2020 Volume 43 Issue 9 Pages 1338-1345
Daptomycin, a cyclic lipopeptide antibiotic, has bactericidal activity against Gram-positive organisms and is especially effective against methicillin-resistant Staphylococcus aureus. Although daptomycin causes unique adverse drug reactions such as elevation of creatine phosphokinase or rhabdomyolysis, the detailed mechanisms underlying these adverse drug reactions in skeletal muscle are unclear. This study aimed to elucidate whether daptomycin causes direct skeletal muscle cell toxicity and investigate the relationship between daptomycin exposure and musculoskeletal toxicity. First, we evaluated the relationship between daptomycin exposure and skeletal muscle toxicity. Of the 38 patients who received daptomycin intravenously, an elevation in creatine phosphokinase levels was observed in five. The median plasma trough concentration of daptomycin in patients with elevated creatine phosphokinase levels was significantly higher than that in patients whose creatine phosphokinase levels were within the normal range, suggesting that increased exposure to daptomycin is related to elevation in creatine phosphokinase levels. In an in vitro study using human rhabdomyosarcoma cells, daptomycin reduced cell viability and increased membrane damage. These effects were more marked under hypoxic conditions. A necroptotic pathway seemed to be involved because phosphorylated mixed lineage kinase domain-like protein expression was enhanced following daptomycin exposure, which was significantly enhanced under hypoxic conditions. These findings indicate that daptomycin elicits cytotoxic effects against skeletal muscle cells via the necroptotic pathway, and the extent of toxicity is enhanced under hypoxic conditions.
Daptomycin (DAP) is a cyclic lipopeptide antibiotic and has bactericidal activity against Gram-positive pathogens, including methicillin-resistant Staphylococcus aureus.1–3) DAP elicits a concentration-dependent bactericidal activity,1) mainly by disrupting bacterial cell-membrane function by forming dimers. It is prescribed for patients with bloodstream infections,2) skin or soft tissue infections,4) and infective endocarditis.4,5) Using a neutropenic murine thigh infection model, Safdar et al. demonstrated that the peak concentration/minimum inhibitory concentration (MIC) ratio and 24 h area under the concentration–time curve (AUC)/MIC are pharmacokinetic (PK)–pharmacodynamic (PD) parameters that best correlate with in vivo DAP efficacy.3)
It has been reported that a DAP trough concentration in plasma (Cmin) of ≥24.3 mg/L is associated with an increase in creatine phosphokinase (CPK) levels. Some patients with DAP Cmin of ≥24.3 mg/L developed rhabdomyolysis.6) CPK is thought to be the most reliable indicator of muscular injury.7) However, this threshold (24.3 mg/L for DAP Cmin) was estimated in only six patients who presented CPK elevation. Moreover, two-thirds of the patients who had developed CPK elevation had severe obesity (more than 100 kg in body weight). Thus, although the package insert of DAP mentions CPK elevation as an adverse drug reaction, the relationship between DAP exposure and CPK elevation needs to be further evaluated.
In a study in Japanese patients with soft skin tissue infection (SSSI), no correlation between DAP Cmin and CPK elevation was observed.8) The dosage of DAP was fixed at 4 mg/kg daily, suggesting that the DAP Cmin was low (median trough level in plasma was 5.9 mg/L). Recently, another observational study suggested that DAP Cmin was significantly associated with CPK elevation based on a logistic regression model.9) The study suggested that frequent CPK monitoring is needed, and therapeutic drug monitoring (TDM) of DAP may be desirable when high-dose DAP is prescribed.
