Synergistic Effects of Commonly Used Intensive Care Unit Drugs and High Temperature on Skeletal Muscle at the Cellular and Tissue Levels

Abstract

Background:

Several drugs are commonly administered to patients with high body temperature in intensive care units. However, previous in vitro studies have investigated only the independent effects of high temperatures or drugs on various cultured cells. This study explored the hypothesis that pharmacologic treatment with representative intensive care unit drugs induces lethal effects on cultured skeletal muscle and engineered muscle tissue at high temperatures.

Methods:

Human skeletal muscle cultures were treated with the representative drugs propofol, dexmedetomidine, and acetaminophen at 37°, 39°, and 41°C for various exposure times. To investigate the effects of the drug treatments, cell viability, lactate dehydrogenase activity, caspase activity, and endoplasmic reticulum (ER) stress were analyzed. Conformational changes in myotubes and functional changes in contractile muscle tissue were also assessed. All experiments were repeated at least three times.

Results:

Dexmedetomidine and acetaminophen had no observable adverse effects at high temperatures, whereas propofol treatment at greater than 200 μM resulted in increased lactate dehydrogenase activity and myotube detachment. Furthermore, this cellular injury was associated with intracellular calcium overload and upregulation of the ER stress–related genes CHOP, GRP78/Bip, and GRP94. Propofol treatment also decreased the contractile ability of muscle tissues at 39°C (vs. 37°C propofol; 95% CI, 30.72 to 114.87%; P < 0.001). Additionally, although tauroursodeoxycholic acid, an ER stress inhibitor, alleviated the increase in caspase-3/7 activity at 39°C (95% CI, 38.10 to 145.22%; P < 0.001) and mitigated myotube detachment, it did not result in notable functional improvement in muscle contraction.

Conclusions:

These results demonstrate that propofol had harmful effects on skeletal muscle cells and tissues at high temperatures in vitro. As these synergistic effects were closely associated with ER stress, tauroursodeoxycholic acid could mitigate propofol-induced apoptosis at high temperatures. These findings could help improve drug treatment for patients, including their functional prognosis in the clinical setting.

The in vitro effects of propofol, dexmedetomidine, and acetaminophen (drugs commonly used in intensive care unit patients) on myoblasts and myotubules were determined at 37°, 39°, and 41°C. At high temperatures, propofol had synergistic cytotoxic effects, including those related to conformational and functional impairments, on human skeletal muscle cells and tissues. Dexmedetomidine and acetaminophen did not have observable adverse cellular effects.

Editor’s Perspective

What We Already Know about This Topic

• Mechanisms underlying intensive care unit–acquired weakness are unclear

• Drug-related apoptosis has been postulated to be a cause of intensive care unit–acquired weakness

• High temperatures can induce apoptosis

What This Article Tells Us That Is New

• The in vitro effects of propofol, dexmedetomidine, and acetaminophen (drugs commonly used in intensive care unit patients) on myoblasts and myotubules were determined at 37°, 39°, and 41°C

• At high temperatures, propofol had synergistic cytotoxic effects, including those related to conformational and functional impairments, on human skeletal muscle cells and tissues

• Dexmedetomidine and acetaminophen did not have observable adverse cellular effects

Body temperature is one of the most important vital signs for patients, including those admitted to the intensive care unit (ICU). Many physiologic and behavioral mechanisms are involved in the regulation of body temperature; of these, physiologic effectors are the naturally occurring, primarily autonomic responses that produce or dissipate heat in a homeostatic manner. Skeletal muscle, which generally comprises greater than 40% of body mass, plays a central role in temperature homeostasis and can be recruited to generate heat through both shivering and nonshivering thermogenesis. Although some in vitro studies have reported that heat stress itself can induce muscle hypertrophy,¹ overall, skeletal muscle mass is strongly associated with the regulation of body temperature.

High body temperature, defined clinically as body temperature above 38.3°C, is frequently observed in the ICU (in 26 to 88% of cases)²; furthermore, approximately 60% of patients requiring mechanical ventilation have high body temperature.³ However, the management of high body temperature above 39.5°C, which can be due to infectious or noninfectious conditions, in ICU patients remains debatable.⁴ Acetaminophen is commonly administered as an antipyretic to patients who decidedly require treatment,⁵ and patients on ventilation require the continuous infusion of sedatives and analgesic drugs. Acetaminophen and dexmedetomidine are also frequently used as adjuncts to limit opioid use in the ICU,⁶ whereas nonsteroidal anti-inflammatory drugs are not commonly used because of concerns regarding adverse events. Furthermore, the Society of Critical Care Medicine (Mount Prospect, Illinois) recommends continuous sedation with either dexmedetomidine or propofol at low concentrations for adults on mechanical ventilation.⁷ Thus, ICU patients routinely receive numerous drugs, especially propofol, dexmedetomidine, and acetaminophen, at high body temperatures.

ICU-acquired weakness, including critical illness myopathy and propofol infusion syndrome, are of significant concern in the context of ICU patient management. However, the mechanisms underlying these conditions remain unclear and may vary considerably.^8,9 Skeletal muscle cell apoptosis in response to certain drugs has been proposed as one of the mechanisms underlying critical illness myopathy and propofol infusion syndrome.^10,11 Additionally, high temperatures themselves are known to induce apoptosis as a hyperthermia treatment—at least 42.5°C to kill tumor cells.¹² However, the synergistic effects of drugs and high temperature on muscles have not been investigated in vitro.

We hypothesized that pharmacologic treatment with some commonly used drugs triggers alterations in muscles at high temperatures, potentially leading to lethal effects at the cellular and tissue levels. Therefore, this study was conducted to investigate the in vitro effects of the representative drugs propofol, dexmedetomidine, and acetaminophen on skeletal muscle cells—specifically, myoblasts and myotubes—at 37°, 39°, and 41°C. We also generated muscle tissues using our recently developed tissue engineering technique to investigate the effects of these drugs and high temperatures on the functionality of muscle tissue, demonstrating how such models can facilitate the assessment of organizational-level toxicity as a viable alternative to animal experiments in preclinical studies.¹³

Material and Methods

Reagents

Propofol (2,6-diisopropylphenol), dexmedetomidine, acetaminophen, and dimethyl sulfoxide (DMSO) were purchased from Sigma-Aldrich (USA). TUDCA and 1,2-bis (2-aminophenoxy) ethane-N,N,N',N'-tetraacetic acid tetrakis (acetoxymethyl ester) (BAPTA-AM) were purchased from Funakoshi Co. Ltd. (Japan) and Abcam (United Kingdom), respectively.

