrane (Hybond-N, Amersham Life Science, Little
Chalfont, UK) by capillary blotting using 20xSSC and cross linked to the
nylon filter using an ultraviolet cross linker (Amersham Life Science) at 70
mJ/cm2 energy
(25).
To produce a caspase-3 cDNA probe, we amplified caspase-3 exonic DNA from rat cDNA by the PCR using the following previously published primers (13): 5'sense ATGGACAACAACGAAACCTCCGTG, 3'antisense CCACTCCCAGTCATTCCTTTAGTG. Amplification reactions were performed with 100 µM of each dNTP in amplification buffer (containing 1.5 mM MgCl2) and 1 unit Taq polymerase at 85°C for 5 min, before 20 pmol of primers were added. Thirty-nine cycles of amplification were completed using the following conditions; 94.6°C for 1 min, 48°C for 1 min, and 72°C for 2 min. The 850-bp PCR product was cloned into the pCR 2.1 vector (Invitrogen, Carlsbad, CA). After bacterial amplification and plasmid purification, the caspase-3 insert was excised with BstXI and EcoRV, separated by electrophoresis on a 1.5% (wt/vol) agarose TAE gel, and purified using Prep-A-Gene DNA Purification Systems (Bio-Rad Laboratories Ltd., Hertfordshire, UK). Product confirmation was by restriction mapping using EcoRI and KpnI. Purified cDNA was random-primed with 32P labeled dCTP (NEN, Boston, MA) using the Prime-a-Gene Labeling System (Promega, Southampton, UK). Unincorporated label was removed using a Sephadex G-50 NICK column (Pharmacia Biotech, St. Albans, UK).
Prehybridization and hybridization were performed using the Church buffer system (0.5 M sodium phosphate and 7% sodium dodecyl sulfate) at 65°C (26). The filter was washed three times in Church wash buffer (40 mM sodium phosphate, 1% sodium dodecyl sulfate) at 65°C for 20 min and then exposed to Kodak Biomax MS film for 24 h. Autoradiographs were quantitatively analyzed by scanning volume density using a Bio-Rad GS-690 densitometer and Molecular Analyst version 4 analysis software (Bio-Rad Laboratories Ltd.). Optical density values for caspase-3 were corrected for loading using the housekeeping gene cyclophilin (27). Results were expressed as percentage of control sample mRNA densities. Transcript size was determined by comparison to RNA molecular weight markers (Promega) using same analysis package and by visual comparison to the ribosomal RNA subunits.
Measurement of Tissue Caspase-3 Protein Level
Tissue levels of caspase-3 protein were determined by immunoprobing of
Western Blots. Twenty µg of protein was separated on a 15% (wt/vol)
polyacrylamide denaturing gel and then electroblotted onto Hybond ECL
nitrocellulose membranes (Amersham Life Science). Membranes were blocked by
the addition of 3% (wt/vol) bovine serum albumin in 0.1% (vol/vol) Tween 20
TBS (TTBS) at 4°C overnight before being probed with a polyclonal rabbit
anti-rat full-length caspase-3 at 1:2000 dilution in TTBS buffer at room
temperature for 2 h. Primary antibody binding was revealed using an
anti-rabbit peroxidase conjugate (Dako) diluted at 1:2000 in TTBS buffer for 1
h and the ECL chemiluminescence detection system (Amersham Life Science).
Recombinant caspase-3 (17 kD and 12 kD subunits; Sigma) was used to verify
antibody efficacy under experimental conditions. Developed films were
semiquantitatively analyzed by volume density using a Bio-Rad GS-690 scanning
densitometer and Molecular Analyst version 4 software. Translation size was
determined by comparison to protein molecular weight markers (Bio-Rad
Laboratories Ltd.) using the same analysis package.
Detection of Caspase-3 Activity in Renal Tissue
The activity of caspase-3 in tissue was detected by the modified
Fluorometric CaspACE Assay System (Promega). Kidney tissue (20 to 50 mg) from
control and NTN rats was ground in liquid nitrogen using a pestle and mortar.
A 1:9 (wt/vol) tissue:buffer extract was prepared in Tris/acetate buffer (pH
7.5) at 30°C (28). The
extract was centrifuged at 12,000 x g for 10 min, and the
supernatant was collected. A volume of supernatant equivalent to 100 µg of
protein was assayed for caspase-3 activity by the ability to cleave the
fluorogenic substrate Ac-DEVD-AMC. The specificity of the assay was determined
using the caspase-3 inhibitor Ac-DEVD-CHO by adding to the sample 30 min
before the substrate. Proteolytic cleavage of the substrates was monitored in
a fluorescence microplate reader (SOFTmax PRO, Molecular Devices Corp.,
Sunnyvale, CA) using an excitation wavelength of 360 nm and an emission
wavelength of 460 nm. The fluorescence intensity was calibrated with standard
concentrations of AMC, and the caspase-3 activity was calculated from the
slope of the recorder trace and expressed in picomoles per minute per µg of
protein.