Although CPK elevation and rhabdomyolysis are unique adverse drug reactions probably associated with exposure to DAP, the detailed mechanism through which DAP exerts toxicity on skeletal muscle cells is not clearly understood. In fact, little is known about the direct toxicity of DAP on skeletal muscle cells. To confirm the direct cytotoxicity of DAP on skeletal muscle cells, we first conducted a clinical retrospective study to verify the relationship between the extent of exposure (DAP Cmin) and incidence of CPK elevation. Second, an in vitro pharmacological study was conducted using cultured myoblast cells to determine whether DAP exerts direct skeletal muscle cytotoxicity. In the pathophysiology of sepsis, a type of severe infection, decreased microvascular blood flow due to systemic inflammatory response syndrome leads to tissue hypoxia.10) In patients with sepsis, decreased ability to extract oxygen in skeletal muscle tissues was observed.10,11) To reflect these points, we thought it was necessary to examine the skeletal muscle toxicity of DAP in hypoxic conditions.
This clinical retrospective study to investigate the relationship between DAP Cmin and CPK elevation was conducted in accordance with the Declaration of Helsinki and its amendments. The study protocol was approved by the ethics committee of the Hokkaido University Hospital (protocol number: 014-0419).
PatientsAn observational clinical research was performed as a retrospective study. Patients admitted to the Hokkaido University Hospital and those who had received DAP parenterally for the treatment or prophylaxis of Gram-positive microorganism infections were considered. Patients who received DAP and had their trough (Cmin) concentration recorded between November 2011 and March 2017 were enrolled. Patients were excluded from this study if 1) they had elevated, i.e., higher than the upper limits of normal (ULN) CPK at baseline (ULN; 248 and 153 U/L for male and female patients, respectively), 2) they were re-administered DAP, 3) the duration of DAP administration was less than 4 d, and 4) their age was less than 20 years. The background characteristics of the patients included in this study are presented in Table 1.
| Patient characteristics | CPK-not elevated group | CPK-elevated group |
|---|---|---|
| Number of patients (male/female) | 33 (21/12) | 5 (1/4) |
| Age (years)† | 55.0 (42.0, 72.0) | 68.0 (39.5, 70.5) |
| Body weight (kg)† at baseline† | 57.6 (44.7, 68.7) | 48.3 (46.0, 56.3) |
| Duration of therapy (days)† | 18.0 (12.0, 38.0) | 13.0 (10.0, 21.0) |
| BMI (kg/m2)† | 20.7 (18.3, 24.8) | 24.5 (20.1, 29.6) |
| ICU admission | 6 (18.2%) | 1 (25.0%) |
| Initial DAP dose (mg)† | 350 (250, 395) | 400 (270, 438) |
| Initial DAP dose per actual body weight (mg/kg)† | 6.00 (5.75, 6.24) | 6.17 (4.96, 7.58) |
| Initial DAP dose per ideal body weight (mg/kg)† | 5.49 (4.78, 6.65) | 8.29 (5.08, 9.20) |
| Comorbidity | ||
| Patients on renal replacement therapy | 6 | 1 |
| Diabetes Mellitus | 9 | 1 |
| Hematological malignancy | 8 | 0 |
| Solid tumor | 8 | 1 |
| Coronary heart diseases | 12 | 2 |
| Laboratory data at baseline | ||
| Serum creatinine (mg/dL)† | 1.07 (0.62, 1.67) | 0.81 (0.71, 0.94) |
| Total bilirubin (mg/dL)† | 0.50 (0.450, 1.25) | 0.60 (0.40, 17.9) |
| serum calcium (mEq/L)† | 8.65 (8.13, 9.13) | 9.10 (7.95, 10.3) |
| ALT (IU/L)† | 17.0 (11.0, 29.0) | 24.0 (10.5, 39.5) |
| LDH (U/L)† | 223 (140, 283) | 213 (191, 394) |
| Eosinophile (%)† | 1.6 (1.0, 5.0) | 1.0 (0.1, 2.7) |
| CPK (U/L)† | 23.0 (13.0, 49.5) | 26.0 (16.5, 35.5) |
| Concomitant medications | ||
| Statins | 4 | 1 |
| Dopamine stimulants or antagonists | 12 | 0 |
| Propofole | 3 | 1 |
| Amiodarone | 3 | 1 |
| Fluoroquinolones | 2 | 1 |
| Source of infection | ||
| Catheter-related/bacteremia/sepsis | 18 | 2 |
| Genitourinary | 1 | 0 |
| Intra-abdominal | 1 | 0 |
| Surgical site infection | 10 | 1 |
| Other or unknown | 3 | 2 |
ALT, alanine transaminase; LDH, lactate dehydrogenase; CRP, C-reactive protein; IQR; interquartile range, †data expressed as the median with IQR.
