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*
Institute for Molecular and Cellular Regulation, Gunma University,
Maebashi, Japan.
Third Department of Internal Medicine, Gunma University School of
Medicine, Maebashi, Japan.
Correspondence to Dr. Itaru Kojima, Institute for Molecular and Cellular Regulation, Gunma University, Maebashi 371-8512, Japan. Phone: 81-27-220-8835; Fax: 81-27-220-8893; E-mail: ikojima{at}showa.gunma-u.ac.jp
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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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Activins are multifunctional cytokines that belong to the transforming growth factor-ß superfamily and regulate the growth and differentiation of cells in various organs (16,17). The actions of activins are modified at several levels by various factors. The most important factor that modulates the actions of activins is an activin-binding protein, namely follistatin (18). This protein stoichiometrically binds to activins and blocks their action (18,19). Follistatin is expressed on the surface of the target cells of activins by binding to the extracellular matrix (20). Activins trapped by follistatin are internalized by endocytosis and subsequently degraded by proteolysis (21). The expression of follistatin is regulated by various factors, including activins themselves in many tissues (22,23,24,25). Hence, the activin-follistatin system is a complex regulatory system that controls diverse cellular functions, including growth and differentiation during development (23,24,25).
Follistatin is abundantly expressed in tubular cells of adult kidneys (26). However, the function of the activin-follistatin system in adult kidneys is not well understood. We recently demonstrated that activin A is an autocrine factor that is produced in tubular cells and tonically inhibits branching tubulogenesis (27). Activin A produced in tubular cells plays a vital role because HGF, which is a morphogen known to regulate tubulogenesis (28), induces branching tubulogenesis mainly by reducing the production of activin A (27). Activin A is expressed in fetal kidneys (29) and was also demonstrated to inhibit branching morphogenesis of ureteric buds in organ culture (30). Activin A may act as a negative regulator of branching morphogenesis during kidney development (27,30,31). These results raise the possibility that the activin-follistatin system, i.e., morphomodulating proteins involved in tubulogenesis during kidney development, may influence the migration, growth, and differentiation of tubular cells after renal ischemia. In this study, we examined this possibility. The results indicate that the production of activin A is upregulated in tubular cells after renal ischemia. Blockade of endogenous activin A action by administration of follistatin protects the kidney from ischemic renal injury by preventing apoptosis and promoting regeneration.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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RNA Extraction and Northern Blot Analyses
Total RNA was extracted from kidney homogenates with the TRIzol reagent
(Life Technologies BRL, Grand Island, NY). Northern blot analyses were
performed as described previously
(27), using cDNA for rat
follistatin (provided by Dr. S. Shimasaki, Salk Institute, La Jolla, CA) and
cDNA for human glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (Clontech,
Palo Alto, CA). The membranes were subjected to autoradiography and analyzed
using a Fuji BAS 2000 (Fuji Photo Film, Tokyo, Japan). For quantification of
the relative follistatin mRNA content, the intensities of the autoradiographic
signals for follistatin and GAPDH were quantified and expressed in arbitrary
density units. The follistatin/GAPDH ratio (based on integrated signals) was
determined for each sample.
Reverse Transcription-PCR
Total RNA was isolated from whole kidneys with the TRIzol reagent (Life
Technologies BRL). First-strand cDNA was made from total RNA using Superscript
Preamplification System (Life Technologies BRL) according to the
manufacturer's instructions. Contaminating genomic DNA was removed with
RNase-free DNase. Five micrograms of DNase-treated RNA was incubated with 1
µl of oligo(dT) at 70°C for 10 min. Two microliters of 10x PCR
buffer, 1 µl of dithiothreitol (0.1 M), 2 µl of dNTP mixture (10 mM),
and 2 µl of MgCl2 (25 mM) were added to each reaction. After
incubation for 5 min at 42°C, 1 µl of reverse transcriptase was added.
Samples were incubated at 42°C for 50 min and then at 70°C for 15 min.
