Propofol may protect neuronal cells from hypoxia re-oxygenation injury, possibly via an antioxidant actions under hypoxic conditions. This study investigated the molecular effects of propofol on hypoxia-induced cell damage using a neuronal cell line. Cultured human IMR-32 cells were exposed to propofol (30 μm) and biochemical and molecular approaches were used to assess cellular effects. Propofol significantly reduced hypoxia-mediated increases in lactate dehydrogenase, a marker of cell damage (mean (SD) for normoxia: 0.39 (0.07) a.u.; hypoxia: 0.78 (0.21) a.u.; hypoxia + propofol: 0.44 (0.17) a.u.; normoxia vs hypoxia, p < 0.05; hypoxia vs hypoxia + propofol, p < 0.05), reactive oxygen species and hydrogen peroxide. Propofol also diminished the morphological signs of cell damage. Increased amounts of catalase, which degrades hydrogen peroxide, were detected under hypoxic conditions. Propofol decreased the amount of catalase produced, but increased its enzymatic activity. Propofol protects neuronal cells from hypoxia re-oxygenation injury, possibly via a combined direct antioxidant effect along with induced cellular antioxidant mechanisms.
Hypoxia and re-oxygenation events occur under various clinical conditions including intra- and peri-operative brain ischaemia [1–5]. In the brain, hypoxia re-oxygenation often leads to severe cellular damage resulting in a loss of neurological function and a poor clinical outcome [6–8]. The underlying mechanisms of hypoxia re-oxygenation injury are multifaceted, but recent studies point towards a central role of reactive oxygen species, which are generated during the hypoxia re-oxygenation  and may harm cells by damaging DNA, lipid peroxidation and oxidation of proteins .
Propofol, a widely used intravenous anaesthetic, has been demonstrated to have neuroprotective properties in several in-vivo [11–14] and in-vitro [15–17] models of cerebral hypoxia re-oxygenation injury. Propofol contains a phenolic hydroxyl group and thus its structure resembles that of α-tocopherol (vitamin E) , a natural antioxidant. The antioxidant activity of propofol results partly from this phenolic chemical structure and may be responsible for its cytoprotective effects . Moreover, propofol has been shown to induce and regulate intracellular signalling pathways [20, 21] and the neuroprotective effects of propofol may also be mediated indirectly via the induction of cell-based antioxidant mechanisms.
In the study presented, we employed an in-vitro model of human neuronal cells to mimic hypoxia re-oxygenation events and evaluate the effects of anaesthetic doses of propofol on hypoxia re-oxygenation induced cell damage and molecular mechanisms using clinically relevant timing.
In-vitro hypoxia was induced employing our recently described enzymatic system in combination with nitrocellulose membranes . Membranes were co-incubated with culture medium for 60 min. During this time, partial pressure of oxygen (pO2) rapidly decreases to levels less than 5 mmHg which is defined as hypoxia . During hypoxia, culture flasks were closed tightly and filled with nitrogen, and pO2 was checked using a tissue oxygen pressure monitor (LICOX® CMP Oxygen Catheter; Integra, Plainsboro, NJ, USA). Cells were maintained in this hypoxic medium for 2 h. Hypoxia was terminated by the addition of fresh oxygenated culture medium. This step represents the re-oxygenation phase in vivo. All molecular, biochemical and cellular analyses were performed with samples taken during the re-oxygenation phase (Fig. 1). Stimulations were performed with anaesthetic concentrations of propofol (30 μm) (Sigma-Aldrich, Steinheim, Germany) dissolved in DMSO (Carl Roth GmbH, Karlsruhe, Germany). Briefly, a stock solution I of 250 mm propofol was prepared in DMSO and diluted 1:100 with the respective cell culture medium to achieve a 2.5-mm propofol stock solution II. The working solution of propofol (30 μm) was then prepared by diluting the stock solution II 1:83 with the respective cell culture medium. Therefore, the final concentration of DMSO in the working solution was 0.012%. Controls were performed by adding DMSO at a final concentration of 0.012%, which does not affect gene expression, cell proliferation or apoptosis [24–26].
