1 Departments of Physiology, Anesthesiology, and Pharmacology/Toxicology, Medical College of Wisconsin and the Department of Veterans Affairs, Veterans Affairs Medical Center, Milwaukee, Wisconsin 53295; and 2 Department of Chemistry, Polytechnic University, Brooklyn, New York 11201
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ABSTRACT |
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Pulmonary arterial endothelial cells possess transplasma membrane electron transport (TPMET) systems that transfer intracellular reducing equivalents to extracellular electron acceptors. As one aspect of determining cellular mechanisms involved in one such TPMET system in pulmonary arterial endothelial cells in culture, glycolysis was inhibited by treatment with iodoacetate (IOA) or by replacing the glucose in the cell medium with 2-deoxy-D-glucose (2-DG). TPMET activity was measured as the rate of reduction of the extracellular electron acceptor polymer toluidine blue O polyacrylamide. Intracellular concentrations of NADH, NAD+, NADPH, and NADP+ were determined by high-performance liquid chromatography of KOH cell extracts. IOA decreased TPMET activity to 47% of control activity concomitant with a decrease in the NADH/NAD+ ratio to 34% of the control level, without a significant change in the NADPH/NADP+ ratio. 2-DG decreased TPMET activity to 53% of control and decreased both NADH/NAD+ and NADPH/NADP+ ratios to 51% and 55%, respectively, of control levels. When lactate was included in the medium along with the inhibitors, the effects of IOA and 2-DG on both TPMET activity and the NADPH/NADP+ ratios were prevented. The results suggest that cellular redox status is a determinant of pulmonary arterial endothelial cell TPMET activity, with TPMET activity more highly correlated with the poise of the NADH/NAD+ redox pair.
lung; endothelium; pyridine nucleotides
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INTRODUCTION |
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TRANSPLASMA MEMBRANE ELECTRON transport (TPMET) systems in various cell types are involved in cell defense (4), cell growth (13, 33), redox signal transduction (17, 20), cellular energetics (5, 23), regeneration of extracellular or plasma membrane antioxidants (34, 40, 42), iron transport (27), activation of redox cycling of xenobiotics (16), and tissue distribution of redox active pharmaceuticals (9, 29, 35). Pulmonary and other endothelial cells possess TPMET systems that transfer intracellular reducing equivalents to blood-borne electron acceptors (3, 6, 9, 10, 30, 31, 37, 38, 44, 45, 47). Thus these endothelial TPMET systems are uniquely situated to influence blood composition and vascular and organ function. However, the endothelial TPMET mechanisms are less well defined than those in some other cell types.
Several endothelial cell TPMET systems have been identified on the basis of different electron acceptor and/or donor specificities (6, 10, 30, 31, 37, 44, 45, 47). At least one such system can utilize the cell membrane-impermeant thiazine electron acceptor, toluidine blue O polyacrylamide (TBOP), as an electron acceptor (3, 10). Given that thiazines are acceptors for a fairly wide array of oxidoreductase systems and that the molecular size of the TBOP polymer precludes cell entry in either the oxidized or reduced forms, TBOP is a particularly useful probe for studying cellular mechanisms involved in TPMET (3, 10). The objective of the present study was to examine the influence of intracellular redox status on pulmonary arterial endothelial cell TBOP reduction via TPMET.
The pyridine nucleotide redox poise is probably the most direct measure of cellular redox status. In the present study, we attempted to manipulate the cytoplasmic redox status of pulmonary arterial endothelial cells and to determine the effects on both intracellular pyridine nucleotides and thiazine reductase TPMET activity. Intracellular NADH, NAD+, NADPH, and NADP+ were quantified by KOH extraction of the cells, followed by high-performance liquid chromatography (HPLC). TPMET was measured using an assay for TBOP reduction. The results indicate that this endothelial cell TPMET system is, in fact, sensitive to cellular redox status. The more highly consistent correlation with NADH/NAD+ than NADPH/NADP+ is consistent with NADH as a likely electron donor for this TPMET system.
