Propofol induces a metabolic switch to glycolysis and cell death in a mitochondrial electron transport chain-dependent manner

Propofol induces a metabolic switch to glycolysis and cell death in a mitochondrial electron transport chain-dependent manner | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Propofol induces a metabolic switch to glycolysis and cell death in a mitochondrial electron transport chain-dependent manner Chisato Sumi , Akihisa Okamoto , Hiromasa Tanaka , Kenichiro Nishi , Munenori Kusunoki , Tomohiro Shoji , Takeo Uba , Yoshiyuki Matsuo , Takehiko Adachi , Jun-Ichi Hayashi , Keizo Takenaga , View ORCID Profile Kiichi Hirota doi: https://doi.org/10.1101/181933 Chisato Sumi 1 Department of Anesthesiology, Kansai Medical University , Hirakata, Japan 2 Department of Human Stress Response Science, Institute of Biomedical Science, Kansai Medical University , Hirakata, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Akihisa Okamoto 2 Department of Human Stress Response Science, Institute of Biomedical Science, Kansai Medical University , Hirakata, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Hiromasa Tanaka 2 Department of Human Stress Response Science, Institute of Biomedical Science, Kansai Medical University , Hirakata, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Kenichiro Nishi 2 Department of Human Stress Response Science, Institute of Biomedical Science, Kansai Medical University , Hirakata, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Munenori Kusunoki 1 Department of Anesthesiology, Kansai Medical University , Hirakata, Japan 2 Department of Human Stress Response Science, Institute of Biomedical Science, Kansai Medical University , Hirakata, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Tomohiro Shoji 1 Department of Anesthesiology, Kansai Medical University , Hirakata, Japan 2 Department of Human Stress Response Science, Institute of Biomedical Science, Kansai Medical University , Hirakata, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Takeo Uba 1 Department of Anesthesiology, Kansai Medical University , Hirakata, Japan 2 Department of Human Stress Response Science, Institute of Biomedical Science, Kansai Medical University , Hirakata, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Yoshiyuki Matsuo 2 Department of Human Stress Response Science, Institute of Biomedical Science, Kansai Medical University , Hirakata, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Takehiko Adachi 3 Department of Anesthesiology, Tazuke Kofukai Medical Institute Kitano Hospital , Osaka, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Jun-Ichi Hayashi 4 University of Tsukuba , Tsukuba, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Keizo Takenaga 5 Department of Life Science, Shimane University Faculty of Medicine , Izumo, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Kiichi Hirota 2 Department of Human Stress Response Science, Institute of Biomedical Science, Kansai Medical University , Hirakata, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Kiichi Hirota Abstract Full Text Info/History Metrics Supplementary material Preview PDF Abstract The intravenous anesthetic propofol (2,6-diisopropylphenol) has been used for the induction and maintenance of anesthesia in operating rooms and for sedation in intensive care units. Although there is no widely accepted definition of propofol infusion syndrome (PRIS), PRIS is defined as the development of metabolic acidosis, rhabdomyolysis, hyperkalemia, hepatomegaly, renal failure, arrhythmia, and progressive cardiac failure. In vitro evidence suggests that PRIS is related to the impaired mitochondrial function. There are indications that preexisting mitochondrial disorders predispose to PRIS. However, the precise molecular mechanisms, including mitochondrial defects and a metabolic conversion by propofol, are largely unknown as yet. To elucidate the underlying cellular and molecular mechanisms of PRIS, we investigated the effects of propofol on the cellular metabolic mode and cell death. We demonstrated that clinically relevant concentrations of propofol, used within a clinically relevant exposure time, suppressed the mitochondrial function, caused the generation of reactive oxygen species, and induced a metabolic switch, from oxidative phosphorylation to glycolysis, by targeting complexes I and III of mitochondria. The data also indicated that a predisposition to mitochondrial dysfunction, caused by a genetic mutation or pharmacological suppression of the electron transport chain by biguanides such as metformin and phenformin, promoted the cell death and caspase activation induced by propofol. Introduction Since its introduction into clinical practice in 1986, propofol (2,6-diisopropylphenol) has been used for the induction and maintenance of anesthesia in operating rooms and for sedation in intensive care units 1 . Although propofol is considered a safe agent for anesthesia and sedation, a rare but severe complication can occur, especially in patients receiving high doses of the anesthetic for prolonged periods. A number of clinical reports have indicated a serious side effect of propofol, propofol infusion syndrome (PRIS) 2 . Since PRIS was first described in 1992 3 , the clinical awareness and research interest into this disorder have continued to grow. The medical literature now includes over 100 published case reports, case series, and reviews. Nonetheless, the exact incidence and etiology of PRIS remain unclear at present. Although there is no widely accepted definition of PRIS, PRIS is defined as the development of metabolic acidosis (lactic acidosis), rhabdomyolysis, hyperkalemia, hepatomegaly, renal failure, arrhythmia, and progressive cardiac failure 2 . There is a strong association between PRIS and propofol infusion at doses greater than 4 mg/kg/h and an exposure longer than 48 h, although the precise molecular mechanisms of PRIS have not been elucidated. In vitro evidence suggests that PRIS is related to impaired mitochondrial function 4 . There are indications that preexisting mitochondrial disorders predispose to PRIS 4 - 6 . Moreover, studies using isolated mitochondria have demonstrated that propofol showed an effect on the