RNase H (1 µl) was added to each reaction, and samples were incubated at
37°C for 20 min. PCR was performed as indicated by the manufacturer
(Perkin-Elmer, Norwalk, CT), with the following primers: rat
ßA subunit: sense, 5'-GGACCTAACTCTCAGCCAGAGATG-3';
antisense, 5'-TCTCAAAATGCAGTGTCTTCCTGG-3'; rat activin type I
receptor: sense, 5'-GGTCTATGAGCAGGGGAAGATGAC-3'; antisense,
5'-ACATT- TTCGCCTTGCCAGC-3'; rat activin type II receptor: sense,
5'-AGATGGAAGTCACACAGCCCAC-3'; antisense,
5'-CAACACTGGTGCCTCTTTTCTCTG-3'; rat GAPDH: sense,
5'-CATGACCACAGTCCATGCCATC-3'; antisense, 5'-CACCCTGTT-
GCTGTAG- CCATATTC-3'. Reactions included 5 µl of 10x PCR
buffer, 2 µl of MgCl2 (50 mM), 1 µl of dNTP mixture, 1 µl
of 3'-primer, 1 µl of 5'-primer, 0.5 µl of Taq
polymerase, and 1 µl of cDNA. Samples were incubated at 94°C for 5 min,
followed by the indicated numbers of cycles of 30 s at 94°C, 30 s at
58°C, and 90 s at 72°C, with final extension at 72°C for 10 min,
in a Perkin-Elmer DNA thermal cycler. PCR used 30 cycles for the
ßA subunit and activin type I and II receptors and 18 cycles
for GAPDH. The levels of transcription of the GAPDH "housekeeping"
gene were found to be similar at all time points examined, enabling analysis
of the relative levels of expression of the desired genes. Reactions without
cDNA were used as negative controls. Rat hepatocyte cDNA was used as a
positive control in each experiment. Reactions were repeated at least
twice.
Measurement of DNA Synthesis
DNA synthesis in renal tubular cells was measured using bromodeoxyuridine
(BrdU) (32). At the indicated
times after reperfusion, BrdU (100 mg/kg), an analogue of thymidine, was
injected intraperitoneally into every animal used in the study. After 1 h,
rats were euthanized, and the kidneys were removed and fixed with 4%
formaldehyde for 24 h. Sections were immunostained using a cell proliferation
kit (Amersham, Tokyo, Japan), as described previously
(33). The number of
BrdU-positive cells was counted in five randomly selected fields of the outer
medulla, using a light microscope at x400 magnification. The
proliferation index was measured in five sections per rat, and the average of
the five determinations was calculated. The results were indicated as the
numbers of BrdU-positive cells per square micrometer.
In Situ Hybridization
The cRNA probe was transcribed from a pBluescript SK(+) vector containing
an approximately 845-bp PstI/XbaI fragment derived from rat
follistatin cDNA (provided by Dr. S. Shimasaki, Salk Institute). Linearized
plasmids were used for in vitro transcription of
digoxigenin-11-UTP-labeled antisense and sense riboprobes with SP6 and T7 RNA
polymerase, respectively, according to the instructions provided by the
manufacturer (Boehringer Mannheim, Mannheim, Germany). Eight-micrometer
sections were mounted on poly-L-lysine-coated slides. After digestion with 5
µg/ml proteinase K at room temperature for 30 min, sections were postfixed
in 0.4% paraformaldehyde at 4°C for 20 min and incubated overnight at
50°C with hybridization buffer containing 1 µg/ml digoxigenin-labeled
cRNA. The buffer contained 50% formamide, 10 mM Tris-HCl (pH 7.5), 600 mM
NaCl, 1 mM ethylenediaminetetraacetate, 0.25% sodium dodecyl sulfate, 1
x Denhardt's solution, 200 µg/ml yeast tRNA, and 10% dextran sulfate.
After hybridization, sections were washed in 2x SSC/50% formamide at
58°C for 30 min, incubated in 1 µg/ml RNase A solution at 37°C for
30 min, and then washed once in 2x SSC and twice in 0.2x SSC at
50°C, for 20 min each time. Sections were then incubated in a 1:500
diluted solution of polyclonal sheep anti-digoxigenin Fab antibody conjugated
with alkaline phosphatase, before washing and detection of the label with
nitroblue tetrazolium chloride and 5-bromo-4-chloro-3-indolylphosphate.
