Unité de Pharmacologie Cellulaire et Moléculaire, Université Catholique de Louvain, B-1200 Brussels, Belgium; and * Service d'Histologie et de Cytologie Expérimentale, Université de Mons-Hainaut, B-7000 Mons, Belgium
Received December 24, 1999; accepted March 27, 2000
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ABSTRACT |
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Key Words: gentamicin; aminoglycosides; apoptosis; renal cell lines; fibroblasts.
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INTRODUCTION |
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In order to gain a better insight into gentamicin-induced apoptosis and to begin unraveling its molecular mechanisms, we have now examined the ability of gentamicin to cause apoptosis in models of cultured cells. The usefulness of the LLC-PK1 [porcine kidney] cells (derived from proximal tubules) to study the cellular fate and toxicity of gentamicin and other aminoglycosides has been highlighted in numerous studies. (Ford et al., 1994; Hori et al., 1984
; Kohlhepp et al., 1994
; Oshima et al., 1989
; Sandoval et al., 1998
; Schwertz et al., 1986
; Steinmassl et al., 1995
). LLC-PK1 cells have also been successfully used to study apoptosis and necrosis induced by cyclosporin A (Healy et al., 1998
). Aminoglycosides have also been shown to cause specific alterations in other renal cell lines such as the MDCK (canine kidney) cells (Stiemer, 1989
), and non-renal cells such as embryonic fibroblasts (Aubert-Tulkens et al., 1979
). In the present study, LLC-PK1 cells have therefore been used in comparison with MDCK cells and fibroblasts to explore the occurrence and specificity of the apoptotic response to gentamicin exposure. We also examined the relation between apoptosis and the capacity of gentamicin to accumulate in cells, on the one hand, and to cause a lysosomal phospholipidosis on the other hand. The latter alteration is one of the earliest and most conspicuous to be detected in proximal tubular cells of animals and man treated with low, clinically relevant doses of aminoglycosides (De Broe et al., 1984
; Tulkens, 1986
).
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MATERIALS AND METHODS |
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Electron microscopy.
Cell pellets were rinsed rapidly with PBS and fixed for 12 h at 4°C with 2% (vol/vol) glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4. Fixation was followed by 3 5-min rinses with 0.1 M sodium cacodylate buffer pH 7.4. Cells were post-fixed with a solution containing 1% OsO4 and 2% K4Fe(CN)6 solution (both wt/vol), stained with 1% uranyl acetate, and pelleted in 2% agar. Pellets were dehydrated in graded ethanol solutions and embedded in Spurr resin. Ultrathin (60 nm nominal) sections were cut on a Reichert Ultracut microtome, collected on rhodanium 400-mesh grids, post-stained with uranyl acetate followed by lead citrate, and rinsed in water. Examination was carried out in a Philips CM-12 electron microscope at 80 kV.
Detection of DNA fragmentation by agarose gel electrophoresis.
Fragmented DNA was isolated from cells as previously described (Wyllie, 1980). In brief, pelleted cells were lysed overnight at 50°C in 0.5% Triton X-1001 mM, Tris-HCl25 mM, and EDTA adjusted to pH 8. The lysates were centrifuged at 10,000 x g for 30 min and the supernatants containing fragmented DNA incubated with proteinase K (20 µg/ml) at 50°C overnight. DNA was extracted with phenol-chloroform-isoamyl alcohol (50:49:1), followed by precipitation with 0.1 vol of 3 M sodium acetate and 2.5 vol of 100% ethanol. The concentration of nucleic acid in each sample was determined by UV absorbance at 260 nm. The same amount of nucleic acid from each sample was treated with RNase (20 µg/ml) at 37°C for 3 h. Two µl of buffer (0.1 M Tris base0.1 M, boric acid0.02 M, and EDTA, pH 8.3 [TBE]) containing 0.02% bromophenol blue and 40% glycerol were added to 10 µg of DNA samples and loaded on 1.8% agarose gels. Samples were then resolved by electrophoresis (150 V for 2 h) using TBE as the running buffer. After electrophoresis, DNA was stained with ethidium bromide (0.5 µg/ml in distilled water) and visualized by fluorescence in UV light.
