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Estimation of H2O2 gradients across biomembranes
Abstract
When cells are exposed to an external source of H2O2, the rapid enzymatic consumption of H2O2 inside the cell provides the driving force for the formation of the gradient across the plasma and other subcellular membranes. By using the concepts of enzyme latency, the following gradients – formed after a few seconds following the exposure to H2O2 – were estimated in Jurkat T-cells: [H2O2]cytosol/[H2O2]peroxisomes=3; [H2O2]extracellular/[H2O2]cytosol=7. The procedure presented in this work can easily be applied to other cell lines and provides a quantitative framework to interpret the data obtained when cells are exposed to an external source of H2O2.
1 Introduction
Signal transduction, development, cell proliferation, apoptosis, and necrosis, among other processes, are believed to be regulated by the redox status of the cell [1-4]. H2O2 is often the oxidant of choice in studies of redox-regulated processes, because it is continuously produced in aerobic metabolism and diffuses easily across cellular compartments [5]. In spite of the high permeability of H2O2, gradients across membranes are indeed formed when a membrane separates the production and consumption sites of H2O2. When cells are exposed to external H2O2, the fast consumption of H2O2 inside the cells provides the driving force for setting up a gradient across the plasma membrane (Fig. 1). The intracellular concentration of H2O2 is lower than the extracellular one and, thus, to establish the actual concentration of H2O2 that is implicated in the intracellular signaling events, this gradient must be determined.
One possible strategy to estimate gradients across biomembranes is based on the well-known fact that enzymes entrapped in compartments show a lower activity than enzymes free in solution, an observation brought forward in the early 1950s by De Duve and co-workers in their seminal studies on subcellular fractionation [6]. In general, the reason for enzyme latency is the permeability barrier constituted by the compartment entrapping the enzyme, which limits the diffusion of the substrate to the enzyme, and not some other factor, such as inhibition of the enzyme when trapped in the compartment.
In this work, we revisited the concepts of enzyme latency to estimate the H2O2 gradients produced in Jurkat T-cells – a cell line widely used in studies of redox regulation – upon exposure to an external source of H2O2. The procedure used can easily be applied to other cell types, thus constituting a general strategy to determine H2O2 gradients.
2 Materials and methods
2.1 Chemicals and biochemicals
Catalase (bovine liver), digitonin, and GSSG reductase (Baker's yeast) were from Fluka (Buchs, Switzerland). Glucose oxidase and NADPH were from Boehringer (Mannheim, Germany). DTPA, GSH, H2O2, NaN3, and Triton X-100 were from Sigma Chemical Co. (St. Louis, MO, USA). All other chemicals were of analytical grade.
2.2 Cell culture
Jurkat cells obtained from ATCC (clone E6-1) were cultured in complete medium (RPMI 1640 medium supplemented with 10% fetal calf serum, L-glutamine, and antibiotics from Life Technologies (Rockville, MD, USA). Cells were incubated at 37°C in humidified air with 5% CO2, and kept in logarithmic phase by routine passage every 2 days. Before use, cells were spun down, resuspended in fresh medium at 1×106 cells/ml, and incubated for at least 1 h. Cell viability was determined by propidium iodide uptake.
2.3 Biochemical measurements
H2O2 was measured with an oxygen electrode following the addition of catalase, which caused a rapid conversion of H2O2 to H2O and O2, the release of the latter being monitored by the oxygen electrode. With a clean and stable electrode, H2O2 concentrations as low as 5 μM could be measured. A calibration curve was made with H2O2 standards before each experiment. Catalase activity was measured as previously described [9] in 0.5 M potassium phosphate buffer, pH 7.0, containing 0.01% digitonin in the presence of 1×106 cells. H2O2 (10 mM initial concentration) consumption was followed at 240 nm at room temperature for 2 min (ϵ 240=43.4 M−1 cm−1). Alternatively, catalase activity was measured with a cell lysate. H2O2 concentrations were plotted on a semi-logarithmic graph against time and the first-order rate constant (catalase activity) was calculated. Titration of catalase activity with digitonin was carried out in a similar manner, but digitonin was added using dimethyl sulfoxide (DMSO) as a vehicle (corrections accounting for the slight inhibition of catalase activity by DMSO at the level added (1% v/v) were considered). Glutathione peroxidase (GPx) activity was measured by studying the kinetics of the enzyme in the whole cell applying a well-established method [10]. The assay mixture contained (final concentrations): 10×106 cells/ml, 0.05 M potassium phosphate buffer, pH 7.0, 1 mM DTPA, 50 μM NaN3, 1.1 U/ml glutathione reductase, 0.1 mM NADPH, 35 μM H2O2, and 1% (v/v) Triton X-100; the concentration of GSH varied between 0.335 and 3.35 mM. Alternatively, glutathione peroxidase activity was measured on a cell lysate without the addition of Triton X-100 to the assay mixture. All reactants, with the exception of H2O2, were pre-incubated at 37°C for 10 min. NADPH consumption was followed at 340 nm (ϵ=6.2×103 M−1 cm−1) at 37°C until all the hydroperoxide was used, recording the absorbance every 0.1 s. For the kinetic analysis, the part of the curve corresponding to 1.6–16 μM H2O2 was used as suggested [10]. Glucose oxidase activity was measured by following O2 consumption with an oxygen electrode.
