Share this article on:

The Effects of Fentanyl on Hepatic Mitochondrial Function

Djafarzadeh, Siamak PhD; Vuda, Madhusudanarao PhD; Jeger, Victor MD, PhD; Takala, Jukka MD, PhD; Jakob, Stephan M. MD, PhD

doi: 10.1213/ANE.0000000000001280
Anesthetic Pharmacology: Original Laboratory Research Report

BACKGROUND: Remifentanil interferes with hepatic mitochondrial function. The aim of the present study was to evaluate whether hepatic mitochondrial function is affected by fentanyl, a more widely used opioid than remifentanil.

METHODS: Human hepatoma HepG2 cells were exposed to fentanyl or pretreated with naloxone (an opioid receptor antagonist) or 5-hydroxydecanoate (5-HD, an inhibitor of mitochondrial adenosine triphosphate (ATP)-sensitive potassium [mitoKATP] channels), followed by incubation with fentanyl. Mitochondrial function and metabolism were then analyzed.

RESULTS: Fentanyl marginally reduced maximal mitochondrial complex–specific respiration rates using exogenous substrates (decrease in medians: 11%–18%; P = 0.003–0.001) but did not affect basal cellular respiration rates (P = 0.834). The effect on stimulated respiration was prevented by preincubation with naloxone or 5-HD. Fentanyl reduced cellular ATP content in a dose-dependent manner (P < 0.001), an effect that was not significantly prevented by 5-HD and not explained by increased total ATPase concentration. However, in vitro ATPase activity of recombinant human permeability glycoprotein (an ATP-dependent drug efflux transporter) was significantly stimulated by fentanyl (P = 0.004).

CONCLUSIONS: Our data suggest that fentanyl reduces stimulated mitochondrial respiration of cultured human hepatocytes by a mechanism that is blocked by a mitoKATP channel antagonist. Increased energy requirements for fentanyl efflux transport may offer an explanation for the substantial decrease in cellular ATP concentration.

Supplemental Digital Content is available in the text.Published ahead of print April 13, 2016

From the *Department of Intensive Care Medicine, Inselspital, Bern University Hospital, University of Bern, Switzerland; and Department of Clinical Research, Graduate School for Cellular and Biomedical Sciences, University of Bern, Bern, Switzerland.

Accepted for publication January 29, 2016.

Published ahead of print April 13, 2016

Funding: Support was provided by the Swiss National Science Foundation (SNSF) (grant no. 32003B_127619/1) and by an MD-PhD scholarship received by Dr. Jeger from the SNSF (grant no. 133901).

The authors declare no conflicts of interest.

Supplemental digital content is available for this article. Direct URL citations appear in the printed text and are provided in the HTML and PDF versions of this article on the journal’s website.

Reprints will not be available from the authors.

Address correspondence to Stephan M. Jakob, MD, PhD, Department of Intensive Care Medicine, Inselspital, Bern University Hospital, University of Bern, CH-3010 Bern, Switzerland. Address e-mail to stephan.jakob@insel.ch.

Drugs can interfere with mitochondrial functions at various levels.1–10 For instance, they can inhibit mitochondrial respiration,2 DNA transcription, and glycolytic and fatty acid β-oxidation enzymatic activities.3 They may also induce the production of free radicals4 and a decrease in cellular endogenous antioxidants. Anesthetics, analgesics, and sedatives are among these drugs. In the mammalian heart, the inhalation anesthetic, isoflurane, inhibits complex I of the mitochondrial electron transport chain.11 In rat brain synaptosomes, propofol inhibits mitochondrial-coupled respiration and ATP production,12 and exposure of macrophages to propofol reduces Δψm and cellular ATP levels.13

Fentanyl is a widely used synthetic lipid-soluble short-acting narcotic analgesic.14–16 It is an opioid that interacts closely with μ-opioid receptors.17,18 It has been shown that fentanyl induces apoptosis of peripheral blood lymphocytes by disrupting Δψm and increasing production of reactive oxygen species.19 Vilela et al20 demonstrated that incubation of isolated brain mitochondria with fentanyl inhibits mitochondrial bioenergetics. Fentanyl is almost exclusively metabolized by the liver,18 and it reportedly impairs basal cellular oxygen consumption of isolated neonatal rat hepatocytes.21,22

Several studies have indicated that the opening of mitoKATP channels can modulate mitochondrial function.23–26 MitoKATP channels are located within the inner mitochondrial membrane and are involved in potassium influx into mitochondria. Under normal conditions, for proper mitochondrial function and cellular calcium homeostasis, the steady state of the mitochondrial matrix volume is maintained by a potassium efflux pathway (potassium-hydrogen antiporter).27 Potassium influx occurs by simple diffusion and mitoKATP channels.28–30 The opening of the mitoKATP channel will increase mitochondrial matrix volume.31,32 Transient-selective mitoKATP channel opening may provide functional protection against cellular injury and preserve cellular energy levels by regulating mitochondrial membrane swelling.29,33 However, continuous and prolonged mitoKATP channel opening induces cytochrome c release from mitochondrial intermembrane spaces, leading to apoptosis.29,33,34 Fentanyl enhances diazoxide-induced mitoKATP channel activity.35 In rats, fentanyl protects the heart against ischemic injury via opioid receptors and mitoKATP channel–linked mechanisms.36

Permeability glycoprotein (Pgp) is a transmembrane glycoprotein that functions as an ATP-dependent drug efflux pump and actively excretes a wide range of clinically important drugs and toxins out of the cell.37,38 Pgp can export structurally diverse hydrophobic compounds from the cell, and drugs that are transported by Pgp are identified as stimulators of its ATPase activity. In mice lacking Pgp, fentanyl-induced analgesia was increased and prolonged.39 Henthorn et al40 reported that fentanyl is a substrate of Pgp in cultured bovine brain microvascular endothelial cells. However, in a porcine kidney–derived cell line expressing human Pgp and transfected with the human multi-drug resistance 1 gene, fentanyl did not behave as a Pgp substrate.41 Thus, the effect of fentanyl on Pgp activity appears to be controversial in both in vitro and in vivo models.

The aim of the present study was to evaluate the effects of fentanyl on cellular and mitochondrial bioenergetics of cultured human hepatocytes and to investigate the potential contributing role of mitoKATP channels in this process. In addition, the in vitro effect of fentanyl on recombinant human Pgp ATPase activity in a cell membrane fraction was evaluated.

Back to Top | Article Outline

METHODS

Chemicals and Reagents

Fentanyl citrate was obtained from Janssen-Cilag AG (Baar, Switzerland) and naloxone from OrPha Swiss GmbH (Küsnacht, Switzerland). All the reagents for cellular respiration, media for cell culture, and 5-hydroxydecanoate (5-HD) were obtained from Sigma-Aldrich (Buchs, Switzerland).

Back to Top | Article Outline

Cell Culture

The human hepatoma cell line HepG2 (American Type Culture Collection) was cultured as described previously.42 All the experiments were performed with cells from the same batch at different passage numbers.

Back to Top | Article Outline

Incubation With Drugs

Quiescent cells were exposed at clinical and therapeutic blood concentrations (0.5 and 2 ng/mL)43–46 or higher (10 and 50 ng/mL) for 1 hour or pretreated with naloxone at 200 and 1000 ng/mL or 5-HD at 50 μM for 30 minutes, followed by incubation with fentanyl at 2 ng/mL for an additional hour at 37°C in a humid atmosphere (5% CO2, 95% air).