The CPK levels of patients who received DAP were reviewed by checking their charts until 3 d after the end of their DAP treatment. In the National Cancer Institute Common Terminology Criteria for Adverse Events (CTC AE) version 4.0, CPK elevation was categorized as follows: grade 1 (> ULN to <≤ 2.5 × ULN), grade 2 (> 2.5 to ≤ 5.0 × ULN), grade 3 (> 5.0 to ≤10.0 × ULN), and grade 4 (> 10.0 × ULN). In this study, the patients with CPK elevation more than “grade 2” during DAP treatment were assigned to the CPK-elevated group. In total, 38 patients and 72 samples for the determination of DAP Cmin were collected, of which the “CPK-elevated group” consisted of 5 patients and the “CPK-not elevated group” consisted of 33 patients.
Determination of Plasma DAP Cmin ConcentrationThe DAP Cmin in patients was determined by using HPLC and UV detection by modifying a previously established method.5) For Cmin, we collected blood samples within 30 min before the next dose of DAP. Next, 180 µL of plasma was spiked with 20 µL of DAP (Toronto Research Chemicals, North York, Canada) working solution (31.3, 62.5, 125, 250, 500, and 1000 mg/L in methanol). This solution was then mixed with 200 µL of methanol and 200 µL of 50 mg/L ethylparaben (Sigma-Aldrich Japan, Tokyo, Japan) in methanol as the internal standard (IS). The plasma samples from patients were mixed with 220 µL of methanol and 200 µL of IS. After centrifugation at 15000 rpm for 15 min, aliquots of the supernatant (50 µL) were analyzed by an HPLC system (LC-10ADVP; Shimadzu, Kyoto, Japan) equipped with a UV detection system at 215 nm and a reversed-phase C18 packed column (TSK-GEL ODS-80Ts, 4.6 × 150 mm; TOSOH, Tokyo, Japan). The mobile phase consisted of ammonium dihydrogen phosphate (40 mM, pH = 4.0) buffer and acetonitrile (63 : 37 (v/v)) at a flow rate of 1.0 mL/min. The column oven was set at 30°C. The analytical measurement range was 3.13–100 mg/L. The calculated values were adjusted because of difference of contents of dissolved DAP in patient sample, multiplied the calculated value by 1.11. The inter-day assay coefficient of variations (CVs) of 6.25, 25.0, and 100 mg/L DAP were 14.0, 12.0, and 11.1%, respectively (n = 6). The relative errors of the inter-day assays (n = 6) of 6.25, 25.0, and 100 mg/L DAP were 3.60, 0.27, and −2.58%, respectively.
Cell Cultures and Their TreatmentsFor in vitro experiments, cultured human rhabdomyosarcoma (RD) cells, derived from spindle-shaped myoblast cell line, were obtained from the Japanese Collection of Research Bioresources (Osaka, Japan). The cells were maintained under standard conditions (37°C, humidified atmosphere, and 5% CO2)12,13) in plastic culture dishes or multiwell-plates (Corning, NY, U.S.A.). These cells were kept in Dulbecco’s modified Eagle’s medium (DMEM) containing high glucose (Sigma-Aldrich) supplemented with 10% fetal bovine serum (FBS), 100 U/mL of penicillin, and 100 µg/mL of streptomycin.