3 Results and discussion
3.1 Consumption of H2O2 by intact Jurkat T-cells
The consumption of H2O2 by intact Jurkat T-cells was examined by two different experimental approaches: (a) exposure of cells to a bolus addition of H2O2 and (b) exposure of cells to a continuous flow of H2O2. Neither approach altered cell viability during the experiment(s).
In the former instances, the decay of H2O2 concentration after supplementing Jurkat T-cells with 100 μM H2O2 followed first-order kinetics (Fig. 2) with a k cell value of 1.0±0.1×10−3 s−1 per 106 cells (n=16). The growth medium (used within a few hours after resuspending cells) did not exhibit significant H2O2 consumption (not shown).
In the latter instances, cells were incubated with a low steady state of H2O2 (9 μM), by simultaneously exposing them to H2O2 and glucose oxidase, an enzyme that reduces O2 to H2O2 during glucose oxidation. Cells were able to maintain the steady state of H2O2 up to 2 h (Fig. 2); hence, their capacity to consume H2O2 did not decrease with time. The k cell obtained with this steady-state incubation approach was similar, within experimental error, to that obtained with a bolus addition of H2O2 and independent of cell density.
It may be surmised that a k cell value of 1×10−3 s−1 per 106cells is a reliable measure of the capacity of Jurkat T-cells to consume H2O2, for low to moderate concentrations of H2O2. Furthermore, the similar k cell values obtained with both experimental approaches, suggest that no substantial changes in GSH levels –sufficient to compromise H2O2 removal by glutathione peroxidase – occurred following a bolus addition of H2O2 to cells.
3.2 Consumption of H2O2 by disrupted Jurkat T-cells
To determine the consumption of H2O2 in disrupted Jurkat T-cells, the enzymatic activities that mainly remove H2O2, catalase and glutathione peroxidase were examined.
3.2.1 Catalase activity
In most tissues, but not all, this enzyme is present in peroxisomes. The subcellular location of catalase in Jurkat T-cells was examined by titration with digitonin, a drug that binds to cholesterol present in membranes forming pores; with this approach, membranes with higher levels of cholesterol are disrupted preferentially, thus allowing the study of enzyme compartmentation and latency [11]. Because only one latency threshold to digitonin was observed (Fig. 3), it may be surmised that in Jurkat T-cells all catalase activity is entrapped within one type of membrane, most likely the peroxisomal membrane.
Catalase activity in fully disrupted cells was 1.65±0.13×10−3 s−1 per 106 cells (n=5). As with the plasma membrane, the peroxisomal membrane constitutes a barrier to H2O2 diffusion and, thus, it is responsible for the latency of catalase activity. In intact cells, or in cells with only the plasma membrane disrupted, catalase activity represented 35% of the total catalase activity measured in fully disrupted cells (Fig. 3). Accordingly, the overall contribution of catalase to the removal of external H2O2 in intact cells was 35% of the value found in disrupted cells: 0.58±0.05×10−3 s−1 per 106 cells.
3.2.2 Glutathione peroxidase activity
Glutathione peroxidase is mainly present in cytosol and mitochondria [13], but also in the nucleus [14]. As mentioned above, glutathione peroxidase requires GSH to complete the catalytic cycle. GSH is not transported into cells [15] and, hence, titration of glutathione peroxidase activity with digitonin is not as useful as for catalase, because the latency of glutathione peroxidase will be determined by both H2O2 and GSH. Because most of the glutathione peroxidase activity is present in cytosol, an approximation of the cytosolic activity was inferred by considering the value found in the cell lysate: 6.0×10−3 s−1 per 106 cells. H2O2 consumption values in intact and disrupted Jurkat T-cells are listed in Table 1.
Parameters measured | |
H2O2 consumption by intact cells | |
k cell (bolus addition) | 1.0×10−3 s−1 per 106 cells |
k cell (continuous flow) | 1.0×10−3 s−1 per 106 cells |
H2O2 consumption by disrupted cells | |
Catalase pathway | |
disrupted peroxisomal membranes | 1.65×10−3 s−1 per 106 cells |
intact peroxisomal membranes | 0.58×10−3 s−1 per 106 cells |
Glutathione peroxidase pathway | 6.0×10−3 s−1 per 106 cells |
Parameters estimated | |
H2O2 catabolism | |
Turnover time | 0.2 s |
Glutathione peroxidase pathway | 4.1 s−1 (91%) |
Catalase pathway | 0.4 s−1 (9%) |
H2O2 profile and gradients | |
P s | 2×10−4 cm s−1 |
Equilibrium [H2O2]in/[H2O2]out | 0.9 s |
[H2O2]cytosol/[H2O2]peroxisomes | 3 |
[H2O2]out/[H2O2]in | 7 |
3.3 H2O2 catabolism
The information gathered on the activities of catalase and glutathione peroxidase above serves two purposes: on the one hand, it is essential for the calculation of an H2O2 gradient and, on the other hand, it makes it possible to characterize the catabolism of H2O2.
