Remifentanil is a potent, injectable μ-agonist approved by the Food and Drug Administration that has only been studied via the traditional IV delivery. Because of its ester-based pharmacophore, it has a short, context-sensitive half-life, which is relatively independent of infusion duration and/or hepatic/renal function.1,2
The efficacy of inhaled remifentanil is unknown. Specifically, the abundance of pulmonary esterases may prevent remifentanil from attaining therapeutic levels in plasma via inhalation. In addition, the safety of exposure to inhaled remifentanil is unknown.
Clinically, inhaled opioids have been investigated for the treatment of pain and dyspnea in end-stage cancer and chronic lung disease.3–10 The few pharmacokinetic studies that have been conducted on inhaled opioids investigated the pharmacokinetics of morphine and fentanyl.11–14 In a study in which they evaluated inhaled fentanyl, the authors concluded that the pharmacokinetic profile of a single dose of inhaled fentanyl was comparable with IV administration,15 demonstrating the feasibility and efficacy of inhaled opioids. No studies, however, have evaluated the safety of repeated administration of inhaled opioids or the pharmacokinetics and pharmacodynamics of the ultrashort-acting opioid, remifentanil.
The aim of this study was to demonstrate, in rodents, that remifentanil could produce analgesia via inhalation while being noninjurious to airway and lung tissue. Our hypothesis was that inhaled remifentanil would produce rapid onset of analgesia, followed by rapid recovery, while being noninjurious and nonirritating to lung tissues and nasal turbinates of rodents.
The rationale for this study was several-fold: the ability to noninvasively induce profound short or potentially even longer-term analgesia and sedation has many potential clinical uses; for example, to place an IV, change bandages, or insert/remove sutures. Inhaled remifentanil also would address several key issues associated with commonly used anesthetics and analgesics; chlorofluorocarbons and nitrous oxide are both greenhouse gases.16 Remifentanil is not a known environmental hazard, and chlorofluorocarbons and nitrous oxide are eliminated only via exhalation. Because of the inertness of chlorofluorocarbons and nitrous oxide to biologic degradation, elimination also can become a safety issue in patients in whom spontaneous respiration is impeded because of residual sedation. Thus, inhaled remifentanil, used alone or in combination with a traditional volatile anesthetic, may decrease the amount of volatile anesthetic needed for a given effect17 or perhaps replace such anesthetics in certain scenarios. In addition, development of a well-defined method to deliver inhaled remifentanil through an anesthesia machine could prevent drug dilution and drug pump errors associated with IV drug delivery and risks inherent to IV delivery of drugs. IV delivery of anesthetics such as remifentanil and other opioids inherently bypasses the body’s ability to regulate and maintain spontaneous respiration, posing potential risks to patients. If remifentanil were administered via inhalation, the dose would be regulated, in part, via inherent respiratory drive: when the patient is more anxious or in pain, he or she is likely to breathe more; when the patient is more relaxed, he or she is likely to breathe less and inhale less drug, while actively eliminating the drug, independent of respiration, renal function, or hepatic function. Thus, we sought to provide the proof of concept that remifentanil is capable of inducing reversible and safe analgesia via inhalation.
All the studies described in this report were approved by the Institutional Animal Use and Care Committee at the University of Utah. All animals were housed 2 per cage (rats) or 5 per cage (mice) in a fully staffed and Association for Assessment and Accreditation of Laboratory Animal Care-approved vivarium. The vivarium was maintained at 22°C to 26°C with a relative humidity of 40% to 50% under 12-/12-hour light/dark cycles. Animals were provided water and standard laboratory chow ad libitum. Dose-response, onset of action, and pharmacokinetic studies were performed with male Sprague−Dawley rats weighing between 200 and 300 g. Pulmonary mechanics measurements were performed with 6-week-old male C57Bl/6 mice weighing 19 to 23 g and a FlexiVent FX-1 instrument (Scireq, Montreal, Quebec, Canada).
Drugs and Reagents
Acetonitrile, methanol, formic acid, sodium thiopental, CalEx II decalcification solution, n-butyl chloride, and formaldehyde were purchased from Fisher Scientific (Fair Lawn, NJ). Ethanol was purchased from Decon Labs (Baltimore, MD). Ketamine/xylazine, methacholine, and meperidine were purchased from Sigma-Aldrich (St. Louis, MO). Remifentanil (Ultiva) was purchased from Mylan Inc. (Canonsburg, PA). Vecuronium was purchased from Sun Pharmaceuticals (Mumbai, Maharashtra, India).