Myoblast Culture

Human skeletal muscle myoblasts (Lonza, USA) were cultured at 37°C in a humidified, 5% CO₂ atmosphere. The myoblasts at passages 2 to 4 were seeded at a density of 4 × 10⁴ cells/cm² and were cultured in skeletal muscle growth medium 2 (SkGM-2; Lonza) for 2 days. This myoblast culture process is hereafter referred to as phase (i).

Induction of Aligned Myotube Formation Using a Micropatterned Culture Substrate

A micropatterned culture substrate was fabricated on normal culture dishes as previously described to induce the formation of biomimetic, aligned myotube structures.¹⁴ Briefly, striped polyacrylamide micropatterns were fabricated on culture dishes through photoinduced polymerization, and human myoblasts were then seeded at a density of 5 × 10⁴ cells/cm² on the micropatterned surface (seeding area, 15 × 15 mm²) and cultured in SkGM-2 for 2 days. After reaching 70 to 90% confluence, the cells were cultured in differentiation medium (Dulbecco’s modified Eagle’s medium with 2% horse serum and 1% penicillin/streptomycin) for 5 to 7 days. This myotube culture process is hereafter referred to as phase (ii).

Production of Contractile Muscle Tissues

To determine the effects of the drugs on muscle tissue functionality, contractile human muscle tissue was generated as previously described.¹⁴ First, micropatterning was performed on thermoresponsive culture dishes (UpCell; CellSeed, Inc., Japan), as on normal culture dishes in phase (ii), to regulate the aligned structure of myofibers in muscle tissues.¹⁴ Human myoblasts were then seeded onto the patterned culture substrate at a density of 12 to 15 × 10⁴ cells/cm² (seeding area, 15 × 15 mm²) and cultured in SkGM-2 for 1 or 2 days. When the cells reached 100% confluence, the medium was changed to differentiation medium supplemented with 5 µM TGF-β RI kinase inhibitor VI (SB-431542; Sigma-Aldrich) for 1 or 2 days. Subsequently, a mixture of bovine plasma–derived fibrinogen (20 mg/ml, 2 ml; Sigma-Aldrich) and thrombin (20 U/ml, 500 µl; Sigma-Aldrich), Matrigel (500 µl; BD Biosciences, USA), and CaCl₂ solution (8 mM, 1 ml) was applied onto the confluent cells at a volume of 900 µl per dish.¹⁴ A square silicon ring was incorporated into the mixture to inhibit gel shrinkage during the maturation of the myoblasts into contractile myofibers, and the dishes were then incubated at 37°C for 30 min to allow gel formation. Finally, differentiation medium containing the anti-fibrinolytic agent aprotinin (20 µg/ml; Sigma-Aldrich) was added to the dish. After 1 week of culture in differentiation medium, the aligned cells were incubated at 20°C for 30 min to detach them from the surface. The muscle tissue on the gel was then placed upside down in the medium on a 6-well plate for electrical pulse stimulation (EPS) applications.

Two carbon electrodes (C-Dish; IonOptix, USA) were immersed in the culture medium, and EPS was applied using an electrical pulse generator (IonOptix) to induce muscle tissue culture by sequential stimulation (voltage, 10 V; frequency, 1 Hz; duration, 3 ms) for 1 h, followed by a 3-h incubation period without stimulation. This procedure started 5 to 7 days after the induction of EPS to promote muscle tissue maturation.¹⁴ The culture medium was replaced with fresh medium every 1 or 2 days. This contractile muscle tissue culture process is hereafter referred to as phase (iii).

Drug Treatment at High Temperature

Samples prepared in phases (i), (ii), and (iii) were cultured in medium containing the appropriate drugs or reagents in an incubator maintained at 37°, 39°, or 41°C for 1 to 48 h. All drugs or reagents, including TUDCA and BAPTA-AM, were administered concurrently. The culture medium was changed every day. Propofol, acetaminophen, TUDCA, and BAPTA-AM were diluted in DMSO, and dexmedetomidine was diluted in Milli-Q water. The same volume of DMSO (0.1% v/v) or Milli-Q water (1% v/v) without reagents was added to one well or dish as the vehicle control sample. The concentrations of the drugs used in each experiment are specified in the corresponding figure, and the treatment exposure time is specified in the figure legend. The concentrations of propofol, acetaminophen, and dexmedetomidine were determined based on previous in vitro and in vivo experiments.^11,15–20

Lactate Dehydrogenase–based Cytotoxicity Assay

Lactate dehydrogenase (LDH) activity, as a measure of extracellular LDH leakage, was assessed using the cytotoxicity LDH assay kit WST (Dojindo Laboratories, Japan) according to the manufacturer’s instructions and a previously reported method.²¹ After drug treatment for 12 h at 37°, 39°, or 41°C, phase (i) samples were assayed in two sets of replicates for each condition: one positive sample and one for the actual assay. Media without cells were used as the blank media for that condition. The positive samples for each condition, including the control group (without drug treatment) and drug treatment groups, were prepared using lysis buffer to release LDH at the maximum from the cells. The absorbance of the medium at a wavelength of 492 nm was measured using a Nivo multimode plate reader. Cytotoxicity was determined using the following equation and normalized to the value for samples without drug treatment (the control group) as 1.0.

$$\text{Relative LDH activity = }\frac{\text{condition sample }-\text{ blank media}}{\text{positive condition sample }-\text{ blank media}}$$

All condition samples, including positive samples, were performed in triplicate. To confirm whether human myoblasts induced cellular damage alone at high temperatures, the initial temperature-only experiments were repeated eight times, followed by subsequent drug treatment experiments, which were repeated five times.

Caspase-3/7, Caspase-9, and Calpain Activity Assays

Caspase-3/7, caspase-9, and calpain activities were assessed using the Caspase-Glo 3/7 assay (Promega, USA), Caspase-Glo 9 assay (Promega), and Calpain-Glo protease assay (Promega) systems, respectively, according to the manufacturer’s instructions. Briefly, after drug treatment at 37°, 39°, or 41°C for 12 or 48 h, 100 µl of the assay reagent was added to each 96-well plate well containing phase (i) samples, and luminescence was measured using a Nivo multimode plate reader, after incubation at room temperature for 30 min or 1 h. The enzyme activities were estimated by comparing the luminescence intensity of the treated cells with that of control untreated cells, which was defined as 100%. All condition samples were performed in triplicate. To confirm whether human myoblasts induced caspase activity alone at high temperatures, the temperature-only experiments were repeated seven times, followed by subsequent drug treatment experiments, which were repeated five or six times.