IMR-32 neuroblastoma cells (LGC Standards, Teddington, UK) were cultured in RPMI-1640 medium (PAA, Coelbe, Germany) supplemented with 20% fetal calf serum (PAA), 4 mm l-glutamine (Seromed, Berlin, Germany), 1% non-essential amino acid (PAA) and 0.1 mg.ml−1 gentamycine (PAA). As high concentrations of fetal calf serum may affect the lactate dehydrogenase (LDH) and hydrogen peroxide levels, DMEM/F12 medium (PAA) containing 1% fetal calf serum was used during the hypoxia re-oxygenation period. Cells were cultured in a humidified incubator at 37 °C and 5% CO2. Confluent cells were detached by trypsinisation (0.025% trypsin-1 mM EDTA; Bioproducts, Ingelheim, Germany) at 37 °C. Cell numbers were evaluated using a haemocytometer and viability was assayed by the trypan-blue dye exclusion technique (Sigma-Aldrich). Cell morphology was judged using bright field microscopy (Leica, DMIL, Germany) combined with an LMscope camera and the XnView v1.95.4 software (Gougelet Pierre-Emmanuel, Reims, France).
For RT-PCR experiments, cells were washed with PBS (Sigma-Aldrich) once before RNA isolation and then lysed in RLT buffer (Qiagen GmbH, Hilden, Germany). The Qiagen RNeasy mini-kit in combination with the manufacturer’s protocol was employed to isolate RNA. 200 ng RNA was transcribed to cDNA using a reverse transcription kit (Applied Biosystems, Foster, CA, USA) with random hexamer primers. Primers (Metabion, Martinsried, Germany) used for PCR amplifications are shown in Table 1. PCR products were loaded onto 2.5% agarose gels containing 0.005% Roti®-Safe GelStain (Carl Roth) for electrophoresis and were visualised using UV-transilumination. Images were analysed with the ImageJ (v1.410, National Institutes of Health, Bethesda, MD, USA) software.
Table 1. PCR primers employed in the study.
Annealing temperature in °C
18s rRNA NR_003286.2
Lactate dehydrogenase released into the medium was measured using a colorimetric cytotoxicity detection kit (Roche, Mannheim, Germany). Samples were prepared according to the manufacturer’s protocol. Briefly, supernatants of the culture medium were collected 24 h after hypoxia and stored at −20 °C. For evaluation of total LDH, cells were lysed with 2% Triton X-100 (Carl Roth) and incubated 15 min at 37 °C. Lactate dehydrogenase activity from medium without cells was used as background control. Samples were measured in 96-well plates at 492 nm using an ELISA reader (Tecan, Crailsheim, Austria) in combination with the Magellan software v1.1 (Tecan Group Ltd, Männedorf, Switzerland).
Hydrogen peroxide levels were quantified with a QuantiChromTM Peroxide Assay Kit (BioAssay Systems, Hayward, CA, USA). Supernatants from cultured cells were collected 15 h after hypoxia and stored at −20 °C. Samples were evaluated in 96-well plates using the manufacturer’s protocol, and absorbance was measured at 585 nm employing an ELISA reader (Tecan).
Cell protein extraction for Western blotting was performed using RIPA buffer composed of 150 mm sodium chloride, 1% NP-40, 1% sodium deoxycholate, 0.1% sodium dodecyl sulphate (SDS), 50 mm Tris-HCl (pH7.6; all from Sigma-Aldrich). Protein concentrations were quantified with Roti®-Quant assays (Carl Roth). The respective amount of protein (30 μg) was mixed with 4× laemmli buffer (8% SDS, 40% glycerol, 20% 2-mercaptoethanol, 0.008% bromphenol blue, 0.25 m Tris-HCl, all from Sigma-Aldrich) and incubated for 3 min at 95 °C. Samples were separated by 10% SDS-PAGE and then transferred to a PVDF membrane (Amersham Pharmacia Biotech, Piscataway, NJ, USA). After blocking (Starting Block T20; Fisher Scientific, Schwerte, Germany) for 1 h at room temperature, the membranes were incubated with the primary antibody over night at 4 °C. Primary antibodies were directed against catalase (Abcam, Cambridge, UK; 1:15 000), caspase-3 (Santa Cruz, Heidelberg, Germany; 1:200) and β-actin (Santa Cruz, Heidelberg, Germany; 1:1000). Membranes were then washed in TBST buffer containing 0.05% Tween 20 (Sigma-Aldrich), and were incubated for 1 h with the secondary antibodies, anti-rabbit (Dako, Hamburg, Germany; 1:10 000) or anti-goat (Santa Cruz, Heidelberg, Germany; 1:2000). After washing in TBST buffer, signals were detected using the ECL kit (ECL-Plus Western blotting Detection Reagents, Amersham Pharmacia Biotech). Intensities of the protein bands were analysed using the ImageJ software (v1.410, NIH).
