Volatile Anesthetics Induce Caspase-dependent, Mitochondria-mediated Apoptosis in Human T Lymphocytes In Vitro 

The following anesthetics were used: sevoflurane, isoflurane (Abbott, Wiesbaden, Germany) and desflurane (Baxter, Unterschleißheim, Germany). All other reagents were purchased from Sigma (Deisenhofen, Germany) unless specified otherwise.

Jurkat T cells (ACC 282; DSMZ, Braunschweig, Germany) and primary human CD3⁺T lymphocytes were isolated and grown in RPMI 1640 medium supplemented with 10% fetal calf serum, 1% glutamine, and 50 mg/ml penicillin and streptomycin (all from Gibco-BRL, Karlsruhe, Germany) in a humidified atmosphere containing 5% carbon dioxide at 37°C as previously described.5Wild-type (clone A3), “Fas-associating protein with death domain”–, and caspase-8–deficient Jurkat T cells were kindly provided by John Blenis, Ph.D. (Professor of Cell Biology, Department of Cell Biology, Harvard Medical School, Boston, Massachusetts).25 

Jurkat T cells or primary CD3⁺T lymphocytes were exposed to either air or a volatile anesthetic–air mixture using a 12-l airtight glass chamber as previously described.5The chamber atmosphere was kept continuously saturated with water at 37°C. Gas was prepared using a gas mixing unit by directing a 95% air–5% carbon dioxide mixture at 6 l/min through calibrated vaporizers (Dräger, Lübeck, Germany) that were placed at the entrance of the chamber. In preliminary experiments, we observed that the different culture conditions used in our experiments did not influence apoptosis in T cells during a period of 48 h, i.e. , cell viability was similar in a standard incubator and in the sealed chamber flushed with a 95% air–5% carbon dioxide mixture. Volatile anesthetic concentrations were monitored at the chamber exit port using a halogen monitor (PM 8050; Dräger). Concentrations of the inhalation anesthetics dissolved in the cell culture medium were measured by a fully automated solid-phase microextraction procedure followed by gas chromatography and mass spectrometry (CTC Combi PAL autosampler, Agilent model 6890 Series plus Gas Chromatograph with Agilent 5973 N Mass Selective Detector; Chromtech, Idstein, Germany). Using an external standard method, calibration curves were achieved for sevoflurane, desflurane, and isoflurane.5 

For each exposure condition, a control sample was obtained simultaneously from T cells cultured in a standard 95% air–5% carbon dioxide incubator without anesthetic exposure. At the end of each experiment, both supernatants and the cells were frozen immediately in liquid nitrogen and stored at −80°C until protein extraction.

Cells were harvested, washed in phosphate-based saline solution, and resuspended in extraction buffer containing 10 mm mannitol, 70 mm sucrose, 20 mm HEPES, 1 mm EDTA, 2 mm dithiothreitol, 100 μm phenyl-methyl-sulfonyl-fluoride, 10 μg/ml leupeptin, 400 ng/ml pepstatin, 10 μg/ml aprotinin, and 5 μg/ml cytochalasin B. The cells were homogenized for 10 s with a Dounce homogenizer and centrifuged at 2,000g  for 5 min to eliminate debris and unbroken cells. The supernatants were centrifuged at 20,000g  for 15 min. The resulting crude mitochondrial pellet was resuspended in lysis buffer containing 20 mm Tris HCl, 2 mm EGTA, 2 mm EDTA, and 6 mm β-mercaptoethanol plus 1% sodium-dodecyl-sulfate. Further centrifugation of the supernatant at 100,000g  for 15 min yielded the cytosolic fraction. The fractions were aliquoted and kept at −80°C.

The amount of cytochrome c released from mitochondria into the cytosol was measured in the cytosolic fraction by Western blotting (a 1:1,000 dilution of cytochrome c; Alexis Corp., Gruenberg, Germany; cat. No. ALX-804-122) and for enzyme-linked immunosorbent assay (Active Motif, Rixensart, Belgium; cat. No. 48006). A 1:500 dilution of a cyclooxygenase-1 antibody (Alexis Corp.; cat. No. ALX-210-710) was used for protein normalization.

