Rapamycin impairs recovery from acute renal failure: role of cell-cycle arrest and apoptosis of tubular cells

Wilfred Lieberthal1, Robert Fuhro1, C. Christopher Andry2, Helmut Rennke3, Vivian E. Abernathy1, Jason S. Koh1, Robert Valeri4, and Jerrold S. Levine5

1 Renal Section, Evans Memorial Department of Clinical Research, Department of Medicine, 4 Naval Blood Research Laboratory, and 2 Department of Pathology, Boston University Medical Center, Boston 02118; 3 Department of Pathology, Brigham and Women's Hospital, Harvard Medical School, Boston, Massachusetts 02115; and 5 Section of Nephrology, Biological Sciences Division, University of Chicago, Chicago, Illinois 60637


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The immunosuppressive effect of rapamycin is mediated by inhibition of interleukin-2-stimulated T cell proliferation. We report for the first time that rapamycin also inhibits growth factor-induced proliferation of cultured mouse proximal tubular (MPT; IC50 ~1 ng/ml) cells and promotes apoptosis of these cells by impairing the survival effects of the same growth factors. On the basis of these in vitro data, we tested the hypothesis that rapamycin would impair recovery of renal function after ischemic acute renal failure induced in vivo by renal artery occlusion (RAO). Rats given daily injections of rapamycin or vehicle were subjected to RAO or sham surgery. Rapamycin had no effect on the glomerular filtration rate (GFR) of sham-operated animals. In rats subjected to RAO, GFR fell to comparable levels 1 day later in vehicle- and rapamycin-treated rats (0.25 ± 0.08 and 0.12 ± 0.05 ml · min-1 · 300 g-1, respectively) (P = not significant). In vehicle-treated rats subjected to RAO, GFR increased to 0.61 ± 0.08 ml · min-1 · 300 g-1 on day 3 (P < 0.02 vs. day 1) and then rose further to 0.99 ± 0.09 ml · min-1 · 300 g-1 on day 4 (P < 0.02 vs. day 3). By contrast, GFR did not improve in rapamycin-treated rats subjected to RAO over the same time period. Rapamycin also increased apoptosis of tubular cells while markedly reducing their proliferative response after RAO. Furthermore, rapamycin inhibited activation of 70-kDa S6 protein kinase (p70S6k) in cultured MPT cells as well as in the renal tissue of rats subjected to RAO. We conclude that rapamycin severely impairs the recovery of renal function after ischemia-reperfusion injury. This effect appears to be due to the combined effects of increased tubular cell loss (via apoptosis) and profound inhibition of the regenerative response of tubular cells. These effects are likely mediated by inhibition of p70S6k.

ischemia; proliferation; 70-kDa S6 protein kinase


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

RAPAMYCIN IS A POTENT IMMUNOSUPPRESSIVE agent that acts by inhibiting the proliferation and clonal expansion of interleukin (IL)-2-stimulated T cells (8, 32, 34, 35, 39). Rapamycin induces growth arrest of proliferating T cells in the mid-to-late G1 phase of the cell cycle (8, 34, 35, 39). This effect of rapamycin is mediated by inhibition of 70-kDa S6 protein kinase (p70S6k), a kinase necessary for cell-cycle progression. Although the role of p70S6k in cell-cycle progression has been most clearly established in lymphocytes, there is limited but convincing evidence of a more generalized role for p70S6k in cell-cycle progression. Thus rapamycin has been shown to inhibit the proliferation of a number of nonimmune cell types, including hepatocytes, vascular smooth muscle cells, and fibroblasts (7, 31, 38, 51).

In addition to its effects on proliferation, rapamycin may also play a role in regulation of apoptosis under certain circumstances (14, 42). All cells express the apoptotic machinery necessary for cell death. Cell viability depends on constant inhibition of this apoptotic pathway by so-called "survival factors" (40, 47). Cell death induced by the loss of these survival factors occurs via the "default pathway" of apoptosis (40, 47). We have recently reported that rapamycin promotes apoptosis of cultured peritoneal macrophages by blocking the survival activity of macrophage survival factors such as lysophosphatidic acid (LPA) (19).

It is important to note that the default pathway of apoptosis is distinct from all other triggers of apoptosis. As opposed to receptor-mediated events (e.g., Fas/FasL) or cytotoxic stimuli, which trigger cell death by their presence, the default pathway of apoptosis is triggered by the absence of survival factors. By showing that rapamycin can partially inhibit the survival activity of known macrophage survival factors, we have provided novel evidence that signaling through p70S6k can modulate the default pathway of apoptosis, at least in macrophages. These data are strengthened by the fact that peritoneal macrophages are fully differentiated and therefore nonproliferative, so that any effects of rapamycin on cell survival must be independent of cell cycle-associated events.

Available data from preclinical studies and human trials of rapamycin have demonstrated little nephrotoxicity (46). This result is not surprising, as the differentiated tubular epithelium of the kidney is normally quiescent, so that proliferation and/or apoptosis of these cells would be uncommon events. During acute renal failure (ARF), however, the severity of tubular injury and the rate of recovery of tubular function depend on a balance between proximal tubular cell death (26, 27) and the ability of sublethally injured tubular cells to enter the cell cycle and proliferate (15, 17, 18, 49). We therefore hypothesized that adverse effects of rapamycin mediated through inhibition of p70S6k may become apparent only when tubular cells are proliferating, as occurs during the recovery phase of ARF (18, 49).

As there is no information on the effects of rapamycin on tubular cells, we first determined whether rapamycin alters the growth and/or fate of cultured mouse proximal tubular (MPT) cells. We provide novel data that rapamycin potently inhibits the proliferation of MPT cells. Moreover, as we reported for macrophages (19), rapamycin also induces apoptosis of MPT cells by inhibiting the survival activity of several renal growth factors. Importantly, rapamycin does not alter the kinetics or severity of apoptosis induced by "positive regulators" of apoptosis such as cytotoxic agents.

Next, using an in vivo model, we show that rapamycin markedly delays the recovery of renal function after experimental ischemic ARF but has no adverse effect on renal function in sham-operated rats. We also provide evidence that the deleterious effect of rapamycin on the course of ARF in vivo is due, at least in part, to the combined effects of impaired regeneration and enhanced apoptosis of renal tubular cells. In light of our in vitro findings, we propose that the enhanced rate of apoptosis seen in rats treated with rapamycin is the result not of cytotoxicity but rather of decreased inhibition of the default pathway of apoptosis. Furthermore, we show that rapamycin inhibits the activation of p70S6k in cultured MPT cells as well as in the renal tissue of rats subjected to ARF. Because rapamycin is a highly specific inhibitor of p70S6k (8, 9, 34, 39), our data strongly suggest that the observed effects of rapamycin on proliferation and apoptosis are mediated by p70S6k inhibition. As ARF is a common complication after transplantation of the kidney (12), heart (12), liver (5), and bone marrow (50), we believe these findings are of substantial clinical relevance.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Reagents

LPA, rapamycin, soybean trypsin inhibitor, trypan blue, culture medium (DMEM and Ham's F-12), penicillin/streptomycin, and 3-(4,5-dimethylthiazol)-2,5-diphenyl tetrazolium bromide (MTT) were obtained from Sigma (St. Louis, MO). Collagenase was purchased from Worthington Biochemical (Freehold, NJ). Epidermal growth factor (EGF) and Hoechst dye (H-33342) were obtained from Calbiochem (San Diego, CA).

Experiments in Primary Cultures of MPT Cells

Culture methods. Cells were cultured from collagenase-digested fragments of proximal tubules obtained from the cortices of C57BL/6 mice using a method we have previously described in detail (20, 41). Briefly, kidneys were carefully dissected to obtain cortical tissue, which was finely minced with a razor blade and incubated at 37°C for 45 min in Hanks' solution containing collagenase and soybean trypsin inhibitor (1.0 mg/ml each). After large undigested fragments were removed by gravity, the suspension was mixed with an equal volume of 10% calf serum in Hanks' solution and then centrifuged at 500 rpm for 2 min at room temperature. The pellets were washed once by centrifugation and then suspended in a defined growth medium. Proximal tubules adhere to the wells, and proximal tubular cells grow out of these tubules. The "growth medium" for MPT cells is a mixture of DMEM and Ham's F-12 (1:1) containing 2 mM L-glutamine, 15 mM HEPES, 5 µg/ml human transferrin, 5 µg/ml insulin, 50 µM hydrocortisone, 500 U/ml penicillin, and 50 µg/ml streptomycin. We have previously identified these cells as being predominantly (>95%) of proximal tubular origin (20, 41). MPT cells growing out of the tubules form confluent monolayers 5 days after plating. The proliferating cells are fed with fresh medium 2 and 5 days after plating. Once MPT cells reach confluence, they are fed three times a week.

Experimental procedure. MPT cells reach confluence 5 days after plating of tubules. All experiments were started 24 h after MPT cells reached confluence, which was denoted day 1 of the experimental period. On day 1 experimental media were added to the cells as described for each experiment (see RESULTS). For the measurement of thymidine uptake and cell proliferation, experiments were ended after 24 h. For experiments assessing cell survival and apoptosis, the experiment was ended after 10 days.