Terminal Deoxynucleotidyl Transferase-Mediated
dUTP-Nick-End-Labeling
For identification of nuclei with DNA strand breaks at the cellular level,
the terminal deoxynucleotidyl transferase-mediated dUTP-nick-end-labeling
(TUNEL) method (34) was
performed, using an apoptosis in situ detection kit (Wako, Tokyo,
Japan). Quantification of TUNEL-positive cells was performed by counting
positive nuclei in tubular cells from five randomly selected fields of the
outer medulla, with a light microscope at x400 magnification. Apoptosis
was measured as a percentage of total tubular cells in five sections per rat
kidney, and the average of the five determinations was used as an apoptotic
index.
Immunohistochemical Analyses
The localization of activin A was examined immunohistochemically with an
avidin-biotin coupling immunoperoxidase technique, using a Vectastain Elite
ABC kit (Vector Laboratories, Burlingame, CA) according to the instructions
provided by the manufacturer. Briefly, the paraffin-embedded sections (4
µm) were deparaffinized and rehydrated in a routine manner. After
inactivation of endogenous peroxidase with 1% metaperiodic acid in
phosphate-buffered saline (PBS) for 10 min at room temperature, sections were
preincubated with normal goat serum for 60 min. The sections were then
incubated with a polyclonal rabbit anti-activin A antibody for 2 h, washed
with PBS, and reacted with a biotinylated goat anti-rabbit IgG for 1 h. After
washing with PBS, sections were reacted with Vectastain Elite ABC reagent. The
antibody was detected with diaminobenzidine tetrahydrochloride in PBS, and the
sections were counterstained with hematoxylin. For immunohistochemical
controls, the primary antibody was replaced with 5% normal goat serum in PBS,
which did not demonstrate positive staining, thus confirming specificity. The
anti-human activin A antibody used in this study recognizes dimers as well as
monomers of the rat ßA subunit
(35).
The localization of type II and type IIB activin receptors was examined in a similar manner. Polyclonal rabbit anti-activin type II receptor and anti-activin type IIB receptor antibodies were generously provided by Dr. K. Miyazono (Cancer Institute, Tokyo, Japan).
Histologic Analyses
Sections were cut at 4 µm and stained with periodic acid-Schiff stain.
Sections were microscopically examined for lesions 1, 2, 3, and 5 d after
reperfusion. The changes observed were limited to the outer medulla, where
tubular damage is most obvious, and were graded as follows: 0, normal; 1,
areas of tubular dilation, necrosis, hemorrhage, and cell desquamation
involving <20% of the fields; 2, similar changes involving >20% but
<40% of the fields; 3, similar changes involving >40% but <60% of the
fields; 4, similar changes involving >60% of the fields. Five sections per
rat were used for analysis. The results were expressed as mean ± SEM
for each experimental group.
Statistical Analysis
The significance of differences between means was compared by t
test. P < 0.05 was considered significant.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Changes in the Expression of Follistatin after Ischemic Renal
Injury
We then examined changes in the expression of follistatin mRNA by using
Northern blotting (Figure 5).
Consistent with previous reports
(26), follistatin mRNA was
abundantly expressed in normal kidneys. Sham operations did not affect
follistatin mRNA expression levels. However, ischemia/reperfusion injury
transiently decreased follistatin mRNA expression in the kidneys
(Figure 5A). The relative
levels of follistatin mRNA in ischemic kidneys were dramatically reduced at 24
and 48 h and returned to basal levels 72 h after reperfusion
(Figure 5B).
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Next, we examined the localization of follistatin mRNA in normal, sham-operated, and ischemic kidneys by in situ hybridization (Figure 6). In normal kidneys, hybridization signals were observed mainly in the inner stripe of the outer medulla, without any detectable hybridization in the cortex or inner medulla (Figure 6, A to C). Follistatin mRNA was distributed in tubular cells (Figure 6D). No staining was observed in any cells of the glomeruli (Figure 6A). Hybridization signals in ischemic kidneys were considerably reduced, compared with normal kidneys (Figure 6E). Hybridization with a control sense probe demonstrated no positive signals (Figure 6F).