Total cell extracts of Jurkat T lymphocytes (30 μg) were boiled in Laemmli sample buffer and subjected to 15% sodium-dodecyl-sulfate-polyacrylamide gel electrophoresis. Before transfer, gels were equilibrated for 15 min in cathode buffer (25 mm Tris, 40 mm glycin, 10% methanol). Using a semidry blotting apparatus (Bio-Rad Laboratories, München, Germany), proteins were transferred at 0.8 mA/cm²for 1 h onto Immobilon P membranes (Millipore Corp., Eschborn, Germany), preequilibrated in methanol (15 s), ddH²O (2 min each side), and anode buffer II (25 mm Tris–10% methanol). Equal loading and transfer were confirmed by Bradford measurement and by amido black staining of the membranes at the end of the procedure. Nonspecific binding sites were blocked by immersing the membrane into blocking solution (10 mm Tris-HCl, pH 8.0, 150 mm NaCl, 0.1% Tween-20 [vol/vol] containing 5% milk powder; Fluka, Buchs, Switzerland) for 1 h at room temperature. Membranes were washed in blocking solution and incubated in a 1:2,000 dilution of anti–p32-caspase-3 antibody (cat. No. 9662) and a 1:1,000 dilution of anti–p17-caspase-3 antibody (cat. No. 9661; all from Cell Signaling Technology Inc., Beverly, MA) in blocking solution plus 5% bovine serum albumin overnight at 4°C, followed by extensive washing with blocking solution. Bound antibody was decorated with goat–anti-rabbit/horseradish peroxidase conjugate (Amersham Pharmacia, Freiburg, Germany), diluted 1:2,000 in blocking solution for 30 min at room temperature. After washing four times (5 min each), the immunocomplexes were detected using ECL Western blotting reagents (Amersham Pharmacia) according to the manufacturer’s instructions. Exposure to Kodak XAR-5 films (Stuttgart, Germany) was performed for 15 s to 1 min.

Total protein cell extracts (10 μl; 50 μg) were mixed with 90 μl assay buffer (100 mm HEPES, pH 7.5, 2 mm dithiothreitol, 2 mm phenyl-methyl-sulfonyl-fluoride). The respective fluorogenic substrate for caspase-3 and Ac-DEVD-AMC (1 μl; 60 μm; Alexis Corp.) was added, and the fluorescence was measured at 30°C for 30 min in a Microplate Spectra Max Gemini XS reader (Molecular Devices, Sunnyvale, CA) at 380/460 nm.

After treatment with anesthetics, suspension CD3⁺primary T cells were washed twice with phosphate-buffered saline and incubated with 3 μg/ml green fluorescent protein (GFP)-annexin V, 2.5 μg/ml propidium iodide, and 2 μg/ml Hoechst 33342 (Molecular Probes, Leiden, The Netherlands) in annexin V binding buffer (10 mm HEPES–NaOH, pH 7.4, 140 mm NaCl, 2.5 mm CaCl²) for 15 min. The cells were then viewed under an Axiovert inverse fluorescence microscope (Zeiss, Jena, Germany), and pictures were taken with a Contax 167 MT camera (YASHICA Kyocera GmbH, Hamburg, Germany) at magnifications of 400× and 1,000×. Nuclear staining by Hoechst stain was revealed using an ultraviolet filter (excitation, 365 nm; emission, 480 nm).