Hypoxic Treatments in RD Cell CultureTo establish hypoxic condition by lowering oxygen, BIONIX-1™ hypoxic culture kit (Sugiyamagen, Tokyo, Japan) was employed. The kit consisted of an oxygen absorber, oxygen concentration monitor, and a pouch.14) By using the kit, the oxygen concentration in the air around cells grown in plastic plates or dishes was maintained at 1.0 ± 0.3%. The RD cells were treated under hypoxic conditions by using this kit.
Cell Viability Assay (3-(4,5-Dimethylthiazol-2-yl)2,5-diphenyl-tetrazolium Bromide (MTT) Assay) and Cytotoxicity Assay (lactate dehydrogenase (LDH)-leakage assay)Confluent RD cells were seeded onto 96-well plastic plates at a density of 4 × 104 cells/well. After 24 h, the cells were incubated with various concentrations of DAP for 24 and 48 h in DMEM without FBS, penicillin, and streptomycin under normoxic or hypoxic conditions. The effect of DAP on cell viability was assessed by the reduction of MTT, which primarily occurs in the mitochondria by the action of succinate dehydrogenase.14) The extent of injury to the outer membrane of the cells was measured by determining the LDH activity leakage from the intracellular to extracellular fluid15) by using the LDH-cytotoxicity kit (Wako, Tokyo, Japan), according to the manufacturer’s instructions.
Detection of Apoptotic and Necrotic Cells and Determination of Caspase ActivityThe activity of the primary effector caspases, caspase-3 and caspase-7, was determined by employing the Caspase-Glo 3/7 assay™ kit (Promega, Madison, WI, U.S.A.).
RD cells were seeded onto 6-well plastic plates at a density of 4 × 105 cells/well. After 24 h, these cells were treated with various concentrations of DAP (10, 100, and 1000 mg/L) or vancomycin (VCM; 1000 mg/L) for 24–48 h in FBS and penicillin-/streptomycin-free DMEM under normoxia or hypoxia. After treatment, the cells were washed twice with ice-cold phosphate buffered saline (PBS) (−) and then cell lysates were collected by scraping with 0.3 mL/well of cell lysis reagent (Luciferase cell culture lysis reagent, Promega). Then, equal volumes of the lysate and substrate solutions from the kit were mixed and incubated for 60 min at room temperature. Caspase activity was estimated based on luminescence by using GloMax™ multiplate reader (Promega).
To detect apoptotic and necrotic cells, we used the real-time apoptosis/necrosis dual assay kit, RealTime-Glo™ Annexin V apoptosis assay (Promega). During the apoptotic process, annexin V conjugated with luciferase in the assay kit binds to phosphatidylserine of the outer leaflet on cellular membranes, generating a luminescence signal. This kit can also detect necrotic cells simultaneously with apoptotic cells, because a DNA-binding dye in the kit generates a fluorescent signal when the cells lose their membrane integrity after undergoing necrosis.16)
Western Blotting AnalysisThe RD cells in 6-well plates were washed in ice-cold PBS (−) and lysed in a buffer containing 50 mmol/L Tris–HCl (pH 6.8), 2% sodium dodecyl sulfate (SDS), 10% glycerol, 1 mmol/L ethylenediaminetetraacetic acid (EDTA), and 10 µL/mL of protease inhibitor cocktail (Sigma-Aldrich). Protein concentrations were quantified with the BCA assay kit™ (Pierce Chemical Co., Rockford, MA, U.S.A.). Samples were separated by SDS-polyacrylamide gel electrophoresis (PAGE) and then the proteins were transferred onto a polyvinylidene difluoride (PVDF)-membrane (Bio-Rad Labs., Richmond, CA, U.S.A.). After one hour blocking procedure, the PVDF membranes were stained overnight at 4°C with primary antibodies such as anti-phospho-mixed lineage kinase domain-like protein (MLKL) monoclonal antibodies from rabbit (phospho S 358, 1 : 1000, Abcam, Cambridge, U.K.) or anti-hypoxia inducible factor (HIF)-1α antibodies (1 : 1000, BD Transduction Laboratories, Lexington, KY, U.S.A.). After washing the membrane, the secondary antibodies were conjugated with horseradish peroxidase (anti-rabbit immunoglobulin G (IgG)) for 45 min at room temperature. The respective proteins were detected by enhanced chemiluminescence reagent (ECL prime Western blotting detection reagent, GE healthcare life sciences, U.K.). Signals of these blots were scanned and analyzed with a computer software (ImageLab software ver. 6.0.1, Bio-Rad Labs).