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METHODS |
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Materials. Potassium hexacyanoferrate (III) [K3Fe(CN)6, ferricyanide], 2-deoxy-D-glucose (2-DG), iodoacetate (IOA), HEPES, lactate dehydrogenase (LDH) assay kit 340-LD, ATP, and Hoechst-33258 were purchased from Sigma Chemical (St. Louis, MO). Trypsin, penicillin-streptomycin, and RPMI 1640 tissue culture medium were from GIBCO (Grand Island, NY). Fetal bovine serum was from Hyclone Laboratories (Logan, UT), and Biosilon beads were from Nunc (Roskilde, Denmark). Type CLS2 collagenase was obtained from Worthington Biochemical (Freehold, NJ), and diiodoindocarbocyanine-acetylated low-density lipoprotein was purchased from Biochemical Technologies (Stoughton, MA). Protein determinations were performed using the Bio-Rad protein assay reagent (Bio-Rad Laboratories, Hercules, CA). NAD+, NADP+, NADPH, and NADH were purchased from Boehringer Mannheim (Indianapolis, IN).
Endothelial cell culture. The bovine pulmonary arterial endothelial cells were cultured in RPMI 1640 medium containing 10% fetal calf serum, 100 U/ml penicillin, 100 µg/ml streptomycin, and 30 mg/ml L-glutamine from segments of calf pulmonary artery obtained from a local meat processing plant as previously described (30). The cells were identified as endothelial cells by cobblestone morphology, which was observed by phase-contrast microscopy and accumulation of diiodoindocarbocyanine-acetylated low-density lipoprotein, as observed by fluorescence microscopy.
To prepare microcarrier beads coated with cells for experiments carried out in spectrophotometric cuvettes, cells between passages 4 and 20 that had grown to confluence in T-75 tissue culture flasks were detached from the flasks by treatment with trypsin (0.05% wt/vol) in Hanks' balanced salt solution (HBSS) containing 5.5 mM glucose. Approximately 5 × 106 cells were seeded onto 3 g (dry wt) of gelatin-coated (2% vol/wt) Biosilon beads (mean diameter 230 µm). The cells on the beads were cultured in biological stirrers (Techne, Princeton, NJ) in the medium described above. Stirring (60 rpm) was intermittent (2 min on, 30 min off) during cell attachment (~6 h) and continuous thereafter. The cells were grown to confluence on the beads as determined by observation with phase-contrast microscopy.Inhibitor treatments. For the inhibitor treatments, experimental and control conditions were carried out using cells from the same culture flask to control for cell culture variables, such as passage number. Inhibitor treatments were carried out in the same manner for studies of TPMET activity and for HPLC measurements of pyridine nucleotides, except as noted. Approximately 0.4- or 0.2-ml packed volume of cell-coated beads were aliquoted from the stirred culture flasks into 10 × 10 × 10-mm polystyrene spectrophotometric cuvettes (Sarstedt, Newton, NC) or conical-bottomed tubes for TPMET activity or HPLC studies, respectively. After the cell-coated beads settled, they were washed four consecutive times by resuspension in 3 ml of HBSS containing 10 mM HEPES, pH 7.4 (HBSS/HEPES), allowing the beads to settle between each wash. The cell-coated beads were resuspended in 3 ml of HBSS/HEPES containing the following treatments: none (control); 0.4 mM IOA; 0.4 mM IOA and 5 mM lactate; glucose-free HBSS/HEPES containing 10 mM 2-DG; or 2-DG and 5 mM lactate. The cells were incubated in the treatment media by mixing on a Nutator mixer at 37° for 30 min before measurements of TPMET activity or pyridine nucleotide and ATP concentrations.
TBOP polymer. To prepare the TBOP polymer, toluidine blue O was incorporated in an acrylamide polymer by copolymerization of toluidine blue O methylacrylamide and acrylamide as previously described (10). Polymer chains of <3.5 kDa were removed by dialysis through a 3.5-kDa cutoff membrane. The amount of reducible toluidine blue O per unit mass of the toluidine blue O polyacrylamide (TBOP+) was ~17 nmol/mg.