In situ detection of DNA fragmentation and quantitative analysis of apoptosis.
A modification of the terminal deoxynucleotidyl transferase (TdT)-mediated dUTP-biotin nick-end labeling (TUNEL) technique of Gavrieli et al. (1992) was used to detect DNA strand breaks in situ. Pelleted cells were rapidly rinsed with PBS containing 1% bovine serum albumin (BSA) and fixed in 4% paraformaldehyde for 30 min. After centrifugation and rinsing with PBS, cells were transferred to 96-well plates and treated with 0.3% H2O2 in methanol for 30 min to quench endogenous peroxidase activity. Apoptotic cells were detected by TdT-mediated extension of 3`OH ends of fragmented DNA, using fluorescein-labeled dUTP as a precursor, according to the instructions from the supplier. DNA-bound fluorescein was detected by reaction with anti-fluorescein antibody conjugated to peroxidase. Peroxidase activity in immunocomplexes was visualized by reaction of diaminobenzidine in the presence of H2O2. Cells were resuspended in PBS, spread on lysine-coated slides and allowed to dry out. Cells were then counterstained with methyl green, and the preparations mounted using permanent medium. Negative controls were performed by substituting distilled water for TdT enzyme in the preparation of the working solution and exhibited no immunostaining. Enumeration of apoptotic nuclei was made on slides picked at random from each experiment, using a Zeiss light microscope with a 40x objective and a 10x eyepiece. For calibration purposes, a square grid was mounted in one eyepiece, determining a square field of 0.04 mm2. We counted all nuclei exhibiting a frank brown labeling. These nuclei most often displayed typical alterations such as pyknosis, crescent-like condensation of chromatin, or segregation into apoptotic bodies. Clusters of apoptotic bodies were given a single count. The incidence of apoptotic nuclei was given as the percentage relative to total nuclei (apoptotic index).
Cell transfection and selection.
An RSV vector containing the human bcl-2 gene and a hygromycin resistance gene (3` phosphotransferase) was introduced into LLC-PK1 and MDCK-cells by means of LipofectAmineTM (cells transfected with the same vector, carrying the resistance gene to hygromycin, but no bcl-2 gene were used as controls). Briefly, 60-mm dishes were seeded with 106 cells in 4 ml of complete culture medium. After 24 h, cells were exposed to a LipofectAmineTM reagent/DNA mixture (2.5 µg of DNA per dish) and incubated for 4 h, after which the medium was replaced with fresh medium for 2 days. Cells were then incubated and grown for 4 weeks in selection media (containing hygromycin sulfate at concentrations spanning from 100 to 1500 µg/ml) to obtain stable transfectants. Cloning was performed from cell populations surviving in 400 µg/ml hygromycin, and resistant populations were expanded in this selection medium. Prior to experiments, hygromycin-resistant cells were cultured in the absence of hygromycin for 1 or 2 passages. In control experiments, no cell survival was observed when non-transfected LLC-PK1 and MDCK-cells were cultured in medium containing 400 µg/ml hygromycin.
Western blotting.