Statistical AnalysisFor the comparison of DAP Cmin between the two groups in the clinical study, the data were presented as the median and interquartile range (IQR), and the differences between the groups were analyzed using the Mann–Whitney U-test. For the analysis of the in vitro study, the data were presented as mean ± standard deviation (S.D.), and statistical significance was analyzed by using ANOVA followed by Dunnett’s multiple comparison test. All the statistical analyses were performed using Prism 7 (GraphPad Software, San Diego, CA, U.S.A.). p < 0.05 indicated statistical significance.
Table 1 presents the patient characteristics at baseline. Of all the eligible patients (n = 38), 33 patients had normal CPK levels throughout the treatment course. However, elevated CPK levels (more than Grade 2 by CTC AE v.4.0) were observed in five patients (13.2%): 600, 1004, 1770, 2668, and 6250 U/L. At these CPK levels, their DAP Cmin in plasma was 12.9, 23.9, 21.0, 24.9, and 42.6 mg/L, respectively (only in patient no 5, CPK level was measured one day after DAP Cmin determination). As shown in Fig. 1, the median DAP Cmin in the CPK-elevated group was 23.9 mg/L (IQR: 16.1–32.6), which was significantly higher than that in the CPK-not elevated group (median 9.21 mg/L, IQR: 6.72–14.2).
DAP Cmin in plasma was determined and compared between the CPK-not elevated group (65 points of plasma samples collected from 33 patients) and CPK-elevated group (7 points of plasma samples collected from 5 patients). Data were expressed as median with interquartile range (IQR) in each group. * p < 0.05 by the Mann–Whitney U-test.
The results of the cytotoxicity assays are shown in Fig. 2. DAP elicited a significant reduction in cell viability (Fig. 2A) and enhanced cellular membrane injury (Fig. 2B) in a concentration-dependent manner. The maximum impact of DAP was observed at 1000 mg/L. Cell viability was reduced only at the highest concentration (1000 mg/L) of DAP under normoxic conditions, whereas the same effect was observed at a lower concentration (100 mg/L) under hypoxic conditions (Fig. 2A). Although VCM (1000 mg/L) influenced cell viability in MTT assay (Fig. 2A), it had no effect on the membrane toxicity of RD cells even under hypoxia (Fig. 2B). DAP elicited cell membrane toxicity at a concentration of 1000 mg/L under both normoxia and hypoxia compared to that in respective control cells (Fig. 2B). In this assay, the hypoxic conditions themselves elicited cell membrane damage. The hypoxic state was verified by the upregulation of HIF-1α protein expression (standardized by the expression of actin protein) via Western blotting. Under hypoxic conditions, the HIF-1α protein expression to actin ratio increased significantly (478 ± 120% vs. normoxic condition in control cells, by Student’s t-test), although DAP exposure did not affect the expression of HIF-1α under both normoxic and hypoxic conditions (Supplementary Fig. 1).