Statistical Analyses
Results are expressed as mean ± SEM. The statistical difference was
assessed by a single factor variance (ANOVA) or t test. Linear
correlation analysis (GraphPad InStat, GraphPad Software, Inc., San Diego, CA)
and multiple linear regression analysis (SSPS; SPS Inc., Chicago, IL) were
applied to determine the correlation and association between parameters.
P < 0.05 was considered to be significant.
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Results |
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Detection of Apoptotic Cells
Apoptotic cells with distinct morphologic motifs were counted from
ISEL-stained sections and confirmed by morphology at light and electron
microscopy levels. Very few apoptotic cells were noted in glomeruli (0.01
± 0.01/glomerulus), tubules (0.01 ± 0.01%), and interstitium
(0.04 ± 0.04/x400 magnification field) of normal rabbit
serum-injected rats. In NTN kidneys, glomerular apoptosis was significantly
increased on days 7, 15, and 30 and peaked on day 7 (0.69 ±
0.15/glomerulus; Figure 1A). In
the tubules and interstitium of NTN kidneys, apoptosis was significantly
increased from day 15 onward, maximally on day 45 (0.83 ± 0.04% and
0.82 ± 0.10/x400 magnification field, respectively;
Figure 1, B and C). Maximal
areas of apoptotic cells were detected in the inflamed or sclerotic glomeruli
(Figure 2A) as well as in the
dilated or atrophied tubules (Figure
2B) and expanded interstitium
(Figure 2C). In positive
control sections that were treated with DNAse I before the TdT reaction,
nearly all of the cells stained, but most of the positive nuclei showed normal
shape and no cytoplasmic condensation (not shown). No staining was present in
the negative control sections using buffer that lacked TdT (not shown).
Electron microscopy confirmed apoptotic cells with distinct morphologic motifs
in some glomeruli, tubules, and interstitium
(Figure 2, D through F).
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Detection of Proliferation (PCNA +) and Inflammation (ED1 +)
In kidneys from nonimmune serum-injected rats, a few PCNA-positive cells
were noted in glomeruli (0.03 ± 0.02/glomerulus), tubules (0.37
± 0.10%), and interstitium (0.09 ± 0.03/x400 magnification
field; Table 1). In contrast,
in NTN kidneys, PCNA staining was dramatically and significantly increased in
the glomeruli throughout the experimental time course and peaked at day 7
(10.88 ± 1.53/glomerulus). In the tubules and interstitium, PCNA
staining was elevated by day 15, peaked on day 30 (3.09 ± 0.43% and
4.40 ± 0.93/x400 magnification field), and remained raised until
the end of the experiment (Table
1). Positive PCNA nuclei were localized in inflamed or sclerotic
glomeruli, dilated tubules, and expanded interstitium
(Figure 3, A through C).
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A small number of ED1-positive cells were seen in the nonimmune serum-injected rat kidneys in glomeruli (1.04 ± 0.32/glomerulus) and interstitium (3.76 ± 0.50/x400 magnification field; Table 1, Figure 3D). ED1 staining in NTN kidneys was significantly increased in glomeruli between day 7 and day 30, peaking on day 7 (37.63 ± 5.24/glomerulus), whereas in the interstitium, staining increased from day 7, reached a maximum on day 15 (27.17 ± 3.52/x400 magnification field), and remained high until the end of experiment (Table 1). Positive ED1 cells were distributed in inflamed glomeruli (Figure 3E), interstitium, and tubular lumen (Figure 3F).
Double Staining for ED1 and -SMA with Apoptosis
ED1 and ISEL double staining positive cells were found in inflamed
glomeruli (Figure 4A) and
tubulointerstitium (not shown). Some cells stained positively for both
apoptosis (ISEL) and -SMA in the interstitium
(Figure 4B).
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Expression of Caspase-3 mRNA
Northern blot analysis revealed a caspase-3 mRNA transcript at 2.7 kb
transcript (Figure 5). In
comparison with the control rat kidneys, the level of caspase-3 mRNA was
increased at all time points, significantly so on days 7, 30, and 45 (173.3%,
228.0%, and 241.7%, respectively; P < 0.05), reaching a peak on
day 45 (Figure 5).
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Tissue Levels of Caspase-3 Protein
A 24 kD band possibly representing a caspase-3 active subunit was gradually
and significantly increased with time in NTN kidneys compared with the
controls, with maximal expression on day 45 (6.08-fold). A 32 kD band,
representing the precursor of caspase-3, was found to be increased on day 45
in NTN kidneys (3.92-fold of controls; P < 0.01). The 29 kD
processing intermediate was also present in all kidneys
(Figure 6).