Back to Top | Article Outline

Cellular Respiration (High-Resolution Respirometry): Permeabilized and Intact Cells

After incubation, the cells were permeabilized to allow the entry of exogenous adenosine diphosphate (ADP) and oxidizable mitochondrial substrates to feed electrons into complexes of the respiratory system (complex I–dependent, complex II–dependent [after rotenone addition], and complex IV–dependent [after antimycin A addition] respiration). In addition, basal, coupled, and uncoupled respiration of intact cells was measured. This method evaluates oxygen consumption of intact cells without the addition of exogenous substrates and ADP, reproducing the respiratory function in the integrated cell. These methods are described in detail in the Supplemental Digital Content 1.

Back to Top | Article Outline

Measurement of Mitochondrial Membrane Potential (Δψm)

Measurement of Δψm in intact cells was performed using the dyes 5,5′,6′,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolocarbocyanine iodide (JC-1) and tetramethylrhodamine methyl ester (TMRM). These methods are described in detail in the Supplemental Digital Content 2 (http://links.lww.com/AA/B401).

Back to Top | Article Outline

Measurement of the HepG2 Cells’ ATP Content, ADP/ATP Ratio, Flavin-Adenine Dinucleotide Levels and ATP Synthase Enzymatic Activity, Reactive Oxygen Species/Reactive Nitrogen Species (ROS/RNS), and Mitochondrial Calcium Levels in HepG2 Cells

For these assessments, commercially available kits were used. These methods are described in detail in the Supplemental Digital Content 3 (http://links.lww.com/AA/B402).

Back to Top | Article Outline

Measurements of Extracellular L-Lactate Levels (Lactate Released to the Culture Medium)

Lactate concentrations in the cell culture supernatants were detected using a 96-well fluorescence-based assay kit. The method is described in the Supplemental Digital Content 4 (http://links.lww.com/AA/B403).

Back to Top | Article Outline

Transmission Electron Microscopy

Transmission electron microscopy was performed as described previously.42

Back to Top | Article Outline

Measurement of HepG2 Cells’ ATPase Activity and Cell-Free In Vitro Fentanyl-Stimulated Pgp ATPase Activity

The ATPase and Pgp ATPase activities of HepG2 cells were measured using commercially available kits (Supplemental Digital Content 5, http://links.lww.com/AA/B404).

Back to Top | Article Outline

Statistical Analysis

The SPSS 21.0 software package (SPSS Inc, Chicago, IL) was used for statistical analysis. Based on our previous findings,47 we set n = 20 per experiment for the initially planned investigations (mitochondrial respiration). To explore potential mechanisms for and consequences of the obtained results, we used sample sizes between 8 and 32, unless performed on 96-well plates, where 24 to 76 experiments were conducted, based on the available space. For the Pgp ATPase activity, 4 experiments per group were performed as recommended by the manufacturer. Only in one investigation were further experiments added after a first statistical analysis (mitochondrial respiration at a fentanyl concentration of 50 ng/mL). In all other experiments, the sample number was fixed before statistical analysis. Because of the relatively low sample sizes in many of the experiments, and after statistical consultation, nonparametric tests were used for all analyses. Comparisons of the slopes of oxygen concentrations (cellular respiration) of each individual experiment (active compound and control) were made using the Wilcoxon matched-pairs signed rank test. For unpaired data (mitochondrial calcium content, ROS/RNS, flavin-adenine dinucleotide [FAD] levels, ADP/ATP ratio, cellular ATPase activity, and lactate levels) the Mann-Whitney test was used. Statistical analysis of mitochondrial ATP synthase enzymatic activity, membrane potential, total cellular ATP content, and Pgp ATPase activities were performed using a Kruskal-Wallis rank sum test, followed by Dunn multiple comparisons. Data are shown as median and interquartile range (IQR). Because of the multiple experiments performed, the addition of experiments after a first statistical analysis for mitochondrial respiration at a fentanyl concentration of 50 ng/mL, and after statistical advice, a conservative approach was taken: results with a P value <0.01 were considered significant and results with a P value >0.15 not significant. Data are indicated as median and IQR in text and figures and displayed as box plots in figures, where boxes indicate median and IQR and whiskers represent minimum and maximum.

Back to Top | Article Outline

RESULTS

Cell and Mitochondrial Morphology Is Not Affected by Fentanyl

Figure 1

Figure 1

To evaluate whether fentanyl induces alterations in cellular and mitochondrial morphology, electron microscopy was performed. Transmission electron microscopic examination revealed no major alterations in cellular or mitochondrial ultrastructure under fentanyl treatment (n = 3) (Figure 1).

Back to Top | Article Outline

Respiratory Function in the Integrated Cell Using Endogenous Substrates Is Not Affected by Fentanyl

Figure 2

Figure 2

To evaluate whether fentanyl stimulation is implicated in hepatic mitochondrial dysfunction, we first investigated the effects of fentanyl exposure on respiratory function in the integrated cell using endogenous substrates and measured maximal capacity of the electron transport chain using carbonyl cyanide-p-trifluoromethoxyphenylhydrazone (FCCP). Compared with our previous investigations of effects of remifentanyl on hepatocytes’ mitochondrial respiration,47 we used a slightly higher number of experiments (20 per condition). A representative diagram of measurement of respiration rates in intact HepG2 using high-resolution respirometry is shown in Figure 2A. The uncoupler FCCP was titrated to an optimum concentration for maximum stimulation of flux. In the absence of exogenous substrates and ADP, fentanyl (2 ng/mL, 1-hour incubation) did not affect basal cellular respiration (Z: 0.000, P = 1.000), oligomycin-insensitive (nonphosphorylating respiration; Z: −0.751, P = 0.452), oligomycin-sensitive (ATP turnover; Z: −0.228, P = 0.819), or FCCP-uncoupled maximal respiration rates (Z: −0.280, P = 0.779; Figure 2B; n = 20 each). Uncoupled respiratory control ratios (the ratio between FCCP and oligomycin-insensitive respiration rates; median [IQR]; control: 8.1 [7.3–8.9]; fentanyl: 9.0 [7.6–9.9]; Z: −1.868, P = 0.062) may have been affected, but coupling efficiency (the ratio between oligomycin-sensitive and basal respiration rates; Z: −0.787, P = 0.431) was not (Figure 2C; n = 20 each).

Back to Top | Article Outline

Cellular Oxygen Consumption Is Impaired After Incubation With Fentanyl Using Exogenous Substrates Specific for Mitochondrial Complexes I, II, and IV