RD cells (2 × 105 cells/cm2) were exposed to daptomycin (DAP, 10–1000 mg/L) or vancomycin (VCM, 1000 mg/L) under the conditions of normoxia or hypoxia (1% oxygen) for 48 h. (A) Cell viability was measured by the MTT assay. Data were expressed as percentage of respective controls (con; for open column, the control was DAP 0 mg/L in normoxia; for closed column, the control was DAP 0 mg/L in hypoxia). Each column represents the mean ± S.D. of 6 determinations. (B) LDH-leakage assay in RD cells. The cells were exposed to various concentrations of DAP for 48 h; then the medium in each well was collected and the activity of LDH was measured. Data were expressed as % of total LDH. Each column represents the mean ± S.D. of 6 determinations. (C) RD cells were exposed to various concentrations of DAP for 48 h, and the cell lysate was used to determine caspase-3/7 activity. The activity of each group was adjusted by the protein contents of cells and expressed as percentage of control in normoxia cells (DAP 0 mg/L in normoxia). Each column represents the mean ± S.D. of three values. con: control. For these figures, * p < 0.05 significantly different from the respective control (DAP 0 mg/L in normoxia or hypoxia condition), and # p < 0.05 indicates significantly different from the control cells in normoxia, by one-way ANOVA with Dunnett’s multiple comparison test.
In contrast to the results of cytotoxicity experiments, the activities of caspase-3 and -7 in DAP-treated RD cells did not change even under hypoxic conditions (Fig. 2C).
Although the effect of DAP at 1000 mg/L tended to inhibit the activities of caspase-3/7, it was not significant (Fig. 3). Moreover, VCM (1000 mg/L) had no effect on the activities of these enzymes.
RD cells (2 × 105 cells/cm2) were exposed to daptomycin (DAP, 10–1000 mg/L) or vancomycin (VCM, 1000 mg/L) under the condition of normoxia or hypoxia (1% oxygen) for 48 h. (A) Intensity of fluorescence, which reflected necrotic cell death, was measured and expressed as percentage of control in normoxia for 24 h. (B) Simultaneous with (A), the intensity of luminescence, which reflected apoptosis, was measured and expressed as percentage of control in normoxia for 48 h. Each column represents the mean ± S.D. of four values. Con; control (DAP 0 mg/L). * p < 0.05 significantly different from the respective control (DAP 0 mg/L in normoxia or hypoxia conditions) and # p < 0.05 indicates significantly different from the control cells in normoxia, by one-way ANOVA with Dunnett’s multiple comparison test. † p < 0.05 significantly different from the control in normoxia by Student’s t-test.
Although apoptosis was significantly increased by hypoxia, DAP exposure itself did not have an effect under both normoxic and hypoxic conditions (Fig. 3B), even at the highest concentration (1000 mg/L). However, necrotic cell death was increased by DAP exposure in a concentration-dependent manner (Fig. 3A), whereas VCM had no effect on RD cell viability. The increase of necrotic cell death was observed from the lower concentration of DAP (10 mg/L or higher) in hypoxic conditions, as compared to that in control cells under normoxia (Fig. 3A).
Evaluation on Involvement of MLKL Phosphorylation in DAP-Induced Necrotic Cell DeathUnder hypoxic conditions, exposure to DAP significantly increased the expression of p-MLKL in a concentration-dependent manner, whereas VCM had no effect on p-MLKL expression (Fig. 4). Under normoxic conditions, no significant increase in p-MLKL expression was observed (Supplementary Fig. 2).
(A) A representative pattern of immunoblot demonstrating (B) analysis of protein expression levels. RD cells were exposed to various concentrations of DAP (10–1000 mg/L) or VCM (1000 mg/L) under hypoxic condition (1% oxygen) for 24 h. * p < 0.05 significantly different from control (DAP 0 mg/L) by one-way ANOVA with Dunnett’s multiple comparison test. Con: control. Each column represents the mean ± S.D. of three values.
The purpose of this study was to clarify whether DAP induces direct skeletal muscle cell toxicity. To this end, we used two different approaches. We first investigated the relationship between DAP Cmin and CPK elevation via a clinical retrospective study. Second, we conducted an in vitro study to examine the direct toxicity of DAP on skeletal muscle cells.