Terminal Deoxynucleotidyl Transferase dUTP Nick End Labeling (TUNEL) Assay

DNA fragmentation associated with apoptosis was assessed using the Click-iT Plus TUNEL assay kits for in situ apoptosis detection (Thermo Fisher Scientific, USA) according to the manufacturer’s instructions. Briefly, after drug treatment at 37° or 39°C for 24 h, phase (i) cells were fixed with 4% paraformaldehyde in phosphate-buffered saline (PBS) for 15 min and then permeabilized with 0.25% Triton-X 100 (Sigma-Aldrich) for 20 min. Thereafter, the samples were processed for the terminal deoxynucleotidyl transferase and Click-iT Plus reactions, washed with 3% bovine serum albumin in PBS, and incubated with Hoechst 33258 (Dojindo Laboratories) for 5 min at room temperature for nuclear staining. Images were acquired using a fluorescence microscope and analyzed using ImageJ software, version 1.53e. The experiments were repeated three times.

Measurement of Cytosolic and Endoplasmic Reticulum (ER) Calcium Concentrations

As Ca²⁺ homeostasis is associated with ER stress, cytosolic and ER Ca²⁺ concentrations were measured using the Fluo-8/AM and Mag-Fluo-4/AM (AAT Bioquest Inc., USA) dyes, which were used for quantification in previous studies,^22–27 respectively, according to the manufacturer’s instructions. Briefly, myoblasts obtained after phase (i) culture in black clear-bottomed 96-well plates were treated with various concentrations of propofol, TUDCA, or BAPTA-AM (a Ca²⁺ chelator and negative control to mitigate Ca²⁺ elevation) at 37°, 39°, or 41°C for 1 h and then with 5 µM Fluo-8/AM or Mag-Fluo-4/AM for 1 h. After washing with Hanks’ balanced salt solution, fluorescence intensity was measured (excitation and emission wavelengths, 492 and 530 nm) using a Nivo multimode plate reader and Nivo system software calibrated at the bottom of the plate. All condition samples were performed in triplicate. The experiments were repeated five or six times.

Quantitative Real-time Polymerase Chain Reaction

Phase (i) and phase (ii) cells were treated at 37° or 39°C for 24 h, after which total RNA was extracted using the RNeasy Plus mini kit (QIAGEN, Germany). The reverse transcription reaction was performed using the high-capacity cDNA reverse transcription kit (Thermo Fisher Scientific) and a T3000 Thermocycler (Biometra, Germany), and the cDNA products were stored at −20°C until further use. Quantitative real-time polymerase chain reaction (PCR) was performed using the TaqMan Fast Advanced Master Mix (Thermo Fisher Scientific) and the StepOnePlus real-time PCR system (Thermo Fisher Scientific). All procedures were performed according to the corresponding manufacturer instructions. The specific primers used with the TaqMan probes are listed in table S1 (Supplemental Digital Content 1, https://links.lww.com/ALN/E101). The quantitative real-time PCR data were analyzed using the delta–delta threshold cycle (ΔΔCt) method with GAPDH as the internal control, and fold-change values were calculated using the 2^−ΔΔCt method. All condition samples were performed in triplicate. The experiments were repeated five or six times.

Immunofluorescence Staining

Myotubes obtained in phase (ii) and treated at 37°, 39°, or 41°C for 24 h were fixed with 4% paraformaldehyde in PBS for 30 min, washed with PBS, incubated in blocking solution (2% bovine serum albumin with 0.5% Triton-X 100) for 1 h, and then treated with a primary antibody against myosin heavy chain (1:1000; R&D Systems, USA) at 4°C overnight. After washing with PBS, the myotubes were then incubated with a secondary antibody (1:1,000; Thermo Fisher Scientific) for 1 h at room temperature. Finally, the myotubes were incubated with Hoechst 33258 for 5 min at room temperature for nuclear staining. Images were acquired using a fluorescence microscope and analyzed using ImageJ. The experiments were repeated five or six times.

Functional Testing of Human Contractile Muscle Tissues

To evaluate the effects of the drugs on muscle tissues, we sequentially observed the contractions of the muscle tissues fabricated in phase (iii) under EPS induction using a CCD camera (Basler Ace; Basler AG, Germany; 30 frames/s) and phase contrast microscopy before and after treatment with 200 µM propofol for 24 h at 37° or 39°C. After drug exposure, the culture medium was replaced with fresh medium to remove excess propofol considering its direct muscle relaxant effect.²⁸ Contraction displacements were measured using a motion analysis tool (ViewPoint; Glenallan Technology Inc., USA), with the contraction decrease rate after drug treatment determined by normalizing against that of untreated tissue as 100%. The experiments were repeated six times. Only the propofol + TUDCA group exhibited poor condition (no movement) before exposure, resulting in the exclusion of the sample once.

Statistical Analysis

All data were analyzed for statistical significance using JMP Pro 16 software. The results were expressed as mean ± SD values. The sample size was defined by the number of repetitions for each experiment. Blinding was not used in this study. For comparing three or more groups, analysis of variance followed by the Tukey–Kramer or Dunnett’s post hoc test was used for statistical analysis. P < 0.05 was considered statistically significant. Due to the large effect size observed in the Supplemental Digital Content (Supplemental Digital Content 1, https://links.lww.com/ALN/E101; figs. S1 and S2 show viability and cytotoxicity with/without drug treatment, fig. S3 shows apoptosis evaluation, fig. S4 shows ER stress without drug treatment, and fig. S5 shows propofol-induced apoptosis evaluation with/without the addition of inhibitors) using immortalized mouse C2C12, the same approach was applied to human myoblasts. However, for the temperature-only experiments, in which it was necessary to confirm whether cell toxicity would occur under high-temperature conditions, the sample size was increased to enhance statistical power. The basic sample size was set to 5 to 6 (α = 0.05, 1 − β = 0.8, Cohen’s f approximately 0.8). For experiments in which a clear difference was observed, the sample size was reduced to three, considering the value of the human myoblast samples.

Results

Effects of Propofol, Acetaminophen, and Dexmedetomidine on Muscle Cells at High Temperatures

The LDH activity assay results indicated that human myoblasts were not affected by culture temperatures of 39° and 41°C [F(2,21) = 1.6296; P = 0.2198; N = 8; fig. 1A]. With regard to the effects of drug treatments on myoblasts cultured at these temperatures, propofol treatment at concentrations greater than or equal to 300 μM resulted in increased LDH activity only at 39° and 41°C (mean difference, 2.27 [95% CI, 1.27 to 3.27] and 4.78 [95% CI, 3.18 to 6.38]; P < 0.0001 and < 0.0001; N = 5; fig. 1B). In the acetaminophen group, LDH activities were elevated only at the high concentration of 5 mM, irrespective of temperature (37°, 39°, and 41°C; mean difference, 1.02 [95% CI, 0.18 to 1.86], 0.89 [95% CI, 0.27 to 1.51], and 1.02 [95% CI, 0.19 to 1.85]; P = 0.0144, 0.0030, and 0.0122; N = 5; fig. 1C). In the dexmedetomidine group, there was no increase in LDH activity even at the maximum concentration of 50 µM (37°, 39°, and 41°C; F(5,24) = 0.5477, 0.4120, and 0.5960; P = 0.7385, 0.8357, and 0.7032; N = 5; fig. 1D). Thus, only propofol treatment over a specific concentration and at high temperature synergistically triggered significant enhancement of cell damage in human myoblasts.