Trypan blue assay. The trypan blue exclusion test was used to determine the number of viable cells remaining after 10 days of incubation under experimental conditions. We have previously described our method for the trypan blue assay (22, 29). Viable cells were defined as cells that remained adherent to the culture dish and excluded trypan blue. Nonadherent cells that, as we have previously shown, are almost all apoptotic (22, 29) were removed by two washes with ice-cold PBS. Adherent cells were harvested by incubation with 0.05% trypsin/0.53 M EDTA · 4 Na for 10 min at 37°C. Trypsin was neutralized by addition of DMEM containing 10% calf serum. Cells were then centrifuged for 5 min at 100 g and resuspended in 1 ml DMEM. Trypan blue (0.04 g/dl) was added for 10 min, and the number of viable cells excluding trypan blue was counted in a hemocytometer.

The effect of rapamycin on MPT cell survival was determined by comparing the number of viable cells present on day 1 of the experiment (control) with that 10 days after incubation in either growth factor-containing or growth factor-free medium plus or minus rapamycin. The number of viable MPT cells after 10 days of incubation is expressed as a percentage of the number of viable MPT cells present on day 1 of the experiment.

Thymidine uptake. At the end of the experiment, [3H]thymidine (2 µCi/mmol; New England Nuclear, Boston, MA) was added to each culture dish. After a 2-h incubation at 37°C, cells were washed three times with ice-cold PBS. Cell DNA was then extracted by incubating cells in ice-cold 5% trichloroacetic acid (TCA) for 1 h at 4°C. After removal of TCA, cells were washed once with fresh TCA, and ice-cold ethanol containing 200 µM potassium acetate was added to each well for 5 min. Then, cells were incubated twice for 15 min in a 3:1 mixture of ethanol and ether. After the monolayers were allowed to air dry, the remaining material (predominantly DNA) was solubilized in 0.1 N sodium hydroxide. Aliquots were added to scintillation fluid and counted in a scintillation counter (model 1600TR Tri-Carb Liquid Scintillation Analyzer, Packard Instruments, Meriden, CT).

Cell proliferation assay. MPT cells were harvested from confluent monolayers and replated in culture wells at a density of ~10,000 cells/well in the presence of 10% serum, EGF (10 nM), or LPA (22 µM). Rapamycin was added to wells containing each growth factor in concentrations ranging from 0 to 100 ng/ml. Four days later, the relative number of cells per well was determined using the MTT assay (19, 36). This assay is based on the ability of mitochondria of viable cells to cleave the tetrazolium rings of the pale yellow MTT dye to a dark blue formazan product. The amount of formazan product produced is directly proportional to the number of viable cells present in a culture dish (36). After removal of medium, 200 µl of MTT (1 mg/ml) were added to each well. After incubation at 37°C for 4 h, the MTT formazan was dissolved by adding 200 µl of 10% SDS in 0.01 N HCl and incubated overnight to solubilize the formazan crystals. Absorbance of aliquots from each well were read by a Dynatech microELISA Plate Reader (Dynatech, Chantilly, VA) with a test wavelength of 570 nm and a reference wavelength of 650 nm. The absorbance of cells incubated in the presence of rapamycin was expressed as a percentage of the absorbance of control wells to which no rapamycin was added.

Identification of apoptosis of MPT cells by nuclear morphology. Nuclear morphology was assessed by fluorescence microscopy of MPT cells stained with the supravital DNA dye H-33342, which enters live as well as dead cells (26, 27). The nuclei of apoptotic cells are readily distinguished from those of viable and necrotic cells on the basis of chromatin condensation, nuclear fragmentation, and increased intensity of H-33342 fluorescence. MPT cells grown on glass slides were incubated in the presence of 10 nM EGF with or without rapamycin (10 ng/ml). After 10 days, the cells were stained with H-33342 by addition of the dye at a concentration of 1 µg/ml. After cells were incubated with the dye at 37°C for 10 min, the cells were fixed with 3.8% paraformaldehyde and mounted on slides with gelvatol. Cells were photographed under fluorescence microscopy at ×100 magnification.

DNA electrophoresis. DNA electrophoresis is a well-established method, independent of nuclear morphology, for determining the presence of apoptosis. In apoptosis, DNA is cleaved into single or multiple nucleosomal fragments (each nucleosomal fragment being ~180 base pairs in length), resulting in the classic "ladder pattern" on DNA electrophoresis (22). MPT cells were incubated for 5 days in EGF-containing medium in the absence or presence of rapamycin. Adherent and detached MPT cells were pooled for DNA extraction and electrophoresis. Cells were lysed in 0.5% Triton X-100, 5 mM Tris · HCl, pH 7.4, and 20 mM EDTA for 30 min at 4°C. After centrifugation at 15,000 g for 20 min, the supernatants were extracted with phenol-chloroform. Then, 3.0 M sodium acetate, pH 5.2 (1/10 vol), and 1.0 M MgCl2 (1/30 vol) were added, and the DNA was precipitated in ethanol. Samples were separated by electrophoresis on a 1.2% agarose gel containing ethidium bromide.

Quantitation of apoptosis using flow cytometry. MPT cells were stained with H-33342, using the same technique as described for immunofluorescent microscopy, and then placed immediately on ice. Flow cytometry was performed on an Epics ESP Flow Cytometer (Coulter Electronics, Hialeah, FL) with an ultraviolet-enhanced argon laser. Hoechst fluorescence was accomplished by excitation with <5 mW of ultraviolet laser light (351- to 364-nm multiline) and detected with a 525-nm band-pass optical filter. Data were analyzed by Epic Elite software (Coulter Electronics). A constant number of events were analyzed for each sample (10,000/sample).

We used two criteria to distinguish normal from apoptotic cells: cell size and the intensity of H-33342 fluorescence. Forward-angle scatter, a measure of relative cell size (shown on the x-axis), was plotted against the intensity (log scale) of H-33342 fluorescence (shown on the y-axis). Viable cells, which have low H-33342 fluorescence and are normal in size, are located predominantly in the bottom right quadrant of the graph. By contrast, apoptotic cells, which are smaller in size (and therefore characterized by a relatively lower forward-angle scatter) and intensely fluorescent, are located predominantly in the top left quadrant of the graph. This approach is a modification of methods used by other investigators (37, 43, 44). We have previously confirmed the reliability of this methodology by separating the two populations by cell sorting using flow cytometry and examining them by electron microscopy (19, 22).

Experiments In Vivo

Model of ARF. Male Sprague-Dawley rats, weighing between 275 and 350 g, were used for all experiments. Rats were fed regular Purina rat chow (Purina Mills, Chicago, IL) and allowed free access to water. Anesthesia was induced with an intraperitoneal injection of Nembutal (pentobarbital sodium; 55 mg/kg). Rats were placed on a thermostatically controlled heated table. Body temperature was monitored via a temperature probe in the rectum and maintained between 36 and 38°C.

ARF was induced in anesthetized rats by removing the left kidney and then clamping the right renal artery for 40 min, as we have previously described (24). The right renal artery was mobilized by careful blunt dissection. A noncrushing vascular clamp was placed across the right renal artery, and the kidney was carefully visualized to ensure that the entire kidney blanched in response to the clamp. The abdomen was then closed. After 40 min of renal artery occlusion (RAO), the abdomen was opened again and the vascular clamp was removed. After removal of the clamp, reperfusion of the kidney was verified visually, and the abdominal wall was then sutured closed. The animals were allowed to regain consciousness and were given free access to food and water. Animals were anesthetized for a second time 1, 3, or 4 days after RAO for measurement of renal function.

In sham-operated rats, the left kidney was removed and the right renal artery was mobilized by blunt dissection using techniques comparable to those used for rats in the ARF groups. However, for sham-operated rats, the renal artery was not clamped. The abdomen was closed for 40 min, then opened and closed again to reproduce the procedure followed in the experimental (ARF) groups of animals. Animals were anesthetized 3 or 4 days later for measurement of renal function.

Administration of rapamycin to rats. Rapamycin (0.05 mg/300 g) or its vehicle (0.1 ml DMSO-EtOH) was given intraperitoneally daily for 3 days before rats were subjected to ARF or sham surgery. On day 4 of rapamycin and/or vehicle administration, the animals were anesthetized and subjected to 40 min of renal ischemia or sham surgery, as described above. Rapamycin or vehicle was continued daily until animals were killed 1, 3, or 4 days after surgery.

Experimental groups. There was a total of 10 experimental groups. Vehicle-treated rats subjected to renal injury were examined 1 day (group 1, n = 8), 3 (group 2, n = 10), and 4 days (group 3, n = 8) after ARF. Rapamycin-treated ARF rats were similarly studied 1 day (group 4, n = 8), 3 (group 5, n = 10), and 4 days (group 6, n = 7) after renal injury. Sham rats treated with vehicle or rapamycin were studied 3 (groups 7 and 9, n = 7) and 4 days (group 8 and 10, n = 7) after sham surgery.

Measurement of renal function. After RAO or sham surgery, rats were reanesthetized 1, 3, or 4 days later with an intraperitoneal injection of Inactin (thiobarbital sodium; 55 mg/kg) and placed on a thermostatically controlled heated table, with body temperature maintained between 36 and 38°C, as described above. A tracheostomy was performed, and PE-240 tubing was placed in the trachea to ensure an adequate airway. A femoral artery was cannulated with PE-50 tubing for blood pressure monitoring and blood sampling. A bladder catheter (PE-90) was placed via a suprapubic incision for urine sampling. The left internal jugular vein was cannulated with PE-50 tubing for infusion of inulin. Mean arterial pressure (MAP) was monitored continuously using a pressure transducer, and body temperature was measured using a rectal probe (24, 25).