Figure 3

Figure 3

We further investigated whether fentanyl exposure induces alteration in mitochondrial respiration using exogenous substrates (maximal stimulated respiration; n = 20 per condition). A representative diagram of measurement of respiration rates in digitonin-permeabilized HepG2 cells using high-resolution respirometry is shown in Figure 3A. Fentanyl at 0.5 ng/mL did not reduce mitochondrial (complex I–dependent [Z: −0.655, P = 0.513] and complex II–dependent respiration [Z: −0.579, P = 0.562]). The effect on complex IV–dependent respiration was not large enough to be significant (control: 78.0 [62.5–89.8]; fentanyl: 70.0 [64.0–76.0] pmol/[seconds × million cells], Z: −1.946, P = 0.052; n = 20 each; Figure 3B). Fentanyl at 2 ng/mL induced a significant reduction in complex IV–dependent respiration (control: 90.0 [78.3–106.0]; fentanyl: 79.0 [68.3–89.5] pmol/[seconds × million cells]; Z: −3.305, P = 0.001), but probably not in complex I–dependent respiration (control: 51.0 [47.0–64.0]; fentanyl: 48.0 [44.3–52.0] pmol/[seconds × million cells]; Z: −1.552, P = 0.121). The effect on complex II–dependent respiration was not large enough to be significant (control: 103.5 [81.3–118.0]; fentanyl: 93.5 [73.5–99.0] pmol/[seconds × million cells]; Z: −2.316, P = 0.021; Figure 3C; n = 20 each). Fentanyl at 10 ng/mL did not significantly impair respiration of complex I–dependent (Z: −0.935, P = 0.350) and complex II–dependent respiration (Z: −0.411, P = 0.681), and the effect on complex IV–dependent respiration was not large enough to be significant (control: 83.0 [71.5–104.5]; fentanyl: 75.5 [65.5–89.5] pmol/[seconds × million cells]; Z: −2.223, P = 0.026; Figure 3D; n = 20 each). The effects of fentanyl at 50 ng/mL were at the limit of statistical significance after 20 experiments. Therefore, 10 experiments per condition were added. This resulted in a reduction in complex I–dependent (control: 35.9 [29.0–43.0]; fentanyl: 32.1 [25.0–39.2] pmol/[seconds × million cells]; Z: −2.985, P = 0.003), complex II–dependent (control: 67.0 [54.4–75.1]; fentanyl: 54.5 [45.8–67.3] pmol/[seconds × million cells]; Z: −3.264, P = 0.001), and complex IV–dependent respiration (control: 68.0 [62.4–77.5]; fentanyl: 60.0 [52.8–70.2] pmol/[seconds × million cells]; Z: −3.094, P = 0.002; Figure 3E, n = 30 each). Citric acid, the inactive ingredient of fentanyl, did not affect cellular respiration (data not shown).

Back to Top | Article Outline

Fentanyl Reduces Respiration Efficiency When Exogenous Fatty Acid Palmitate Is Used as a Substrate

In our experiments, the effect of fentanyl on stimulated mitochondrial respiration was in the magnitude of 10% to 20%. In studies in hepatocytes from suckling rats, morphine inhibited oxygen consumption by up to 25% and fentanyl by up to 40%21 when palmitate was used as a substrate for mitochondrial respiration. To evaluate whether fentanyl affects mitochondrial respiration to a greater extent when exogenous fatty acids are used as a substrate, we evaluated the effect of fentanyl on palmitate-dependent respiration. Fentanyl at 50 ng/mL reduced the efficiency of mitochondrial respiration (control: 5.8 [4.8–7.1]; fentanyl: 4.7 [4.4–5.2]; Z: −2.654, P = 0.008; n = 10; Figure 4C). The reduction at the other concentrations (fentanyl 2 ng/mL: 5.4 [5.05–5.9]; fentanyl: 4.5 [4.1–5.0]; Z: −2.449, P = 0.012; fentanyl 10 ng/mL: control: 5.7 [5.1–6.1]; fentanyl: 4.6 [4.0–5.1]; Z: −1.990, P = 0.047; n = 10 each; Figure 4, A and B) was not large enough to be significant. Neither dose affected state 3 respiration significantly (fentanyl 2 ng/mL: Z: −1.020, P = 0.308; fentanyl 10 ng/mL: Z: −0.255, P = 0.799; fentanyl 50 ng/mL: Z: −0.764, P = 0.445; n = 10 each; Figure 4, A–C). The effect on state 4o was not large enough to be significant (fentanyl 50 ng/mL: control: 5.1 [4.0–5.7]; fentanyl: 6.5 [4.9–7.6] pmol/[seconds × million cells]; Z: −2.142, P = 0.032; Figure 4, A–C).

Figure 4

Figure 4

All effects of palmitate disappeared if cells were pretreated with 5-HD (50 μM, 30 minutes) before the addition of fentanyl (2 ng/mL, 1 hour; state 3: Z: −0.534, P = 0.594; state 4o: Z: −0.474, P = 0.635; respiratory control ratio calculated as state 3/state 4o.: Z: −1.192, P = 0.233; n = 9 each; Figure 4D).

Back to Top | Article Outline

Fentanyl-Induced Reduction in Cellular Oxygen Consumption Is Prevented by Preincubation With Naloxone

We further evaluated whether antagonism with naloxone (an opioid receptor antagonist) can prevent fentanyl-induced reduction in cellular oxygen consumption. Cells were preincubated with naloxone for 1.5 hours either alone (Figure 5, B and D) or preincubated with naloxone (200 and 1000 ng/mL) for 30 minutes before the addition of fentanyl (2 ng/mL, 1-hour incubation; n = 20 each) (Figure 5, A and C). Controls for each series were incubated with medium alone. Antagonism with naloxone at 1000 ng/mL abolished the effect of fentanyl: naloxone 1000 ng/mL, complex I, Z: −0.981, P = 0.326; complex II, Z: −0.282, P = 0.778; complex IV, Z: −0.907, P = 0.364; n = 20 each; Figure 5, A and C). Naloxone at 200 ng/mL abolished the effect of fentanyl on complex I, Z: −0.841, P = 0.400, and was likely to abolish the effects on complex II (control: 76.5 [59.6–104.5]; fentanyl + naloxone: 71.5 [60.3–87.2] pmol/[seconds × million cells]; Z: −1.609, P = 0.108) and complex IV (control: 77.5 [68.0–102.0]; fentanyl + naloxone: 77.5 [65.0–85.8] pmol/[seconds × million cells]; Z: −1.681, P = 0.093).

Figure 5

Figure 5

Incubation of the cells with naloxone alone at nanogram per milliliter had no significant effect (naloxone 1000 g/mL, complex 1, Z: −1.252, P = 0.211; complex II, Z: −0.131, P = 0.896; complex IV, Z: −0.168, P = 0.867; n = 20 each; Figure 5, B and D). Incubation of the cells with naloxone at 200 ng/mL had no significant effect either on complex II (Z: −1.410, P = 0.159) or on complex IV (Z: −0.897, P = 0.370). The effect on complex I was not large enough to be significant (naloxone 200 ng/mL, control: 41.0 [38.0–45.0]; naloxone: 47.5 [42.3–53.0] pmol/[seconds × million cells]; Z: −2.207, P = 0.027).

Back to Top | Article Outline

Fentanyl-Induced Reduction in Cellular Respiration Is Abolished by 5-HD (an Inhibitor of Mitochondrial ATP-Sensitive Potassium [mitoKATP] Channels)

Because fentanyl enhances the effects of mitoKATP channel–opening drugs,35 we further evaluated whether 5-HD abolishes fentanyl-induced reduction in cellular respiration. Cells pretreated with the mitoKATP channel inhibitor 5-HD (50 μM, 30 minutes) before the addition of fentanyl (2 ng/mL, 1 hour) exhibited no significant changes in complex activities in comparison with controls (complex I, Z: −1.008, P = 0.313; complex II, Z: −1.269, P = 0.204; complex IV, Z: −1.344, P = 0.179; n = 20 each; Figure 6A). Incubation of the cells with 5-HD alone did not affect cellular respiration (complex I, Z: −0.859, P = 0.391; complex II, Z: −0.448, P = 0.654; complex IV, Z: −0.859, P = 0.391; n = 20 each; Figure 6B).