After treatment with anesthetics, Jurkat T cells were washed in phosphate-buffered saline and resuspended in 50 μl binding buffer (10 mm HEPES, 140 mm NaCl, 2.5 mm CaCl², pH 7.9) containing 12.27 μg/ml GFP-annexin V and 0.5 μg/ml propidium iodide. Samples were incubated in the dark at room temperature for 15 min. Subsequently, 450 μl binding buffer was added, and the percentage of early apoptotic lymphocytes was measured using a flow cytometer (FACS®-CaliburFCM; Becton Dickinson, Heidelberg, Germany). Lymphocytes were gated using forward scatter and side scatter, and fluorescence intensity was measured in 2 × 10⁵lymphocytes. The fluorescence intensity of GFP-annexin V was measured at the fluorescence 1 channel, and the fluorescence intensity of propidium iodide was measured at the fluorescence 3 channel.

The mitochondrial membrane potential was analyzed using the fluorochrome stain DiOC⁶(3) (Molecular Probes). In brief, after sevoflurane exposure, 10⁵cells were stained in a solution with 40 nm DiOC⁶(3) for 30 min. Staining was quantified by fluorescence 1 and scatter characteristics using a FACS®-Calibur flow cytometer (Becton Dickinson). The fluorescence was measured by flow cytometry.

Differences in measured variables between the experimental conditions were assessed using one-way analysis of variance on ranks followed by a nonparametric Student-Newman-Keuls test for multiple comparisons. Results were considered statistically significant if P  was less than 0.05. The tests were performed using the SigmaStat software package (Jandel Scientific, San Rafael, CA).

Exposure of freshly isolated CD3⁺T lymphocytes to sevoflurane (8 vol%) or the known apoptosis inducer MG132 (1 μm, positive control) for 24 h provoked the cell surface exposure of phosphatidylserine (GFP-annexin V staining) and nuclear fragmentation (Hoechst staining) (fig. 1A), two typical features of apoptosis. Necrosis was excluded by propidium iodide negativity (fig. 1A, lower panel). To quantify apoptosis in response to sevoflurane, we performed a flow cytometric analysis of the GFP-annexin V/propidium iodide–stained CD3⁺T cells. As shown in a representative experiment in figure 1B, 8 vol% sevoflurane exposure increased the number of GFP-annexin V–positive T cells (2.57 to 8.01%; lower right quadrant) without seriously perturbing membrane integrity (propidium iodide staining, 0.12 to 1.02%; upper right quadrant). Although this increase was small, it was significant (fig. 1C; median [range]: 16% [7–26%]*vs.  4% [3–12%] untreated control cells; n = 10; *P < 0.05). To determine whether induction of apoptosis was specific for a particular anesthetic, we performed additional GFP-annexin V/propidium iodide flow cytometric experiments using Jurkat T lymphocytes exposed to either desflurane (18 vol%; fig. 2A) or isoflurane (5 vol%; fig. 2B) for 24 h. In contrast to sevoflurane (fig. 1B), desflurane did not trigger any phosphatidylserine exposure or plasma membrane disruption (fig. 2A). As with sevoflurane (fig. 1B), isoflurane increased the number of GFP-annexin V–positive, apoptotic cells from 0.7 to 4.8%, although more propidium iodide–positive necrotic cells were detected as well (3.5 to 8.1%; fig. 2B). These data indicate that two types of T cells, primary CD3⁺T lymphocytes and Jurkat T lymphocytes, underwent apoptosis after exposure to sevoflurane or isoflurane.

To investigate whether the apoptotic effects of the anesthetics were dose dependent and associated with the activation of the crucial effector caspase, caspase 3, we exposed Jurkat T cells to various doses of sevoflurane (2.5, 5, and 8 vol%), isoflurane (1.5, 2.5, and 5 vol%), and desflurane (6, 12, and 18 vol%) and studied the processing of p32 procaspase 3 to its active p19/17 forms and the cleavage of its major substrate poly-ADP-ribose-polymerase (PARP) by immunoblotting. As shown in figure 3A, desflurane induced neither caspase-3 processing nor a significant cleavage of PARP from the p116 to the p85 form. In contrast, processed p19/17 caspase 3 and increased abundance of the p85 PARP product were detected at 5 vol% isoflurane and 5 and 8 vol% sevoflurane (figs. 3B and C). To confirm and quantify caspase-3 activation, we performed a caspase-3 activity assay of the cytosols of these T cells by using a specific fluorogenic caspase-3 substrate (DEVD-AMC). Jurkat T cells exposed to desflurane showed activity comparable to that of untreated control cells (fig. 3D; n = 10 each). In contrast, cells exposed to sevoflurane or isoflurane showed a significant increase in caspase-3 activity (fig. 3D; P < 0.05 vs.  untreated cells, n = 10 each). Therefore, T-cell death induced by sevoflurane and isoflurane was associated with caspase-3 activation, which confirms that it proceeded via  apoptosis.