Glomerular filtration rate (GFR) was measured using classic inulin clearance techniques (23, 25). Briefly, [3H]inulin was infused at a rate of 0.06 µCi/min. Blood and urine samples were obtained during three consecutive 20-min clearance periods. The blood samples were centrifuged, and the hematocrit was measured. Samples of plasma and urine (5 µl each) were added separately to 3 ml of liquid scintillation fluid and counted for 10 min in a Packard 1600TR Tri-Carb Liquid Scintillation Counter. Sodium and potassium concentrations were measured in plasma and urine samples using ion-specific electrodes. Inulin clearance and the fractional (FENa) and absolute excretion of sodium and potassium were calculated using standard formulas (25).

Kidney morphology. Sections of kidney were immersion fixed in 10% formaldehyde (in PBS) and imbedded in paraffin. Coronal sections of paraffin-imbedded tissue were cut so that the cortex, outer medulla, and inner medulla were present in each section. The sections were stained with periodic acid-Schiff and examined by light microscopy. One of the coauthors (H. Rennke) examined coded slides of kidney sections from each of the 10 experimental groups in blinded fashion, quantified the proportion of necrotic tubules within the outer medulla of each section, and expressed the data as a percentage of the total number of tubules per microscopic field. The data were decoded and analyzed statistically by another coauthor (Lieberthal W).

Immunohistochemical staining for proliferating cell nuclear antigen. Kidney tissue sections were randomly chosen from 3 rats within each of the 10 experimental groups (30 tissue sections altogether) and coded by one of the coauthors (Lieberthal W). Paraffin-embedded sections of the coded kidney tissue samples were stained with antibody to proliferating cell nuclear antigen (PCNA). The paraffin-embedded sections of kidney were incubated at 37°C for 30 min. They were treated sequentially with xylene, absolute alcohol (EtOH), 80% EtOH, 70% EtOH, and finally ddH2O. For antigen (PCNA) retrieval, the sections were immersed in a buffer (3.75 g/l glycine and 10 mM EDTA, pH 3.6) and microwaved twice for 5 min. The slides were allowed to cool to room temperature and then washed with PBS, followed by 1% egg albumin at room temperature for 30 min. The sections were then incubated with a murine monoclonal anti-PCNA antibody (1:50 dilution in 1% egg albumin), followed by peroxidase-conjugated anti-rabbit antibody (Jackson Laboratories) and stained with the Dako EnVision system. PCNA-positive nuclei stained bright red. The sections were not counterstained. The slides were mounted using an aqueous-based medium. The immunohistochemically stained sections, still coded, were randomly photographed using phase-contrast microscopy (×40 magnification). Fields restricted to the outer medulla of the kidney sections were photographed. A coauthor of this study (Abernathy VE) counted the number of PCNA-positive cells per coded photograph. The data were then decoded and analyzed by another coauthor (Lieberthal W).

Quantitation of apoptotic cells in kidney tissue. Apoptotic cells were quantified in kidney sections by light microscopy of formaldehyde-fixed kidney sections stained with hematoxylin and eosin (H & E). One of the coauthors (Levine JS) randomly chose slides of three kidney sections from each experimental group and coded the slides so they could be photographed in blinded fashion. Another coauthor (Lieberthal W) took 36 photographs of the cortical region of each kidney section at random. Thus 108 photographs were taken of each experimental group. Sections were photographed at ×1,000 magnification, which allowed identification of apoptotic cells and bodies. Cells containing condensed or fragmented nuclei (which have a characteristic appearance on H & E stain) were considered apoptotic. When the condensed nuclei of apoptotic cells had fragmented into multiple pieces, all the fragments were considered as having originated from a single apoptotic cell. Only apoptotic cells and bodies that appeared to be of tubular origin were counted. Because we could not be certain that free apoptotic cells in the tubular lumen were tubular cells, these were excluded from the analysis. After all the photographs had been quantitated in this fashion for each tissue section, the results were given to a coauthor (Levine JS), who broke the code and analyzed the data. The final data are expressed as the number of apoptotic cells per five microscopic fields.

Effect of Rapamycin on p70S6k Activity

The state of activation of p70S6k was assessed by immunoblotting, using two anti-p70S6k antibodies of different specificity. Although activation of p70S6k requires phosphorylation at a number of serine and theronine residues, the activity of p70S6k is most closely related to the state of phosphorylation of the Thr412 residue (1, 48). We used an antibody (Upstate Biotechnology, Lake Placid, NY) that only recognizes p70S6k when phosphorylated at Thr412. In addition, we used an antibody (Upstate Biotechnology) that recognizes both the inactive and activated (total) forms of the kinase.

Harvesting of tissue. Cultured cells were washed once with ice-cold PBS and then lysed in ice-cold lysis buffer. The lysis buffer consisted of RIPA buffer containing anti-proteases and kinase inhibitors. The RIPA buffer contained 20 mM Tris base, pH 7.4; 140 mM NaCl; 0.5 g/dl deoxycholate; 1 g/dl SDS; 1 g/dl Triton X-100; and 10 g/dl glycerol. Immediately before use, we added the following to the lysis buffer: 1 mM DTT, 1 mM PMSF, 10 µg/ml aprotinin, and 10 µg/ml pepstatin.

To measure p70S6k activity in vivo, samples of kidney tissue (obtained 2 days after sham surgery or RAO) were snap-frozen in liquid nitrogen and stored at -80°C. On the day of immunoblotting, ice-cold lysis buffer (identical to that used for cultured cells) was added to the samples. Each sample was subjected to sonication, followed by homogenization with a Dounce homogenizer. The protein concentration of each sample was measured.

Imunoblotting procedure. Lysates containing comparable amounts of protein (15 µg) were boiled for 15 min in sample buffer and loaded onto a 10% polyacrylamide SDS gel. Proteins were transferred electrophoretically to Immobilon-P membranes (Millipore, Bedford, MA), which were washed in 150 mM NaCl, 100 mM Tris · HCl, pH 7.5, and 0.05% Tween 20 (TBST) and blocked for 1 h at room temperature in TBST containing 10% wt/vol nonfat powdered milk (MTBST). The membranes were then incubated with the primary antibody that recognizes only p70S6k phosphorylated at Thr412 (1:1,000 dilution in TBST) at 4°C overnight and then washed three times with TBST. The membranes were then incubated with the secondary antibody (horse anti-rabbit antibody conjugated to horseradish peroxidase; 1:10,000 dilution in TBST; Amersham). After a 2-h incubation at room temperature, filters were washed three times with TBST. Bound secondary antibody was detected using the ECL enhanced chemiluminescence system (Amersham Pharmacia Biotech, Piscataway, NJ).

To determine the amount of total p70S6k present in each lane, the membranes were stripped by incubation at 60°C for 7 min in a solution containing Tris · HCl (pH 6.8), 10% SDS, and 0.7% basal Eagle's medium. We then probed for total p70S6k using the same methods described above except that a different primary antibody that recognizes the combination of both activated and inactive forms of p70S6k was used.

Calculations

GFR (ml · min-1 · 300 g-1) was determined by calculating the inulin clearance as previously described (23) and then correcting the values for body weight. FENa was calculated as the ratio of urine to plasma concentrations of sodium divided by the ratio of urine to plasma concentrations of inulin × 100.

Statistical Analysis

All data are expressed as means ± SE. Measurements of GFR and electrolyte excretion were obtained during three consecutive "clearance periods" (23). Variability of all parameters between clearance periods was <5%, with no significant temporal trend. We therefore averaged data from the three periods for each variable to obtain one value per animal for each variable. All comparisons of two groups were made with the Student's t-test. The Bonferroni correction was used when more than two groups were compared. A P value <0.05 was considered significant.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Experiments in Cultured MPT Cells

Rapamycin inhibits growth factor-induced proliferation of MPT cells. Confluent monolayers of MPT cells were incubated for 24 h in medium containing either 10% serum, 10 nM EGF, or 22 µM LPA. Rapamycin was added in concentrations ranging from 0 to 100 ng/ml. After 24 h of incubation, thymidine uptake was measured. Rapamycin inhibited thymidine uptake by MPT cells in a dose-dependent fashion (Fig. 1A).


View larger version (20K):
[in this window]
[in a new window]
 
Fig. 1.   Effect of rapamycin on mouse proximal tubular (MPT) cell thymidine uptake and proliferation. A: thymidine uptake. Confluent MPT cell monolayers were incubated in DMEM containing 1 of 3 growth factors: serum (10%; ), epidermal growth factor (EGF; 10 nM; open circle ), or lysophosphatidic acid (LPA; 22 µM; down-triangle). Rapamycin was added in concentrations varying from 0.1 to 100 ng/ml. After incubation at 37°C for 24 h, tritiated thymidine uptake was measured over a 1-h period. The effect of rapamycin is expressed as %inhibition of thymidine uptake. B: MPT cell poliferation. MPT cells were harvested from confluent monolayers and replated at a density of ~10,000 cells/well in the presence of serum (10%; ), EGF (10 nM; open circle ), or LPA (22 µM; triangle ). Four days later, the relative number of cells was determined by an 3-(4,5-dimethylthiazol)-2,5-diphenyl tetrazolium bromide (MTT) assay. Rapamycin inhibited both thymidine uptake (A) and cell growth (B) with a comparable IC50 of 1-10 ng/ml. *P < 0.01 for 100 vs. 10 ng/ml rapamycin. dagger P < 0.01 for 5 vs. 0.1 ng/ml rapamycin.