TPMET activity. TBOP+ reduction by the cells was measured in two ways. First, TBOP+ (2 mg/ml of HBSS/HEPES) was added to the cell-coated beads in spectrophotometric cuvettes, as previously described (10). Once the cell-coated beads had settled to the bottom of the cuvettes (~25 s), the initial absorbance of the medium above the settled beads was recorded at 590 nm with a Beckman DU 7400 spectrophotometer. The cuvette contents were then gently mixed with the Nutator mixer, and at 2.5-min intervals, the cuvettes were placed in the spectrophotometer where the cell-coated beads were allowed to settle. Absorbance was measured again, and the cuvettes were returned to the mixer.
The second means of measurement of TBOP+ reduction involved the use of the secondary electron acceptor, ferricyanide, to simplify the assay. The ferricyanide was used to oxidize the reduced TBOP (TBOPH), which is otherwise subject to autooxidation. In contrast to TBOPH, the reduced form of ferricyanide, ferrocyanide, does not autooxidize. The assay was carried out as previously described (30) by washing the cell-coated beads and adding 3 ml of fresh HBSS/HEPES containing TBOP+ and ferricyanide (0.2 mg/ml and 600 µM, respectively) to the cells in cuvettes. The wash solutions and the assay mixture also contained the same inhibitor treatments to which the cells were exposed during the previous 30-min treatment incubations. The absorbance of the medium surrounding the cells was measured, as described above, at 421 nm. Ferrocyanide, TBOP+, and TBOPH do not absorb at these wavelengths, allowing the concentration of ferricyanide to be determined using an extinction coefficient of 1 mMPyridine nucleotide HPLC. To determine the effects of the treatments on intracellular pyridine nucleotides and ATP, an HPLC method adapted from Stocchi et al. (39) was used. The cells on the Biosilon beads (200-µl packed bead volume) were treated as described in Inhibitor treatments. After 30 min of treatment, the cells on the beads were allowed to settle to the bottom of the tubes. The medium was removed from the tubes with a fine-tipped micropipette tip to remove as much of the residual extracellular medium as possible, and 0.3 ml of ice-cold 0.5 N KOH and 0.1 ml of HBSS/HEPES were added simultaneously to the cells on the microcarrier beads. The samples were mixed vigorously for 10 s with a vortex mixer, placed in an ice bath for 3 min, mixed vigorously after 1.5 min, followed by the addition of ice-cold water (0.225 ml). After being mixed vigorously, the contents of the tubes were centrifuged for 1 min at 2,100 g at 4°C. The cell extract was removed from the beads, which were dried and weighed to determine the cell culture surface area. The pH of 0.4 ml of the supernatant was adjusted to ~6.0 by the addition of 10 µl of 6 N HCl and 20 µl of 1 M ammonium acetate (pH 4.7). The pH was measured, and, if necessary, additional ammonium acetate solution was added to obtain a pH as close as possible to 6.0. After centrifugation at 2,100 g for 1 min at 4°C, the cell extract supernatant was filtered through a 0.22-µm cellulose acetate syringe filter (µStar syringe filters; Costar, Corning, NY), and 100 µl of the filtrate was injected into the HPLC system.
The HPLC system consisted of an autosampler fitted with a 100-µl sample loop, a dual piston pump, a variable wavelength detector, and a helium degasser (Hewlett Packard 1050, models 79865A, 79852A, 79853C, and 79856A, respectively). A Supelcosil octadecylsilane LC-18-T (3-µm particle size; 150 × 4.6 mm) column (Supelco, Bellefonte, PA) was used with a guard column (Upchurch, Oak Harbor, WA) that was packed with a pellicular medium composed of octadecyl groups chemically bonded to 37- to 53-µm glass beads (Whatman, Clifton, NJ). The components of the cell extract were separated at room temperature at a flow rate of 1.0 ml/min using a mobile phase consisting of two solutions, solutions A and B, that were made with reagent grade H2O (18.0 M
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Additional measurements. LDH activity in the treatment medium and in the cells was determined after each TBOP-ferricyanide reduction measurement, as previously described (30). The extracellular medium was removed from the cells, the cells were lysed, and LDH activity in the extracellular medium and in the lysed cells was measured. The fraction of the total cell LDH released into the medium during the study period was 4.5 ± 0.3 %mean (± SE) for all experiments with control and treated cells (n = 39), and no significant differences in LDH release between control and treated cells (one-way ANOVA, P > 0.05) were detected.