For reactive oxygen species measurements (H2DCFDA assays) cell numbers were evaluated with trypan-blue exclusion and 10 000 cells per well were cultured in collagen-coated black 96-well plates (Greiner Bio-One, Frickenhausen, Germany). After hypoxia re-oxygenation, cells were washed with pre-warmed PBS and cultured in a phenol red-free DMEM/F12 medium (Gibco®-invitrogen, NY, USA) with 1% fetal calf serum (PAA) for 15 h. Intracellular reactive oxygen species were measured using 10 μm 2′7′-dichlorodihydrofluorescein diacetate (Sigma-Aldrich) which is oxidised to fluorescent dichlorofluorescein (DCF) by intracellular reactive oxygen species. DCF-loaded cells were kept in the dark and were immediately placed in a fluorescence ELISA reader (Genios FL; Tecan) with the temperature set to 37 °C. Fluorescence was measured at an excitation wavelength of 485 nm and an emission wavelength of 535 nm. Data were acquired at time points 0 min and 30 min and results were calculated as the increase of fluorescence per well [(Ft30 − Ft0) / Ft0 * 100], where Ft30 = fluorescence at 30 min and Ft0 = fluorescence at 0 min .
For catalase activity assays, cells were detached from the growth surface 15 h after hypoxia using a rubber policeman followed by a centrifugation step at 400 × g for 10 min at room temperature. The cell pellet was sonicated and homogenised on ice in 300 μl cold PBS containing 1 mm EDTA (Sigma-Aldrich) and then centrifuged at 10 000 × g for 15 min at 4 °C. The supernatant was removed and the protein concentration was quantified using the Roti®-Quant assay. All samples were stored at −80 °C. Enzymatic activity of catalase was evaluated by a modification of the QuantiChromTM Peroxide Assay Kit (BioAssay Systems). Briefly, catalase (Sigma-Aldrich) was dissolved in water and the following standards were prepared: 50 units.ml−1, 25 units.ml−1, 12.5 units.ml−1, 6.25 units.ml−1, 3.125 units.ml−1, 1.563 units.ml−1, and 0 units.ml−1; 20 μl of each sample (1.25 μg protein) and catalase standards were added to 96-well plates. To start the reaction, 20 μl hydrogen peroxide (150 μm) solution was added to each well followed by 10 min incubation at room temperature. For the detection of catalase activity, 200 μl detection reagent (prepared as described by the manufacturer’s protocol) was added to each well. Plates were incubated at room temperature for 30 min and read at 585 nm using an ELISA reader (Tecan). Catalase activity in the samples was calculated from the standard curve.
All experiments were independently performed 3–4 times using at least duplicate samples. Statistics were performed by using the statistics software GraphPad Prism version 5.01 for Windows (GraphPad Software Inc., San Diego, CA, USA). Data were analysed by one-way analysis of variance, and in cases of significant differences, adjusted for multiple comparisons (Tukey).