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To validate antibody efficacy further, we performed Western blots using recombinant active caspase-3 protein, kidney tissue from Wistar rats with apoptosis induced by SNx (5), rat proximal tubule cells with cisplatin-induced apoptosis, and normal human renal tissue (Figure 7). The full-length caspase-3 antibody strongly bound with 12 and 17 kD recombinant caspase-3 protein. In SNx Wistar rats, the antibody reacted with both 24 kD and 17 kD bands at a greater intensity than in a control animal. In a rat proximal tubule cell line treated with cisplatin, the antibody revealed a similar upregulation of a 24 kD but not a 17 kD band. In normal human renal tissue, a 20 kD band was detectable.
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Activity of Caspase-3
There was a significant increase of caspase-3 protease activity in NTN rat
kidneys at all time points compared with the controls
(Figure 8). On day 7, this
increase was 2.12-fold in controls and dropped to 1.85-fold on day 15 before
rising again on day 30 (2.29-fold) and reaching a peak on day 45 (2.38-fold).
The specific and competitive tetrapeptide inhibitor of caspase-3, Ac-DEVD-CHO,
almost fully inhibited the caspase-3 activity in the assays, demonstrating
assay specificity.
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Correlation among Renal Histology, Apoptosis, and Caspase-3
Cellular apoptosis correlated closely with inflammation and proliferation
in glomeruli (r = 0.676 and 0.656), tubules (r = 0.658, no
inflammation), and interstitium (r = 0.653 and 0.624) (all P
< 0.01). Apoptosis also correlated with GS (r = 0.412; P
< 0.05), TA, and IF (r = -0.917 and 0.917; P < 0.01)
in NTN kidneys. Multiple regression analysis showed that apoptosis was more
closely associated with TA and IF (Std ß coefficients = -0.674 and 0.612;
P < 0.01) than inflammation or proliferation and that tubular
apoptosis was a better predictor of changes in SCr and proteinuria (Std ß
coefficients = 0.448 and 0.577) than either glomerular or interstitial
apoptosis.
There was a parallel expression of caspase-3 mRNA (c3m), 24 kD protein (C3p-24), and activity (C3a) (r = 0.645, 0.742, and 0.757; P < 0.01). The levels of c3m, C3p-24, and C3a correlated with the cellular apoptosis in glomeruli (r = 0.565 and 0.432, except C3p-24 and Gapo), tubules (r = 0.622, 0.890, and 0.669), and interstitium (r = 0.629, 0.911, and 0.693) (all P < 0.01), which also correlated with the cellular inflammation in glomeruli (r = 0.365, P < 0.05, not C3p-24 and Ged1, and r = 0.529, P < 0.01) and interstitium (r = 0.722, 0.765, and 0.756; P < 0.01). Multiple regression analysis showed that caspase-3 activity was the best predictor of inflammation (Std ß coefficient = 0.773), whereas caspase-3 protein was the best predictor of apoptosis (Std ß coefficient = 0.759) (all P < 0.01).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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However, elevated apoptosis within a diseased kidney should not always be viewed as harmful. The apoptotic deletion of infiltrating neutrophils (myeloperoxidase positive) in glomerulonephritis limits neutrophil-mediated glomerular injury and may play a role in the resolution of glomerular inflammation. Failure of these mechanisms might lead to disintegration of neutrophils within the inflamed glomerulus and the development of persistent inflammation leading to scarring (37). Furthermore, the role of macrophage apoptosis may be a central regulator of the progression and resolution of macrophage-mediated tissue injury (34).
In this study, the peak in glomerular apoptosis (day 7) is likely to result
from the appropriate clearance of the inflammatory cells recruited into the
glomeruli during the initial immune response, as demonstrated by double
staining of ISEL and ED1. The later peak of apoptosis occurring from days 30
to 45 coincides with the severe TA and interstitium injury that characterizes
the late stages of this experimental model. However, determining the cellular
origin of apoptotic cells is complicated by changes in cell surface proteins
that occur early in apoptosis and complicating identification by double
staining for interstitial inflammatory cells (ISEL and ED1 positive) or
myofibroblasts (ISEL and -SMA positive), thus using location for
general identification is essential.
Given these two peaks of apoptosis, we demonstrated for the first time at the mRNA, protein, and activity levels significant increases in caspase-3 that coincide with elevated apoptosis. During the NTN time course, caspase-3 mRNA transcription was increased, particularly in the early inflammatory phase and late fibrotic phases, indicating the requirement for de novo synthesis of caspase-3 in both phases. This increase resulted in both elevated precursor and active caspase-3 protein, which were associated with a parallel change in the enzyme activity within NTN rat kidneys.