The turnover time of H2O2 may be calculated from the sum of the pseudo first-order rate constants for H2O2 consumption. Assuming that glutathione peroxidase and catalase represent the main H2O2 sinks, the capacity of Jurkat T-cells to consume H2O2 is 6.6×10−3 s−1 per 106 cells, which, after conversion to units referred to the cell volume 2 , corresponds to 4.5 s−1, thus yielding a turnover time of 0.2 s (1/4.5) (Table 1). This turnover time is slower than that estimated for hepatocytes (3×10−3 s) [12], an observation that may be explained by the very high activities of glutathione peroxidase and catalase present in the liver. Nevertheless, a turnover time of 0.2 s indicates that when Jurkat T-cells are exposed to extracellular H2O2 or to an agent that increases intracellular H2O2 production, an intracellular steady state will be reached in a few seconds.
The relative importance of catalase and glutathione peroxidase for the elimination of H2O2 can be calculated from the pseudo first-order rate constants obtained for these enzymes. Glutathione peroxidase activity (6×10−3 s−1 per 106 cells=4.1 s−1) is responsible for most of the H2O2 consumption in cytosol (91%), whereas catalase (0.58×10−3 s−1 per 106 cells=0.4 s−1) appears to play a minor role (9%), as also observed in other cell types, such as hepatocytes and endothelial cells [17, 18]. Therefore, most of the H2O2 added to Jurkat T-cells is probably removed via glutathione peroxidase with the concomitant change of the thiol/disulfide status inside the cell within a few seconds.
3.4 Profile of H2O2 in Jurkat T-cells
In addition to the gradient across the plasma membrane, a gradient is expected across the membrane of cellular organelles that consume significant amounts of H2O2, like peroxisomes [5] (which contain catalase), nucleus [14], mitochondria [13], and endoplasmic reticulum [13] (all of which contain glutathione peroxidase). For peroxisomes, results shown in Fig. 3 allow the estimation of the gradient across the peroxisomal membrane: in fact, the ratio measured between catalase activities in intact and in lysed peroxisomes was 0.35, which, according to Eq. 1, is the gradient between cytosolic and peroxisomal H2O2 concentrations upon incubation with an external source of H2O2. H2O2 gradients were not estimated for organelles containing glutathione peroxidase, but upon exposure to an external source of H2O2, the concentration of H2O2 in these organelles may be anticipated to be lower than in the cytosol. In organelles like lysosomes, that do not contain H2O2-consuming enzymes, a concentration similar to that in cytosol may be expected. Fig. 5 shows the H2O2 profile that is expected to form in Jurkat T-cells upon incubation with an external source of H2O2. Because H2O2 permeation and catabolism are fast, this profile will form, within a few seconds, whether the source is a steady state or a bolus addition.
4 Concluding remarks
When cells are exposed to external sources of H2O2, gradients across cell membranes are established, with a magnitude that depends on the intensity of the intracellular consumption of H2O2 and on the permeability characteristics of the membrane to H2O2. Once the profile of H2O2 concentration is known, the actual intracellular concentration of H2O2 that triggers the process under study can be easily estimated, which is useful for several reasons. First, the physiological relevance of the observations obtained can be discussed on a basis of comparing the concentration that was found to trigger the process in vitro with the concentration of H2O2 in vivo. For example, in Jurkat T-cells, a bolus addition of 50 μM of H2O2 is sufficient to induce apoptosis [21]. According to the results presented in this work, the maximal intracellular concentration of H2O2 reached inside the cells was around 7 μM, and in organelles like mitochondria it was probably around 2 μM. These values are much more likely to be reached in vivo than the 50 μM added, thereby supporting the physiological relevance of the findings. Second, different cell types, with distinct properties in terms of H2O2-consuming enzymes and of permeability to H2O2, form different gradients and consequently, the knowledge of the actual intracellular concentrations of H2O2 that elicit the same process in the various cell types helps to compare results between cell lines. It may be concluded that the procedure presented in this work provides a quantitative framework to interpret the data obtained upon incubation of cells with an external source of H2O2.
The data presented here provide further understanding on the bactericidal and bacteriostatic activities of H2O2 in body fluids [22-25]: on the one hand, bacteria – being small organisms – have a large ratio between cellular area and volume and, hence, the gradient established between extra- and intracellular concentrations of H2O2 is small or negligible [24]; on the other hand, the larger host cells would be subjected to a gradient as described in this work. As a corollary, for the same extracellular concentration of H2O2, the intracellular medium of bacteria will be subjected to a higher concentration of H2O2 than that of host cells, thus contributing to a selective toxicity of the peroxide towards pathogens.
Acknowledgements
F.A. acknowledges Grant BPD/11778/97 from PRAXIS XXI/FCT. Research supported by NIH Grant 1RO1-AG16718.