mitochondrial respiratory chain. A decrease in the transmembrane electrical potential (ΔΨ) has been reported in liver mitochondria isolated from rats incubated with propofol 7 . An increase of the oxygen consumption rate (OCR) has suggested that propofol acts as an uncoupler in oxidative phosphorylation (OXPHOS) 8 . It has also been indicated that incubation of isolated mitochondria from rats with high concentrations of propofol (100 to 400 μM) resulted in a strong inhibition of the activity of complex I and, to a lesser degree, of complexes II and III 9 . Other reports have also demonstrated a reduction of complex IV activity in skeletal muscles, which led to a hypothesis that a propofol metabolite causes a disruption of the respiratory chain 10 . However, the precise molecular mechanisms, including the relationship between mitochondrial defects and metabolic reprogramming in the pathophysiology of PRIS, are largely unknown as yet. To investigate the underlying cellular and molecular mechanisms of PRIS, we investigated the effects of propofol on cell life and death, oxygen metabolism, and mitochondrial function using cells of various origins, including transmitochondrial cybrid cells harboring a mitochondrial DNA defect and mutations. We demonstrated that clinically relevant concentrations of propofol, used within a clinically relevant exposure time, suppressed the mitochondrial function, caused the generation of reactive oxygen species (ROS), and induced the metabolic reprogramming 11 , from OXPHOS to glycolysis, by targeting complexes I and III of mitochondria. The data also indicated that a predisposition to mitochondrial dysfunction, caused by a genetic mutation or pharmacological suppression of the electron transport chain (ETC) in mitochondria by biguanides such as metformin and phenformin, promoted the cell death and caspase activation induced by propofol. Results Propofol induced cell death and activation of caspases in a concentration‐ and time-dependent manner To determine whether propofol induces the cell death, we examined the concentration‐ and time-response relationship between propofol and cell death. Neuronal SH-SY5Y cells were treated with the indicated concentrations of propofol and for the indicated times. Cells were stained with propidium iodide (PI) and recombinant fluorescein isothiocyanate (FITC)-conjugated annexin V, and the proportion of dead cells was evaluated by flow cytometry. Concentrations of propofol equal to or greater than 50 μM induced the cell death within 6 h ( Figs. 1a and 1b ). Interestingly, 25 μM propofol induced the cell death at a longer incubation, 12 h ( Fig. 1c ). Next, activities of caspase-9 and caspase-3/7 were evaluated. Propofol at concentrations equal to or greater than 50 μM activated caspase-9 ( Fig. 2a ). As in the case of cell death and caspase-9, caspase-3/7 was activated by the treatment with 50 μM or greater propofol concentrations within 6 h. Importantly, 25 μM propofol, which did not induce caspase-3/7 activation within 6 h ( Fig. 2b ), induced the activation at 12 h ( Fig. 2c ). Next, a release of lactate dehydrogenase (LDH) was investigated ( Fig. 2d ). Within 6 h, only propofol at a concentration of 150 μM increased the LDH release. On the other hand, during 12-h incubation, not only 150 μM but also 50 and 100 μM propofol statistically significantly increased the LDH release. Measurement of the mitochondrial membrane voltage (ΔΨm) showed that propofol at concentrations equal to or greater than 50 μM decreased ΔΨm within 6 h ( Fig. 2e ). In addition, at concentrations higher than 50 μM propofol suppressed the cell viability, measured by an MTS [3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2 H -tetrazolium] assay, within 6 h ( Supplementary Fig. 1a ). 2,4-Diisopropylphenol, commonly known as 2,4-propofol, is an isomeric form of propofol, which does not show a hypnotic effect. We tested the effects of 2,4-diisopropylphenol on caspase-3/7 activation in SH-SY5Y cells. Similar to propofol, 2,4-diisopropylphenol induced caspase-3/7 activation within 6 h ( Supplementary Fig. 1b ). Interestingly, 25 μM 2,4-diisopropylphenol could activate caspase-3/7 within 6 h, suggesting that 2,4-diisopropylphenol is more toxic than propofol. Finally, the effects of propofol were investigated in cells of different origins, including mouse myoblast C2C12 cells ( Supplementary Fig. 1c ), human cervical cancer HeLa cells ( Supplementary Fig. 1d ), and Lewis lung carcinoma P29 cells ( Supplementary Fig. 1e ). Similar to its effects on SH-SY5Y cells, 50 and 100 μM propofol but not 12.5 or 25 μM propofol induced caspase-3/7 activation in the other cell lines within 6 h. Download figure Open in new tab Figure 1 Propofol induced cell death in a concentration‐ and time-dependent manner SH-SY5Y cells were exposed to the indicated concentrations (12.5, 25, 50, 100, and 150 μM) of propofol for 6 h (a and b) and 3, 6, and 12 h (c). Cells were harvested, and percentages of cell death were measured by flow cytometry. The ratio of PI-positive and/or annexin V-positive cells [(Q1 + Q2 + Q4)/(Q1 + Q2 + Q3 + Q4)] was used to calculate the percentage of dead cells (a and b) (n = 3). Data presented in (b and c) are expressed as the mean ± SD. Differences between treatment groups were evaluated by one-way ANOVA, followed by Dunnett’s multiple comparison test (b), or by two-way ANOVA, followed by Dunnett’s multiple comparison test (c). * p < 0.05 compared to the control cell population (incubation for 0 h, no treatment). Download figure Open in new tab Figure 2 Propofol induced caspase activation in a concentration‐ and time-dependent manner SH-SY5Y cells were exposed to the indicated concentrations (12.5, 25, 50, 100, and 150 μM) of propofol for 6 h (a and b) and 3, 6, and 12 h (c). Caspase-9 (n = 5) (a) and caspase-3/7 (n = 5) (b and c) activities in each treatment group at different time points.