Fig. 1.

Lactate dehydrogenase (LDH) activity in human myoblasts after 12 h of drug exposure at different temperatures (37°, 39°, and 41°C). (A) No drug treatment. (B) Propofol treatment (50 to 400 µM). (C) Acetaminophen treatment (1 to 5 mM). (D) Dexmedetomidine treatment (0.1 to 50 µM). The data represent relative LDH activity in treated groups compared with that in the control group (without drug treatment). Each bar represents the mean ± SD value (N = 5–8). *P < 0.05; **P < 0.01; ***P < 0.001 compared with control.

Effects of Drug Treatments and High Temperatures on Caspase Activities and Apoptosis Induction

Treatment with propofol or acetaminophen is reported to induce apoptosis in some cell types at normal culture temperature (37°C).^29,30 In this study, we assessed the activities of caspase-3/7 and caspase-9, key regulators of apoptosis, to investigate apoptosis in treated myoblasts. High temperature alone (without drug treatments) did not affect caspase-3/7 and caspase-9 activities (caspase-3/7: F(2,18) = 0.3317, P = 0.7220, N = 7; caspase-9: F(2,18) = 2.3420, P = 0.1247, N = 7; fig. 2A). Furthermore, acetaminophen did not activate caspase-3/7 and caspase-9 even at the high concentration of 5 mM (caspase-3/7 at 37°, 39°, and 41°C: F(3,16) = 1.3978, 0.5769, and 0.1760; P = 0.2798, 0.6385, and 0.9111; N = 5; and caspase-9 at 37°, 39°, and 41°C: F(3,16) = 0.5384, 0.3369, and 0.1515; P = 0.6627, 0.7989, and 0.9272; N = 5; fig. 2B). Conversely, treatment with 300 μM propofol at high temperatures increased caspase-3/7 and caspase-9 activities (caspase-3/7 at 39° and 41°C: mean difference, 101.22% [95% CI, 40.34 to 162.10] and 128.68% [95% CI, 59.27 to 198.09]; P = 0.0015 and 0.0005; N = 5; and caspase-9 at 39° and 41°C: mean difference, 61.96% [95% CI, 24.82 to 99.10] and 121.09% [95% CI, 47.86 to 194.32]; P = 0.0014 and 0.0016; N = 5; fig. 2C).

Fig. 2.

Apoptosis in human myoblasts after drug treatments at different temperatures (37°, 39°, and 41°C). (A to C) Caspase-3/7 and -9 activities after 12 h of culture without (A) and with (B and C) drug treatments. (B) Acetaminophen (1 to 5 mM). (C) Propofol (1 to 300 µM). (D) Caspase-3/7 activities after prolonged propofol exposure (48 h) at 37° or 39°C. (E) TUNEL-positive myoblast counts after control and 200 µM propofol treatment at 37° or 39°C for 24 h. Blue, nuclei; green, TUNEL-positive cells. Scale bar, 200 µm. The data represent mean ± SD values (N = 3–7). *P < 0.05; **P < 0.01; ***P < 0.001 compared with control (or 37°C control). †††P < 0.001 compared with 37°C propofol treatment. ###P < 0.001 compared with 39°C control. TUNEL, terminal deoxynucleotidyl transferase dUTP nick end labeling.

Prolonged exposure to propofol can induce cell death, even at lower concentrations.²⁹ As reported previously, propofol at concentrations greater than or equal to 5 μM activated caspase-3/7 in myoblasts after 48 h of exposure (four times as long as in fig. 2C) at 39°C (control vs. 5 μM: mean difference, 38.70% [95% CI, 6.87 to 70.53]; P = 0.0135; N = 5; fig. 2D). Furthermore, there was a substantial increase in the number of TUNEL-positive myoblasts after treatment with 200 μM propofol at 39°C (vs. 37°C control, 200 μM propofol at 37°C, and 39°C control: mean difference, 78.47 [95% CI, 45.56 to 111.39], 84.35 [95% CI, 51.43 to 117.26], and 84.58 [95% CI, 51.67 to 117.50]; P = 0.0003, 0.0002, and 0.0002; N = 3; fig. 2E). The discrepancy between the results in fig. 2C and fig. 2E with 200 μM propofol at 39°C may reflect the potential effects of prolonged exposure (twice as long as in fig. 2C).

Collectively, these results indicate that propofol treatment at high temperatures synergistically caused cytotoxicity and apoptosis in human myoblasts and that prolonged propofol exposure, even at reduced dosages, could trigger apoptosis in these cells; conversely, although acetaminophen treatment alone was cytotoxic at high concentrations, high temperatures did not result in a synergistic increase in cytotoxicity and apoptosis. Therefore, apoptosis may have been the key mechanism underlying the synergistic effects of propofol treatment at high temperatures.

Elevation of ER Stress and Cytosolic Ca²⁺ Concentrations by Propofol Treatment at High Temperatures

Heat stress has been reported to induce ER stress in rat muscles.³¹ To confirm that propofol exposure at high temperatures exacerbated ER stress in myoblasts, the expression levels of the ER stress activation markers CHOP, GRP78/binding immunoglobulin protein (Bip), and GRP94 were assessed after treatment. The expression levels of all the markers were elevated after treatment with 200 μM propofol at 39°C (CHOP, GRP78/Bip, and GRP94: mean difference, 60.12 [95% CI, 53.27 to 66.97], 16.24 [95% CI, 13.30 to 19.19], and 7.75 [95% CI, 6.95 to 8.55]; P < 0.0001 for all; N = 5), but not at 37°C (CHOP, GRP78/Bip, and GRP94: mean difference, 0.67 [95% CI, −6.18 to 7.51], 0.06 [95% CI, −2.89 to 3.01], and 0.32 [95% CI, −0.48 to 1.12]; P = 0.9921, 0.9999, and 0.6772; fig. 3A).

Fig. 3.

Evaluation of ER stress and Ca²⁺ concentrations in human myoblasts after propofol treatment at high temperatures. (A) Expression of ER stress markers after propofol treatment at 39°C for 24 h. Relative expression levels were determined by normalizing against GAPDH as the internal control. (B) Cytosolic Ca²⁺ concentrations after 1 h of propofol exposure at different temperatures (37°, 39°, and 41°C). (C) ER Ca²⁺ concentrations after propofol treatment for 1 h. (D) Calpain activity in cells after propofol treatment (100 to 300 µM) at different temperatures (37°, 39°, and 41°C) for 12 h. The data represent mean ± SD values (N = 5 to 6). *P < 0.05; **P < 0.01; ***P < 0.001 compared with control (or 37°C control). †††P < 0.001 compared with propofol treatment at 37°C. ###P < 0.001 compared with 39°C control. Bip, binding immunoglobulin protein; CHOP, C/EBP homologous protein; ER, endoplasmic reticulum; FI, fluorescence intensity; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; GRP78, 78-kDa glucose-regulated protein; GRP94, 94-kDa glucose-regulated protein.