As an independent measure of the effect of rapamycin on tubular cell growth, MPT cells were also harvested from confluent monolayers and replated at 10,000 cells/well in medium containing either 10% serum, 10 nM EGF, or 22 µM LPA in the presence of varying doses of rapamycin. The number of cells per well was determined 24 h after the experiment was initiated. As shown in Fig. 1B, rapamycin also inhibited MPT proliferation in a dose-dependent fashion.

The IC50 of rapamycin for both thymidine uptake and cell growth was comparable (1-10 ng/ml) (Fig. 1) and corresponds to published values for the IC50 of rapamycin on inhibition of p70S6k activity.

Rapamycin promotes apoptosis of MPT cells by inhibiting the survival effect of renal growth factors. We have previously shown that rapamycin inhibits the activity of survival factors (such as LPA) on peritoneal macrophages, thereby inducing apoptosis in these cells (19). Macrophages, unlike MPT cells, are terminally differentiated, nonproliferating cells (19). We therefore determined whether rapamycin would similarly inhibit the survival activity of renal growth factors for MPT cells, a proliferative cell type. The viability of MPT cells, after incubation in growth factor-free medium for 10 days, was reduced to 30 ± 2% of control (freshly confluent) monolayers (Fig. 2). Two renal growth factors, EGF (10 nM) and LPA (22 µM) (22), both markedly improved MPT survival over the same 10-day period to 91 ± 2 and 96 ± 2% of control, respectively (Fig. 2). Rapamycin significantly reduced cell viability in the presence of EGF and LPA to 60 ± 3 and 51 ± 2% of control, respectively (P < 0.05). In contrast, rapamycin had no effect on the survival of MPT cells incubated for 10 days in the absence of growth factors (Fig. 2).


View larger version (18K):
[in this window]
[in a new window]
 
Fig. 2.   Effect of rapamycin on cell viability. Confluent MPT cell monolayers were incubated in growth factor-free medium or in medium containing either 10 nM EGF or 22 µM LPA. Rapamycin (10 ng/ml) or vehicle was added to each condition. After 10 days, cell viability was determined and expressed as a percentage of viability of freshly confluent cells. Cell viability was markedly reduced in cells incubated in the absence of growth factors. Rapamycin had no effect on the viability of MPT cells in the absence of growth factors but markedly reduced viability of MPT cells incubated with serum, EGF, or LPA. *P < 0.05 compared with growth factor-free medium with or without rapamycin. dagger P < 0.05 compared with cells incubated with EGF in the absence of rapamycin. Dagger P < 0.05 compared with LPA in the absence of rapamycin.

The reduction in cell viability of MPT cells in the presence of rapamycin could theoretically be due to two processes: decreased proliferation and/or increased apoptosis of MPT cells. We used three distinct methods to determine whether rapamycin promoted apoptosis of MPT cells: nuclear morphology (via immunofluorescence microscopy), DNA electrophoresis, and flow cytometry.

First, MPT cells were grown on coverslips in the presence of EGF alone (10 nM) or EGF+rapamycin (10 ng/ml). After 10 days, cells were stained with H-33342 and examined under fluorescence microscopy. The nuclei of MPT cells incubated with EGF appear normal; the intensity of H-33342 fluorescence is low; and a normal chromatin pattern is seen (Fig. 3). By sharp contrast, MPT monolayers incubated for the same period of time in EGF+rapamycin contain many cells whose nuclei show characteristic changes of apoptosis. These include increased intensity of fluorescence, chromatin condensation, and nuclear fragmentation (Fig. 3).


View larger version (83K):
[in this window]
[in a new window]
 
Fig. 3.   Fluorescence microscopy of MPT cells incubated in EGF with (right) and without rapamycin (left). Confluent MPT cells were incubated for 10 days in medium containing 10 nM EGF in the absence or presence of rapamycin (10 ng/ml). Cells were then stained with H-33342 dye (see MATERIALS AND METHODS) and photographed by fluorescence microscopy (×400 magnification). The nuclei of MPT cells incubated with EGF in the absence of rapamycin show normal morphology. By contrast, many of the cells incubated with EGF in the presence of rapamycin show features typical of apoptosis. These include increased intensity of fluorescence, nuclear condensation (arrows), and, in some cases, fragmentation of chromatin (arrowheads). Thus rapamycin induces apoptosis of MPT cells in the presence of EGF.

Second, electrophoresis of DNA from MPT cells incubated for 10 days in the presence of EGF alone shows minimal degradation, consistent with fully viable cells (Fig. 4). In contrast, DNA from MPT cells incubated with EGF+rapamycin shows degradation in a distinct ladder pattern, indicative of the oligonucleosomal cleavage of DNA that is characteristic of apoptosis (Fig. 4).


View larger version (77K):
[in this window]
[in a new window]
 
Fig. 4.   Electrophoresis of DNA from vehicle-treated and rapamycin-treated MPT cells. Confluent MPT cells were incubated in EGF-containing medium in the presence of rapamycin (10 ng/ml) or vehicle for 10 days. DNA was then extracted from cells and subjected to electrophoresis. DNA from cells cultured with EGF in the absence of rapamycin shows little fragmentation. However, in cells incubated with EGF+rapamycin, DNA fragmentation is evident in a ladder pattern characteristic of apoptosis.

Finally, apoptosis induced by rapamycin was quantified using flow cytometry (Fig. 5). After incubation of MPT cells in the presence of EGF alone for 10 days, a small proportion of the cells were apoptotic (~10%) (Fig. 5, top). Apoptosis was increased to 24% when cells were incubated with EGF+rapamycin (Fig. 5, middle). In the absence of EGF and rapamycin, the proportion of apoptotic cells was the highest among the groups studied (~44%) (Fig. 5, bottom). The presence of rapamycin in cells incubated without EGF did not change the number of apoptotic cells compared with cells that were deprived of EGF in the absence of added rapamycin (data not shown). Comparable results were obtained when the same studies were done with 22 µM LPA instead of EGF (data not shown).


View larger version (70K):
[in this window]
[in a new window]
 
Fig. 5.   Quantification of apoptosis by flow cytometry. Confluent MPT were cultured for 10 days with either EGF alone (top), EGF+rapamycin (middle), or growth factor-free medium (bottom). Cells were harvested, stained with H-33342 and analyzed by flow cytometry on the basis of size (forward scatter) and the fluorescent intensity of nuclei stained with H-33342. Viable cells are located in the bottom right quadrant of the graph (normal size, low H-33342 fluorescence), whereas apoptotic cells are found predominantly in the top left quadrant (small size, intense H-33342 fluorescence). Thus rapamycin partially inhibits the antiapoptotic (survival) activity of EGF.

These data demonstrate for the first time that rapamycin induces apoptosis in cultured renal proximal tubular cells by partially inhibiting the survival activity of growth factors.

Rapamycin does not alter cell death induced by cytotoxic agents such as ATP depletion or cisplatin toxicity. To determine whether rapamycin enhances apoptosis induced by ischemic or toxic injury to tubular cells, we examined the effect of rapamycin on apoptosis induced by ATP depletion or cisplatin (28, 30). The viability of MPT cells exposed to the mitochondrial inhibitor antimycin (2 µM) for 5 days was comparable in the presence and absence of rapamycin (75 ± 2 and 80 ± 3% of control, respectively). Similarly, the viability of MPT cells exposed to cisplatin (25 µM) for 5 days was comparable in the presence and absence of rapamycin (50 ± 5 and 45 ± 2%, respectively). Thus, in sharp contrast to the proapoptotic effect of rapamycin on MPT cells cultured in the presence of renal growth factors, rapamycin had no effect on the kinetics or severity of apoptosis induced by an ischemic or toxic trigger. These data indicate that the proapoptotic effects of rapamycin are limited to inhibition of the survival activity of growth factors, with consequent activation of the default pathway of apoptosis.

In Vivo Experiments in Rats

Effect of rapamycin on GFR after sham surgery or induction of ARF. We next determined whether rapamycin would have a similar effect on renal tubular cell proliferation and survival in vivo. Rats were subjected to either sham surgery or RAO for 40 min as a model of ischemic ARF. For sham-operated rats given vehicle, GFR at 3 and 4 days after surgery was 1.54 ± 0.07 and 1.89 ± 0.07 ml · min-1 · 300 g-1, respectively. Treatment of sham-operated rats with rapamycin did not significantly alter GFR at either 3 or 4 days (1.72 ± 0.07 and 1.75 ± 0.22 ml · min-1 · 300 g-1, respectively). We conclude that rapamycin does not have a deleterious effect on GFR in animals with normal kidneys over the 4 days of observation.