In the clinical retrospective study, data of 38 patients who received an intravenous dose of DAP were examined. CPK elevation was observed in 5 patients (13.2%). The median DAP Cmin in patients with elevated CPK was significantly higher than that in patients without elevated CPK, indicating that DAP Cmin is a useful marker for predicting musculoskeletal toxicity of this antibiotic.
Recently, a retrospective clinical observational study conducted in a Japanese university hospital demonstrated that DAP Cmin was a substantial predictor of CPK elevation.9) In that study, CPK elevation was observed in 4/20 patients with a Cmin > 19.5 mg/L, which was similar to our results in patients with elevated CPK (median DAP Cmin = 23.9 mg/L). TDM of DAP might also be beneficial to avoid treatment failure,17) as multivariate analysis demonstrated that lower Cmin of DAP (< 3.18 mg/L) was related to poor treatment outcomes. Although the approved dose regimen for DAP is up to 6 mg/kg daily, Urakami et al. pointed out the necessity of a high-dose regimen of above 8 mg/kg to ensure its effectiveness in patients with normal renal function.18) Thus, TDM for DAP is necessary and useful for maximizing its efficacy and safety.
The direct cytotoxicity of DAP was observed in the in vitro study using cultured skeletal muscle RD cells. The toxicity of DAP on skeletal muscle was observed in early clinical trials with a 12 h dosing interval. Moreover, adverse skeletal muscle reactions such as CPK elevation19,20) were observed. We also attempted to apply the hypoxia model in cultured RD cells because severe infectious diseases such as sepsis cause severe topical circulatory failure, which leads to tissue hypoxia resulting in functional failure of organs.21)
In these in vitro studies, DAP itself was shown to cause direct skeletal muscle toxicity at normal oxygen concentration (normoxic condition). In addition, according to our hypothesis, DAP exerted significantly more musculoskeletal toxicity under hypoxic condition than under normoxic condition. Even in normoxia, DAP caused a reduction in cellular viability and cellular membrane injury (Figs. 2A, B, 3A). These effects of DAP under normoxia condition support the fact that DAP elicited CPK elevation even in healthy volunteers (Phase I study).22)
Among patients with elevated CPK levels, one patient attained a CPK level of 6250 IU/L after DAP treatment. The patient started receiving DAP (7.1 mg/kg every 48 h) on suspected prosthetic heart valve infection, had an underlying coronary heart disease (as described in Table 1), with circulatory failure and impaired blood flow to the lower limbs. Tissue hypoxia might have occurred in this patient, resulting in increased skeletal muscle injury, with significant contributions from DAP. Although heart disease itself is a condition associated with elevated CPK,23) this patient had also undergone renal replacement therapy; therefore, it is considered that musculoskeletal toxicity was induced as a result of DAP accumulation. Although the baseline CPK in this patient was 35 IU/L, significantly elevation of CPK was observed as DAP administration progressed. Finally, DAP Cmin in this patient was elevated up to 38.4 mg/L on 13 d after treatment of DAP, and on the following day the CPK value exceed 6000 IU/L).
DAP primarily induced cytotoxicity through necroptosis and not by apoptosis. DAP-induced elevation of LDH leakage from RD cells (Fig. 2B) implied that the cellular membrane injury had occurred because of DAP exposure. These activities were not observed in VCM-treated RD cells. In fact, there is no description on CPK elevation as an adverse drug reaction of VCM in its package insert.
In the primary culture of skeletal muscle cells (differentiated myotubes) from rats, DAP did not cause any membrane damage at 1000 mg/L.24) In that study, damage to skeletal muscle plasma membrane was quantified using histochemical methods, and a DAP concentration of more than 2000 mg/L was required. In the present study, we employed RD myoblast cells of human origin; they differ from rat myotube cells in that they are undifferentiated. In fact, DAP-induced cytotoxicity was observed at lower concentrations in RD cells (Figs. 2–4) than in the rat myotube primary culture. This difference in sensitivity to DAP toxicity could be due to the difference in cell differentiation conditions or the species of origin. Moreover, as shown in the results of Figs. 2A and 4, the hypoxic conditions that we established might have increased the susceptibility of cells to DAP toxicity.