Next, we evaluated cytosolic Ca²⁺ and ER Ca²⁺ concentrations. Temperatures of 39° and 41°C, even without the drug treatment, increased cytosolic Ca²⁺ concentrations compared with those at 37°C (37°C vs. 39° and 41°C: mean difference, 1.09 × 10⁶ [95% CI, 0.40 × 10⁶ to 1.78 × 10⁶] and 2.13 × 10⁶ [95% CI, 1.00 × 10⁶ to 3.26 × 10⁶]; P = 0.0037 and < 0.0001; N = 6; fig. 3B). Furthermore, while propofol treatment at 37°C caused no elevation in cytosolic Ca²⁺ concentration, the concentration was increased by treatment at 39° and 41°C (39° and 41°C + 100 μM: mean difference, 1.54 × 10⁶ [95% CI, 0.10 × 10⁶ to 2.98 × 10⁶] and 2.52 × 10⁶ [95% CI, 1.00 × 10⁶ to 4.04 × 10⁶]; P = 0.0350 and 0.0002; fig. 3B). Additionally, ER Ca²⁺ concentration decreased after propofol treatment only at high temperatures and in a concentration-dependent manner (39° and 41°C + 100 μM: mean difference, 2.65 × 10⁶ [95% CI, −0.60 × 10⁶ to 5.85 × 10⁶] and 2.93 × 10⁶ [95% CI, 0.90 × 10⁶ to 4.96 × 10⁶]; P = 0.1156 and 0.0062; and 39° and 41°C + 200 µM: mean difference, 4.93 × 10⁶ [95% CI, 2.00 × 10⁶ to 7.86 × 10⁶] and 4.84 × 10⁶ [95% CI, 3.00 × 10⁶ to 6.68 × 10⁶]; P = 0.0051 and 0.0001; N = 5; fig. 3C). The Mag-Fluo-4 dye, chosen for ER Ca²⁺ quantification in this study, may have limitations, such as issues with organelle localization and nonspecific changes due to other cellular alterations caused by inflammation. However, previous studies have demonstrated its localization to the ER in various cell types, and it has been used for quantitative assessments, making it a useful reference.^23–27

Finally, we evaluated calpain activity in relation to both ER stress and cytosolic Ca²⁺ elevation. Propofol treatment at greater than or equal to 200 μM activated calpain only at 41°C as compared with that in the no treatment group (mean difference, 223.91% [95% CI, 47.88 to 399.94]; P = 0.0121; N = 5; fig. 3D). Treatment at 37°C did not induce any enhancement of calpain activity. Unexpectedly, while Ca²⁺ concentrations increased with propofol treatment at 39°C (fig. 3B), the increase in calpain activity was not substantial at 39°C. This may be due to the sensitivity of each measurement and the possibility that the Ca²⁺ increase did not reach the threshold necessary for a significant increase in calpain activity.³² In summary, only propofol treatment at high temperatures elicited substantial ER stress and cytosolic Ca²⁺ overload in human myoblasts.

Effects of TUDCA Supplementation on Propofol-induced Apoptosis at High Temperatures

As shown in figures 2 and 3, propofol-induced apoptosis was initiated by ER stress at high temperatures. Therefore, in this study, the effects of TUDCA, an ER stress inhibitor, on treated myoblasts were assessed by analyzing caspase-3/7 activity. TUDCA alleviated caspase-3/7 activity in the cells after propofol treatment at high temperatures (propofol at 39° and 41°C vs. propofol + TUDCA at 39° and 41°C: mean difference, 91.66% [95% CI, 38.10 to 145.22] and 72.87% [95% CI, 26.02 to 119.71]; P = 0.0008 and 0.0016; N = 5–6; fig. 4A). It also mitigated the cytosolic Ca²⁺ overload after propofol treatment at 39°C (propofol at 39°C vs. propofol + TUDCA at 39°C: mean difference, 2.98 × 10⁶ [95% CI, 1.29 × 10⁶ to 4.67 × 10⁶]; P = 0.0002; N = 5; fig. 4B) as well as the increased expression of ER stress markers (CHOP, GRP78/Bip, and GRP94; propofol vs. propofol + TUDCA: mean difference, 35.83 [95% CI, 25.58 to 46.08], 7.87 [95% CI, 4.46 to 11.28], and 2.98 [95% CI, 0.10 to 5.85]; P < 0.0001, < 0.0001, and = 0.0409; N = 5; fig. 4C). Thus, TUDCA alleviated the apoptosis and attenuated the cytosolic Ca²⁺ overload and upregulation of ER stress markers induced by the synergistic effect of propofol treatment and high temperature.

Fig. 4.

Effects of TUDCA on human myoblasts treated with propofol at high temperatures. (A) Caspase-3/7 activities in the cells after propofol treatment at 39° or 41°C for 12 h with and without the addition of TUDCA. (B) Cytosolic Ca²⁺ concentrations after 1 h of propofol treatment at 39° or 41°C with and without the addition of TUDCA or BAPTA (a Ca²⁺ chelator used as the negative control). (C) Expression of ER stress markers after propofol treatment at 39°C for 24 h with and without the addition of TUDCA (200 µM propofol and 1 mM TUDCA and/or 10 µM BAPTA). Relative expression levels were determined by normalizing against GAPDH as the internal control. Propofol, TUDCA, and/or BAPTA were administered concurrently. The data represent the mean ± SD values (N = 5 to 6). *P < 0.05; **P < 0.01; ***P < 0.001 compared with control. ††P < 0.01; †††P < 0.001 compared with TUDCA control. #P < 0.05; ##P < 0.01; ###P < 0.001 compared with propofol and TUDCA treatment. ‡‡‡P < 0.001 compared with BAPTA control. §§§P < 0.001 compared with propofol and BAPTA treatment. BAPTA, 1,2-bis (2-aminophenoxy) ethane-N,N,N',N'-tetraacetic acid tetrakis (acetoxymethyl ester); Bip, binding immunoglobulin protein; CHOP, C/EBP homologous protein; ER, endoplasmic reticulum; FI, fluorescence intensity; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; GRP78, 78-kDa glucose-regulated protein; GRP94, 94-kDa glucose-regulated protein; TUDCA, tauroursodeoxycholic acid.