(d) SH-SY5Y cells were exposed to the indicated concentrations (12.5, 25, 50, 100, and 150 μM) of propofol for 6 and 12 h. The levels of LDH activity were assayed in culture supernatants (n = 3). Treatment with lysis buffer served as a control. (e) Average mitochondrial membrane potential (ΔΨm) of untreated cells and cells treated with the indicated concentrations (25, 50, and 100 μM) of propofol (n = 3) for 6 h. Values indicate the ratio [Q2/(Q2 + Q4)] of green JC-1 monomers (527 nm emission) to red aggregates (590 nm emission). Data presented in (a–g) are expressed as the mean ± SD. Differences between treatment groups were evaluated by one-way ANOVA, followed by Dunnett’s multiple comparison test (a, b, and e), or by two-way ANOVA, followed by Dunnett’s multiple comparison test (c and d). * p < 0.05 compared to the control cell population (incubation for 0 h, no treatment). Propofol suppressed oxygen metabolism and induced ROS generation We investigated the effects of propofol on oxygen metabolism and glycolysis in SH-SY5Y cells by assaying OCR and the extracellular acidification rate (ECAR), which is a surrogate index for glycolysis. SH-SY5Y cells were preincubated with the indicated concentrations of propofol for the indicated periods. The mitochondrial OCR was significantly suppressed by the treatment with 50 μM propofol for 6 h ( Figs. 3a and 3b , Supplementary Figs. 2a and 2c–f). Accordingly, the ECAR levels were significantly higher upon the treatment with 50 μM propofol ( Figs. 3c and 3d , Supplementary Fig.2b ). Propofol at a concentration of 25 μM, which exerted no significant effects within 6 h, suppressed OCR ( Fig. 3e ) and enhanced ECAR ( Fig. 3f ) after 12 h of incubation. Thus, the evidence indicates that propofol affects the oxygen metabolism in mitochondria. It has been reported that the disturbance of the mitochondrial ETC leads to the generation of ROS by cells 12 , 13 . ROS generation was observed in SH-S5Y5 cells in response to propofol exposure within 3 and 6 h ( Fig. 3g ). Cell death was suppressed by the treatment with 10 mM N -acetylcysteine ( Fig. 3h ). Download figure Open in new tab Figure 3 Oxygen metabolism and ROS generation in SH-SY5Y cells treated with propofol OCR (a, b, and e) and ECAR (c, d, and f) in SH-SY5Y cells exposed to the indicated concentrations of propofol (12.5, 25, 50, and 100 μM) for 6 h (b and d) or 0, 3, 6, and 12 h (e and f). Data presented in (b, d–f) are expressed as the means ± SD. Differences between treatment groups were evaluated by one-way ANOVA, followed by Dunnett’s multiple comparison test (b and d), or by two-way ANOVA, followed by Dunnett’s multiple comparison test (e and f). (g) ROS production in SH-SY5Y cells exposed to 25, 50, and 100 μM propofol (n = 3) for 3 h. (h) SH-SY5Y cells were exposed to the indicated concentrations (50 and 100 μM) of propofol for 6 h with or without treatment with 10 mM N -acetylcysteine. Cells were harvested, and percentages of cell death were measured by flow cytometry. MFI: median fluorescence intensity; NAC: N -acetylcysteine. * p < 0.05 compared to the control cell population. Involvement of mitochondria in propofol-induced cell death and caspase activation As shown in figure 3 , propofol affected the mitochondrial ETC and intracellular oxygen metabolism in SH-SY5Y cells. To examine the involvement of mitochondria in propofol-induced cell death, we used the P29 cell line and its derivative ρ0P29, which lacks the mitochondrial DNA (mtDNA) 14 - 16 . P29 and ρ0P29 cells were exposed to the indicated concentrations of propofol for 6 h, and then cell death ( Fig. 4a ) and the activity of caspase-3/7 ( Fig. 4b ) were assayed. Both cell death and caspase-3/7 assays indicated that, unlike P29 cells, ρ0P29 cells were completely resistant to 50 μM and partially resistant to 100 μM propofol after 6 h of incubation ( Figs. 4a and 4b ). Together with the results showing that propofol affects the ETC function of mitochondria, this evidence strongly suggests that mitochondria play a critical role and are one of the targets in propofol-induced cell death. Download figure Open in new tab Figure 4 Involvement of functional mitochondria in propofol-induced caspase activation and cell death P29 cells and cells of the ρ0P29 derivative lacking mtDNA were exposed to the indicated concentrations (25 and 50 μM) of propofol for 6 h. (a) Cells were harvested, and percentages of cell death were measured by flow cytometry. The ratio of PI-positive and/or annexin V-positive cells [(Q1 + Q2 + Q4)/(Q1 + Q2 + Q3 + Q4)] was used to calculate the percentage of dead cells ( Supplementary Fig. 1a ) (n = 3). (b) Caspase-3/7 activity in each treatment group (n = 3) at 6 h. Differences between treatment groups were evaluated by one-way ANOVA, followed by Dunnett’s multiple comparison test. * p < 0.05 compared to the control cell population; # p < 0.05 compared to the indicated experimental groups. Effects of propofol on the ETC complex-dependent OCR Next, we examined the oxygen consumption, which depends on the activity of mitochondrial respiratory chain complexes I–IV in membrane-permeabilized and intact cells, using an Extracellular Flux Analyzer™ ( Supplementary Fig. 3 ). OCR traces of mitochondrial respiration were detected using protocol A ( Fig. 5a , Supplementary Fig. 3a ) and protocol B ( Fig. 5b , Supplementary Fig. 3b ). The results indicated that propofol significantly suppressed the complex I‐ and complex III-dependent OCR but not the complex II‐ or complex IV-dependent OCR ( Figs. 5c-5d ). Download figure Open in new tab Figure 5 Effects of propofol on OCR driven by each complex of the mitochondrial electron transport chain Representative OCR traces of mitochondrial respiration using protocol A ( Supplementary Fig. 4a ) and protocol B ( Supplementary Fig. 4b ). Mitochondrial ETC-mediated OCR, driven by complexes I (c), II (d), III (e), and IV (f), were assayed by a flux analyzer-based protocol. SH-SY5Y cells were exposed to 100 and 200 μM propofol for 6 h and subjected to the assay. Differences between treatment groups were evaluated by one-way ANOVA, followed by Dunnett’s multiple comparison test. * p < 0.05 compared to the control cell population. Mitochondrial ETC inhibitors synergistically enhanced propofol toxicity We investigated the effects of several ETC inhibitors on the propofol-induced cell death in SH-SY5Y cells. Cells were exposed to rotenone (100 nM), antimycin A (25 μg/mL), or oligomycin (4 μM), with or without the indicated concentrations of propofol, for 6 h, and cell death was assayed by flow