Back to Top | Article Outline

Mitochondrial Membrane Potential (Δψm) Is Not Affected by Fentanyl Using the Dyes TMRM and JC-1

Figure 6

Figure 6

Figure 7

Figure 7

Figure 8

Figure 8

Because reduced and inefficient mitochondrial respiration can result from lowered Δψm, we next investigated whether fentanyl induces alterations in mitochondrial membrane potential. For these experiments, we used 2 different dyes, JC-1 and TMRM (Figure 7 and 8). Fentanyl at 2 ng/mL for 1 hour did not induce a significant reduction in mitochondrial membrane potential measured with TMRM (control: 43.2 [34.9–56.3]; fentanyl: 44.2 [36.4–52.8]; 5-HD: 43.9 [33.5–55.7]; fentanyl + 5-HD: 46.8 [36.7–57.3] relative fluorescence units per nanogram cellular protein; Kruskal-Wallis rank sum test:

CV

CV

, P = 0.868; n = 45 each; Figure 7) and also did not appear to induce a significant reduction if measured with JC-1 (control: 0.43 [0.40–0.54] [n = 73]; fentanyl: 0.43 [0.38–0.47] [n = 74]; fentanyl + 5-HD: 0.44 [0.39–0.49] 590/530 fluorescence ratio [n = 76]; Kruskal-Wallis rank sum test:

CV

CV

, P = 0.094; Figure 8).

Back to Top | Article Outline

Cellular ATP Content but Not ATP Synthase Activity (Complex V or F1F0 ATPase) Is Reduced After Incubation With Fentanyl

Because [mitoKATP] channel stimulation and reduced mitochondrial respiration can decrease ATP content, this was evaluated in the next experiment. Although fentanyl at 2 ng/mL for 1 hour reduced cellular ATP content, this effect was not significantly attenuated by preincubation with 5-HD (50 μM) (control: 6.1 [5.3–7.4]; fentanyl: 3.6 [2.7–4.2]; fentanyl + 5-HD: 4.0 [3.4–4.8]; Kruskal-Wallis rank sum test:

CV

CV

, P < 0.001, followed by Dunn multiple comparisons test: control versus fentanyl: P < 0.001, fentanyl versus 5-HD + fentanyl: P = 0.126; n = 54 each; Figure 9A). The effect of fentanyl at 2 ng/mL for 1 hour was not large enough to alter cellular ATP synthase activity (control: 4.3 [3.9–4.7] mM/min/mg cellular protein; fentanyl: 4.1 [3.9–4.2] mM/min/mg cellular protein; fentanyl + 5-HD: 4.5 [4.0–4.9] mM/min/mg cellular protein; Kruskal-Wallis rank sum test:

CV

CV

, P =0.081; Figure 9B).

Figure 9

Figure 9

The ATP synthase inhibitor, oligomycin, at 500 nM inhibited the activity of the F1FO ATPase by >90% (data not shown), indicating the specificity of the reaction to the F1FO ATPase complex.

Back to Top | Article Outline

Lactate Released to the Culture Medium Is Not Affected by Fentanyl

Figure 10

Figure 10

To evaluate the significance of the decreased ATP content, we evaluated whether it was compensated for by increased glycolysis, which should increase lactate concentration in the cell culture medium. Incubation of the cells with fentanyl for 1 hour at 2 ng/mL did not significantly affect extracellular lactate levels (control: 1.8 [1.6–2.3] μM; fentanyl: 2.9 [2.2–3.8] μM; 5-HD: 3.3 [2.7–3.4] μM; fentanyl + 5-HD: 2.5 [2.4–3.8] μM; Kruskal-Wallis rank sum test:

CV

CV

; P = 0.032; n = 6 each; Figure 10).

Back to Top | Article Outline

ATPase Activity and ADP/ATP Ratio Are Not Affected by Exposure to Fentanyl

Figure 11

Figure 11

To characterize alterations in ATP metabolism suggested by the reduced ATP content, the activity of mitochondrial ATP synthase and the ADP/ATP ratio were studied. Treatment of the cells with fentanyl at 2 ng/mL for 1 hour neither induced an increase in total cellular ATPase activity (Mann-Whitney U test: Z: −1.172, P = 0.247; n = 10 each; Figure 11A) nor changed the ADP/ATP ratio (Mann-Whitney U test: Z: −0.988, P = 0.327; n = 20 each; Figure 11B).

Back to Top | Article Outline

Fentanyl Stimulates Cell-Free In Vitro Pgp ATPase Activity

Figure 12

Figure 12

5-HD did not seem to abolish fentanyl-induced reduction in cellular ATP content, and total ATPase activity of cells treated with fentanyl was unchanged. Measurements of total cellular ATPase activity can be challenging because each specific cellular ATPase may need specific conditions for optimal ATPase activation.48,49 Thus, we cannot exclude the possibility that there were masked changes in specific cellular ATPase activities under our assay conditions. Because it has been suggested that drugs such as morphine,50 fentanyl,40 and verapamil51 are substrates of Pgp, which is an ATP-dependent cellular transport or efflux ATPase, we next investigated whether fentanyl affects Pgp ATPase activity.37,38 For these experiments, recombinant human Pgp membrane fractions were treated with fentanyl at 2, 10, and 40 ng/mL and excess ATP for 30 minutes, and ATP consumption rates were measured. Fentanyl stimulated the rate of ATP consumption similarly to the positive control verapamil (Kruskal-Wallis rank sum test:

CV

CV

, P = 0.004; n = 4 each, according to the Pgp-Glo™ Assay System manufacturer’s instructions [Promega, Madison, WI]; Figure 12) and as found by others.40

Back to Top | Article Outline

Fentanyl Reduces Cellular ATP Content in a Dose-Dependent Manner

Figure 13

Figure 13

Because Pgp ATPase activity demonstrated dose responsiveness to fentanyl, additional experiments were performed to determine the cellular ATP content with escalating doses of fentanyl. For these experiments, cells were treated with fentanyl at 0.5, 2, 10, and 50 ng/mL for 1 hour, and cellular ATP content was measured. Fentanyl reduced cellular ATP content in a dose-dependent manner (Kruskal-Wallis rank sum test:

CV

CV

, P < 0.001; n = 24 each; Figure 13).

Back to Top | Article Outline

Mitochondrial Calcium Content, ROS/RNS, and FAD Levels

Finally, to elucidate further mechanisms and consequences of fentanyl-induced impairment of mitochondrial respiration and decreased ATP content—potentially, but not necessarily, related to mitoKATP channel stimulation—we measured intramitochondrial calcium concentration and ROS/RNS and FAD levels.

Figure 14

Figure 14

Because an excessive mitochondrial calcium level can impair respiratory capacity,52 we first investigated mitochondrial calcium content. Fentanyl had no effect on intramitochondrial calcium concentration (Mann-Whitney U test: Z: −1.116, P = 0.265; n = 32/16 [controls/fentanyl]; Figure 14A). Inefficiency in the mitochondrial respiratory capacity and ATP production may induce alterations in mitochondrial ROS generation. Depending on the condition, mitochondrial ROS generation can increase or decrease.53–55 For example, increased mitochondrial ROS generation has been shown when mitochondrial proton motive force increases,54 whereas, under low oxygen levels, impaired ROS generation has been reported.55 Therefore, we further evaluated cellular ROS/RNS levels. Fentanyl did not alter ROS/RNS levels significantly (control: 4.6 [4.3–4.8] vs fentanyl: 4.3 [4.1–4.6] nM/μg; Mann-Whitney U test: Z: −2.083, P = 0.037; n = 24 each; Figure 14B). To further evaluate and correlate mitochondrial electron transport chain substrate availability to respiration, we measured mitochondrial FAD levels. The effect of fentanyl cellular FAD levels was not large enough to be significant (control: 0.11 [0.10–0.11] vs fentanyl: 0.10 [0.10–0.11] nM/μg; Mann-Whitney U test: Z: −2.205, P = 0.028, n = 8 each; Figure 14C).

Back to Top | Article Outline

DISCUSSION

In the present study, we demonstrate that fentanyl slightly reduces cultured human hepatocytes’ mitochondrial respiration by a mechanism that is blocked by a mitoKATP channel antagonist. 5-HD seems to abolish a fentanyl-induced reduction in cellular ATP content only marginally if at all. No major visible ultrastructural alterations were found after incubation with fentanyl.