Effects of Drug Treatments on Conformational Changes in Human Myotubes at High Temperatures

During muscle development, myoblasts differentiate into myotubes through the fusion of muscle progenitor cells. Thus, in addition to the effects of drug treatments at high temperatures at the cellular level, we also assessed the effects on myotubes differentiated from myoblasts. For this purpose, we used our previously developed method to regulate the alignment of myotubes using a micropatterned culture substrate such that they mimic the aligned structure of native muscle.¹⁴ When the aligned myotubes were cultured without propofol or acetaminophen, the high temperatures alone induced no conformational changes in the myotubes [F(2,15) = 0.6718, P = 0.5255; N = 6; fig. 5A]. However, immunofluorescence staining showed significant detachment of myotubes from the culture substrate after treatment with 5 mM acetaminophen regardless of culture temperature (vs. 37°, 39°, and 41°C control: mean difference, 48.34% [95% CI, 30.48 to 66.2], 50.33% [95% CI, 30.95 to 69.71], and 44.59% [95% CI, 26.26 to 62.92]; P < 0.0001; N = 5–6; fig. 5B). This indicated that acetaminophen treatment alone could severely damage myotubes. Conversely, while there were no conformational changes in myotubes after treatment with 300 μM propofol at 37°C [F(3,17) = 1.4633, P = 0.2598; N = 5–6; fig. 5C], significant cell detachment was observed after propofol treatment even at relatively low concentrations (200 and 300 μM) at high temperatures (39°C + 200 and 300 μM: mean difference, 11.17% [95% CI, −0.82 to 23.16] and 21.61% [95% CI, 9.61 to 33.61]; P = 0.0707 and 0.0006; 41°C + 200 and 300 μM: mean difference, 24.28% [95% CI, 12.78 to 35.78] and 34.80% [95% CI, 23.31 to 46.29]; P = 0.0001 and < 0.0001; N = 5 to 6; fig. 5C). This suggests that high temperatures synergistically enhanced the effect of propofol treatment.

Fig. 5.

Effects of drug treatments on human myotubes at different temperatures (37°, 39°, and 41°C). (A to C) Conformational changes in aligned myotubes after 24 h of culture without (A) and with (B and C) drug treatments. (B) Acetaminophen (3 to 5 mM). (C) Propofol (50 to 300 µM). Blue, nuclei; green, MHC. Scale bar, 200 µm. (D) Expression levels of MYH7b and CHOP in myotubes after the drug treatments. Relative expression levels were determined by normalizing against GAPDH as the internal control. The data represent mean ± SD values (N = 5 to 6). *P < 0.05; **P < 0.01; ***P < 0.001 compared with control (or 37°C control); †P < 0.05; ††P < 0.01; †††P < 0.001 compared with 39°C control. #P < 0.05; ##P < 0.01; ###P < 0.001 compared with propofol treatment at 39°C. CHOP, C/EBP homologous protein; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; MHC, myosin heavy chain; MYH7b, myosin heavy chain 7b.

Temperatures of 39° and 41°C have been reported to induce increased expression of slow myosin heavy chain (MHC) in cultured myoblasts because of a shift from the fast to slow myofiber type.³³ Therefore, in the current study, we evaluated the expression of myosin heavy chain 7b (MYH7b), a representative protein expressed in slow muscles, in myotubes treated with propofol. Unexpectedly, unlike in the previous study,³³ a temperature of 39°C alone did not result in a clear increase in MYH7b expression (vs. 37°C control: mean difference, 0.31 [95% CI, −0.05 to 0.67]; P = 0.1034; N = 6; fig. 5D). Furthermore, although propofol treatment at 37°C did not reduce expression (vs. 37°C control: mean difference, 0.18 [95% CI, −0.17 to 0.54]; P = 0.4851; fig. 5D), treatment at 39°C dramatically decreased MYH7b expression (vs. 37°C control: mean difference, 0.92 [95% CI, 0.57 to 1.28]; P < 0.0001; fig. 5D). This result corresponds with that of a previous study indicating that skeletal muscle apoptosis causes slow MHC downregulation in a rat model.³⁴ Additionally, CHOP expression levels increased significantly after propofol treatment at 39°C (vs. 37°C control: mean difference, 4.50 [95% CI, 3.20 to 5.80]; P < 0.0001; fig. 5D) but not at 37°C (vs. 37°C control, mean difference 0.12 [95% CI, −1.18 to 1.42]; P = 0.9942; fig. 5D).

Thus, propofol treatment resulted in conformational changes and downregulation of slow MHC expression in myotubes only at high temperatures, indicating that the synergistic effects specifically affected the maintenance of myotubes. In contrast, high-dose acetaminophen induced myotube detachment irrespective of temperature. These results suggest that the two drugs induced different conformational changes in human myotubes.

Effects of TUDCA on Human Myotubes under Propofol Treatment at High Temperatures

Activation of muscle cell death pathways is reported to contribute to a significant loss of muscle fibers in a rat model.³⁵ As it was expected that the myotube detachment induced by the drug treatments would involve apoptosis associated with ER stress, and TUDCA was observed to alleviate propofol-induced apoptosis at the cellular level (fig. 4), we next investigated whether TUDCA could prevent the detachment of myotubes induced by propofol treatment. As expected, TUDCA mitigated myotube detachment (propofol + TUDCA vs. propofol: mean difference, 33.06% [95% CI, 20.79 to 45.33]; P < 0.0001; N = 5; fig. 6A) and suppressed the downregulation of MYH7b after propofol treatment at 39°C (propofol + TUDCA vs. propofol: mean difference, 0.54 [95% CI, 0.19 to 0.89]; P = 0.0020; N = 5; fig. 6B), which indicates a correlation between skeletal muscle apoptosis and the downregulation of slow MHC, as depicted in fig. 5D. Furthermore, TUDCA attenuated the upregulation of CHOP expression induced by propofol treatment at 39°C (propofol vs. propofol + TUDCA: mean difference, 4.29 [95% CI, 1.80 to 6.77]; P = 0.0008; fig. 6B). Collectively, these results suggest that TUDCA was effective in attenuating propofol-induced apoptosis at high temperatures, preventing the conformational changes in myotubes, and suppressing MYH7b downregulation.

Fig. 6.

Effects of TUDCA on myotubes after propofol treatment for 24 h at high temperatures. (A) Conformational changes in myotubes after propofol treatment (200 µM) at 41°C with and without the addition of 1 mM TUDCA. Blue, nuclei; green, MHC. Scale bar, 200 µm. (B) MYH7b and CHOP expression levels in myotubes after propofol treatment at 39°C with and without the addition of TUDCA. Relative expression levels were determined by normalizing against GAPDH as the internal control. Propofol and/or TUDCA were administered concurrently. The data represent mean ± SD values (N = 5). *P < 0.05; **P < 0.01; ***P < 0.001 compared with control. †P < 0.05; ††P < 0.01; †††P < 0.001 compared with TUDCA control. #P < 0.05; ##P < 0.01; ###P < 0.001 compared with propofol and TUDCA treatment. CHOP, C/EBP homologous protein; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; MHC, myosin heavy chain; MYH7b, myosin heavy chain 7b; TUDCA, tauroursodeoxycholic acid.