Pelleted cells were rinsed with PBS and cell lysis was obtained by a 1-h incubation on ice in 30 mM TrisHCl buffer (pH 7.4) containing 150 mM NaCl, 1 mM EDTA, 0.1% Triton X-100, and protease inhibitors (1 mM phenylmethylsulfonyl fluoride (PMSF), 2 µg/ml pepstatin, and 2 mg/ml leupeptin). After sonication and centrifugation, 100-µg protein aliquots of lysates were separated on 12% SDSPAGE (80 V at 4°C, overnight) using 2.5 mM TrisHCl, 19.2 mM glycine, and 1% SDS, pH 8.3 as a running buffer. Proteins were transferred onto a nitrocellulose membrane and blocked for 1 h with 3% BSA in 10 mM TrisHCl, 150 mM NaCl, and 0.05% TWEEN®-20, pH 8 (TBST). The membranes were then incubated for 1 h with monoclonal mouse anti-Bcl-2 antiserum and for 1 h with alkaline phosphatase-conjugated goat anti-mouse IgG. Color was developed by incubation with nitro blue tetrazolium/5-bromo-4-chloro-3-indolyl phosphate (NBT/BCIP) according to the manufacturer's instructions (Promega Corp., Leiden, The Netherlands).
Determination of the activity of cysteine aspartatespecific proteases (caspases) activity.
Pelleted cells were rinsed with PBS and suspended in lysis buffer (100 mM N-2-hydroxyethylpiperazine-N`-2-ethanesulfonic acid, N-2-hydroxyethylpiperazine-N`-2-ethanesulfonic acid (HEPES), pH 7.5, containing 10% sucrose, 0.1% 3-[(cholamidopropyl) dimethylammonio]-1-propane-sulfonate (CHAPS), 1 mM EDTA, 1 mM PMSF, 10 µg/ml pepstatin A and 10 µg/ml leupeptin) and left on ice for 30 min. Following sonication, lysates were centrifuged at 10,000 x g for 10 min (4°C) and supernatants were stored at 20°C. Activity was then determined in the supernatants by fluorimetric assay using the substrate DEVDAMC, which releases the fluorochrome 7-amino-4-methyl coumarin (AMC). This substrate was originally described as being specifically cleaved by caspase 3; however, recent data and manufacturer's information shows that it can be also cleaved by caspases 6, 7, and 8). The appropriate amount of protein cell extracts (50 µg) was incubated in 100 mM HEPES, pH 7.5, 10% sucrose, 0.1% CHAPS, 1 mM EDTA, 1 mM PMSF, 10 µg/ml leupeptin at 30°C for 30 min with 20 mM acetyl-Asp-Glu-Val-Asp-7-amino-4-methylcoumarin (DEVD-AMC) in a total volume of 0.5 ml. At the end of the incubation, the released fluorochrome was measured in a Perkin-Elmer LS-30 fluorescence spectrophotometer (Perkin-Elmer, Beaconsfield, UK) with an excitation wavelength of 380 nm and an emission wavelength of 460 nm. Enzyme activity was expressed as arbitrary units relative to protein concentration.
Biochemical studies.
Cell phospholipids were determined by assaying lipid phosphorus as described previously (Aubert-Tulkens et al., 1979). Proteins were measured according to Lowry's method or by Bradford's assay (using the Bio-Rad® dye reagent according to manufacturer's instructions).
Assay of gentamicin cellular uptake.
Washed cell pellets were resuspended in 0.01% Triton X-100 and subjected to brief sonication to achieve complete dispersion. Gentamicin was then assayed by a disc-plate microbiological assay, which had been fully validated for the quantitative assay of aminoglycosides in lysates of cultured fibroblasts (Tulkens and Trouet, 1978). In the present study, the limit of detection was 0.8 µg/ml and the range of linearity of the zone diameter vs. the log of the antibiotic concentration extended from 0.8 to 10 µg/ml. Samples were prepared and appropriately diluted to systematically contain gentamicin concentrations within this range.
Sources of products.