Based on the current findings, it was interesting to investigate whether sevoflurane-induced apoptosis was dependent on caspases or could also be executed in a caspase-independent manner. For that purpose, we performed quantitative flow cytometric analysis after GFP-annexin V/propidium iodide staining and caspase-3 and PARP processing by immunoblotting in the presence of absence of the pan-caspase inhibitor N -benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone (Z-VAD.fmk). As shown by a representative experiment in figures 4A–C, sevoflurane (8 vol%) induced apoptosis (fig. 4B; lower right quadrant; 5.3% vs.  1.5% control) and secondary necrosis (fig. 4B; upper right quadrant; 9.1% vs.  3.3%) that was fully blocked by 100 μm Z-VAD.fmk treatment (fig. 4C; 1.1% lower right and 3.9% upper right). Statistical analysis of the sum of apoptotic and secondary necrotic cells of 10 independent experiments revealed that the changes were highly significant (comparison sevoflurane-treated vs.  untreated T cells: 15% [10–23%]*vs.  4% [3–12%] and comparison sevoflurane- vs.  Z-VAD.fmk/sevoflurane-treated: 15% [10–23%]*vs.  6% [5–6%], n = 10; *P < 0.05). Z-VAD.fmk (100 μm) alone had no influence on the GFP-annexin V/propidium iodide staining (data not shown). Concomitant with this finding, Z-VAD.fmk prevented the formation of the p19/p17 active forms of caspase 3 and the p85 cleavage product of PARP induced by sevoflurane (fig. 4D). These data show that sevoflurane requires caspases for the induction of apoptosis in Jurkat T cells.

We next wanted to know which apoptosis signaling pathway is used by sevoflurane. To assess the extrinsic Fas/CD95 receptor–induced pathway, we preincubated Jurkat T cells with a neutralizing anti-Fas/CD95 receptor antibody (anti-Fas/CD95; 1 μg/ml; cat. No. MD-11-3; MBL, Woburn, MA) 1 h before sevoflurane exposure and performed GFP-annexin V/propidium iodide flow cytometric analysis. These experiments demonstrated that sevoflurane-induced apoptosis and secondary necrosis could not be abolished by pretreatment with the anti-Fas/CD95 receptor antibody (fig. 5A,vs.  B vs.  C). By contrast, as expected, FasL-induced cell death was entirely blocked by preincubating with the neutralizing anti-Fas antibody (fig. 5D,vs.  E). The anti-Fas antibody alone did not affect cell death at all (fig. 5F). To confirm our results and investigate whether death receptor signaling in general had any impact on sevoflurane-induced apoptosis, we analyzed phosphatidylserine exposure and propidium iodide positivity in Jurkat T cells deficient for “Fas-associating protein with death domain” or caspase 8, two components that are absolutely required for death receptor signaling.25,26As shown in figure 6, similar numbers of GFP-annexin V– and/or propidium iodide–positive cells were counted irrespective of whether untreated or sevoflurane-treated cells were wild type, “Fas-associating protein with death domain” deficient, or caspase-8 deficient (6% [2–8%] of the untreated cells vs.  16% [10–21%] of the sevoflurane-treated wild-type T cells [fig. 6A,vs.  B], 9% [5–12%]vs.  23% [17–29%] of the “Fas-associating protein with death domain”–deficient T cells [fig. 6C,vs.  D], and 8% [5–12%]vs.  27% [19–33%] of the caspase-8–deficient T cells [fig. 6E vs.  F] expressed phosphatidylserine on their surface). Therefore, sevoflurane-induced apoptosis is not mediated via  the death receptor signaling pathway.