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Effect of rh-Follistatin on Renal Function after Ischemic Renal
Injury
To assess the role of endogenous activin A in renal regeneration, we
administered rh-follistatin (25 µg/kg) or saline solution to rats with
ischemic renal injuries. First, we evaluated parameters of renal function
(Figure 7). In control
(saline-treated) rats, the serum creatinine levels peaked 12 h after
reperfusion and rh-follistatin significantly reduced the peak creatinine
concentration (Figure 7A). The
serum BUN levels peaked 24 h after reperfusion and returned to normal levels
72 h after reperfusion. Peak BUN levels were significantly lower for
rh-follistatin-treated rats than for control rats
(Figure 7B). Electrolyte
concentrations were not affected by rh-follistatin administration (data not
shown). No significant differences in body weights between the control and
rh-follistatin-treated groups were observed at the indicated times
(Figure 7C).
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Effect of rh-Follistatin on Histologic Changes Induced by Renal
Ischemia
We next examined whether exogenous follistatin affects the histologic
changes induced by ischemia. Control (saline-treated) kidneys exhibited signs
of congestion, hemorrhage, tubular dilation, cast formation, and tubules
plugged with desquamated epithelial cells after ischemic renal injury
(Figure 8, A and B). In
contrast, rh-follistatin-treated kidneys exhibited no cast formation and fewer
tubules were dilated (Figure
8D). Apoptotic tubular cells with highly condensed nuclear
chromatin were also observed in control kidneys
(Figure 8C) but not in
rh-follistatin-treated kidneys. Neutrophil infiltration was observed in both
control kidneys and rh-follistatin-treated kidneys (data not shown).
Semiquantitative analysis demonstrated that the damaged tubular areas in
rh-follistatin-treated kidneys were smaller than those in control kidneys
(Figure 8E).
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Effect of rh-Follistatin on Apoptosis Induced by Ischemic Renal
Injury
We next examined the effect of rh-follistatin on renal ischemia-induced
apoptosis by using the TUNEL method (Figure
9). TUNEL-positive cells were not observed in normal or
sham-operated kidneys (Figure 9, A and
B). In contrast, TUNEL-positive cells were predominantly localized
in the outer medulla of control (saline-treated) kidneys after ischemic renal
injury (Figure 9C); only a few
positive cells were observed in the cortex and inner medulla (data not shown).
Consistent with light-microscopic examination results, the number of
TUNEL-positive cells in rh-follistatin-treated kidneys was markedly lower than
that in control kidneys (Figure
9D). Reduction of the number of TUNEL-positive cells by exogenous
follistatin was observed 24 and 48 h after reperfusion
(Figure 9E).
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Effect of rh-Follistatin on DNA Synthesis in Tubular Cells after
Ischemic Renal Injury
Proliferation of tubular cells is the hallmark of early regeneration after
ischemic renal injury (1). To
investigate the effect of exogenous rh-follistatin on cell proliferation after
ischemic renal injury, we measured DNA synthesis in renal tubular epithelial
cells (Figure 10). In normal
(Figure 10A) and sham-operated
(data not shown) kidneys, BrdU-positive cells were rarely observed.
Twenty-four hours after ischemic renal injury, BrdU-positive cells were
observed predominantly in the outer medulla of control (saline-treated)
kidneys (Figure 10B), where
damage to the tubular cells was most obvious, but not in other regions (data
not shown). In contrast, the number of BrdU-positive cells in
rh-follistatin-treated kidneys was significantly greater than that in control
kidneys (Figure 10, C and D). Quantitative analysis demonstrated that intravenous infusion of rh-follistatin
enhanced renal tubular epithelial cell proliferation
(Figure 11). Enhancement of
DNA synthesis by rh-follistatin was observed at concentrations of 
15
µg/kg.
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Effect of rh-Activin A on DNA Synthesis in Tubular Cells after
Ischemic Renal Injury
To further assess the role of the activin-follistatin system in renal
regeneration, we administered rh-activin A (12.5 µg/kg) to rats with
ischemic renal injuries. Exogenously administered activin A did not affect
renal function after ischemic injury (data not shown) but inhibited DNA
synthesis in tubular cells. The number of BrdU-positive cells in ischemic
kidneys treated with rh-activin A was significantly lower than that in
ischemic kidneys treated with saline solution
(Figure 11).