In marked contrast, rapamycin markedly impaired the recovery of GFR in rats subjected to ischemic ARF. One day after RAO, the GFR of vehicle-treated and rapamycin-treated rats was reduced to a comparably severe degree (0.25 ± 0.08 and 0.12 ± 0.05 ml · min-1 · 300 g-1, respectively) (Fig. 6A). Thereafter, vehicle-treated rats showed progressive improvement in their GFR, with values increasing to 0.61 ± 0.08 ml · min-1 · 300 g-1 by day 3 (P < 0.02 vs. day 1) and to 0.99 ± 0.09 ml · min-1 · 300 g-1 by day 4 after surgery (P < 0.02 vs. day 3) (Fig. 6A). Although GFR on day 4 was substantially improved compared with that on day 1 in vehicle-treated ARF rats, it was still lower than the GFR measured on the same day in vehicle-treated sham-operated rats (P < 0.02). Thus GFR does not fully recover within 4 days in vehicle-treated rats. By marked contrast to the vehicle-treated group, GFR measured 3 and 4 days after RAO in rapamycin-treated ARF rats showed no improvement and was not significantly different from the GFR on day 1. The GFR of rapamycin-treated rats was 0.39 ± 0.10 ml · min-1 · 300 g-1 on day 3 after RAO (P = not significant vs. day 1) and 0.48 ± 0.11 ml · min-1 · 300 g-1 on day 4 (P < 0.02 vs. day 1) (Fig. 6A). The GFR in vehicle- and rapamycin-treated animals became statistically different on day 4 after RAO (Fig. 6A) (P < 0.02). We conclude from these data that rapamycin impairs recovery of GFR after ischemic injury.


View larger version (32K):
[in this window]
[in a new window]
 
Fig. 6.   Effect of rapamycin on renal function after renal ischemia-reperfusion injury in rats. A: glomerular filtration rate (GFR) in rats treated with vehicle (open bars) or rapamycin (hatched bars) was measured 1, 3, and 4 days after induction of acute renal failure (ARF). After 1 day, GFR is reduced to a comparable degree in vehicle- and rapamycin-treated rats. GFR improves progressively in vehicle-treated rats 3 and 4 days postsurgery, whereas there is no recovery of GFR in rapamycin-treated rats over the same period. B: fractional sodium excretion (FENa) in rats treated with vehicle (open bars) or rapamycin (hatched bars) was measured 1, 3, and 4 days after ARF. After 1 day, FENa is increased to a comparable degree in vehicle- and rapamycin-treated rats. FENa improves progressively in vehicle-treated rats after 3 and 4 days, whereas there is no change in FENa in rapamycin-treated rats over the same period. *P < 0.02 compared with vehicle-treated animals 1 day after ischemia-reperfusion. dagger P < 0.02 compared with vehicle-treated animals examined 3 days after renal injury. Dagger P < 0.02 compared with vehicle-treated animals examined 4 days after renal injury.

Effect of rapamycin on FENa after sham surgery and ARF. FENa was normal in vehicle-treated, sham-operated rats 3 and 4 days after sham surgery (0.06 ± 0.02 and 0.05 ± 0.02%, respectively). Rapamycin did not significantly alter FENa in sham-operated rats (0.08 ± 0.03 and 0.04 ± 0.01% at 3 and 4 days, respectively). As expected, induction of ARF significantly increased FENa on day 1 after surgery in both vehicle- and rapamycin-treated rats (3.33 ± 0.47 and 4.14 ± 1.18%, respectively) (Fig. 6B). These values are not statistically different. In vehicle-treated ARF rats, FENa improved progressively to 0.59 ± 0.10% by day 3 (P < 0.01 vs. day 1) and to 0.09 ± 0.01% by day 4 (P < 0.01 vs. day 3). FENa in vehicle-treated ARF rats on day 4 was not different from that of sham-operated rats 4 days after surgery (P = not significant). By contrast, FENa in rapamycin-treated rats remained elevated 3 (2.72 ± 0.68%) and 4 days (2.50 ± 0.46%) after surgery (both P = not significant vs. day 1). FENa of rapamycin-treated rats was still markedly elevated at 4 days after surgery (P < 0.02 vs. vehicle-treated rats on the same day) (Fig. 6B).

Renal morphology. The morphology of the outer stripe of the medulla was normal in sham vehicle-treated and rapamycin-treated rats studied 2 and 3 days after sham surgery (data not shown). In rats subjected to RAO, the proportion of necrotic tubules 1 day after surgery was not as increased to a comparable extent in vehicle-treated and rapamycin-treated rats (34 ± 5 and 40 ± 7%, respectively) (Fig. 7). Representative micrographs of renal tissue on day 1 in both groups are provided in Fig. 8. In vehicle-treated animals, the proportion of necrotic tubules fell by days 3 and 4 to 6 ± 2 and 3 ± 1%, respectively (P < 0.01 vs. day 1 for both days 3 and 4) (Fig. 7). In sharp contrast, the proportion of necrotic tubules in rapamycin-treated rats 3 and 4 days after ARF remained elevated and was comparable to that on day 1 post-RAO (26 ± 6 and 28 ± 4%, respectively) (Fig. 7). Representative micrographs of renal tissue on day 4 show substantial improvement in renal morphology in vehicle-treated rats (less tubular injury, more patent tubules) but no improvement in the morphology from kidneys of rapamycin-treated rats (Fig. 8).


View larger version (33K):
[in this window]
[in a new window]
 
Fig. 7.   Quantitation of necrosis of tubular cells in vehicle- and rapamycin-treated rats after ischemia-reperfusion injury. One day after induction of ARF in vehicle- and rapamycin-treated rats (open and hatched bars, respectively), a large proportion of necrotic tubular cells were evident in the outer renal medulla. The degree of necrosis was comparable in the 2 groups 1 day after surgery. By 3 and 4 days after surgery, the severity of necrosis diminished markedly in vehicle-treated rats. By contrast, the extent of tubular cell necrosis was unchanged over the same period of observation in rapamycin-treated rats. *P < 0.01 compared with vehicle-treated animals one day after renal ischemia-reperfusion injury. dagger P < 0.02 compared with vehicle-treated animals on the same day after renal injury.



View larger version (112K):
[in this window]
[in a new window]
 
Fig. 8.   Morphology of kidneys from vehicle (A)- and rapamycin-treated rats (B) after ischemia-reperfusion injury. Sections of kidney stained with periodic acid-Schiff and photographed at ×400 magnification are shown. One day after ischemia-reperfusion, severe morphological injury is evident in kidneys from both vehicle-treated (top left) and rapamycin-treated rats (top right). In both groups there is extensive necrosis of tubular cells and obstruction of the tubular lumen by casts. In vehicle-treated ARF rats 4 days after surgery, renal morphology was considerably improved, with less necrosis and reduced intratubular cast formation (bottom left). By contrast, renal morphology in rapamycin-treated rats 4 days after surgery (bottom right) shows no improvement compared with day 1.

Proliferation of renal tubular cells after sham surgery and ARF. Our next set of studies addressed the mechanism of this deleterious effect of rapamycin on recovery from ischemic ARF. Our in vitro data using MPT cells suggested two potential contributory mechanisms, namely, inhibition of proliferation of recovering tubular cells and induction of apoptosis through blockade of the survival activity of renal growth factors. We used PCNA staining to determine whether rapamycin's inhibitory effect on proliferation extended to an in vivo setting. Three days after sham surgery, there were few PCNA-positive cells in kidney sections from vehicle-treated and rapamycin-treated kidneys [7 ± 2 and 2.5 ± 1/high-power field (hpf), respectively] (Fig. 9).


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 9.   Quantitation of proliferating cell nuclear antigen (PCNA)-positive cells in kidney tissue from vehicle- and rapamycin-treated rats after ischemia-reperfusion injury (open bars and hatched bars, respectively). Rapamycin reduces the number of PCNA-positive cells in rats subjected to renal artery occlusion (RAO). *P < 0.01 compared with vehicle-treated ARF rats on the same day post-ARF.

In vehicle-treated rats, the number of PCNA-positive cells was increased to 74 ± 20, 94 ± 17, and 44 ± 13/hpf on days 1, 3, and 4, respectively, after RAO (Figs. 9 and 10). In rapamycin-treated rats subjected to RAO, the number of PCNA-positive cells was substantially lower than in vehicle-treated rats on days 1 (3.1 ± 1/hpf), 3 (27 ± 12/hpf), and 4 (4.7 ± 1/hpf) (Fig. 9). Representative micrographs taken 3 days after sham surgery show few PCNA-positive cells in kidney tissue from vehicle- and rapamycin-treated rats. In rats subjected to RAO, there were many more PCNA-positive cells 3 days after surgery in vehicle-treated than in rapamycin-treated animals (Fig. 10). Thus rapamycin inhibits proliferation of renal tubular cells both in vitro and in vivo.


View larger version (91K):
[in this window]
[in a new window]
 
Fig. 10.   Immunohistochemistry of PCNA-stained kidney sections from vehicle (A) and rapamycin-treated rats (B) after ischemia-reperfusion injury. Representative light microscopic fields of kidney sections obtained 3 days after surgery and stained for PCNA are shown. There are few PCNA-positive nuclei in kidney sections from sham-operated animals, whether vehicle-treated (top left) or rapamycin-treated (top right). The number of PCNA-positive nuclei in rats subjected to RAO is greater in those treated with vehicle (bottom left) than for those treated with rapamycin (bottom right).