Effects of Propofol Treatment at High Temperature on the Functionality of Engineered Human Muscle Tissue

Our group has previously reported the development of a tissue engineering method to produce muscle tissues with biomimetic structures and functions of native muscle.^14,36 This functionally mature muscle tissue contracts in response to electrical stimulation, and in this study, its contractile abilities were determined to evaluate the effects of propofol treatment by comparing muscle contractions before and after the drug treatments. Supplemental video 1 (Supplemental Digital Content 2, https://links.lww.com/ALN/E102) showed twitch contractions of the engineered muscle tissue in response to electrical stimulation (frequency, 1 Hz) at 37°C. The muscle tissue similarly contracted before and after propofol treatment. On the other hand, the muscle contractions significantly reduced after the same treatment at 39°C, compared with that before the treatment (Supplemental Digital Content 3, supplemental video 2, https://links.lww.com/ALN/E103). As shown in figure 7A, contraction displacements were quantitatively measured before and after propofol treatment at 37° and 39°C, and the propofol treatment at 37°C caused no change in the muscle contractions (37°C control vs. 37°C propofol: mean difference, 4.66% [95% CI, −37.42 to 46.73]; P = 0.9974; fig. 7B). On the other hand, although muscle contraction was not affected by incubation at 39°C alone (vs. 37°C control: mean difference, 13.06% [95% CI, −29.01 to 55.13]; P = 0.8885; fig. 7B), the propofol treatment at 39°C induced the remarkable reduction in contraction displacement (39°C control vs. 39°C propofol: mean difference, 64.39% [95% CI, 22.32 to 106.46]; P = 0.0012; fig. 7B). These data indicated that the synergistic effect of propofol and high temperature significantly affected the contractile ability of the engineered muscle tissues.

Fig. 7.

Effects of drug treatments on the contraction of the engineered human muscle tissue. The tissues were treated with 200 µM propofol and/or 1 mM TUDCA for 24 h at 37° or 39°C, and pre- and posttreatment muscle contractions were compared. (A) Representative displacement profiles of muscle contractions induced by EPS (frequency, 1 Hz). (B) Decrease rates (%) of muscle contraction displacements before and after drug treatments. The data represent mean ± SD values (N = 5 to 6). *P < 0.05; **P < 0.01; ***P < 0.001 compared with 37°C control. ††P < 0.001; †††P < 0.001 compared with 37°C propofol treatment. #P < 0.05; ##P < 0.01 compared with 39°C control. EPS, electrical pulse stimulation; TUDCA, tauroursodeoxycholic acid.

Notably, although we expected that TUDCA would prevent this reduction based on the results of the previous experiments (figs. 4 and 6), it was not able to mitigate the reduction in muscle contraction after propofol treatment at 39°C (vs. 39°C propofol: mean difference, 12.40% [95% CI, –31.73 to 56.52]; P = 0.9194; fig. 7B). As this result did not agree with the results obtained using myoblasts and myotubes, further experiments will be required to understand the underlying mechanisms, such as myopathy and mitochondrial dysfunction, caused by propofol treatment at high temperatures. Nonetheless, our functional testing of the engineered human muscle tissue showed contraction impairment only with propofol treatment at 39°C, indicating that the synergistic effect of treatment at high temperature significantly affected its functionality.

Discussion

The aim of this study is to assess the effects of sedatives, analgesics, and antipyretics—specifically, propofol, dexmedetomidine, and acetaminophen—on muscle cells and tissues at high temperatures in vitro. Previous studies have reported the effects of high temperatures on cultured cells, with the slope of the cell survival curve differing above and below 42.5°C, depending on cell type.¹² In a mouse model, heat stress induced more apoptosis in muscle than in the liver,³⁷ suggesting that heat tolerance may also differ among different organs. Given their role in body temperature regulation, we focused on the effects of drug treatments on cultured muscle cells. Our findings demonstrated that a 24-h exposure to 39°C did not affect the contraction of our engineered human muscle tissue; similar experiments were conducted using C2C12 mouse muscle myoblasts to investigate the differences between them and human cells (Supplemental Digital Content 1, supporting information; figs. S1 to S5, https://links.lww.com/ALN/E101), which showed that high temperatures did activate GRP78/Bip in C2C12 cells (Supplemental Digital Content 1; fig. S4, https://links.lww.com/ALN/E101). As it has been previously reported that a temperature of 40°C can induce an ER stress response in human AD293 cells, whereas severe heat stress (43°C) did not cause an increase in GRP78/Bip level,³⁸ our results in this study suggest that high temperatures activate some ER stress responses while having no cytotoxic effects on skeletal muscle cells and inducing no decrease in muscle contraction.

While several studies have examined the effect of acetaminophen overdose on hepatic cells,^18,30 only a few have reported its effect on nonhepatic cells.¹⁷ The mechanisms underlying acetaminophen toxicity involve its metabolism into N-acetyl-p-benzoquinone imine by cytochrome P450 enzymes. In a rat model, cytochrome P450 in skeletal muscle had the potential to metabolically activate drug-dependent toxicity,³⁹ although no study has investigated the effects of acetaminophen overdose on muscle cells. This study revealed that, irrespective of temperature, treatment with acetaminophen had a concentration-dependent cytotoxic effect on cultured muscle cells and tissues. Acetaminophen can trigger cell death in various cell types by activating either apoptotic or necrotic pathways.^18,30 Notably, 5 mM acetaminophen did not enhance caspase activity in this study, indicating the lack of involvement of an apoptotic death mechanism. Moreover, acetaminophen treatment resulted in a significant loss of myotubes in a concentration-dependent manner irrespective of temperature. Several studies have reported the occurrence of rhabdomyolysis after acetaminophen overdose.⁴⁰ Therefore, our in vitro study could result in a new approach to understanding the relationships between acetaminophen administration and skeletal muscles in the future. Although acetaminophen concentrations at which we could confirm myotube loss were 2.5 to 5 times higher than the reported plasma concentrations in overdose patients (1 to 2 mM)⁴¹ and may be further elevated due to interstitial concentration being approximately half of the tissue concentration,⁴² previous studies have reported similar dosages of acetaminophen affecting cultured hepatic cells in vitro.¹⁸ Overall, this study demonstrated that acetaminophen and high temperatures did not have synergistic effects.