RT-PCR experiments showed that addition of propofol significantly reduced the mRNA expression of caspase-3 (hypoxia: 0.23 (0.08) a.u.; hypoxia + propofol: 0.04 (0.05) a.u.; p < 0.05 vs hypoxia) whereas the expression of bax and bcl-2 was not significantly influenced by the addition of propofol (Fig. 2a). Although gene expression of caspase-3 was regulated by propofol, evaluation of protein levels of inactive pro-caspase-3 and active-caspase-3 did not show any differences among the groups (pro-caspase-3 hypoxia: 1.42 (0.04) a.u., pro-caspase-3 hypoxia + propofol: 1.51 (0.12) a.u., pro-caspase-3 normoxia control: 1.47 (0.14) a.u.; p > 0.05; active-caspase-3 hypoxia: 0.64 (0.08) a.u., active-caspase-3 hypoxia + propofol: 0.64 (0.05) a.u., active-caspase-3 normoxia control: 0.57 (0.04) a.u.; p > 0.05; Fig. 2b). Nevertheless, morphologically IMR-32 cells that received propofol before and during hypoxia showed reduced signs of cell damage such as cell rounding and detachment from the growth surface (Fig. 2c). This observation was also confirmed by the evaluation of released LDH as a marker for cell damage. Hypoxia increased the release of LDH (hypoxia: 0.78 (0.21) a.u., normoxia control: 0.39 (0.07) a.u.; p < 0.05) and addition of propofol reduced the hypoxia-mediated cell damage (hypoxia + propofol: 0.44 (0.17) a.u.; p < 0.05 vs hypoxia; Fig. 2d).
Hypoxia increased intracellular reactive oxygen species (hypoxia: 267.70 (28.91) a.u., normoxia control: 206.60 (27.99) a.u.; p < 0.01) and addition of propofol significantly reduced the generation of reactive oxygen species (hypoxia + propofol: 234.00 (33.39) a.u.; p < 0.05 vs hypoxia; Fig. 3a). Moreover, hydrogen peroxide released into the culture medium was significantly increased 15 h after hypoxia (hypoxia: 17.38 (2.70) μm, normoxia control: 1.68 (0.14) μm; p < 0.001) whereas addition of propofol resulted in diminished levels of hydrogen peroxide (hypoxia + propofol: 13.22 (1.29) μm; p < 0.05 vs hypoxia; Fig. 3b).
Catalase regulates the decomposition of hydrogen peroxide to water and oxygen and plays a central role in hydrogen peroxide metabolism. Gene expression of catalase was not changed by the addition of propofol (hypoxia: 0.20 (0.04) a.u., hypoxia + propofol: 0.11 (0.12) a.u.; p > 0.05; Fig. 4a). Western blotting results showed increased amounts of catalase under hypoxic conditions (hypoxia: 1.29 (0.04) a.u., normoxia control: 0.92 (0.04) a.u.; p < 0.001) and addition of propofol decreased the amount of catalase (hypoxia + propofol: 1.04 (0.03) a.u.; p < 0.01 vs hypoxia; Fig. 4b). However, measurements of catalase activity employing hydrogen peroxide based enzymatic assays revealed a statistically significant increase in the enzymatic activity after addition of propofol (hypoxia: 2.45 (0.46) U.ml−1, hypoxia + propofol: 5.17 (1.68) U.ml−1; p < 0.05; Fig. 4c).
Hypoxia and re-oxygenation are crucial events in physiological as well as pathophysiological processes and numerous clinical conditions are associated with hypoxia re-oxygenation . Especially in the brain, intra- or peri-operative ischaemia can lead to hypoxia re-oxygenation injury which is associated with severe neuronal cell damage [29, 30]. Propofol has been reported to exert neuroprotective effects in the setting of hypoxia re-oxygenation injury [31, 32], but the detailed cellular and molecular mechanisms for this are still under discussion .