Apoptosis of renal tubular cells after sham surgery and ARF. To examine the effects of rapamycin on apoptosis in vivo, we used nuclear condensation and fragmentation as markers of cells undergoing apoptosis (Fig. 11). As expected, very few apoptotic cells were detected in sham-operated rats. At most, 1 apoptotic nucleus (out of a total of 108 fields) was detected in either vehicle- or rapamycin-treated rats at 3 and 4 days post-sham surgery (data not shown).


View larger version (138K):
[in this window]
[in a new window]
 
Fig. 11.   Morphological features of apoptotic tubular cells in kidney sections. Light microscopic fields of kidney sections stained with hematoxylin and eosin and photographed at ×1,000 magnification are shown. Representative photographs of kidney sections are given to demonstrate the typical features of apoptosis on light microscopy. Top left: 2 apoptotic cells (arrows) are present opposite one another within the same tubule. Both cells show the same morphological features of apoptosis, including condensation of nuclear chromatin, cell shrinkage, and loss of adhesion to adjacent tubular cells. Top middle: a single apoptotic cell (arrow) in which the condensed nucleus has undergone fragmentation into multiple fragments of varying size. Top right and bottom left and right: apoptotic bodies that have been phagocytosed by adjacent tubular cells. Each arrow points to fragments of condensed chromatin considered to have originated from a single apoptotic cell.

However, as has been previously reported (26, 27), apoptotic tubular cells were present in kidney sections of rats subjected to ARF (Fig. 12). On day 1 after RAO, the number of apoptotic cells was comparable in kidney sections from vehicle- and rapamycin-treated rats (0.22 ± 0.02 and 0.21 ± 0.01/5 microscopic fields, respectively) (Fig. 12). When examined 3 and 4 days after RAO, the number of apoptotic cells in vehicle-treated ARF rats remained unchanged (0.17 ± 0.02 and 0.14 ± 0.03/5 microscopic fields, respectively). However, in kidneys of rapamycin-treated rats subjected to ARF, the number of apoptotic cells was substantially higher on days 3 (0.35 ± 0.03/5 microscopic fields) and 4 (0.35 ± 0.0203/5 microscopic fields) post-RAO than in vehicle-treated rats at the same time points postsurgery (P < 0.01) (Fig. 12). Thus rapamycin increased the number of cells dying by apoptosis in the kidneys of rats subjected to ischemic ARF. We therefore conclude that the in vivo effects of rapamycin on apoptosis are comparable to those observed in vitro.


View larger version (32K):
[in this window]
[in a new window]
 
Fig. 12.   Quantitation of apoptosis of renal tubular cells in the kidneys from vehicle- and rapamycin-treated rats subjected after ischemia-reperfusion injury. Apoptosis is increased in kidneys of rapamycin-treated rats during the recovery period after ischemia-reperfusion injury. One day after RAO, the number of apoptotic cells in vehicle- and rapamycin-treated rats (open and hatched bars, respectively) was comparable. However, at 3 and 4 days after injury, the number of apoptotic cells in rapamycin-treated ARF rats increased and was statistically greater than that in vehicle-treated ARF rats at the same time points. There were approximately twice the number of apoptotic cells in kidney sections from rapamycin-treated vs. vehicle-treated rats at 3 and 4 days after ARF. *P < 0.01 compared with vehicle-treated ARF rats on the same day.

Taken together, our data suggest that rapamycin impairs recovery of GFR after ischemic ARF through at least two mechanisms, namely, inhibition of proliferation and induction of apoptosis.

Body weight. Body weight before surgery was comparable in all groups (data not shown). Rats treated with vehicle or rapamycin lost a comparable amount of weight after sham surgery (Table 1). Weight loss was greater in vehicle-treated ARF rats than in vehicle-treated sham rats. Rapamycin increased weight loss in ARF rats (Table 1).

                              
View this table:
[in this window]
[in a new window]
 
Table 1.   Effect of rapamycin on body weight in sham-operated and ARF rats

Effect of rapamycin on p70S6k activity in MPT cells and renal tissue in vivo. Activation of p70S6k requires phosphorylation at a number of serine and theronine residues (1). However, phosphorylation of Thr412 is essential for activation, and the activity of p70S6k is most closely related to the state of phosphorylation at Thr412 (1, 48). We therefore used an antibody specific for p70S6k that is phosphorylayed at the Thr412 residue to assess the effect of rapamycin on its activity. An antibody that recognizes total p70S6k was used to establish that equal amounts of p70S6k were added to each lane during SDS-PAGE electrophoresis (see MATERIALS AND METHODS)

Confluent MPT cell monolayers were incubated in growth factor-containing medium with varying concentrations of rapamycin (0-50 ng/ml). The cells were lysed 24 h later, and p70S6k activity was determined by immunoblotting. Rapamycin inhibited activation of p70S6k cultured MPT cells over concentrations ranging from 0.5 to 50 ng/ml (Fig. 13).


View larger version (37K):
[in this window]
[in a new window]
 
Fig. 13.   Effect of rapamycin on the state of activation of 70-kDa S6 protein kinase (p70S6k). In vitro (A), rapamycin inhibited the activation of p70S6k in cultured MPT cells. In vivo (B), activated p70S6k is undetectable in kidney tissue from sham-operated animals. The quantity of activated p70S6k is increased in the kidneys of vehicle-treated rats subjected to ARF. In contrast, activated p70S6k is markedly reduced in the kidneys of rapamycin-treated rats subjected to ARF. In both immunoblots, the amount of total kinase loaded in each lane was comparable. (Immnoblots shown are representative of 3 separate experiments.)

In kidney tissue from sham-operated rats subjected to ARF, the activated form of p70S6k was not detectable (Fig. 13). The activity of p70S6k was substantially increased in the kidneys of vehicle-treated rats subjected to ARF (Fig. 13). In rapamycin-treated rats subjected to ARF, activity of p70S6k was substantially reduced compared with the vehicle-treated ARF rats (Fig. 13).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Rapamycin is a macrolide-triene antibiotic with potent immunosuppressive activity (32). The mechanism of action of rapamycin is distinct from that of other immunosuppressive agents such as calcineurin antagonists (cyclosporin or FK-506) and antimetabolites (azathioprine or mycophenalate). Although calcineurin antagonists inhibit both IL-2 production and upregulation of the high-affinity IL-2 receptor in response to T cell activation, rapamycin has no effect on IL-2 production or on affinity of the IL-2 receptor. Instead, rapamycin acts downstream of the calcineurin antagonists by inhibiting signaling events triggered by interaction of IL-2 with its receptor. Rapamycin therefore inhibits IL-2-induced activation of T cells, thereby blocking the proliferation and clonal expansion of these cells (8, 11, 34, 35, 39). Rapamycin achieves these effects via inhibition of p70S6k, a kinase necessary for cell-cycle progression in T cells. Inhibition of p70S6k by rapamycin causes mid-to-late G1-phase arrest of IL-2-stimulated T cells (7, 8, 21, 34, 35, 39).

In preclinical studies, no substantial nephrotoxicity of rapamycin has been demonstrated, with the exception of exacerbation of cyclosporin-induced renal injury (2-4). Clinical studies in humans after transplantation have also not yet revealed any adverse renal effects (46).

In this report, we demonstrated that rapamycin inhibits the proliferation of cultured MPT cells, with an IC50 of ~1 ng/ml (Fig. 1). We also demonstrated that rapamycin induces apoptosis of confluent MPT cells by inhibiting the survival activity of several renal growth factors (Figs. 3-5). Importantly, although rapamycin induces apoptosis in MPT cells through activation of the default pathway, it has no effect on apoptosis induced by cytotoxic triggers such as ATP depletion (28) or cisplatin (30). These data are consistent with our previous report of the effects of rapamycin on primary cultures of peritoneal macrophages (19). We also showed directly that rapamycin inhibits activation of p70S6k in MPT cells (Fig. 13). Taken together, these data suggest that, in primary cultures of mouse renal tubular cells, signaling through p70S6k contributes to inhibition of the default pathway of apoptosis without affecting apoptosis induced by other pathways.

Because recovery from ischemic ARF depends, at least in part, on proliferation of the remaining viable tubular cells (15-18, 45, 49), we hypothesized that rapamycin, by inhibiting this regenerative process, would slow or prevent the recovery of kidneys subjected to acute ischemia. In addition, we hypothesized that rapamycin would slow or prevent recovery by also promoting apoptotic tubular cell loss through interference with the survival activity of renal growth factors (26, 27). We tested both of these novel hypotheses in a RAO model of ischemic ARF.

The effects of rapamycin on recovery of renal function after ischemic ARF revealed a similar pattern for three separate indices of renal injury, namely, GFR (Fig. 6A), FENa (Fig. 6B), and injury of the outer medulla (Figs. 7 and 8). For all three indexes, no statistical differences were observed on day 1 after RAO between vehicle-treated and rapamycin-treated rats (Figs. 6-8). However, although GFR, FENa, and renal morphology all improved toward normal in vehicle-treated rats by days 3 and 4 after RAO, recovery in rapamycin-treated rats at these same time points was either significantly reduced (GFR and FENa) (Fig. 6) or completely absent (renal morphology) (Figs. 7 and 8). Thus the deleterious effects of rapamycin on recovery of renal function after ischemic ARF were most evident during the recovery phase of ARF rather than during its induction. We therefore conclude that rapamycin severely impairs the repair of renal injury and the recovery of renal function after acute ischemic injury.