We measured mitochondrial respiration using specific exogenous substrates for mitochondrial complexes I, II, and IV, and, in addition, maximal respiration due to oxidation of the exogenous fatty acid palmitate. Therapeutic blood levels of fentanyl43–46 decreased complex II–dependent and complex IV–dependent mitochondrial and palmitate-dependent respiration although the effect was small.

In 2 studies, in neonatal rat hepatocytes, it has already been shown that analgesic doses of fentanyl reduced palmitate-dependent cellular oxygen consumption.21,22

In our study, naloxone, which is an opioid inverse agonist,17,18 attenuated fentanyl-induced reduction in mitochondrial respiration, indicating that the fentanyl effect was mediated by an opioid receptor mechanism. We incubated the cells with fentanyl for only 1 hour because fentanyl has a rapid onset and short duration of action, with a plasma half-life of 90 minutes and an elimination half-time of a few hours.56

The mitochondrial respiration impairment was accompanied by reductions in cellular ATP content. The reduction in ATP levels in the presence of fentanyl may be due to an effect of the drug on oxygen consumption or ATP synthase activity or on other components of mitochondrial/cellular function. In isolated rat brain mitochondria, high concentrations of fentanyl interfered with the mitochondrial electron transport chain (complexes II and IV).20 Reduced mitochondrial respiration and ATP production may induce the acceleration of glycolysis and lactate production to provide additional ATP. However, we found no increase in extracellular lactate levels with fentanyl treatment. Surprisingly, despite a reduction in mitochondrial ATP content, the ADP/ATP ratio remained unchanged after fentanyl exposure. Although the kit we used for total ATP concentration is more sensitive and inactivates ATPases in contrast to the kit for the assessment of the ADP/ATP ratio, we suggest that decreased ATP availability due to fentanyl-induced impairment of mitochondrial respiration, combined with increased ATP use for the Pgp efflux pump, can result in a decrease in total adenosine store and thereby cause a decrease in both ATP and ADP.

Fentanyl did not induce a reduction in mitochondrial membrane potential, as measured with TMRM, and also did not appear to induce a significant reduction if measured with JC-1. Changes in the mitochondrial respiratory capacity and ATP production may alter (increase or decrease) mitochondrial ROS generation.53,54 Increased mitochondrial ROS generation has been shown when mitochondrial proton motive force increases,54 and under hypoxic conditions, ROS generation has been reported to be decreased.55 In 1 study, incubation of peripheral blood lymphocytes with fentanyl induced apoptosis with disrupted Δψm and increased production of ROS.19 In our study in HepG2 cells, extracellular ROS concentrations were not altered significantly, and we did not find evidence that fentanyl uncoupled mitochondrial respiration.

MitoKATP channels may contribute to the regulation of mitochondrial function,23–26,57 and the presence of mitoKATP channels in hepatocytes has been shown.58 In rabbit ventricular myocytes, diazoxide, a mitoKATP channel opener, induced reversible oxidation of flavoproteins (an index of mitochondrial redox state).59 In isolated cardiac mitochondria, mitoKATP channel openers decreased mitochondrial membrane potential and ATP production and produced swelling.60 It has been suggested that potassium influx into the mitochondrial matrix increases matrix volume59 and reduces the potential across the inner mitochondrial membrane.30

In our present study, 5-HD (a mitoKATP channel antagonist) did not seem to abolish fentanyl-induced reduction in cellular ATP content. An alternative explanation for the reduction of cellular ATP content is the activation of cellular ATPase activity. Therefore, we first investigated whether fentanyl increases total cellular ATPase activity. The effect of fentanyl was not large enough to significantly alter total cellular ATPase activity. However, measurement of total cellular ATPase activity is challenging because each specific cellular ATPase may have different critical conditions for optimal ATPase activation.48,49 Thus, it is possible that there are masked specific cellular ATPase activities under our assay conditions. We then investigated whether fentanyl increases a specific cellular ATPase activity. It has been reported that opioids are Pgp substrates.61–63 Drugs, such as morphine, which stimulate Pgp ATPase activity are usually compounds for cellular transport or efflux.50 However, the effect of fentanyl on Pgp activity is somewhat controversial.39–41 In an in vitro study assessing transcellular movement of fentanyl in porcine kidney epithelial cells expressing human Pgp,41 fentanyl did not behave as a Pgp substrate. In contrast, Henthorn et al40 reported that, in cultured bovine brain microvascular endothelial cells, fentanyl is a Pgp substrate. The 2 different cell types used in these studies might have differed in terms of Pgp expression and activity, as well as in cell membrane composition and characteristics.40,41 In addition, different methods have been used to evaluate fentanyl as a substrate for Pgp. In an in vivo study in mice lacking Pgp, fentanyl-induced analgesia was increased and prolonged, suggesting that Pgp alters the efficiency of fentanyl.39 However, the authors did not perform detailed pharmacokinetic studies for fentanyl in plasma, and the effect of Pgp on fentanyl concentration in the brain was not evaluated.

In our present study using an in vitro cell-free system assay, we were able to demonstrate that fentanyl increases Pgp ATPase activity, suggesting that it is indeed a Pgp substrate. Protein expression of Pgp in HepG2 cells has been reported,64 and increased Pgp ATPase activity upon fentanyl treatment might account for the observed reduced ATP levels in our experiments. Because the Pgp ATPase activity demonstrated a dose responsiveness to fentanyl, we evaluated the effect of escalating fentanyl doses on cellular ATP content. We were able to demonstrate a dose-dependent reduction in ATP content. We suggest that this represents an increased need for ATP to power the Pgp efflux pump as the fentanyl concentration is increased. Our combined data indicate that ATP content is affected at fentanyl concentrations between 2 and 10 ng/mL, probably also dependent on methodological factors such as type of plates used to culture cells and the cell passage number.

Schematic representation of the proposed effect of fentanyl on mitochondrial function and Pgp activity is shown in Figure 15. μ-Opioid receptors are transmembrane proteins that couple to inhibitory G-proteins.65 After activation of μ-opioid receptors by fentanyl, the Gα and Gβγ subunits dissociate from one another and subsequently act on intracellular effector pathways such as tyrosine kinase and protein kinase C (PKC) pathways.65–67 It has been suggested that PKC activators, such as phorbol esters, enhance the opening of mitoKATP channels,68,69 and the opening of the mitoKATP channel by diazoxide (a mitoKATP channel opener) is mediated by the PKC signaling pathway.70

Figure 15

Figure 15

In our study, fentanyl reduced cultured human hepatocytes’ mitochondrial respiration by a mechanism that was blocked by a mitoKATP channel antagonist (5-HD). Opening of the mitoKATP channel was probably mediated by the activation of the PKC pathway. Antagonism with naloxone abolished a fentanyl-induced decrease in cellular respiration, indicating that fentanyl exerts these pharmacologic actions via receptor-mediated mechanisms. Fentanyl is highly lipid soluble and can pass across the cell membranes. 5-HD could not prevent fentanyl-induced reduction in cellular ATP content, indicating activation of cellular ATPases. Using an in vitro assay, we show that fentanyl is a substrate for efflux pump Pgp and directly stimulates its ATPase activity. In intact cells, PKC activators have been shown to increase phosphorylation and activity of the Pgp71–74 and decrease cellular drug accumulation.73,75,76 Therefore, fentanyl might exert its effect on the Pgp efflux pump either by direct interaction with it, by μ-opioid receptor–mediated activation of the PKC pathway or by a combination of both.