cytometry. Neither 100 nM rotenone, 25 μg/mL antimycin, 4 μM oligomycin, nor 12.5 μM or 25 μM propofol alone induced the cell death within 6 h ( Fig. 6a ). On the other hand, 12.5 and 25 μM propofol induced the cell death in the presence of rotenone, antimycin A, and oligomycin ( Fig. 6a ). Similarly, 12.5 and 25 μM propofol with rotenone, antimycin A, and oligomycin induced the caspase-3/7 activity within 6 h ( Fig. 6b ). The evidence indicates that cooperative inhibition of mitochondria by both propofol and the ETC inhibitors induces cell death at even clinically relevant concentrations of propofol within 6 h. Download figure Open in new tab Figure 6 Synergistic effects of propofol and mitochondrial ETC inhibitors on caspase activity and cell death Levels of caspase-3/7 activity and cell death of SH-SY5Y cells treated with propofol and mitochondrial ETC inhibitors. Cells were treated with 12.5, 25, or 50 μM propofol and with either 100 nM rotenone, 4 μM oligomycin, or 25 μg/mL antimycin A and subjected to (a) a cell death assay and (b) a caspase-3/7 activity assay (n = 3).Percentages of cell death were measured by flow cytometry. The ratio of PI-positive and/or annexin V-positive cells [(Q1 + Q2 + Q4)/(Q1 + Q2 + Q3 + Q4)] was used to calculate the percentage of dead cells ( Supplementary Fig. 1a ) (n = 3). All data are expressed as the means ± SD. * p < 0.05 compared with control cells (no treatment); #p < 0.05 compared with the indicated groups. rot: rotenone; olig: oligomycin; anti: antimycin A. Genetic predisposition to mitochondrial dysfunction increased propofol-induced caspase activation and cell death For further study, we used transmitochondrial cybrids carrying mtDNA with defined pathogenic mutations. mtDNA donors and ρ0P29 cells were used to obtain transmitochondrial cybrids P29mtA11, P29mtB82M, P29mtCOIM, and P29mtΔ ( Tables 2 and 3 ) 16 , and oxygen metabolism profiles of these clones were examined ( Figs. 7a and 7b ). These transmitochondrial cybrids were exposed to the indicated concentrations of propofol for 6 h to investigate the cell death and caspase-3/7 activation. Interestingly, in contrast to parental P29 cells, the cell death flow cytometry assay indicated that 25 μM propofol induced the cell death within 6 h in P29mtA11, P29mtB82M, and P29mtCOIM cells ( Fig. 7c ). Caspase-3/7 activation was also induced in P29mtA11, P29mtB82M, and P29mtCOIM cells even with 25 μM propofol ( Fig. 7d ). In contrast, P29mtΔ cells, similar to ρ0P29 cells, were completely resistant to 50 μM propofol and partially resistant to 100 μM propofol ( Fig. 7d ). Thus, our experimental results indicated that cells harboring genetic mutations in mtDNA ( ND6 in complex I or COI in complex IV) were more susceptible to propofol. Download figure Open in new tab Figure 7 Effects of propofol on caspase activity and cell death in various transmitochondrial cybrid cells (a) OCR and (b) ECAR of P29, its cybrid cells, and ρ0 cells. (c and d) P29, its cybrid cells, and ρ0 cells were exposed to the indicated concentrations (12.5, 25, and 50 μM) of propofol for 6 h. Cells were harvested, and percentages of cell death were measured by flow cytometry. The ratio of PI-positive and/or annexin V-positive cells [(Q1 + Q2 + Q4)/(Q1 + Q2 + Q3 + Q4)] was used to calculate the percentage of dead cells (n = 3) (c). Caspase-3/7 activity in each treatment group (n = 3) at 6 h (d). (e) P29, P29mtA11 and ρ0 cells were exposed to 25 and 50 μM propofol for 6 h and subjected to ROS assay. (f) P29mtA11 cells were exposed to 25 μM propofol with or without 10 mM NAC for 6 h. Cells were harvested, and percentages of cell death were measured by flow cytometry. Data presented in (a–f) are expressed as the means ± SD. Differences between treatment groups were evaluated by one-way ANOVA, followed by Dunnett’s multiple comparison test (a, b, e and f) or by two-way ANOVA, followed by Dunnett’s multiple comparison test (c and d). * p < 0.05 compared to the control cell population. A11: P29mtA11 cells; B82M: P29mtB82M cells; COIM: P29mtCOIM cells; mtΔ: P29mtΔ cells. ROS generation was investigated in P29 cells, P29mtA11 cells and ρ0P29 cells in response to propofol exposure. Cells were exposed to 25 μM propofol for 6 h. P29mtA11 cells produced more ROS compared to P29 cells. 25 μM propofol, which did not produce ROS in P29 cells, induced generation of more ROS in P29mtA11 cells. In contrast, ρ0P29 cells did not produce ROS even under 50 μM propofol. Accordingly, cell death induced by 25 μM propofol was suppressed by treatment with 10 mM NAC. Pharmacological suppression of mitochondrial ETC increased propofol-induced cell death and caspase activation The biguanides metformin and phenformin are widely used to reduce high blood sugar levels caused by diabetes 17 . Metformin and phenformin have also been shown to suppress complex I of ETC, which is used by cells to generate energy 18 - 22 . To confirm the effect of blockade of ETC, SH-SY5Y cells were pretreated with 2.5–20 mM metformin or 5–15 μM phenformin for 6 h, with or without the indicated concentrations of propofol, and then the cells were tested using the OCR and ECAR assays. Incubation for 6 h with either 2.5 mM metformin or 5 μM phenformin did not affect OCR ( Fig. 8a and Supplementary Fig. 4a ) and ECAR ( Fig. 8b and Supplementary Fig. 4b ) in SH-SY5Y cells. In contrast, not only 25 μM but also 12.5 μM propofol, which did not affect OCR or ECAR alone, significantly decreased OCR ( Fig. 8a ) and increased ECAR ( Fig. 8b ) in the presence of 2.5 mM metformin. Next, we investigated ROS generation upon cell exposure to 25 μM propofol with or without 5 mM metformin. Exposure to 25 μM propofol did not induce ROS generation without metformin. With metformin, however, 25 μM propofol induced ROS generation ( Fig. 8c ). Then, SH-SY5Y cells were tested for cell death ( Fig. 8d ) and caspase 3/7 activation ( Fig. 8e ) after treatment with 5 mM metformin or 5 μM phenformin ( Supplementary Figs. 4c and 4d ) for 6 h, with or without the indicated concentrations of propofol. Metformin at 5 mM and phenformin at 5 μM increased the propofol-induced cell death and caspase-3/7 activation. Download figure Open in new tab Figure 8 Synergistic effects of propofol and metformin