The in vivo administration of DAP to male rats at 150 mg/kg twice daily for 3 d led to a reduction in the number of myofibers with a loss of plasma membrane integrity, suggesting that the sarcolemma may be the target of DAP-induced musculoskeletal toxicity.25)
Necroptosis is a programmed cell death that is distinct from apoptosis. It leads to the disruption of cellular membrane and release of cellular contents, which is executed by receptor-interacting protein kinase (RIPK)-1 and/or RIPK-3 when caspase is inhibited.26,27) The phosphorylation of MLKL by RIPK-3 resulted in translocation of MLKL to the cellular membrane and cell rupture.27) In our present study, DAP showed its direct cellular membrane toxicity as demonstrated in the LDH-leakage assay (Fig. 2B) or real-time necrosis fluorescence assay (Fig. 3A). This suggested the involvement of p-MLKL in the mechanism of cellular membrane injury and toxicity, more significantly under hypoxic condition.
Our results obtained by using RD cells showed that DAP treatment at concentrations of 10 mg/L or higher in hypoxic conditions resulted in a significant increase in necrotic cells compared to that observed with control cells in normoxia conditions (Fig. 3A). The minimum concentration (10 mg/L) exerted in in vitro cytotoxic effects under hypoxic condition was not quite different from the median Cmin in patients with elevated-CPK described in Fig. 1. However, whether hypoxic conditions actually accelerate the DAP-induced rise in CPK requires further investigation.
Nevertheless, our study had several limitations. First, for some results of cellular experiments, there was a discrepancy in the plasma concentrations (Cmin) of DAP at which toxicity developed based on the clinical and in vitro study. In particular, the onset of toxicity in DAP treated RD cells under normoxic conditions required a higher concentration of DAP than the Cmin (estimated 23.9 mg/L in CPK-elevated patients in Fig. 1) in patients with CPK elevation.
On the other hand, DAP is known to be incorporated into some immune cells and might be involved in the secretion of pro-inflammatory cytokines.28) Depending on the immunological mechanism, indirect cytotoxicity, in addition to the direct toxicity observed in RD cells, could be one of the mechanisms of skeletal muscle toxicity in vivo.
As another reason of this discrepancy, we speculated that reduced blood flow in skeletal muscle tissue led to the topical accumulation of DAP in patients with severe systemic infection.
Second, necrostatin-1, an RIPK1-inhibitor, showed no significant effect on the prevention of DAP-induced increase in necroptotic cells under hypoxic conditions (data not shown). Similarly, Weinlich et al. reported the involvement of RIPK1-independent necroptotic pathway.27) In RD cells, there could be other mechanisms that do not mediate the activation of RIPK-1 during DAP-induced necrotic musculoskeletal cell death. Necroptosis is also known to involve a pathway that directly activates RIPK-3 without RIPK-1,26) however, the involvement of a more detailed mechanism could be elucidated using useful inhibitors for RIPK-3 such as GSK’ 872. This is an issue that needs to be resolved in the future.
In conclusion, DAP induces direct skeletal muscle toxicity by inducing necroptosis via MLKL activation. This toxicity was enhanced under hypoxic conditions. Under conditions such as severe sepsis, topical hypoxic conditions in skeletal muscle tissues and overexposure to DAP may lead to and accelerate musculoskeletal toxicity. The modification of necroptotic pathway may also contribute to the regulation of musculoskeletal toxicity induced by DAP.
This work was supported by JSPS KAKENHI, Grant Number JP16K08901.
The authors declare no conflict of interest.
The online version of this article contains supplementary materials.