Propofol has been shown to have both cytoprotective or cytotoxic effects in both in vitro and in vivo models depending on exposure time and concentration.^{11,15,16,29,43} Propofol-induced apoptosis is reported to be triggered by mitochondrial respiratory chain failure and autophagy.^11,16,29,43 In this study, supraclinical concentrations of propofol (greater than or equal to 200 μM) had no cytotoxic effects at 37°C but induced significant apoptosis at higher temperatures (39° and 41°C) after 12 and 24 h; furthermore, clinical concentrations of propofol triggered caspase activity after 48 h of prolonged exposure at 39°C. These findings indicate that prolonged exposure to even clinical concentrations of propofol may pose greater risks to muscle cells at high temperatures than at a normal temperature. Moreover, a 24-h exposure to lower concentrations of propofol at high temperatures resulted in reduced myosin levels and slow MHC downregulation after myotube formation; the contraction ability of muscle tissues also decreased significantly after propofol treatment at 39°C. As our tissue model demonstrated the impact on muscle functionality, it could also prove helpful in understanding other important phenomena in clinical situations.

Our results also demonstrated that propofol-induced apoptosis via ER stress and intracellular Ca²⁺ overload occurred only at high temperatures, as propofol treatment at high temperatures depleted ER Ca²⁺ concentrations and elevated cytosolic Ca²⁺ concentrations. Although Ca²⁺ alterations can induce the ER stress response, the precise molecular mechanisms regulating ER stress–triggered apoptosis remain unclear. Currently, the CHOP pathway is the most well characterized ER stress–induced apoptosis pathway.⁴⁴ The activating transcription factor 6 and PKR-like eukaryotic initiation factor 2α kinase (PERK) pathways promote the production of CHOP and growth arrest and DNA damage 153 (GADD153) in response to ER stress. Consistent with this mechanism, the synergistic effects observed resulted in increased expression of CHOP, GRP78/Bip, and GRP94, which are linked to the activating transcription factor 6 and PERK pathways, in this study.

Caspase-12, the first enzyme identified as being significantly involved in ER stress–induced apoptosis, is activated by calcium-activated calpains, which are cytoplasmic proteases. After activation in the ER, caspase-12 cleaves procaspase-9, resulting in the activation of caspase-3, which is dependent on caspase-9. Thus, caspase-12 triggers apoptosis directly rather than through the mitochondrial cytochrome C/Apaf-1 pathway.⁴⁵ However, the function of caspase-12 in human cells remains uncertain because of the presence of several mutations that impair its activity, and other caspases, notably caspase-4,⁴⁶ may also contribute to the ER stress–triggered apoptosis of human cells. Although this study did not investigate caspase-4, it showed the activation of calpain, caspase-9, and caspase-3/7 after propofol treatment at high temperatures in human cells. These findings could provide clues for understanding the phenomenon of ER stress–induced apoptosis.

TUDCA is known to inhibit ER stress, regulate inflammation, and suppress apoptosis. Recent studies have also indicated that TUDCA can inhibit cell death in neurodegenerative diseases such as Parkinson’s disease and Alzheimer’s disease, metabolic disorders such as diabetes, and inflammatory bowel diseases such as Crohn’s disease and ulcerative colitis.^47,48 Therefore, in the current study, we examined the effects of TUDCA on cells treated with propofol at high temperatures. Although TUDCA’s multifaceted effects may influence Ca²⁺ dysregulation, TUDCA inhibited the increase in caspase-3/7 activity due to the drug treatment, resulted in maintenance of the Ca²⁺ balance, reduced the expression of ER stress markers (CHOP, GRP78/Bip, and GRP94), and alleviated myotube detachment after treatment; however, contrary to expectation, it was unable to prevent the decrease in contraction force caused by propofol treatment. The reasons for this remain unclear, and further experiments will be required to elucidate other possible functional dysfunctions, such as muscular dystrophies, atrophies, myopathies, autophagy, and mitochondrial dysfunction caused by propofol treatment at high temperatures.

Dexmedetomidine showed no cytotoxic effects on cultured human muscle cells in this study, even at high temperatures. Some previous studies have demonstrated that dexmedetomidine has protective effects on various cells and organs^19,20,49; for example, it was reported to have organ-protective effects against heat stress (above 42.5°C) in a mouse model.⁴⁹ From this perspective, it is reasonable that dexmedetomidine treatment had no synergistic effects at high temperatures, and these results suggest that dexmedetomidine administration is highly safe for patients with elevated body temperature.

The differences between in vitro drug concentrations and clinical levels, along with potential limitations in ER Ca²⁺ imaging, such as the lack of specific localization of Mag-Fluo-4 to the ER, warrant cautious interpretation. Our muscle tissue model demonstrated propofol’s effects on tissue-level functionality but does not completely replicate in vivo muscle physiology. For example, the unloaded (no resting tension) state of stimulated muscle cells differs from in vivo conditions, and muscle tissues inherently participate in interorgan interactions, as well as cell-to-cell interactions with blood vessel and nerve cells.⁵⁰ Therefore, to determine the effects of the drugs more precisely, muscle tissue models that mimic native muscle more closely will be required. Nonetheless, our study findings highlighted the benefits of in vitro experiments, including sustained drug concentrations and controlled culture temperatures, essential for understanding the investigated phenomena. Furthermore, tissue engineering technologies may also have important implications for clinical practice, particularly in the context of critical illness myopathy.

In conclusion, our findings indicate that propofol had synergistic cytotoxic effects, including those related to conformational and functional impairments, on human skeletal muscle cells and tissues at high temperatures, whereas dexmedetomidine and acetaminophen did not have observable adverse cellular effects. This suggests that selecting dexmedetomidine as the first-line sedative for patients with high body temperatures would be a reasonable approach and that when propofol is necessary, its exposure time and bolus administration should be minimized to prevent a sudden increase in concentration, even in accordance with concurrent clinical guidelines. Since drug selection can affect functional prognosis, the findings of this study could allow us to address currently unknown issues in the field of anesthesia and ICU management from new perspectives. In future work, synergistic effects of clinically relevant doses of acetaminophen and propofol should also be investigated since they are commonly coadministered.

Acknowledgments

The authors thank Editage for English language editing.

Research Support

Supported in part by CellSeed Inc. (Tokyo, Japan).

Competing Interests

Tokyo Women’s Medical University (Tokyo, Japan) receives research funding from CellSeed Inc. (Tokyo, Japan). Dr. Shimizu is a shareholder of CellSeed Inc. The other authors declare no competing interests.

Supplemental Digital Content

Supporting information, https://links.lww.com/ALN/E101

Supplemental Methods and Results

Supplemental Figures and Tables

Supplemental Video 1, https://links.lww.com/ALN/E102

Supplemental Video 2, https://links.lww.com/ALN/E103

Supplementary Material

aln-143-999-s001.pdf ^{(1.4MB, pdf)}

Download video file ^{(1.3MB, mp4)}

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