Renal tubular cells that survive an acute injury have been shown to leave G0, enter the cell cycle, and begin proliferation. The ability of tubular cells to proliferate is believed to be essential for repair of injured tubules, replacement of lost tubular cells, and return of renal function after ARF (15-18, 45, 49). To test our hypothesis that rapamycin inhibits this proliferative response, we quantitated PCNA staining in kidney sections from experimental and sham groups. Minimal PCNA staining was present in sham-operated rats treated with either vehicle or rapamycin. This finding is consistent with the fact that tubular cells of the normal kidney are largely quiescent. Although PCNA staining was present 1, 3, and 4 days after ARF in kidney sections of vehicle-treated rats, the number of PCNA-positive nuclei was markedly reduced at all three time points after ARF in rapamycin-treated rats (Figs. 9 and 10). These data support the hypothesis that an important mechanism by which rapamycin delays recovery from ARF is through inhibition of tubular cell proliferation.

Multiple lines of evidence suggest that renal tubular cells die by apoptosis as well as by necrosis after tubular cell injury and that apoptosis contributes substantially to the loss of renal function observed in ARF (26-28, 30). In light of our in vitro data showing that rapamycin interferes with the survival activity of several renal growth factors, we tested the hypothesis that an increase in apoptosis might also contribute to the delayed recovery of renal function found in rapamycin-treated rats subjected to ARF.

Identification of apoptotic cells in tissue sections is difficult for several reasons. First, on induction of apoptosis, cells rapidly shrink and fragment into even smaller apoptotic bodies that can be clearly identified only under high-power magnification. Second, phagocytic cells very rapidly clear apoptotic cells and bodies from tissues, so that while many cells may be dying by apoptosis, very few may be identifiable at any one point in time in a given tissue section. Finally, the terminal deoxynucleotide transferase dUTP nick-end labeling (TUNEL) assay, while commonly used to identify apoptotic cells, is not specific and can stain necrotic nuclei as well (10). Hence, in situations like ARF, in which both necrosis and apoptosis are present, TUNEL staining may exaggerate the apparent number of apoptotic cells. For these reasons, we chose to quantify apoptotic cells by examining kidney sections stained by H & E. Apoptotic cells and apoptotic bodies can be identified by the presence of condensed and/or fragmented nuclei (Fig. 11). The identification of these nuclear morphological features is specific for apoptosis. In our analysis, we excluded apoptotic cells or bodies that did not appear to be of tubular origin or that were present in the lumen of tubules.

As expected, very few apoptotic cells were seen in kidneys of sham-operated animals. After RAO, apoptosis of tubular cells was clearly present. Although the number of apoptotic tubular cells was quantitatively comparable in vehicle- and rapamycin-treated rats on day 1 after RAO, the number of apoptotic cells was markedly increased in rapamycin- vs. vehicle-treated rats by 3 and 4 days after RAO (Figs. 8 and 9). As for GFR, FENa, and renal morphology, the deleterious effects of rapamycin were limited to the recovery phase of ARF and did not seem to affect the initial severity of renal injury. This result is consistent with our in vitro data showing that rapamycin has no effect on apoptosis induced by cytotoxic stimuli and suggests that the observed effects of rapamycin on apoptosis after ARF may be attributable to inhibition of p70S6k and increased activation of the default pathway.

The activated form of p70S6k was undetectable in kidney tissue from sham-operated animals but was substantially increased after ischemia-reperfusion injury (Fig. 13). Treatment with rapamycin markedly diminished the amount of activated p70S6k in the renal tissues of rats subjected to ARF (Fig. 13). These data, in conjunction with the known extremely high degree of specificity of rapamycin as an inhibitor of p70S6k (9), make it very likely that the observed effects of rapamycin on proliferation, apoptosis, and recovery of renal function are all mediated by inhibition of p70S6k. Furthermore, our observation that p70S6k is activated in the kidneys of rats subjected to RAO is entirely novel and suggests an important role for this kinase in the process of recovery from ARF.

ARF, like many other catabolic states, including sepsis, burns, and severe trauma, is well known to be associated with weight loss (33). We show that rapamycin exacerbates the loss of weight associated with ARF while having no effect on the weight of sham-operated animals (Table 1). The muscle wasting seen in catabolic states is related, at least in part, to a disorder in insulin-like growth factor-I (IGF-I) metabolism within skeletal muscle (6, 13). Rapamycin has been shown to interfere with IGF-I-mediated effects in skeletal muscle. Because p70S6k is an important component of the IGF-I signaling pathway (6, 13), it is possible that exacerbation of weight loss by rapamycin in ARF may be the result of its effects on the IGF-1 axis in muscle.

In summary, rapamycin inhibits proliferation of renal tubular cells and induces apoptosis of these cells by inhibition of the survival activity of renal growth factors. Our data suggest that p70S6k activates downstream targets that play a key role in modulating the default pathway of apoptosis. Furthermore, rapamycin impairs the recovery of renal function after ischemia-reperfusion injury. This effect appears to be due, at least in part, to the combined effects of impaired regeneration and increased apoptosis of tubular cells. Finally, we provide novel evidence that implicates p70S6k as an important modulator of events necessary for recovery after ARF. Because ischemic ARF is a common complication after transplantation of kidney (12), heart (12), liver (5), and bone marrow (50), our findings are highly relevant to clinical transplantation.


    ACKNOWLEDGEMENTS

We are grateful to Dr. Yuhui Xu, Dana Farber Cancer Institute, Boston, who processed and stained kidney tissue for PCNA.


    FOOTNOTES

This work was supported by National Institutes of Health Grants DK-375105, DK-52898, DK-59793, and AR/AI-2732, a Clinical Scientist Award from the National Kidney Foundation, and a Career Enhancement Award from the American Society of Nephrology. The work was also supported by the US Navy (Office of Naval Research Contract N00014-94-C-0149, with the funds provided by the Naval Medical Research and Development Command). The opinions or assertions contained herein are those of the authors and are not to be construed as official or reflecting the views of the Navy Department or Naval Service at large.

Address for reprint requests and other correspondence: W. Lieberthal, Evans Biomedical Research Center, Renal Section, 5th Fl., Rm. 537, 650 Albany St., Boston, MA 02118.

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.

Received 10 July 2000; accepted in final form 15 May 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Alessi, DR, Kozlowski MT, Weng QP, Morrice N, and Avruch J. 3-Phosphoinositide-dependent protein kinase 1 (PDK1) phosphorylates and activates the p70 S6 kinase in vivo and in vitro. Curr Biol 8: 69-81, 1998[ISI][Medline].

2.   Andoh, TF, Burdmann EA, and Bennett WM. Nephrotoxicity of immunosuppressive drugs: experimental and clinical observations. Semin Nephrol 17: 34-45, 1997[ISI][Medline].

3.   Andoh, TF, Burdmann EA, Franceschini N, Houghton DC, and Bennett WM. Comparison of acute rapamycin nephrotoxicity with cyclosporine and FK506. Kidney Int 50: 815-822, 1996.

4.   Andoh, TF, Mattos DA, Meyer MM, Andoh T, and Barry JM. Chronic cyclosporine nephropathy in renal transplanation. Transplant Proc 28: 2100-2103, 1996[ISI][Medline].

5.   Bilbao, I, Charco R, Balsells J, Lazaro JL, Hidalgo E, Llopart L, Murio E, and Margarit C. Risk factors for acute renal failure requiring dialysis after liver transplantation. Clin Transplant 12: 123-129, 1998[ISI][Medline].

6.   Botfield, C, Ross RJ, and Hinds CJ. The role of IGFs in catabolism. Baillieres Clin Endocrinol Metab 11: 679-697, 1997[ISI][Medline].

7.   Chou, M, and Blenis J. The 70kDa S6 kinase: regulation of a kinase with multiple roles in mitogenic signalling. Curr Opin Cell Biol 7: 806-814, 1995[ISI][Medline].

8.   Chung, J, Kuo CJ, Crabtree GR, and Blenis J. Rapamycin-FKBP specifically blocks growth-dependent activation of and signaling by the 70 kd S6 protein kinases. Cell 69: 1227-1236, 1992[ISI][Medline].

9.   Davies, SP, Reddy H, Caivano M, and Cohen P. Specificity and mechanism of action of some commonly used protein kinase inhibitors. Biochem J 351: 95-105, 2000[ISI][Medline].

10.   Didenko, VV, and Hornsby PJ. Presence of double-strand breaks with single-base 3' overhangs in cells undergoing apoptosis but not necrosis. J Cell Biol 135: 1369-1376, 1996[Abstract].

11.   Dumont, FJ, and Su QS. Mechanisms of action of the immunopressant rapamycin. Life Sci 58: 373-395, 1996[ISI][Medline].