Dulbecco's modification of Eagle's minimum essential medium (DMEM), trypsin (1:250), LipofectAmineTM reagent, hygromycin sulfate, and foetal bovine serum were purchased from Life Technologies, Rockville, MD. Gentamicin sulfate (mixture of C1, C1a, and C2/C2a components in a weight ratio of 29.2:23.3:47.6) was kindly provided as powder for microbiological standard by Schering-Plough, Inc., Kenilworth, NJ. DEVD-AMC and N-benzyloxycarbonyl-Val-Ala-Asp-fluoromethyl-ketone (Z-VAD.fmk) were purchased from Calbiochem-Novabiochem, San Diego, CA. In situ cell death detection kit®, mouse anti-Bcl-2, and alkaline phosphatase-conjugated goat anti-mouse immunoglobulin came from Boehringer-Mannhein, Mannheim, Germany (presently Roche Diagnostics, Basle, Switzerland). The plasmid containing the human bcl-2 cDNA was a generous gift from Professor D.L. Vaux (Walter and Eliza Hall Institute of Medical Research, Melbourne, Victoria, Australia). Unless stated otherwise, all other products were purchased from Sigma Chemical Co., St-Louis, MO.
Statistical analyses.
Two-group comparisons were performed by Student's t-test. When more than 2 groups were involved, a first statistical analysis was done by ANOVA, followed by Tukey (all groups) or Dunnett (test vs. control) post hoc tests.
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RESULTS |
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Effect of Protein Synthesis Inhibition
To test for the role of protein synthesis in gentamicin-induced apoptosis, cells were incubated for 3 days in the presence of cycloheximide (1 µM), gentamicin (2 mM), or their combination. As shown in Table 1, cycloheximide alone caused no significant increase of apoptosis in rat fibroblasts or MDCK cells. Cycloheximide also completely prevented the apoptosis induced by gentamicin in these 2 types of cells, bringing the apoptotic index to a level very close to that seen with cycloheximide alone (fibroblasts) or even somewhat lower (MDCK cells). In sharp contrast, LLC-PK1 cells displayed a marked sensitivity to cycloheximide, which, by itself, caused a larger increase in the apoptotic index than that observed with gentamicin alone. The combination of cycloheximide and gentamicin was more than additive in these cells.
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DISCUSSION |
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The data presented in this paper suggest that gentamicin acts at a level where Bcl-2 can still block or delay this cascade, which would rank the drug as an early initiator of apoptosis. This first conclusion is reinforced by our observation that gentamicin-induced apoptosis requires active protein synthesis in at least 2 of the 3 types of cells studied (fibroblasts and MDCK cells). Although the elements involved in the execution phase indeed preexist within the cell (Weil et al., 1996) and only need to be triggered by appropriate initiation, effective gene transcription, and/or translation of the corresponding mRNA are necessary to produce the factors that allow the process to move from the early steps of the commitment phase (initiation) to the onset of the execution phase (Umanskii, 1996
). Inhibition of protein synthesis has also been shown to suppress apoptosis induced in proximal tubular cells by nephrotoxins other than gentamicin, such as cisplatin (Takeda et al., 1998
), and in streptomycin-induced apoptosis in vestibular hair cells (Nakagawa et al., 1998
).
The proteins that must be produced in rat fibroblasts, MDCK cells, or proximal tubule cells for apoptosis to occur remain to be determined. However, the fact that cycloheximide acts as a protectant against gentamicin-induced apoptosis in rat fibroblasts and MDCK cells virtually excludes the possibility that the cytotoxicity of gentamicin could be related to an inhibition of protein synthesis, as suggested from studies with the atypical aminoglycoside hygromycin, G-418 (Buchanan et al., 1987); G-418, however, also causes apoptosis by itself in a Bcl-2-preventable manner (Chen et al., 1995
). We have no simple explanation to offer for the unanticipated finding that LLC-PK1 cells undergo apoptosis in the presence of cycloheximide alone, especially since these cells have been claimed to show marked protein synthesis inhibition when exposed to gentamicin (Okuda et al., 1992
). Possibly, LLC-PK1 cells constitutively express the elements of apoptosis machinery and cycloheximide as well as gentamicin could act by abolishing the production of short-lived proteins acting as apoptosis antagonists.