In the present study, we employed a two-enzyme based hypoxia model in combination with human neuronal cells (IMR-32) to mimic hypoxia re-oxygenation in vitro and to evaluate the effects of anaesthetic doses of propofol on hypoxia re-oxygenation induced cell damage and molecular mechanisms. In contrast to other hypoxia models such as hypoxic chambers, in our enzymatic system, hypoxic conditions are established within minutes [22, 34], making it an ideal model for the investigation of clinically relevant scenarios and mechanisms that are associated with a rapid onset of hypoxia. Here we show that hypoxia significantly increased the damage of IMR-32 cells (LDH measurements, cell morphology), but did not change the activity of caspase-3 which is a key effector molecule in apoptosis . Anaesthetic doses of propofol (30 μm) [36, 37] decreased the hypoxia re-oxygenation induced cell damage without attenuating caspase-3 activity. Similar results have been obtained by Tu et al. who investigated the effects of propofol on neuropathogenesis in newborn rats under hypoxic conditions and showed that neither propofol nor hypoxia influenced apoptosis . Employing the neuroblastoma cell line SH-SY5Y, Wu et al. demonstrated that 10 μm propofol improved cell proliferation and inhibited dynorphin A induced cytotoxicity , an observation that is in concordance with our in vitro results obtained with 30 μm propofol.
Hypoxia re-oxygenation injury is associated with the generation of reactive oxygen species, which have various, mostly deleterious, effects on affected tissue [39–41]. Several authors suggest that cerebral hypoxia re-oxygenation leads to cytotoxic damage and that these events can be attenuated by the antioxidant activity of propofol [42, 43]. In our study, we showed that compared to normoxic conditions, reactive oxygen species and hydrogen peroxide concentrations are increased under hypoxia and that this effect is significantly attenuated by the addition of propofol. Several studies propose that the rise in intracellular reactive oxygen species is due to a loss of the mitochondrial membrane potential (Ψm) by the absence of oxygen. Under physiological conditions, the mitochondrial membrane is polarised and has a Ψm, which is maintained by ATP and proton gradients across the membrane. These gradients are in turn dependent on oxygen availability. Maintenance of Ψm keeps proteins such as cytochrome c and reactive oxygen species within the confines of the mitochondria. Periods of hypoxia, on the other hand disturb the proton gradients and have the potential to reduce Ψm, which in turn results in an increase of reactive oxygen species in the cytoplasmic compartment [28, 44]. Under physiological conditions cytoplasmic reactive oxygen species are scavenged by superoxide dismutase, glutathione peroxidase, catalase as well as other small molecular antioxidants, including glutathione, ascorbic acid and α-tocopherol. These endogenous antioxidants are unable to cope with the excess reactive oxygen species generated during hypoxia re-oxygenation, resulting in an increased oxidation of macromolecules (e.g. lipids, proteins and nucleic acids) and cell damage .
Although our findings confirm the above-mentioned studies suggesting antioxidant properties of propofol as at least partly responsible for the observed neuroprotective effects, they do not verify whether direct antioxidant effects of propofol mediated by its phenolic chemical structure, indirect actions of propofol (e.g. induction of an antioxidant response in the target cells) or a combination of both are responsible for the antioxidant activity of the anaesthetic.
Catalase catalyses the decomposition of hydrogen peroxide to water and oxygen and may therefore be a potential candidate for indirect, target cell-mediated cytoprotective effects of propofol . Our Western blotting experiments show a reduction of the amount of catalase protein in IMR-32 cells after the addition of propofol. In contrast, activity measurements revealed an increased catalase activity in cells that received propofol, pointing towards a possible augmentation of the catalytic function by direct interactions of the anaesthetic with the catalase molecule. Another explanation for our findings could be the fact that the catalase activity assay employed in this study uses hydrogen peroxide as substrate and may also detect other hydrogen peroxide degrading peroxidases. This hypothesis is further supported by our preliminary studies performed using a catalase-specific blocker that revealed that 5–12% of the measured activity is not due to catalase but to other peroxidases. That propofol may indeed increase the activities of other hydrogen peroxide degrading enzymes has also been demonstrated by groups that detected elevated activities of glutathione peroxidase in serum and plasma of pigs and adult patients during propofol anaesthesia [47, 48].
Taken together, our results show that propofol can protect neuronal cells from hypoxia re-oxygenation injury and that the associated effects may at least partly be due to the capability of the anaesthetic to induce cellular antioxidant mechanisms.
The authors would like to thank C. Heinrich and C. Rodde for technical assistance and advice.
No external funding and no competing interests declared.