on caspase activity and cell death (a) OCR and (b) ECAR of SH-SY5Y cells exposed to the indicated concentrations of metformin (2.5, 5, 10, and 20 mM) for 6 h. (c) SH-SY5Y cells were exposed to 25 μM propofol with or without 5 mM metformin, and ROS production was determined (n = 3). (d and e) SH-SY5Y cells were exposed to the indicated concentrations (12.5, 25, 50, and 100 μM) of propofol with or without 5 mM metformin for 6 h. (d) Cells were harvested, and percentages of cell death were measured by flow cytometry. The ratio of PI-positive and/or annexin V-positive cells [(Q1 + Q2 + Q4)/(Q1 + Q2 + Q3 + Q4)] was used to calculate the percentage of dead cells (n = 3). (e) Caspase-3/7 activity in each treatment group (n = 3). Data presented in (a–e) are expressed as the means ± SD. Differences between treatment groups were evaluated by one-way ANOVA, followed by Dunnett’s multiple comparison test (a, b, and c), or by two-way ANOVA, followed by Dunnett’s multiple comparison test (d and e). * p < 0.05 compared to the control cell population; # p < 0.05 compared to the indicated experimental groups. Discussion In this study, we demonstrated for the first time that propofol at clinically relevant concentrations and within clinically relevant incubation times altered the oxygen metabolism by targeting mitochondrial complexes I and III and induced a cellular metabolic switch, from OXPHOS to glycolysis, and generation of ROS. The suppression of mitochondria elicited the cell death in cell lines of various origins, including transmitochondrial cybrids carrying mtDNA with defined pathogenic mutations. The concentrations of propofol tested in this study varied from 12.5 to 150 μM. It has been reported that the plasma concentrations of propofol during anesthesia and sedation are from 2 μg/mL (11 μM) up to 5 μg/mL (27.5 μM) 23 . The concentration of propofol was also measured in tissue samples from rats treated with propofol at a dose of 20 mg/kg/h, and the study indicated that the tissue concentration of propofol could reach 200 μM under certain conditions 24 . Thus, the evidence indicates that the concentrations of propofol used in this study were clinically relevant. The duration of exposure used in this study ranged from 3 to 12 h, which was also within the clinically relevant period of exposure. Although predictive factors for PRIS have not been established, there is a consensus that exposure to high doses of propofol for prolonged periods is the most critical risk factor for PRIS 1 , 2 , 6 , 25 . In this study, we demonstrated that propofol at concentrations equal to or greater than 50 μM but not at or below 25 μM showed cell toxicity within 6 h. On the other hand, even 25 μM propofol significantly increased the cell death and caspase activity during incubation for 12 h ( Figs. 1b and 1f ). In addition, propofol at 50 and 25 μM significantly suppressed OCR and increased ECAR within 6 and 12 h, respectively. At 100 μM, propofol inhibited both OCR and ECAR, which could be due to a rapid cell death induced by 100 μM propofol. The evidence indicates that propofol at clinically relevant concentrations suppresses OXPHOS and induces a metabolic switch, from OXPHOS to glycolysis, resulting in the enhancement of lactate production. This metabolic conversion can be one of the most critical cellular mechanisms of lactic acidosis observed in PRIS 1 , 2 . As clearly demonstrated in this and other studies 26 - 29 , propofol induces cell death. In this study, we demonstrated that not only neuronal SH-SY5Y cells but also cells of other origins, such as C2C12 muscle cells, HeLa cervical carcinoma cells, and P29 lung cancer cells, are susceptible to propofol. Although there is no consensus on the target organs or tissues in the context of PRIS, our results suggest that propofol can exert toxicity against a wide range of tissues. There are at least two known modes of cell death, apoptosis and necrosis. Apoptosis is a strictly regulated or programmed process involving the activation of specific proteases, which are responsible for organized removal of damaged cells 30 . Necrosis is a pathophysiologically different form of cell death, which is accompanied by a loss of Δψm and impairment of OXPHOS. In this study, propofol treatment promoted the caspase-9 and caspase-3/7 activation. These data indicate that propofol at least activates the apoptosis pathway. Meanwhile, the flow cytometry analysis demonstrated that treatment of cells with propofol concentrations greater than 50 μM for 12 h also resulted in an LDH release ( Fig. 1d ) and increased the PI + /annexin V − cell population, in addition to the PI + /annexin V + cell population ( Fig. 1a ). These data strongly suggest that propofol elicits both types of cell death, resulting in cell membrane injury at a 50 μM concentration and 12-h exposure. As demonstrated in figures 2 and 3 , propofol inhibited the mitochondrial oxygen consumption in a time‐ and concentration-dependent manner. In this study, we measured OCR, which is dependent on the activity of each ETC complex in intact cells, using an extracellular flux analyzer. Several studies have used isolated mitochondria, and some have suggested that propofol inhibits the enzymatic activity of complexes II and IV 31 . However, our study using intact cells indicated that propofol does not affect the complex II‐ or complex IV-dependent OCR. Another finding was that propofol induced ROS generation ( Fig. 2g ). While the precise origin of ROS remains unclear, our findings strongly suggest that mitochondria play a critical role in this process. ρ0P29 cells and P29mtΔ cells did not produce ROS in response to propofol treatment (data not shown). It has been demonstrated that the disturbance of mitochondria leads to increased production of ROS by ETC 12 , 13 , 32 - 34 . In fact, cybrid cells such as P29mtA11 and P29mtB82M with a mutation in the mitochondrial NADH dehydrogenase subunit 6 ( ND6 ) gene generate ROS under 20% O 2 conditions 16 , 35 , which implies that propofol-mediated cell death is dependent on ROS from mitochondria. Mitochondrial disease, once thought to be a rare clinical entity, is now recognized as an important cause of a wide range of neurological, cardiac, muscle, and endocrine