Where and how gentamicin initiates apoptosis was not established by our experiments. Yet, the fact that apoptosis develops slowly over time and in strict correlation with cellular drug levels strongly suggests that it is the cell-associated drug which is acting as the initiator. Aminoglycosides have been previously shown to slowly enter both fibroblasts and LLC-PK1 cells and to accumulate in their lysosomes (Ford et al., 1994; Tulkens and Trouet, 1978
). Because the clearance of gentamicin by all 3 cell types was essentially similar in our conditions, we suspect that the drug enters these cells by a common mechanism. For fibroblasts, this mechanism has been shown to be fluid-phase pinocytosis, i.e., a constitutive process of uptake of non-diffusible solutes occurring in all eucaryotic cells (Mukherjee et al., 1997
). Gentamicin uptake in fibroblasts is indeed non-saturable (viz. the present experiments and Tulkens and Trouet, 1978), and its rate of influx is very similar to that of an authenticated marker of fluid-phase endocytosis in the same cells (Cupers et al., 1994
). Cell fractionation and morphometric studies have shown that gentamicin pinocytosed by fibroblasts and accumulated in lysosomes reaches, therein, concentrations at least 30-fold larger than in the extracellular medium (Aubert-Tulkens et al., 1979
).
Less is known about the concentration of gentamicin in the lysosomes of LLC-PK1 cells, where part of the drug may traffic to the Golgi vesicles (Sandoval et al., 1998), but animal studies have revealed a huge accumulation of gentamicin in lysosomes of proximal tubular cells (Giurgea-Marion et al., 1986
), suggesting that this property is probably very general. Gentamicin stored in lysosomes, as well as in Golgi vesicles, is expected to become fully protonated and therefore strongly polycationic, due to the acidic pH (~5 to ~6) prevailing in these compartments (Anderson and Pathak, 1985
; Ohkuma and Poole, 1978
). The pKa's of the ionizable groups in aminoglycosides span from ~5.5 to 9.5 (Claes et al., 1977
). It is probably in this context that the role of intracellular gentamicin as an initiator of apoptosis must be examined.
One of the most early and conspicious alterations related to the accumulation of gentamicin in lysosomes was the development of a phospholipidosis which is directly related to the capacity of the intralysosomal polycationic drug to bind to phospholipid bilayers (see Laurent et al, 1990 for review, Mingeot-Leclercq et al., 1988 and Piret et al., 1992 for a discussion of the molecular mechanisms). This phospholipidosis was clearly observed here for fibroblasts and MDCK cells but developed only to a minimal extent in LLC-PK1 cells in our conditions, suggesting that it is probably not the key initiator of apoptosis. Lysosomal phospholipidosis may, however, develop also in LLC-PK1 cells if incubation is prolonged (Schwertz et al., 1986). This set of data therefore suggests that it is rather the drug accumulation per se which is most critical. It may be envisioned that the large concentrations of gentamicin in lysosomes (30 mM or more [see above]) will eventually cause a destabilization of their membrane resulting in their in situ disruption, which, by itself, is likely to trigger apoptosis. We showed indeed that millimolar concentrations of gentamicin permeabilizes liposomes at pH 5.5 (Van Bambeke et al., 1993
). We know of at least one clear example where disruption of lysosomes in situ (by photooxidation) causes apoptosis (Brunk et al., 1997
), probably by the release of the lysosomal proteases and endonucleases in the cytosol (Ohsawa et al., 1998
). We also know that the relocation of the lysosomal proteases, cathepsins B and D in the cytosol, triggers apoptosis (Ohsawa et al., 1998
), possibly by direct activation of caspases (Vancompernolle et al., 1998
). In this context, it may even be speculated that phospholipidosis, which entails a partial binding of gentamicin to the intralysosomally accumulated phospholipids, may actually be protective, since it will reduce the amount of free drug available for binding to the lysosomal membrane. This could be the basis for the higher susceptibility of LLC-PK1 cells compared to the other 2 cell types studied here.