12.   Finn, W. Prevention of ischemic injury in renal transplantation. Kidney Int 37: 171-182, 1990[ISI][Medline].

13.   Frost, RA, and Lang CH. Differential effects of insulin-like growth factor 1 (IGF-1) and IGF-binding protein-1 on protein metabolism in human skeletal muscle cells. Endocrinology 140: 3962-3970, 1999[Abstract/Free Full Text].

14.   Grammer, TC, and Blenis J. The serine protease inhibitors, tosylphenylalanine chloromethyl ketone and tosyllysine chloromethyl ketone, potently inhibit pp70s6k activation. J Biol Chem 271: 23650-23652, 1996[Abstract/Free Full Text].

15.   Hammerman, MR. Potential role of growth factors in the prophylaxis and treatment of acute renal failure. Kidney Int 64: S19-S22, 1998.

16.   Hammerman, MR, and Miller SB. Therapeutic use of growth factors in renal failure. J Am Soc Nephrol 5: 1-11, 1994[Abstract].

17.   Harris, RC. Growth factors and cytokines in acute renal failure. Adv Ren Replace Ther 4: 43-53, 1997[Medline].

18.   Humes, HD. Acute renal failure: growth factors, cell therapy, gene therapy. Proc Assoc Am Phys 109: 547-557, 1997[ISI][Medline].

19.   Koh, JS, Lieberthal W, Heydrick S, and Levine JS. Lysophosphatidic acid is a major serum noncytokine survival factor for murine macrophages which acts via the phosphatidylinositol 3-kinase signaling pathway. J Clin Invest 102: 716-727, 1998[Abstract/Free Full Text].

20.   Kroshian, VM, Sheridan A, and Lieberthal W. Functional and cytoskeletal changes induced by sublethal injury in proximal tubular epithelial cells. Am J Physiol Renal Fluid Electrolyte Physiol 266: F21-F30, 1994[Abstract/Free Full Text].

21.   Lane, HA, Fernandez A, Lamb NJ, and Thomas G. p70s6k function is essential for G1 progression. Nature 363: 170-172, 1993[ISI][Medline].

22.   Levine, JS, Koh JS, Triaca V, and Lieberthal W. Lysophosphatidic acid: a novel growth and survival factor for renal proximal tubular cells. Am J Physiol Renal Physiol 273: F575-F585, 1997.

23.   Levinsky, N, and Lieberthal W. Clearance Techniques. New York: Plenum, 1986, p. 227-247.

24.   Lieberthal, W, Fuhro R, Andry C, and Valeri CR. Effects of hemoglobin-based oxygen-carrying solutions in anesthetized rats with acute ischemic renal failure. J Lab Clin Med 135: 73-81, 2000[ISI][Medline].

25.   Lieberthal, W, Fuhro R, Freedman JE, Toolan G, Loscalzo J, and Valeri CR. O-raffinose crosslinking markedly reduces the systemic and renal vasoconstrictor effects of unmodified human hemoglobin. J Pharm Exp Ther 288: 1278-1999, 1999[Abstract/Free Full Text].

26.   Lieberthal, W, Koh JS, and Levine JS. Necrosis and apoptosis in acute renal failure. Semin Nephrol 18: 505-518, 1998[ISI][Medline].

27.   Lieberthal, W, and Levine J. Mechanisms of apoptosis and its potential role in renal tubular epithelial cell injury. Am J Physiol Renal Fluid Electrolyte Physiol 271: F477-F488, 1996[Abstract/Free Full Text].

28.   Lieberthal, W, Menza SA, and Levine JS. Graded ATP depletion can induce apoptosis or necrosis of cultured mouse proximal tubular cells. Am J Physiol Renal Physiol 274: F315-F327, 1998[Abstract/Free Full Text].

29.   Lieberthal, W, Triaca V, Koh JS, Pagano PJ, and Levine JS. Role of superoxide in apoptosis induced by growth factor withdrawal. Am J Physiol Renal Physiol 274: F691-F702, 1998.

30.   Lieberthal, W, Triaca V, and Levine J. Mechanisms of death induced by cisplatin in proximal tubular epithelial cells: apoptosis vs. necrosis. Am J Physiol Renal Fluid Electrolyte Physiol 270: F700-F708, 1996[Abstract/Free Full Text].

31.   Liu, Y, Gorospe M, Kokkonen GC, Boluyt MO, Younes A, Mock YD, Wang X, Roth GS, and Holbrook NJ. Impairments in both p70 S6 kinase and extracellular signal-regulated kinase signaling pathways contribute to the decline in proliferative capacity of aged hepatocytes. Exp Cell Res 240: 40-48, 1998[ISI][Medline].

32.   Martel, RR, Klicius J, and Galet S. Inhibition of the immune response by rapamycin, a new antifungal antibiotic. Can J Physiol Pharmacol 55: 48-51, 1977[ISI][Medline].

33.   Mitch, WE. Mechanisms causing loss of muscle in acute uremia. Ren Fail 18: 389-394, 1996[ISI][Medline].

34.   Morice, WG, Brunn GJ, Wiederrecht G, Siekierka JJ, and Abraham RT. Rapamycin-induced inhibition of p34cdc2 kinase activation is associated with G1/S-phase growth arrest in T lymphocytes. J Biol Chem 268: 3734-3738, 1993[Abstract/Free Full Text].

35.   Morice, WG, Wiederrecht G, Brunn GJ, Siekierka JJ, and Abraham RT. Rapamycin inhibition of interleukin-2-dependent p33cdk2 and p34cdc2 kinase activation in T lymphocytes. J Biol Chem 268: 22737-22745, 1993[Abstract/Free Full Text].

36.   Mossman, T. Rapid colorimetric assay for cell growth and survival: application to proliferation and cytotoxicity assays. J Immunol Methods 65: 55-63, 1983[ISI][Medline].

37.   Ormerod, M, Sun XM, Snowden R, Davies R, Fearnhead H, and Cohen G. Increased membrane permeability of apoptotic thymocytes: a flow cytometric study. Cytometry 14: 595-602, 1993[ISI][Medline].

38.   Powers, JF, Tischler AS, and Cherington V. Discordant effects of rapamycin on proliferation and p70S6 kinase phosphorylation in normal and neoplastic rat chromaffin cells. Neurosci Lett 259: 137-140, 1999[ISI][Medline].

39.   Price, DJ, Grove JR, Avruch J, and Bierer BE. Rapamycin-induced inhibition of the 70-kilodalton S6 protein kinase. Science 257: 973-977, 1992[ISI][Medline].

40.   Raff, M. Social controls on cell survival and cell death. Nature 356: 397-400, 1992[ISI][Medline].

41.   Sheridan, AM, Schwartz JH, Kroshian VM, Tercyak AM, Laraia J, Masino S, and Lieberthal W. Renal mouse proximal tubular cells are more susceptible than MDCK cells to chemical anoxia. Am J Physiol Renal Fluid Electrolyte Physiol 265: F323-F350, 1993[Abstract/Free Full Text].

42.   Shi, Y, Frankel A, Radvanyi LG, Penn LZ, Miller RG, and Mills GB. Rapamycin enhances apoptosis and increases sensitivity to cisplatin in vitro. Cancer Res 55: 1982-1988, 1995[Abstract].

43.   Sun, XM, Snowden R, Skilleter D, Dinsdale D, Ormerod M, and Cohen G. A flow-cytometric method for the separation and quantitation of normal and apoptotic thymocytes. Anal Biochem 204: 351-356, 1992[ISI][Medline].

44.   Swat, W, Ignatowicz L, and Kisielow P. Detection of immature CD4+8+ thymocytes by flow cytometry. J Immunol Methods 137: 79-87, 1991[ISI][Medline].

45.   Toback, FG. Regeneration after acute tubular necrosis. Kidney Int 41: 226-246, 1992[ISI][Medline].

46.   Vasquez, EM. Sirolimus: a new agent for prevention of renal allograft rejection. Am J Health Syst Pharm 57: 437-448, 2000[ISI][Medline].

47.   Weil, M, Jacobson M, Davies HC, Gardner R, Raff K, and Raff M. Constitutive expression of the machinery for programmed cell death. J Cell Biol 133: 1053-1059, 1996[Abstract].

48.   Weng, QP, Kozlowski M, Belham C, Zhang A, Comb MJ, and Avruch J. Regulation of the p70 S6 kinase by phosphorylation in vivo. Analysis using site-specific anti-phosphopeptide antibodies. J Biol Chem 273: 16621-16629, 1998[Abstract/Free Full Text].

49.   Witzgall, R, Brown D, Schwartz C, and Bonventre JV. Localization of proliferating cell nuclear antigen, vimentin, c-fos and clusterin in postischemic kidney. Evidence for a heterogeneous genetic response among nephron segments and a large pool of mitotically active and dedifferentiated cells. J Clin Invest 93: 2175-2188, 1994[ISI][Medline].

50.   Zager, R. Acute renal failure in the setting of bone marrow transplantation. Kidney Int 14: 341-344, 1994.

51.   Zhu, J, Wu J, Frizell E, Liu SL, Bashey R, Rubin R, Norton P, and Zern MA. Rapamycin inhibits hepatic stellate cell proliferation in vitro and limits fibrogenesis in an in vivo model of liver fibrosis. Gastroenterology 117: 1198-1204, 1999[ISI][Medline].


Am J Physiol Renal Fluid Electrolyte Physiol 281(4):F693-F706