A limitation of the present study is the use of human hepatoma cell line HepG2. Because HepG2 cells contain high levels of mitochondria and mitochondrial DNA content, they are an excellent choice to study mitochondrial toxicity.77 However, HepG2 shows a different phenotype and certain dissimilarities when compared with primary human hepatocytes.78,79 For example, HepG2 cells secrete lower amounts of triglycerides78 and contain low levels of cytochrome P450 enzymes, which are involved in drug metabolism.80 Therefore, further experiments are warranted with primary hepatocytes or whole organs, before the results of our study can be translated to the human condition.

Although the effects of fentanyl on mitochondrial function that we documented are rather small, opioids—especially at high doses—can cause liver failure. As an example, the agonist/antagonist opioid buprenorphine, if injected IV, results in 80 times higher peak concentrations than after sublingual administration.81 This has been associated with cytolytic hepatitis.82 The mechanism related to this was evaluated in rat hepatocytes and in living mice, where high doses of buprenorphin were associated with impaired mitochondrial respiration and ATP formation.82 Similarly, street heroin, composed of heroin, 6-monoacetylmorphine, and morphine, induced mitochondrial dysfunction and apoptosis in cortical neuron cells.83 In rat liver mitochondria, morphine decreased mitochondrial ATPase, activity,84 and in SH-SY5Y cells, methadone caused concentration-dependent depletion of ATP levels.85

In patients receiving fentanyl for pain control, plasma concentrations are certainly lower than in drugs addicts. Nevertheless, in patients treated with fentanyl after cardiac arrest, fentanyl blood concentrations of up to 18 ng/mL were recorded,44 and in patients with cancer exposed to transdermal fentanyl, plasma concentrations of up to 14.7 ng/mL have been documented.45,46 Fentanyl plasma concentrations were highest in cachectic patients and in those with reduced renal function, signs of liver injury and/or inflammation.46 Therefore, our results suggest that mitochondrial dysfunction may play a significant role in patients exposed to very high fentanyl doses, such as in drug addicts and cachectic patients with liver and/or renal failure and infection. It has been proposed that, in the latter group of patients, plasma fentanyl concentrations could be measured and fentanyl administration reduced if the concentration is high.46 It is not known whether patients with intensive care unit-acquired weakness, significant muscle loss, organ failure, and inflammation achieve similarly high or possibly higher fentanyl concentrations. If these patients develop energetic failure, drug-induced mitochondrial dysfunction may be considered.

In summary, our data suggest that fentanyl reduces stimulated cultured human hepatocytes’ mitochondrial respiration by a mechanism that is blocked by a mitoKATP channel antagonist. 5-HD does not seem to completely abolish a fentanyl-induced reduction in cellular ATP content. This could not be explained by alterations in total cellular ATPase activity; however, fentanyl markedly stimulated the in vitro ATPase activity of recombinant human Pgp.

Back to Top | Article Outline

DISCLOSURES

Name: Siamak Djafarzadeh, PhD.

Contribution: This author helped design the study, conduct the study, collect the data, analyze the data, and prepare the manuscript.

Name: Madhusudanarao Vuda, PhD.

Contribution: This author helped conduct the study, collect the data, and analyze the data.

Name: Victor Jeger, MD, PhD.

Contribution: This author helped conduct the study, collect the data, and analyze the data.

Name: Jukka Takala, MD, PhD.

Contribution: This author helped design the study, analyze the data, and prepare the manuscript.

Name: Stephan M. Jakob, MD, PhD.

Contribution: This author helped design the study, conduct the study, collect the data, analyze the data, and prepare the manuscript.

This manuscript was handled by: Markus W. Hollmann, MD, PhD, DEAA.