At first glance, the hypothesis of gentamicin-induced release of lysosomal proteases and of a direct activation of caspases may seem to fail to account for the protective effect, which acts upstream of the caspases involved in the execution phase. Yet, we cannot exclude that lysosomal disruption activates caspase 8, which is involved in the initiation phase (the substrate used to assay for caspase activity is cleaved by this enzyme as well as by caspases 3, 6, and 7, and the inhibitor Z-VAD.fmk is a non-specific inhibitor of most caspases). We may also envisage that lysosomal destabilization triggers mitochondrial events such as the so-called "permeability transition" (Lemasters et al., 1998) and/or the release of the apoptosis-inducing-factor (AIF) (Susin et al., 1999
), which will then cause apoptosis in a Bcl-2-preventable or -retardable manner. Other alternative explanations than a lysosomal origin in the triggering of gentamicin-induced apoptosis must, however, also be envisioned. Gentamicin, indeed, has been shown to modulate membrane enzyme activities, to cause changes in membrane fluidity and increases in cytosolic Ca++ concentration, and to directly alter the pericellular membrane and mitochondrial membranes (Holohan et al., 1988
; Jutila et al., 1998
; Okuda et al., 1992
; Van Bambeke et al. 1993
). All these changes are known to trigger apoptosis (Lang and Liu, 1997
; Umankskii, 1996).
The data presented here provides, however, no direct support for any of these potential mechanisms although they do not rule them out either. Conversely, the data plead against a potential role for gentamicin binding to megalin or other cell surface constituents. Binding to megalin and acidic phospholipids undoubtedly takes place in proximal tubular cells in vivo and is responsible for preferential uptake of aminoglycosides by these cells (Moestrup et al., 1995). Yet, the rates of gentamicin influx observed here in LLC-PK1 and in MDCK cells, which are slightly lower than those of fibroblasts, preclude a major role of membrane binding in gentamicin uptake in these cells, under our conditions. In addition, there is no evidence of the presence of megalin in fibroblasts, and we show here that these cells are only slightly less sensitive to gentamicin when compared to LLC-PK1, as far as apoptosis is concerned.
The toxicological implications of our findings need to be critically assessed. We see that a ~20% apoptotic index is reached in LLC-PK1 cells when these contain 20 µg of gentamicin per mg of protein. In comparison, animals treated for 10 days with a clinically relevant dose of gentamicin (10 mg/kg), and which demonstrate an extensive apoptotic process in their proximal tubule epithelium, contain ~10 µg of drug per g of cortex (El Mouedden et al., 2000; see the discussion in this paper for estimates of the equivalents of human and animal doses of aminoglycosides). Because the drug that accumulated in kidney in vivo was found almost exclusively in proximal tubules (Kuhar et al., 1979
), where it localizes mainly in lysosomes (Giurgea-Marion et al., 1986
), these cortical levels will translate into local drug concentrations of ~15 to 50 mM (see Laurent et al., 1983 for estimates of the fractional volume of proximal tubules and Wilmotte et al., 1983 for measurements of the aggregated volume of lysosomes in the kidney cortex of rats treated for 10 days with gentamicin at 10 mg/kg). Thus, the lysosomal concentrations of gentamicin at the level of both the target cells (proximal tubules) and in the cultured cells are of the same order of magnitude. This clearly supports the validity of the cultured cells model to study the in vivo situation. The fact that gentamicin-induced apoptosis is observed in the 2 renal cell lines to a similar extent, i.e., disregarding their cytological origin, as well as in fibroblasts, coupled with the observation that apoptosis has been recognized as an important determinant in aminoglycoside-induced toxicity towards hair cells in the cochlea (Lang and Liu, 1997
) also indicates that we are probably dealing with an intrinsic toxicological property of aminoglycosides of potentially broad significance.
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ACKNOWLEDGMENTS |
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NOTES |
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