Loss of mitochondrial transmembrane potential and release of mitochondrial intermembrane proteins are important effectors of apoptosis. The incubation of Jurkat T lymphocytes with the fluorochrome DiOC⁶(3) allows the determination of the mitochondrial transmembrane potential. Flow cytometric analysis of DiOC⁶(3)-stained Jurkat T cells demonstrated that only 6% (3–10%, n = 10; P < 0.05) of untreated cells (fig. 7A) versus  18% (9–23%, n = 10; P < 0.05) of sevoflurane-treated cells had a significantly lower mitochondrial transmembrane potential. Because sevoflurane significantly decreased the mitochondrial transmembrane potential in Jurkat T lymphocytes, we hypothesized that cytochrome c would be released from the mitochondrial intermembrane space. As expected, MG132-treated Jurkat T cells exhibited cytochrome c release from mitochondria into the cytosol, with a resultant decrease in the mitochondrial content (fig. 7B, upper panel). In addition, semiquantitative analysis via  enzyme-linked immunosorbent assay revealed a significant decrease in mitochondrial cytochrome c content after MG132 treatment (fig. 7C). Interestingly, sevoflurane (8 vol% for 24 h) exposure of T cells resulted also in a smaller, although significant decrease of cytochrome c content in the mitochondrial fraction, with a concomitant increase in the cytosol as shown by both immunoblotting (fig. 7B, upper panel) and enzyme-linked immunosorbent assay (fig. 7C). Equal loading of the mitochondrial fraction was assured by immunodetection of the mitochondrial marker cyclooxygenase 1 (fig. 7B, bottom panel). These data clearly indicate that sevoflurane uses the mitochondrial signaling pathway for apoptosis induction in T cells.

The inflammatory stress response and the postoperative immunosuppression after anesthesia and major surgery are characterized by peripheral T cell lymphopenia and leukocytosis.6,13,27–29It has recently been shown that general anesthetics can modulate the inflammatory stress response.1,6This may result in an increased susceptibility to infectious complications, systemic inflammatory response syndrome, and sepsis.30Programmed cell death is an essential part of life in multicellular organisms and represents the predominant process responsible for the termination of surgical injury–induced inflammation and immunologic homeostasis.13,27However, evidence regarding immunomodulatory effects of volatile anesthetics due to apoptosis is conflicting. This may result from differences in study design as well as the inherent limitations of in vitro  models. The work presented here supports the hypothesis that volatile anesthetics, in particular sevoflurane, may act as triggers of apoptosis in primary CD3⁺and established Jurkat T lymphocytes. This effect may contribute to perioperative T-cell apoptosis after major surgery performed during general anesthesia and may play a crucial role in the defense against nosocomial infections.18 

Our data demonstrate that apoptosis induced by sevoflurane and, to a lesser extent, by isoflurane was caspase dependent and mitochondria mediated. We think that these effects were consistent and physiologically significant for the following reasons: (1) Both agents acted in a specific manner because another volatile anesthetic, desflurane, did not exert any apoptotic effect. (2) They provoked caspase-3 processing and apoptosis in a dose-dependent, pharmacologically relevant manner. (3) Caspase-3 processing was accompanied by increased caspase-3 activity and the cleavage PARP, a major caspase-3 substrate. (4) Apoptosis and caspase-3 processing was caspase dependent, i.e. , blocked by the general caspase inhibitor Z-VAD.fmk. (5) Sevoflurane-induced apoptosis was independent of the death receptor signaling pathway but involved mitochondrial signaling, i.e. , the decrease in the mitochondrial membrane potential and the release of cytochrome from the intermembrane space into the cytosol.