To normalize the data for comparisons between studies, the protein content of the cells in each experiment was measured as previously described (30), and cell DNA was measured using the assay of Labarca and Paigen (22). TPMET activity is expressed as picomoles of TBOPH-mediated ferricyanide reduction per minute per cm2 of cell surface area (pmol · min ![]() |
RESULTS |
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The reduction of TBOP+ to TBOPH by the bovine
pulmonary arterial endothelial cells is shown in Fig.
2. When TBOP+ was added to
the medium surrounding the cell-coated beads, the concentration of the
polymer-oxidized dye moieties in the medium decreased until a steady
state was reached (Fig. 2). The steady state represents the opposing
rates of reduction by the cells and autooxidation within the medium
(10). When the cells were removed from the medium (at the
time represented by the dotted line in Fig. 2), the medium
TBOP+ concentration returned to its initial value as the
result of autooxidation unopposed by reduction by the cells. The
complete recovery of TBOP+ in the medium upon reoxidation
confirmed that the disappearance of TBOP+ was due to
reduction by the cells.
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Figure 3 shows that ferricyanide
reduction by the cells depended on the presence of TBOP in the assay
medium. There was almost no reduction of ferricyanide by the cells on
this time scale in the absence of TBOP. The reduction of ferricyanide
in the presence of cells and TBOP indicates reduction of
TBOP+ to TBOPH by the cells. In the presence of TBOP, the
ferricyanide reduction was zero order until the ferricyanide was
essentially exhausted, as expected of a secondary electron acceptor
that does not affect the primary reaction rate. Thus the rate of
ferricyanide disappearance was used to calculate the TBOP+
reduction rate, as indicated in METHODS.
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Figure 4 shows examples of the effects of
the IOA and 2-DG treatments, respectively, on the rate of
TBOP+ reduction by the cells. Both inhibitors decreased the
rate of TBOP+ reduction (Fig. 4). When lactate was included
in the medium along with either IOA or 2-DG, reduction was not
inhibited (Fig. 4). The rate of TBOP+ reduction in the
absence and presence of lactate but without either 2-DG or IOA was
215 ± 5 and 230 ± 10 pmol · min1 · cm
2,
respectively, indicating that lactate alone did not substantially influence the rate of TBOP+ reduction by the cells in the
absence of the inhibitors.
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The pyridine nucleotide and ATP concentrations in control cells and
cells exposed to IOA and 2-DG in the absence or presence of lactate are
shown in Table 1. NADH concentrations
were lower in extracts of cells treated with IOA or 2-DG than in
control cells. This effect of the inhibitors was not observed when
lactate was included in the medium along with the inhibitors. No
significant differences were detected among the mean values of
NAD+, NADP+, and NADPH concentrations under
control and inhibitor treatment conditions without or with lactate. ATP
concentrations were lower in cells treated with IOA or 2-DG than in
control cells. Lactate prevented the effect of IOA, but not of 2-DG, on
ATP concentrations.
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The effects of the treatment conditions on the TBOP+
reduction rate and intracellular NADH/NAD+ and
NADPH/NADP+ ratios are shown in Figs.
5 and 6.
In general, changes in reduction rate and the poise of the
NADH/NAD+ and NADPH/NADP+ pairs were in the
same direction. However, the correlation was greater for the
NADH/NAD+ ratio than for the NADPH/NADP+ ratio,
which can be appreciated by the representation of the data in Fig.