disorders 4 , 5 . It has been reported that complex I is uniquely sensitive to many anesthetic agents 36 . To approach the issue, we used cells harboring mtDNA mutations 15 , 16 , 37 . As shown in figures 3 and 6 , ρ0P29 cells, which do not have mtDNA, are resistant to 50 μM propofol. P29mtΔ cells, carrying the nuclear genome of P29 cells and the mitochondrial genome of ΔmtDNA4696, with a 4,696-bp deletion 15 , are also resistant to propofol. In contrast, P29mtA11 cells with the G13997A mutation in mtDNA and P29mtB82M cells with the 13885insC insertion, both of which affect the NADH-ubiquinone oxidoreductase chain 6 (ND6) protein of complex I, are more sensitive to propofol than P29 cells are. Propofol at 12.5 and 25 μM induced the cell death and caspase-3/7 activation in both mutant cell lines within 6 h ( Fig. 6c ). This is consistent with the evidence showing that 25 μM propofol induced the cell death in the presence of a sublethal concentration of rotenone. P29mtCOIM cells with a missense mutation in cytochrome c oxidase I (COI) are as sensitive to propofol as P29mtA11 and P29mtB82M cells. This is also consistent with the result showing that propofol induced the cell death in the presence of oligomycin ( Fig. 5 ). This evidence indicates that cells with mutations in mtDNA are more sensitive to propofol. In addition to the genetic mutations affecting mitochondrial function, pharmacological disturbance of mitochondria with biguanides, including metformin and phenformin, also synergistically suppressed the mitochondrial function and induced caspase activation and cell death. The primary effect of biguanides is generally thought to be the inhibition of respiratory complex I (NADH:ubiquinone oxidoreductase), which leads to the energy stress by decreasing ATP synthesis by OXPHOS 18 , 19 , 38 , 39 . In addition, a number of studies have indicated that biguanides also affect complexes II, III, and IV and F1F0-ATPase 20 , 40 . Metformin and phenformin are used as antidiabetic drugs and are associated with lactic acidosis 20 , 41 , 42 . As demonstrated in this study, both metformin and phenformin increased ECAR and decreased OCR, which indicates a metabolic switch. In addition to biguanides, other clinically used drugs, including chloramphenicol 43 , 44 , aspirin 45 , 46 , statins 18 , and local anesthetics 47 , also inhibit mitochondrial functions. Our results indicate that pre-exposure to mitochondrial inhibitors may increase the toxicity of propofol. There are some limitations in this study. Thus, we only used cultured cells. Because PRIS is a systemic syndrome, using cells is not sufficient to completely understand the pathophysiology of PRIS. An in vivo investigation using experimental animals may be warranted to confirm our findings. In conclusion, clinically relevant concentrations of propofol used within a clinically relevant exposure time suppressed the mitochondrial function, caused the generation of ROS, and induced metabolic reprogramming, from OXPHOS to glycolysis, by targeting complexes I and III of mitochondria. Our data also indicated that a predisposition to mitochondrial dysfunction, caused by genetic mutations and pharmacological suppression of ETC by biguanides such as metformin and phenformin, promoted the cell death and caspase activation by propofol. The process is likely to constitute the molecular basis of PRIS. Materials and Methods Reagents Propofol (2,6-diisopropylphenol), 2,4-diisopropylphenol, and dimethyl sulfoxide were obtained from Sigma–Aldrich (St. Louis, MO, USA). Rotenone, oligomycin, and antimycin A were obtained from Abcam, Inc. (Cambridge, MA, USA). The Key Resources Table is provided as Table 1 . View this table: View inline View popup Download powerpoint Table 1 Key Resources Table Cell lines and cell culture Established cell lines derived from human neuroblastoma SH-SY5Y cells and cervical carcinoma HeLa cells were maintained in RPMI 1640 medium supplemented with 10% fetal bovine serum, 100 U/mL penicillin, and 0.1 mg/mL streptomycin. The mouse cell lines and their characteristics are listed in Tables 2 and 3 . P29 cells originated from Lewis lung carcinoma (C57BL/6 mouse strain), and B82 cells are fibrosarcoma cells derived from the L929 fibroblast cell line (C3H/An mouse strain) 15 , 16 , 48 . Parental P29 cells, ρ0 cells, and the transmitochondrial cybrids were grown in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with pyruvate (0.1 mg/mL), uridine (50 mg/mL), and 10% fetal bovine serum. We isolated ρ0 cells by treating parental P29 cells with 1.5 mg/mL ditercalinium, an antitumor bis -intercalating agent. Enucleated cells of mtDNA donors were prepared by pretreatment with cytochalasin B (10 μg/mL) for 2 min, followed by centrifugation at 7,500 × g for 10 min 16 . The resultant cytoplasts were fused with ρ0 cells using polyethylene glycol. The transmitochondrial cybrids (see Table 3 ) were isolated in a selection medium that allows exclusive growth of the cybrids 16 . View this table: View inline View popup Download powerpoint Table 2 Identification of pathogenic mutations in mtDNA sequences View this table: View inline View popup Download powerpoint Table 3 Genetic characteristics of parent cells and their transmitochondrial cybrids Cell growth MTS assay Cell growth was assessed using a CellTiter 96 AQueous One Solution Cell Proliferation Assay™ with MTS (Promega, Madison, WI, USA) 47 , 49 . Briefly, cells were seeded into 96-well plates (2 × 10 4 cells/well) and cultured overnight. On the following day, the cells were treated with the indicated concentrations of the appropriate drug(s) for varying times. After the treatment, 20 μL of the CellTiter 96 AQueous One Solution™ reagent was added to each well, the plates were incubated at 37 °C for 1 h, and the absorbance of each sample was measured using an iMark™ microplate reader (Bio-Rad, Hercules, CA, USA) at a wavelength of 490 nm. Cell viability was calculated by comparing the absorbance of treated cells with that of the control cells incubated without drugs, which was defined as 100%. All samples were tested in triplicate or quadruplicate in each experiment. Caspase-3/7 and caspase-9 activity assays Activity levels of caspase-3/7 and caspase-9 were