A whole-body small animal inhalation chamber was constructed essentially as described by Schroeder et al.,18 with the following modifications: a 6.5-quart Hefty® bin was fitted with a low-volume MicroMist Nebulizer (Hudson RCI, Morrisville, NC). The forced air nebulizer was attached at one end, with air flowing through the vaporizer into the chamber. A vacuum system was not incorporated. A small hole was made at the opposite end of the bin to allow for pressure balance and access to the rat’s tail for real-time analgesic testing during the exposure. Rats were restrained using a Broome style small rodent restrainer (Plas-labs, Lansing, MI) within the chamber with their noses located proximal to the nebulizer. Varying concentrations (100–2000 μg/mL) of remifentanil were diluted in 0.9% saline and administered through the nebulizer with filtered air as the carrier.
Analgesia was assessed using an IITC Tail Flick Analgesia Meter, model 336G (IITC Life Science, Woodland Hills, CA). This is an established outcome measure to evaluate analgesia in rodents.19 A 4 × 6 mm heat source generated a tail stimulus. A built-in sensor on the tail groove detected time to tail flick, with 0.01-second accuracy. Tails were tested 2 cm from the tip using 50% light intensity and a preprogrammed cutoff time of 20 seconds to prevent tissue damage/surface burn injury.
Fifty-three rats (n = 5–11/group) were exposed to increasing concentrations of remifentanil aerosols or 0.9% saline (control) for 5 minutes. Analgesia testing was performed by the use of tail flick after 5 minutes of exposure to inhaled remifentanil. Time to tail flick in drug-exposed groups was compared with time to tail flick in pretest baseline (naive) and inhaled saline control groups. Each rat was tested once. Aerosols were generated from 0.9% saline and solutions containing remifentanil at concentrations of 0, 100, 250, 500, 750, 1000, and 2000 μg/mL. Concentrations of remifentanil were increased until the cutoff was achieved; specifically, a lack of tail flick within 20 seconds of heat exposure; 2000 μg/mL was tested first, as researchers were unsure of the concentration required to produce analgesia via inhalation in rats. Because of profound effect, the dose was dramatically lowered to 100 μg/mL and increased incrementally until 5 of 5 rats showed a time to tail flick of 20 seconds. A Student t test was performed to identify doses that elicited a statistically significant difference in the time to tail flick compared with the pretest baseline (naive) and inhaled saline control groups. Additional details are provided below in the Statistical Analysis section.
Onset of Action and Recovery Study
To measure the kinetics of analgesia, 50 rats (n = 4–9/group) were exposed to an aerosol from a 1000 μg/mL solution of remifentanil, which was the concentration that caused the maximal measurable level of analgesia in 5 minutes in the dose−response study. Rats were exposed for up to 5 minutes, and then the chamber was vented and opened to allow the rats to recover. Animals were tested for tail flick response at the following time points during (0, 1, 2, 3, 4, and 5 minutes) and following 5-minute exposure (6, 7, 8, 9, and 10 minutes), demonstrating both onset of and recovery from analgesia. Each rat was tested once.
Measurement of Remifentanil in Blood
Two milliliters of blood was collected per rat after decapitation at the following time points: 0, 1, 3, 5 (during exposure), 8, and 20 minutes (postexposure) after whole-body chamber exposure to 1000 μg/mL aerosolized remifentanil. Time points for blood collection were predetermined using a D-optimal sampling strategy (Supplemental Digital Content 1, http://links.lww.com/AA/B382). The blood was collected into 6 mL of n-butyl chloride to stop esterase-mediated degradation of remifentanil.20 The organic fraction containing remifentanil was separated and dried, and the residues were stored at −80°C until analysis. A validated quantitative liquid chromatography-tandem mass spectroscopy assay was used to measure the amount of remifentanil in rat blood. The analysis was performed on a TSQ Quantum AM liquid chromatography-tandem mass spectroscopy instrument using an XBridge C8 50 × 2.1 column eluted at room temperature with 15% acetonitrile: 85% aqueous formic acid (0.1% v/v) at 0.25 mL/min to chromatographically separate the analytes. The quantitative range of this assay was 0.25 to 2500 ng/mL with intra-assay accuracy and precision within 15% of all target concentrations. Here, data are represented as the ratio of analyte to internal standard (meperidine, 50 ng/sample).21 Absolute quantitative values were deemed unreliable because of the potential of contamination from the fur during the blood collection procedure, as well as the goal of relating blood concentrations to effect, versus establishing specific pharmacokinetic parameters for rats using this specific exposure system.