7.
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DISCUSSION |
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The results suggest that the pulmonary arterial endothelial cell surface thiazine reductase is sensitive to the cytoplasmic redox status as reflected in the ratios of oxidized/reduced pyridine nucleotides. Thus it would appear that within the range of ratios achieved in this study, the intracellular supply of reducing equivalents is an important factor controlling the rate of electron transport to the extracellular electron acceptor.
The concept behind the manipulations carried out is that both IOA and 2-DG result in inhibition of the cytosolic source of NADH, but at two different sites along the glycolytic pathway, glyceraldehyde-3-phosphate dehydrogenase and hexokinase, respectively (18). Inhibition at either site can be overridden by the addition of lactate as an alternative source of NADH via the lactate dehydrogenase reaction. These manipulations have been used to examine the role of glycolysis in TPMET in endothelial and other cell types, as well as in other endothelial cell functions (7, 14, 25, 28, 32, 46, 48). However, their effects on endothelial intracellular NAD+, NADH, NADP+, NADPH, and ATP concentrations have not been previously determined.
In fact, there is little information available on endothelial cell
pyridine nucleotide concentrations in general. In the one study
available for comparison in which human umbilical vein endothelial cell
NAD+, NADP+, NADH, NADPH, and ATP were all
quantified, the concentrations were 78.4 ± 4.3, undetectable,
9.5 ± 1.5, 7.6 ± 1.1, and 595 ± 13 pmol · µg1 DNA (means ± SE,
n = 18), respectively, compared with 45.6 ± 2.5, 0.98 ± 0.1, 3.29 ± 0.2, 5.8 ± 0.3, and 427 ± 23 pmol · µg
1 of DNA (means ± SE),
respectively, in the present study (2). Considering
differences in cell source, culture conditions, etc., the baseline
levels of intracellular pyridine nucleotides and ATP in this study were
reasonably close to those measured in the human umbilical vein
endothelial cells. The comparison implies some consistency in related
metabolic functions of the two endothelial cell types in their
respective culture conditions and between the different extraction and
HPLC techniques.
As is generally true with the use of inhibitors, unintended effects are potentially confounding. IOA, a sulfhydryl alkylating agent, is not a highly specific inhibitor. While 2-DG appears to be rather specific for the hexokinase reaction, hexokinase is also a source of glucose-6-phosphate for NADPH via the pentose phosphate pathway, which is consistent with the 2-DG effects on the NADPH/NADP+ ratio. However, the common effects observed using the two inhibitors and the reversal of the common effects by lactate suggest that it is their common effects on the glycolytic source of NADH that is most important for their effects on this TPMET activity. It also appears to confirm that neither inhibitor effect was a result of direct TPMET inhibition. This further distinguished this endothelial thiazine reductase from that of at least one other TPMET system, the NADH-ascorbate free radical reductase of rat liver cells, which is apparently directly inhibited by IOA (41).
The results are also consistent with NADH actually being an electron donor for this TPMET system, which has been the interpretation of the results with similar metabolic inhibitors in other TPMET systems. For example, TPMET reduction of the tetrazolium WST-1 in various cell lines, including Jurkat, HeLa, and isolated spleen cells, was inhibited by IOA and 2-DG and enhanced by cyanide (7, 8). These results were interpreted as indicating that TPMET reduction of WST-1 in these cells was dependent on glycolytic NADH production, with sparing of cytoplasmic NADH by cyanide. A somewhat similar pattern was observed in a neuroblastoma cell dichloroindolphenol acceptor TPMET system that was stimulated by glucose, inhibited by 2-DG, but unaffected by cyanide (48). The correlation between NADH levels and oxygen reduction by TPMET systems, in some cell types, has been considered to be a demonstration of TPMET in cellular NADH/NAD+ balance when glycolysis is the dominant cellular energy source of ATP (5, 7, 8, 23).