assessed using an Apo-ONE™ homogeneous caspase-3/7 assay kit (Promega) and a Caspase-Glo™ 9 assay kit (Promega), respectively, according to the manufacturer’s protocols 47 , 49 . Briefly, cells were seeded into 96-well plates (2 × 10 4 cells/well) and incubated overnight. On the following day, the cells were treated with the indicated concentrations of the appropriate drug(s) for varying times. After the treatment, 100 μL of the Apo-ONE™ caspase-3/7 reagent was added to each well. The plates were incubated at room temperature for 1 h, and the luminescence of each well was measured using an EnSpire™ multimode plate reader (PerkinElmer, Waltham, MA, USA). Caspase activity was calculated by comparing the levels of luminescence of the treated cells with that of the control cell population incubated without drugs, which was defined as 100%. The assays were performed in triplicate at least twice. Data were expressed as the mean ± standard deviation (SD). Analysis of cell death The protocol was described previously 47 , 49 . Briefly, the levels of cell apoptosis were measured using an Annexin V–FITC apoptosis detection kit (BioVision, Milpitas, CA, USA), according to the manufacturer’s instructions. For the analysis, cells were seeded into 6-well plates (3 × 10 5 cells/well) and incubated overnight. On the following day, the cells were treated with the indicated concentrations of the appropriate drug(s) for varying times and harvested by centrifugation at 1,200 rpm for 3 min. The cell pellets were resuspended in a mixture comprised of 500 μL of binding buffer, 5 μL of annexin V–FITC, and 5 μL of PI (50 μg/mL). The suspensions were incubated for 5 min at room temperature in the dark and analyzed using a FACSCalibur flow cytometer (BD Biosciences, San Jose, CA, USA) equipped with the CellQuest Pro™ software. The data were evaluated using the FlowJo™ version 9.9.4 software (TreeStar, Ashland, OR, USA), then exported to Excel spreadsheets, and subsequently analyzed using the statistical application GraphPad™ Prism 7. Determination of mitochondrial membrane potential The mitochondrial membrane potential (ΔΨm) was determined by flow cytometry using a MitoPT™ JC-1 assay kit (ImmunoChemistry Technologies, Bloomington, MN, USA), according to the manufacturer’s instructions 49 . For the analysis, cells were seeded into 6-well plates (3 × 10 5 cells/well) and cultured overnight. On the following day, the cells were treated with the indicated concentrations of the appropriate drug(s) for varying times and then pelleted by centrifugation at 1,200 rpm for 3 min. The cells were then resuspended in JC-1, incubated at 37 °C for 15 min in the dark, and collected by centrifugation at 1,200 rpm for 3 min. The cell pellets were resuspended in 500 μL of assay buffer. The samples were subsequently analyzed using a FACSCalibur flow cytometer (BD Biosciences) equipped with the CellQuest Pro™ software for the detection of red JC-1 aggregates (590 nm emission) or green JC-1 monomers (527 nm emission). The data were evaluated using the FlowJo version 7.6.3 software (TreeStar), then exported to Excel spreadsheets, and subsequently analyzed using the statistical application GraphPad™ Prism 7. LDH-based cytotoxicity assay Cytotoxicity was measured using a CytoTox-ONE™ kit (Promega) as described previously 47 , 49 . Briefly, cells were cultured overnight in 96-well plates (2 × 10 4 cells/well) and treated with the indicated drug(s) for varying lengths of time. Twenty microliters of the CytoTox-ONE™ reagent was added to each well, and the plates were incubated at 22 °C for 10 min. The reaction was terminated by adding 50 μL of the stop solution, and the fluorescence was recorded at an excitation wavelength of 560 nm and an emission wavelength of 590 nm using an EnSpire™ multimode plate reader (PerkinElmer). The percentage of cell death was determined by comparing the release of LDH (fluorescence value) in each treatment group with that of the positive control treated with the lysis solution, which was defined as 100%. Meanwhile, the level of LDH release from untreated cells (negative control) was defined as 0%. Each sample was assayed in triplicate. Measurement of ROS generation ROS generation was detected with 2′,7′-dichlorofluorescin diacetate (DCFH-DA) (Molecular Probes, Eugene, OR, USA). Briefly, cells cultured in 35-mm-diameter glass-bottom culture dishes (MatTek, Ashland, MA, USA) were incubated with 10 μM DCFH-DA for 10 min at 37 °C in a serum-free DMEM, then washed twice with Dulbecco’s phosphate-buffered saline, and analyzed using a flow cytometer (Beckton Dickinson). The mean fluorescence intensity was analyzed using the CellQuest software (Becton Dickinson). Cellular oxygen consumption and extracellular acidification measurement Cellular OCR and ECAR were determined with the XF Cell Mito Stress Test™ and XF Glycolysis Stress Test™, respectively, using an XFp Extracellular Flux Analyzer™ (Seahorse Bioscience, USA) 47 . Cells (2 × 10 5 cells/well) were seeded into an XFp cell culture microplate, and OCR was assessed in glucose-containing XF base medium according to the manufacturer’s instructions. The sensor cartridge for the XFp analyzer was hydrated in a 37 °C non-CO 2 incubator on the day before the experiment. For the OCR assay, injection port A on the sensor cartridge was loaded with 1.5 μM oligomycin (complex V inhibitor), port B was loaded with 2 μM carbonyl cyanide-4-(trifluoromethoxy) phenylhydrazone (FCCP), and port C was loaded with 0.5 μM rotenone/antimycin A (inhibitors of complex I and complex III). During the sensor calibration, cells were incubated in a 37 °C non-CO 2 incubator in 180 μL of assay medium (XF base medium with 5.5 mM glucose, 1 mM pyruvate, and 2 mM L-glutamine, pH 7.4). The plate was immediately placed into the calibrated XFp extracellular flux analyzer for the Mito Stress test ( Supplementary Fig. 3a ). The assay parameters were calculated as follows: OCR (basal) = (last rate measurement before oligomycin injection) − (minimum rate measurement after rotenone/antimycin-A injection); OCR (maximal) = (maximum rate measurement after FCCP injection) − (minimum rate measurement after rotenone/antimycin A injection); OCR (non-mitochondrial respiration) = (minimum