Back to Top | Article Outline

REFERENCES

1. Neustadt J, Pieczenik SR. Medication-induced mitochondrial damage and disease. Mol Nutr Food Res 2008;52:780–8.
2. Chan K, Truong D, Shangari N, O’Brien PJ. Drug-induced mitochondrial toxicity. Expert Opin Drug Metab Toxicol 2005;1:655–69.
3. Fromenty B, Pessayre D. Impaired mitochondrial function in microvesicular steatosis. Effects of drugs, ethanol, hormones and cytokines. J Hepatol 1997;26(Suppl 2):43–53.
4. Dong H, Haining RL, Thummel KE, Rettie AE, Nelson SD. Involvement of human cytochrome P450 2D6 in the bioactivation of acetaminophen. Drug Metab Dispos 2000;28:1397–400.
5. Chitturi S, George J. Hepatotoxicity of commonly used drugs: nonsteroidal anti-inflammatory drugs, antihypertensives, antidiabetic agents, anticonvulsants, lipid-lowering agents, psychotropic drugs. Semin Liver Dis 2002;2:169–84.
6. Mansouri A, Haouzi D, Descatoire V, Demeilliers C, Sutton A, Vadrot N, Fromenty B, Feldmann G, Pessayre D, Berson A. Tacrine inhibits topoisomerases and DNA synthesis to cause mitochondrial DNA depletion and apoptosis in mouse liver. Hepatology 2003;38:715–25.
7. Ezoulin MJ, Li J, Wu G, Dong CZ, Ombetta JE, Chen HZ, Massicot F, Heymans F. Differential effect of PMS777, a new type of acetylcholinesterase inhibitor, and galanthamine on oxidative injury induced in human neuroblastoma SK-N-SH cells. Neurosci Lett 2005;389:61–5.
8. Balijepalli S, Boyd MR, Ravindranath V. Inhibition of mitochondrial complex I by haloperidol: the role of thiol oxidation. Neuropharmacology 1999;38:567–77.
9. Souza ME, Polizello AC, Uyemura SA, Castro-Silva O, Curti C. Effect of fluoxetine on rat liver mitochondria. Biochem Pharmacol 1994;48:535–41.
10. Sarah M, Poonam K. Diazepam induced early oxidative changes at the subcellular level in rat brain. Mol Cell Biochem 1998;178:41–6.
11. Hanley PJ, Ray J, Brandt U, Daut J. Halothane, isoflurane and sevoflurane inhibit NADH:ubiquinone oxidoreductase (complex I) of cardiac mitochondria. J Physiol 2002;544:687–93.
12. Marian M, Parrino C, Leo AM, Vincenti E, Bindoli A, Scutari G. Effect of the intravenous anesthetic 2,6-diisopropylphenol on respiration and energy production by rat brain synaptosomes. Neurochem Res 1997;22:287–92.
13. Wu GJ, Tai YT, Chen TL, Lin LL, Ueng YF, Chen RM. Propofol specifically inhibits mitochondrial membrane potential but not complex I NADH dehydrogenase activity, thus reducing cellular ATP biosynthesis and migration of macrophages. Ann N Y Acad Sci 2005;1042:168–76.
14. Soliman HM, Mélot C, Vincent JL. Sedative and analgesic practice in the intensive care unit: the results of a European survey. Br J Anaesth 2001;87:186–92.
15. Shapiro BA, Warren J, Egol AB, Greenbaum DM, Jacobi J, Nasraway SA, Schein RM, Spevetz A, Stone JR. Practice parameters for intravenous analgesia and sedation for adult patients in the intensive care unit: an executive summary. Society of Critical Care Medicine. Crit Care Med 1995;23:1596–600.
16. Walder B, Tramèr MR. Analgesia and sedation in critically ill patients. Swiss Med Wkly 2004;134:333–46.
17. Reisine T, Pasternak GW. Hardman JG, Limbird LE. Opioid analgesics and antagonists. Goodman & Gilman’s The Pharmacological Basis of Therapeutics.1996:Washington, DC: McGraw-Hill, 521–56.
18. Mather LE. Clinical pharmacokinetics of fentanyl and its newer derivatives. Clin Pharmacokinet 1983;8:422–46.
19. Delogu G, Moretti S, Antonucci A, Marandola M, Tellan G, Sale P, Carnevali R, Famularo G. Apoptogenic effect of fentanyl on freshly isolated peripheral blood lymphocytes. J Trauma 2004;57:75–81.
20. Vilela SM, Santos DJ, Félix L, Almeida JM, Antunes L, Peixoto F. Are fentanyl and remifentanil safe opioids for rat brain mitochondrial bioenergetics? Mitochondrion 2009;9:247–53.
21. Zamparelli M, Eaton S, Quant PA, McEwan A, Spitz L, Pierro A. Analgesic doses of fentanyl impair oxidative metabolism of neonatal hepatocytes. J Pediatr Surg 1999;34:260–3.
22. Zamparelli M, Eaton S, Spitz L, Pierro A. Amino acids counteract the inhibitory effect of fentanyl on hepatocyte oxidative metabolism. J Pediatr Surg 2000;35:736–9.
23. Debska G, Kicinska A, Skalska J, Szewczyk A, May R, Elger CE, Kunz WS. Opening of potassium channels modulates mitochondrial function in rat skeletal muscle. Biochim Biophys Acta 2002;1556:97–105.
24. Das M, Parker JE, Halestrap AP. Matrix volume measurements challenge the existence of diazoxide/glibencamide-sensitive KATP channels in rat mitochondria. J Physiol 2003;547:893–902.
25. Grover GJ, Garlid KD. ATP-Sensitive potassium channels: a review of their cardioprotective pharmacology. J Mol Cell Cardiol 2000;32:677–95.
26. O’Rourke B. Myocardial K(ATP) channels in preconditioning. Circ Res 2000;87:845–55.
27. Inoue I, Nagase H, Kishi K, Higuti T. ATP-sensitive K+ channel in the mitochondrial inner membrane. Nature 1991;352:244–7.
28. Mitchell P, Moyle J. Translocation of some anions cations and acids in rat liver mitochondria. Eur J Biochem 1969;9:149–55.
29. Garlid KD, Paucek P. Mitochondrial potassium transport: the K(+) cycle. Biochim Biophys Acta 2003;1606:23–41.
30. Garlid KD. Cation transport in mitochondria—the potassium cycle. Biochim Biophys Acta 1996;1275:123–6.
31. Ardehali H, O’Rourke B. Mitochondrial K(ATP) channels in cell survival and death. J Mol Cell Cardiol 2005;39:7–16.
32. Hanley PJ, Daut J. K(ATP) channels and preconditioning: a re-examination of the role of mitochondrial K(ATP) channels and an overview of alternative mechanisms. J Mol Cell Cardiol 2005;39:17–50.
33. Pomerantz BJ, Robinson TN, Heimbach JK, Calkins CM, Miller SA, Banerjee A, Harken AH. Selective mitochondrial KATP channel opening controls human myocardial preconditioning: too much of a good thing? Surgery 2000;128:368–73.
34. Costa AD, Quinlan CL, Andrukhiv A, West IC, Jabůrek M, Garlid KD. The direct physiological effects of mitoK(ATP) opening on heart mitochondria. Am J Physiol Heart Circ Physiol 2006;290:H406–15.
35. Zaugg M, Lucchinetti E, Spahn DR, Pasch T, Garcia C, Schaub MC. Differential effects of anesthetics on mitochondrial K(ATP) channel activity and cardiomyocyte protection. Anesthesiology 2002;97:15–23.
36. Kato R, Ross S, Foëx P. Fentanyl protects the heart against ischaemic injury via opioid receptors, adenosine A1 receptors and KATP channel linked mechanisms in rats. Br J Anaesth 2000;84:204–14.
37. Borst P, Schinkel AH. Genetic dissection of the function of mammalian P-glycoproteins. Trends Genet 1997;13:217–22.
38. Higgins CF. ABC transporters: from microorganisms to man. Annu Rev Cell Biol 1992;8:67–113.
39. Thompson SJ, Koszdin K, Bernards CM. Opiate-induced analgesia is increased and prolonged in mice lacking P-glycoprotein. Anesthesiology 2000;92:1392–9.
40. Henthorn TK, Liu Y, Mahapatro M, Ng KY. Active transport of fentanyl by the blood-brain barrier. J Pharmacol Exp Ther 1999;289:1084–9.
41. Wandel C, Kim R, Wood M, Wood A. Interaction of morphine, fentanyl, sufentanil, alfentanil, and loperamide with the efflux drug transporter P-glycoprotein. Anesthesiology 2002;96:913–20.
42. Regueira T, Lepper PM, Brandt S, Ochs M, Vuda M, Takala J, Jakob SM, Djafarzadeh S. Hypoxia inducible factor-1 alpha induction by tumour necrosis factor-alpha, but not by toll-like receptor agonists, modulates cellular respiration in cultured human hepatocytes. Liver Int 2009;29:1582–92.
43. Singleton MA, Rosen JI, Fisher DM. Plasma concentrations of fentanyl in infants, children and adults. Can J Anaesth 1987;34:152–5.
44. Bjelland TW, Klepstad P, Haugen BO, Nilsen T, Salvesen O, Dale O. Concentrations of remifentanil, propofol, fentanyl, and midazolam during rewarming from therapeutic hypothermia. Acta Anaesthesiol Scand 2014;58:709–15.
45. Bista SR, Haywood A, Hardy J, Norris R, Hennig S. Exposure to fentanyl after transdermal patch administration for cancer pain management. J Clin Pharmacol 2015 2015; [Epub ahead of print].