Several studies have already reported anesthetic-induced, caspase-dependent apoptotic cell death. For example, treatment of various immune cells with propofol resulted in growth inhibition, formation of apoptotic bodies, DNA laddering and fragmentation, and activation of caspases 3, 6, 8, and 9.19,20Morphine and various opioids potently induce opioid receptor–dependent apoptosis of human T cells, macrophages, microglia, and neurons.31–35This could in part be responsible for recurrent infections often seen with patients who are addicted to opiates. In addition, phenobarbital and diazepam cause apoptotic neurodegeneration in the developing rat brain at plasma concentrations relevant for seizure control in humans.36Anesthesia-related drugs, such as neuromuscular blocking agents, in particular pancuronium, seem to regulate apoptosis by increased expression and activation of the Fas/CD95 death signaling pathway.21A previous report also demonstrated that sevoflurane and isoflurane induced apoptosis in normal peripheral blood mononuclear cells in vitro .24However, compared with this latter study, our analysis here differed in the experimental setup and the obtained results. First, we studied CD3⁺T lymphocytes instead of peripheral blood mononuclear cells. Second, we administered volatile anesthetics by gaseous application, not by liquid injection. The former application may lead to an artificial initial peak of the volatile anesthetic concentration in the medium, a situation that is not comparable to clinical application.24With this protocol, the authors observed a higher percentage of apoptotic cells in the isoflurane as compared with the sevoflurane groups and concluded that the toxicity of isoflurane was higher than that of sevoflurane at equimolar aqueous concentrations.24This is in marked contrast to our results, where sevoflurane showed a higher apoptosis rate in T cells than isoflurane did.

The following lines of evidence suggest that our in vitro  findings of sevoflurane- and isoflurane-mediated apoptosis of T cells may be of biologic relevance and therefore crucial in a clinical setting. First, the apoptotic effect of these volatile anesthetics could be observed in human primary CD3⁺T lymphocytes that had been obtained from healthy donors. Second, the caspase-3 activation occurred at in vitro  concentrations comparable to those attained in the plasma of patients during administration of sevoflurane for general anesthesia.5,37,38Finally, it was previously shown that exposure of T cells to sevoflurane caused a diminished production of interleukin 3, a cytokine that plays a functional role in T-cell survival, growth differentiation, and leukocyte adhesion.5,39,40Likewise, sevoflurane was reported to inhibit the activation of the transcription factor activator protein 1 (AP-1) in human T lymphocytes in vitro .5AP-1 is known to be down-regulated after withdrawal of interleukin 2, interleukin 3, or granulocyte-monocyte colony-stimulating factor, and this is associated with decreased cell cycle progression and apoptotic cell death.24,41,42Therefore, the inhibition of AP-1 and interleukin 3 by sevoflurane observed could be responsible for the induction of apoptosis seen in our study.

Characteristically, cytotoxic drug–induced apoptosis involves permeabilization of mitochondrial outer membranes and provokes the release of mitochondrial components, such as cytochrome c (“intrinsic” pathway).43Therefore, it was not surprising that sevoflurane activated this pathway rather than death receptor signaling, to induce apoptosis. The release of cytochrome c, the sole water-soluble component of the electron transfer chain, leads to the failure of maintaining mitochondrial membrane potential and adenosine triphosphate synthesis. This would increase the production of reactive oxygen species with subsequent lipid peroxidation. We found that sevoflurane, at concentrations that induced apoptosis, provoked reactive oxygen species production (data not shown). These findings are consistent with the view that impaired functionality and induction of altered mitochondrial metabolism is an early key step of initiator caspase activation.12Alterations in mitochondrial energy metabolism after surgical trauma and general anesthesia are in accord with our in vitro  findings of organelle-specific initiation of cell death by volatile anesthetics.44We therefore conclude that mitochondria are central mediators of sevoflurane-induced apoptosis.

Taken together, our data suggest that sevoflurane and isoflurane are specific inducers of caspase-dependent, mitochondria-mediated apoptosis in human T cells. Therefore, our data provide a potential molecular mechanism for the apoptotic effects associated with general anesthesia.