Caspase-3 is translated as an inactive 32 kD precursor that is proteolytically processed to become a functionally active enzyme (7,38,39,40). Activation of caspase-3 requires two proteolytic cleavage events: removal of the NH2 terminal prodomain generating a 29 kD processing intermediate that is subsequently cleaved into 17 kD and 11 kD or 12 kD subfragments (7,39,40). However, other active fragments, such as 20 kD and 18 kD, have also been reported (13,41). These subfragments then heterodimerize to form the activated protease (39,40). Western blot analysis of caspase-3 in this study revealed a 24 kD band that we believe is also an active caspase-3 protein. Interestingly, this band is approximately 4 kD bigger than the recognized largest active caspase-3 (13,41) and larger still than other reported active caspase-3 proteins (7,39,40). However, this was the smallest immunoreactive fragment that we could detect in renal tissue from Wistar Kyoto rats, and its expression correlates well with the increase in caspase-3 activity, whereas the other immune reactive bands do not. It therefore may represent a specific renal or Wistar Kyoto rat renal isoform of caspase-3. The fact that this full-length caspase-3 antibody binds strongly to recombinant 17 kD and 12 kD human caspase-3 protein and the recognized 20 kD band in human renal tissue (41) confirms the antibody's efficacy. Any question of species selectivity is answered by the ability of antibodies to bind to 17 kD caspase-3 in renal tissue from Wistar rats in which apoptosis was induced by SNx; however, the 24 kD band was also shown to increase in these remnant kidneys. Furthermore, the 24 kD band was the only nonprecursor/intermediate band detectable in rat proximal tubule cells (42) when apoptosis was induced by cisplatin (43), which suggests a renal- rather than a strain-specific isoform. Detection of this unusual caspase-3 subunit may be due to the use of this unique full-length caspase-3 antibody, rather than the more commonly used antibodies to the "active" subunits. In our hands, these antibodies, e.g., polyclonal rabbit anti-caspase-3 antibodies (Pharmingen, San Diego, CA) react with the 17 kD band in remnant kidneys from SNx Wistar rats but fail to show any active band in the NTN Wistar Kyoto rat kidneys even though activity and mRNA levels are comparable. This promotes the 24 kD band as an active caspase-3 subunit. However, without isolation and characterization of this 24 kD protein, its active status cannot be ratified.
Western blot analysis also revealed some interesting findings with regard to the 32 kD pro-caspase-3. Despite early progressive increases in activity and mRNA, there was little change in pro-caspase-3 until day 45. This suggests that all de novo pro-caspase-3 is immediately processed to the active form until this time. The accumulation of pro-caspase-3 may indicate that caspase processing becomes a limiting factor in the level of apoptosis or an attempt to prevent further increases in apoptosis despite increases in caspase-3 mRNA.
Although increased caspase-3 in this model of renal scarring is novel, it is not surprising given the well-documented role of caspases in the execution phase of apoptosis (8,44,45). Caspase-3 is potentially the most important effector enzyme in apoptosis, providing a common pathway to both death receptor- and mitochondria-dependent apoptotic mechanisms (8,46). Caspase-3 has also been linked to the pathogenesis of other models of renal injury associated with apoptosis. For instance, it was found to be upregulated at both mRNA and protein levels during reperfusion in a rat model of acute renal ischemia (13), whereas increased activity was reported after the administration of nephrotoxic doses of cyclosporin A in salt-depleted rats (14).
The pivotal role of caspase-3 in the apoptosis machinery makes it an
attractive target to regulate apoptosis-related cell death. In vitro,
the induction of apoptosis in mouse proximal tubule cells by cisplatin has
been inhibited by Ac-Asp-Glu-Val-Asp-H, a known caspase-3 inhibitor
(43). Application of this
therapeutic approach in vivo is clearly more problematic, although
there has been some success. For example, elevated apoptosis in hepatic
parenchymal cells during endotoxemia (tumor necrosis factor- mediated)
was prevented by injection of Z-VAD, providing a caspase-3 inhibition and
effective disease treatment
(47). It has been also been
reported that the administration of B-D-FMK (pan caspase inhibitor) was
significantly caspase-3 inhibiting and neuroprotective when given by
intracerebral or systemic injection after cerebral hypoxiaischemia
(16). More recent, it has been
reported that Z-VAD-FMK reduced the caspase-3 activity and prevented the early
onset of not only renal apoptosis but also inflammation and tissue injury in a
mouse model of renal ischemia
(17). Given these findings, a
similar blockade of caspase-3 in progressive renal scarring may provide a
novel therapeutic approach to the treatment of renal scarring by controlling
inappropriate apoptosis.
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J. Rossert, W. M. McClellan, S. D. Roger, and D. L. Verbeelen Epoetin treatment: what are the arguments to expect a beneficial effect on renal disease progression? Nephrol. Dial. Transplant., March 1, 2002; 17(3): 359 - 362. [Full Text] [PDF]
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