Although this study does not directly impute NADH as the direct electron donor to the thiazine TPMET system, it is not obvious what other intermediate electron donor would be likely to link NADH levels to this TPMET activity. Ascorbate has been ruled out (30), and NADPH rather than NADH is generally considered to be the cytoplasmic source of electrons for regenerating intracellular electron donors such as glutathione (36). One possible intermediate between NADH and the TPMET system might be coenzyme Q10, which is a plasma membrane constituent in at least some cell types where it is apparently maintained in the hydroquinone form by a plasma membrane NADH-cytochrome b5 reductase (1, 19, 42).
The greater correlation between NADH/NAD+ and TBOP reduction rate than between NADPH/NADP+ and TBOP reduction rate does not rule out a role for NADPH either as a direct electron donor or as member of a more complicated electron transfer chain. The experimental conditions targeted glycolysis. Thus the proportionate range achieved for NADPH concentrations was not as great as for NADH. It would be interesting to expand that range in future studies. However, metabolic inhibitors and other tools available for manipulating intracellular NADPH tend to provide a more complex spectrum of cellular effects. Thus a more complex protocol will probably have to be devised to further address this question.
The measurements made in the current study were of total cytosolic and mitochondrial KOH-extractable cellular pyridine nucleotides, with no distinction between protein-bound and protein-free forms. However, the correlation between the TBOP+ reduction rate and the cellular NADH or NADPH concentrations or reduced/oxidized pyridine nucleotide ratios in the presence of the glycolytic inhibitors is consistent with the supposition that the differences observed in total cell NADH or NADPH reflected differences in available cytosolic levels. The observation that the poise of the NADH/NAD+ and NADPH/NAD+ pairs are so different, despite having virtually identical midpoint potentials, may be at least partly attributable to compartmentation and protein binding. However, this discrepancy has been seen even in the cytosolic pool, where it is apparently due in part to the reaction kinetics of several reversible, highly active cytosolic dehydrogenases and their substrate and product concentrations (43). Other factors involved in regulating the poise of the redox pairs include the rate of transfer of reducing equivalents between cellular compartments, the rate of utilization of NADH in the respiratory chain (43), and the rate of utilization of NADPH, predominately for regeneration of intracellular antioxidants and biosynthetic reactions.
The pulmonary endothelial thiazine reductase in the intact lung and in cells in culture reduces monomeric thiazine drugs to lipophilic cell-permeant compounds (9, 10). This physicochemical change dramatically influences the tissue distribution of this drug class (11, 15, 21, 24, 26). The results of the present study reveal that the redox status of the tissue, as reflected in the poise of the reduced/oxidized pyridine nucleotides, is likely to have an impact on the tissue distribution of thiazines as well as other compounds that are electron acceptors for this TPMET system. Because reduction commonly increases the lipophilicity and, therefore, the membrane permeation of such electron acceptors, their use in studying intracellular mechanisms involved in TPMET can be complicated by their own effects on intracellular metabolism (7, 29). Because TBOP cannot enter the cells in the oxidized or reduced forms, this complexity in interpretation is avoided.
In conclusion, these results indicate that cytoplasmic redox status, as reflected in the NADH/NAD+ and NADPH/NADP ratios, affects TPMET to thiazine compounds and also indicate that the NADH/NAD+ appears to be more directly correlated with TPMET activity. In addition, these studies suggest that future studies evaluating the possibility that TBOP+ reduction may provide a nondestructive index of changes in cell redox status may be worthwhile.
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ACKNOWLEDGEMENTS |
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This work was supported by the American Heart Association, National Heart, Lung, and Blood Institute Grants HL-24349 and HL-65537, and the Department of Veterans Affairs.
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FOOTNOTES |
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Address for reprint requests and other correspondence: M. P. Merker, Veterans Affairs Medical Center, Research Service 151, Milwaukee, WI 53295 (E-mail: mmerker{at}mcw.edu).
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.
First published October 12, 2001; 10.1152/ajplung.00283.2001
Received 24 July 2001; accepted in final form 11 October 2001.
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