rate measurement after rotenone/antimycin A injection); proton leak = (minimum rate measurement after oligomycin injection) – (non-mitochondrial respiration). For the ECAR assay, injection port A on the sensor cartridge was loaded with 10 mM glucose. During the sensor calibration, cells were incubated in a 37 °C non-CO 2 incubator in 180 μL of assay medium (XF base medium with 2 mM L-glutamine, pH 7.4). The plate was immediately placed into the calibrated XFp Extracellular Flux Analyzer™ for the Glycolysis Stress test ( Supplementary Fig. 3b ). Oligomycin (1 μM) and 50 mM 2-deoxy-D-glucose were loaded for the measurement. ECAR was normalized for the total protein/well and calculated as follows: ECAR (glycolysis) = (maximum rate measurement after glucose injection) − (last rate measurement before glucose injection). Measurement of oxygen consumption in permeabilized cells The activity of individual respiratory chain complexes was evaluated in permeabilized cells 50 , 51 . Briefly, cells were washed with MAS buffer (220 mM mannitol, 70 mM sucrose, 10 mM KH 2 PO 4 , 5 mM MgCl 2 , 2 mM HEPES, 1 mM EGTA, 0.2% fatty acid-free bovine albumin, adjusted to pH 7.2 with KOH), and the medium was replaced with MAS buffer supplemented with 10 mM pyruvate, 1 mM malate, 4 mM ADP, and 1 nM plasma membrane permeabilizer ™ . The cells were then loaded into the XFp analyzer to measure respiration rates using cycles of 30 s mixing/30 s waiting/2 min measurement. After measurement of pyruvate-driven respiration, rotenone (final concentration: 2 μM) was injected through port A to halt the complex I-mediated respiratory activity. Next, succinate (10 mM) was injected through port B to donate electrons at complex II, bypassing complex I inhibition. Addition of antimycin A (2 μM) via port C inhibited complex III, and N,N,N′,N′-tetramethyl- p -phenylenediamine (0.1 mM), combined with ascorbate (10 mM), was subsequently injected through port D to measure complex IV activity. As an alternative approach, cells were initially provided with pyruvate to measure complex I activity. After injection of rotenone, duroquinol was injected to stimulate complex III-mediated respiration. Statistical analysis All experiments were repeated at least twice, and each sample was evaluated in triplicate. Representative data, expressed as the mean ± SD, are shown. Differences between treatment groups were evaluated by one-way analysis of variance (ANOVA) or two-way ANOVA, followed by Dunnett’s multiple comparison test or t -test using GraphPad Prism 7. P -values of < 0.05 were considered statistically significant. Author contributions statement C.S. performed the experiments, analyzed the data, and co-wrote the manuscript. A.O., M.T., M.K., H.S., T.U., Y.M., and T.A. contributed to the data analysis and discussion. J.H. and K.T. provided experimental materials and contributed to the discussion. K.H. designed and supervised the study, analyzed the data, and co-wrote the manuscript. Additional information Competing financial interests The authors declare no competing financial interests. Acknowledgments This work was supported by the Japan Society for the Promotion of Science KAKENHI, grants #26670693 and #24592336 to K.H., #25462457 to K.N., and #15K15577 to T.A, a research grant from Katano Kai to K.H. and A.O., research grant B from Kansai Medical University to A.O., and a KMU consortium grant from Kansai Medical University to K.H. We would like to thank Editage ( www.editage.jp ) for English language editing. References ↵ Kam , P. C. & Cardone , D. Propofol infusion syndrome . Anaesthesia 62 , 690 - 701 , doi: 10.1111/j.1365-2044.2007.05055.x ( 2007 ). OpenUrl CrossRef PubMed Web of Science ↵ Bray , R. J. Propofol infusion syndrome in children . Paediatr Anaesth 8 , 491 - 499 ( 1998 ). OpenUrl CrossRef PubMed Web of Science ↵ Parke , T. J. et al. Metabolic acidosis and fatal myocardial failure after propofol infusion in children: five case reports . BMJ 305 , 613 - 616 ( 1992 ). OpenUrl Abstract / FREE Full Text ↵ Finsterer , J. & Frank , M. 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Share Propofol induces a metabolic switch to glycolysis and cell death in a mitochondrial electron transport chain-dependent manner Chisato Sumi , Akihisa Okamoto , Hiromasa Tanaka , Kenichiro Nishi , Munenori Kusunoki , Tomohiro Shoji , Takeo Uba , Yoshiyuki Matsuo , Takehiko Adachi , Jun-Ichi Hayashi , Keizo Takenaga , Kiichi Hirota bioRxiv 181933; doi: https://doi.org/10.1101/181933 Share This Article: Copy Citation Tools Propofol induces a metabolic switch to glycolysis and cell death in a mitochondrial electron transport chain-dependent manner Chisato Sumi , Akihisa Okamoto , Hiromasa Tanaka , Kenichiro Nishi , Munenori Kusunoki , Tomohiro Shoji , Takeo Uba , Yoshiyuki Matsuo , Takehiko Adachi , Jun-Ichi Hayashi , Keizo Takenaga , Kiichi Hirota bioRxiv 181933; doi: https://doi.org/10.1101/181933 Citation Manager Formats BibTeX Bookends EasyBib EndNote (tagged) EndNote 8 (xml) Medlars Mendeley Papers RefWorks Tagged Ref Manager RIS Zotero Tweet Widget Facebook Like Google Plus One Subject Area Pharmacology and Toxicology Subject Areas All Articles Animal Behavior and Cognition (7820) Biochemistry (18262) Bioengineering (14430) Bioinformatics (43195) Biophysics (22023) Cancer Biology (19140) Cell Biology (26220) Clinical Trials (138) Developmental Biology (13669) Ecology (20452) Epidemiology (2067) Evolutionary Biology (24905) Genetics (15880) Genomics (23046) Immunology (18246) Microbiology (41426) Molecular Biology (17585) Neuroscience (91077) Paleontology (680) Pathology (2914) Pharmacology and Toxicology (4961) Physiology (7899) Plant Biology (15550) Scientific Communication and Education (2072) Synthetic Biology (4440) Systems Biology (10026) Zoology (2323) window.__CF$cv$params={r:'a1f74403ade1e726',t:'MTc4NDc3Mzg1Mg==',u:'019f8ccfbf357841abe076612299b739',ut:'jDpXT1pzM9Yjo4ZqyvmH_m6tfoI06l9RQXOuHsmPDyw-1784773852-1.2.1.1-rnFfl4QfEcZ8wRBX2ihHaJneNnybov17gy5skv5RzdLrZS8EBqpII4z49OsAj6tPyoo8EN3Cq13FqSNKZfo1dqve3bADQFlSxDbl4VrxM98',i:60};(function(){if(!document.body)return;var s=document.createElement('script');s.src='/cdn-cgi/challenge-platform/scripts/precursor/main.js';document.head.appendChild(s);})();