46. Suno M, Endo Y, Nishie H, Kajizono M, Sendo T, Matsuoka J. Refractory cachexia is associated with increased plasma concentrations of fentanyl in cancer patients. Ther Clin Risk Manag 2015;11:751–7.
47. Djafarzadeh S, Vuda M, Takala J, Jakob SM. Effect of remifentanil on mitochondrial oxygen consumption of cultured human hepatocytes. PLoS One 2012;7:e45195.
48. Vásárhelyi B, Szabó T, Vér A, Tulassay T. Measurement of Na+/K+-ATPase activity with an automated analyzer. Clin Chem 1997;43:1986–7.
49. Torlińska T, Grochowalska A. Age-related changes of NA(+), K(+) - ATPase, Ca(+2) - ATPase and Mg(+2) - ATPase activities in rat brain synaptosomes. J Physiol Pharmacol 2004;55:457–65.
50. Ambudkar SV, Dey S, Hrycyna CA, Ramachandra M, Pastan I, Gottesman MM. Biochemical, cellular, and pharmacological aspects of the multidrug transporter. Annu Rev Pharmacol Toxicol 1999;39:361–98.
51. Orlowski S, Mir LM, Belehradek J Jr, Garrigos M. Effects of steroids and verapamil on P-glycoprotein ATPase activity: progesterone, desoxycorticosterone, corticosterone and verapamil are mutually non-exclusive modulators. Biochem J 1996;317 (Pt 2):515–22.
52. Griffiths EJ, Rutter GA. Mitochondrial calcium as a key regulator of mitochondrial ATP production in mammalian cells. Biochim Biophys Acta 2009;1787:1324–33.
53. Sena LA, Chandel NS. Physiological roles of mitochondrial reactive oxygen species. Mol Cell 2012;48:158–67.
54. Echtay KS, Murphy MP, Smith RA, Talbot DA, Brand MD. Superoxide activates mitochondrial uncoupling protein 2 from the matrix side. Studies using targeted antioxidants. J Biol Chem 2002;277:47129–35.
55. Packer L, Fuehr K. Low oxygen concentration extends the lifespan of cultured human diploid cells. Nature 1977;267:423–5.
56. Frederic S, Bongard I, Darryl Y, Sue Janine RE.Current Diagnosis and Treatment, Critical Care. 20083rd ed. New York, McGraw Hill Medical Cop., .
57. Leanza L, Biasutto L, Managò A, Gulbins E, Zoratti M, Szabò I. Intracellular ion channels and cancer. Front Physiol 2013;4:227.
58. Nakagawa Y, Yoshioka M, Abe Y, Uchinami H, Ohba T, Ono K, Yamamoto Y. Enhancement of liver regeneration by adenosine triphosphate-sensitive K+ channel opener (diazoxide) after partial hepatectomy. Transplantation 2012;93:1094–100.
59. Liu Y, Sato T, O’Rourke B, Marban E. Mitochondrial ATP-dependent potassium channels: novel effectors of cardioprotection? Circulation 1998;97:2463–9.
60. Holmuhamedov EL, Jovanović S, Dzeja PP, Jovanović A, Terzic A. Mitochondrial ATP-sensitive K+ channels modulate cardiac mitochondrial function. Am J Physiol 1998;275:H1567–76.
61. Letrent SP, Polli JW, Humphreys JE, Pollack GM, Brouwer KR, Brouwer KL. P-glycoprotein-mediated transport of morphine in brain capillary endothelial cells. Biochem Pharmacol 1999;58:951–7.
62. Dagenais C, Graff CL, Pollack GM. Variable modulation of opioid brain uptake by P-glycoprotein in mice. Biochem Pharmacol 2004;67:269–76.
63. Dagenais C, Ducharme J, Pollack GM. Interaction of nonpeptidic delta agonists with P-glycoprotein by in situ mouse brain perfusion: liquid chromatography-mass spectrometry analysis and internal standard strategy. J Pharm Sci 2002;91:244–52.
64. Kano T, Wada S, Morimoto K, Kato Y, Ogihara T. Effect of knockdown of ezrin, radixin, and moesin on P-glycoprotein function in HepG2 cells. J Pharm Sci 2011;100:5308–14.
65. Al-Hasani R, Bruchas MR. Molecular mechanisms of opioid receptor-dependent signaling and behavior. Anesthesiology 2011;115:1363–81.
66. Childers SR, Snyder SH. Guanine nucleotides differentiate agonist and antagonist interactions with opiate receptors. Life Sci 1978;23:759–61.
67. Childers SR, Creese I, Snowman AM, Synder SH. Opiate receptor binding affected differentially by opiates and opioid peptides. Eur J Pharmacol 1979;55:11–8.
68. Sato T, O’Rourke B, Marbán E. Modulation of mitochondrial ATP-dependent K+ channels by protein kinase C. Circ Res 1998;83:110–4.
69. Liu Y, Ytrehus K, Downey JM. Evidence that translocation of protein kinase C is a key event during ischemic preconditioning of rabbit myocardium. J Mol Cell Cardiol 1994;26:661–8.
70. Wang Y, Ashraf M. Role of protein kinase C in mitochondrial KATP channel-mediated protection against Ca2+ overload injury in rat myocardium. Circ Res 1999;84:1156–65.
71. Chambers TC, McAvoy EM, Jacobs JW, Eilon G. Protein kinase C phosphorylates P-glycoprotein in multidrug resistant human KB carcinoma cells. J Biol Chem 1990;265:7679–86.
72. Hamada H, Hagiwara K, Nakajima T, Tsuruo T. Phosphorylation of the Mr 170,000 to 180,000 glycoprotein specific to multidrug-resistant tumor cells: effects of verapamil, trifluoperazine, and phorbol esters. Cancer Res 1987;47:2860–5.
73. Blobe GC, Sachs CW, Khan WA, Fabbro D, Stabel S, Wetsel WC, Obeid LM, Fine RL, Hannun YA. Selective regulation of expression of protein kinase C (PKC) isoenzymes in multidrug-resistant MCF-7 cells. Functional significance of enhanced expression of PKC alpha. J Biol Chem 1993;268:658–64.
74. Bates SE, Lee JS, Dickstein B, Spolyar M, Fojo AT. Differential modulation of P-glycoprotein transport by protein kinase inhibition. Biochemistry 1993;32:9156–64.
75. Dong ZY, Ward NE, Fan D, Gupta KP, O’Brian CA. In vitro model for intrinsic drug resistance: effects of protein kinase C activators on the chemosensitivity of cultured human colon cancer cells. Mol Pharmacol 1991;39:563–9.
76. Tsuruo T. Mechanisms of multidrug resistance and implications for therapy. Jpn J Cancer Res 1988;79:285–96.
77. Höschele D. Cell culture models for the investigation of NRTI-induced mitochondrial toxicity. Relevance for the prediction of clinical toxicity. Toxicol In Vitro 2006;20:535–46.
78. Dixon JL, Ginsberg HN. Regulation of hepatic secretion of apolipoprotein B-containing lipoproteins: information obtained from cultured liver cells. J Lipid Res 1993;34:167–79.
79. Pinti M, Troiano L, Nasi M, Ferraresi R, Dobrucki J, Cossarizza A. Hepatoma HepG2 cells as a model for in vitro studies on mitochondrial toxicity of antiviral drugs: which correlation with the patient? J Biol Regul Homeost Agents 2003;17:166–71.
80. Donato MT, Lahoz A, Castell JV, Gómez-Lechón MJ. Cell lines: a tool for in vitro drug metabolism studies. Curr Drug Metab 2008;9:1–11.
81. Berson A, Gervais A, Cazals D, Boyer N, Durand F, Bernuau J, Marcellin P, Degott C, Valla D, Pessayre D. Hepatitis after intravenous buprenorphine misuse in heroin addicts. J Hepatol 2001;34:346–50.
82. Berson A, Fau D, Fornacciari R, Degove-Goddard P, Sutton A, Descatoire V, Haouzi D, Lettéron P, Moreau A, Feldmann G, Pessayre D. Mechanisms for experimental buprenorphine hepatotoxicity: major role of mitochondrial dysfunction versus metabolic activation. J Hepatol 2001;34:261–9.
83. Cunha-Oliveira T, Rego AC, Garrido J, Borges F, Macedo T, Oliveira CR. Street heroin induces mitochondrial dysfunction and apoptosis in rat cortical neurons. J Neurochem 2007;101:543–54.
84. Cunha-Oliveira T, Silva L, Silva AM, Moreno AJ, Oliveira CR, Santos MS. Acute effects of cocaine, morphine and their combination on bioenergetic function and susceptibility to oxidative stress of rat liver mitochondria. Life Sci 2013;92:1157–64.
85. Perez-Alvarez S, Cuenca-Lopez MD, de Mera RM, Puerta E, Karachitos A, Bednarczyk P, Kmita H, Aguirre N, Galindo MF, Jordán J. Methadone induces necrotic-like cell death in SH-SY5Y cells by an impairment of mitochondrial ATP synthesis. Biochim Biophys Acta 2010;1802:1036–47.
86. Gross GJ, Peart JN. KATP channels and myocardial preconditioning: an update. Am J Physiol Heart Circ Physiol 2003;285:H921–30.
    87. Gross GJ, Fryer RM. Sarcolemmal versus mitochondrial ATP-sensitive K+ channels and myocardial preconditioning. Circ Res 1999;84:973–9.

      Supplemental Digital Content

      Back to Top | Article Outline
      © 2016 International Anesthesia Research Society