Effect of rh-Follistatin on DNA Synthesis in Tubular Cells in Intact
Kidneys
To elucidate the mechanism of DNA synthesis enhancement by exogenous
follistatin, we administered rh-follistatin to normal rats and examined
whether tubular cell proliferation in intact kidneys was induced by
rh-follistatin administration. Exogenously administered follistatin did not
induce tubular cell proliferation (Figure
11) or affect tubular structure or renal function (data not
shown).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Apoptosis triggered by ischemia potentially contributes to the development of postischemic tubular cell death (38). A previous study demonstrated a biphasic pattern of tubular cell apoptosis after renal ischemia, suggesting that apoptosis observed in the early phase after renal ischemia is mainly attributable to the ischemic injury and that late-phase apoptosis is attributable to the programmed removal of tubular cells for control of excessive proliferation (39). In this study, the expression of activin A was observed 12 h after reperfusion, which preceded the appearance of TUNEL-positive tubular cells. Exogenous follistatin reduced the number of TUNEL-positive cells 24 and 48 h after reperfusion but not thereafter (Figure 9E). This result suggests that, after renal ischemia, upregulated endogenous activin A may be involved in the induction of early-phase apoptosis of tubular cells.
Inflammation is also considered to be one of the most important causes of
tissue injury in organs subjected to ischemia. A series of recent studies
suggested a novel role for activin in inflammation and repair processes in
various organs
(36,40,41,42).
For example, increased expression of activin was observed in various types of
inflammatory processes, including cutaneus wound repair and inflammatory
arthropathies. The level of activin expression was correlated with the degree
of inflammation in inflammatory bowel disease. Strong expression of activin A
was induced in vitro by proinflammatory cytokines such as
interleukin-1 and tumor necrosis factor-
, which are known to be
released from macrophages and stromal cells at the sites of tissue injury and
inflammation (43). Therefore,
proinflammatory cytokines are possible inducers of activin expression in this
model. With respect to the mechanism of action of activin, the release of
activin into the circulation precedes the release of proinflammatory cytokines
after lipopolysaccharide treatment, suggesting a proinflammatory action of
activin (41). In contrast,
activin A produces anti-inflammatory effects by blocking the action of
interleukin-6 (42). The action
of activin in inflammatory processes probably depends on the type of cell or
tissue. It was recently reported that inhibition of ischemia-induced apoptosis
prevents inflammation and subsequent tissue injury
(44). Our results demonstrated
that rh-follistatin reduced the number of apoptotic cells
(Figure 9). Therefore, it is
possible that rh-follistatin exerts an anti-inflammatory action. Although we
could not observe rh-follistatin modulation of neutrophil infiltration in this
study, we cannot completely exclude the possibility that endogenous activin
regulates the degree of inflammation after renal ischemia, including
myeloperoxidase activity, the degree of monocyte/macrophage infiltration, and
the tissue levels of proinflammatory cytokines. This issue remains to be
addressed.
Our results demonstrate for the first time that blockade of the action of endogenous growth inhibitors is an alternative to the infusion of growth factors for acceleration of renal regeneration. Follistatin probably potentiates multiple types of endogenous growth factors involved in renal regeneration. Follistatin may also enhance the effects of any exogenous growth factors. The advantage of using follistatin is that it accumulates in the kidneys after intravenous administration (45). In the kidneys, follistatin binds to heparan sulfate proteoglycan (20) and remains in the extracellular matrices for several days (45). The stability of follistatin may be beneficial for the treatment of acute renal failure after a variety of insults.
In summary, the activin-follistatin system is altered in kidneys after ischemic injury. Endogenous activin A may function to promote apoptosis and inhibit regeneration of renal tubular cells. Blockade of the actions of activin with the administration of follistatin accelerates recovery from ischemic renal injuries. Follistatin has therapeutic potential for the prevention and treatment of tubular damage leading to acute renal failure.
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Acknowledgments |
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) or saline solution ([UNK])
at the time of reperfusion, and serum creatinine levels (A), blood urea
nitrogen (BUN) levels (B), and body weight (C) were measured at the indicated
time points. Values are mean ± SEM. *, P < 0.01
versus saline solution.