Three rats were exposed to 1000 μg/mL aerosolized remifentanil, a dose-eliciting maximal analgesia, for 5 minutes every other day for a total of 3 exposures, to evaluate the potential adverse effects of remifentanil on respiratory tissue after repeated exposure. Rats were killed by lethal injection of sodium thiopental. The lungs were fixed using 10% neutral-buffered formalin delivered through a tracheal cannula at a constant pressure of 25 cm H2O for 30 minutes before excision, followed by fixation for an additional 5 days. Skulls were dissected and fixed in CalEx II fixative/decalcifier for 14 days to allow for sectioning and assessment of the nasal turbinates. Before processing, the tissues were placed in 70% ethanol. Serial sections of 5 μm were prepared and stained with eosin and hematoxylin by the University of Utah Research Histology core. Nasal turbinates were evaluated at 3 levels: level 1, immediately posterior to the upper incisor teeth; level 2, at the first and second palatal ridge; and level 3, at the first upper molar teeth, according to the National Institute of Environmental Health Sciences standards for histologic analysis of rat nasal turbinates. A board-certified veterinary pathologist evaluated all samples.
Twenty mice (n = 5/group) were used to assess pulmonary function after acute and repeated exposure to aerosolized remifentanil using a FlexiVent FX -1 small animal ventilator (Scireq). Forced expiratory maneuvers using FlexiVent often are used to assess airway responsiveness in mice.22–26 Specifically, changes in lung resistance, airway resistance, tissue resistance, lung compliance, lung elastance, and tissue elastance were determined by the use of a constant-phase model that has been extensively and successfully used to assess lung mechanics in mice.27,28 Mice were anesthetized with intraperitoneal ketamine/xylazine (50/8 mg/kg), tracheotomized with an 18-g metal cannula, and connected to the FlexiVent. Mouse default ventilation pattern was initiated (rate 150/min, tidal volume 10 mL/kg, 30 cm H2O max, 3 cm H2O positive end-expiratory pressure). Mice were then paralyzed with vecuronium (0.5 mg/kg, intraperitoneally). An electrocardiogram was monitored continuously to ensure proper anesthesia and viability throughout the procedures. Body temperature was monitored constantly and maintained by the use of a heat lamp. Normalization of lung volumes was then performed by initiating a large amplitude perturbation (deep inflation), as described previously.25,26 After normalization, baseline measurements were assessed using broadband low-frequency forced oscillations. Pulmonary mechanics were then evaluated after exposure to an aerosol using an Aeroneb vaporizer (Aerogen Ltd, Galway, Ireland) that creates an aerosol of 2.5 to 4.0 μm particle volume mean diameter. This small particle size facilitates deep airway deposition. Lung mechanics were tested on each mouse 6 times. Control mice were exposed to aerosols of 0.9% saline for 5 treatments followed by a methacholine challenge of 25 mg/mL. Methacholine is a synthetic, nonselective, muscarinic receptor agonist that is used widely to evaluate for airway hyperresponsiveness. For remifentanil exposure, the aforementioned procedure was performed with 1 dose of saline followed by 4 treatments of increasing solution concentrations of remifentanil (25, 50, 100, and 200 μg/mL in saline), followed by a methacholine challenge (25 mg/mL), providing an assessment of any irritating acute effects of remifentanil. For repeated exposure, the mice were exposed to 1000 μg/mL aerosolized remifentanil or saline for 5 minutes every other day for 3 treatments via the whole-body exposure chamber; 1000 μg/mL was chosen, as it elicited the maximal measurable level of analgesia in the dose-response study and was used for the histopathology studies. Forty-eight hours after the third exposure, pulmonary mechanics measurements were performed as described previously for the acute exposures. Hereafter, these 4 exposures over 7 days will be referred to as “subacute administration.”
The experiments featured in this study were powered to achieve 80% power with 1-way analysis of variance and 2-sample t tests. The Shapiro-Wilk test was used to assess the normality of the data and/or the residuals before performing any statistical comparisons. Data are expressed as medians (interquartile range), and comparisons between groups at a single point in time were performed using the t test or the nonparametric Mann-Whitney U test, as appropriate.
For the dose−response and the onset of action tests, we expected a mean time to tail flick of 3 seconds in the control group and a mean of 20 seconds in the high-dose remifentanil groups with an SD of 4 seconds expected for each group. This yielded >99% and 87% power to detect a statistically significant difference using the Student t test for the dose−response and onset of action tests, respectively. The acute pulmonary mechanics experiments were performed using a 1-way analysis of variance with a mean lung elastance of 30 cm H2O/mL expected in the saline control group. Expected lung elastances in the remifentanil group ranged from 30 to 45 cm H2O/mL across the dose range. The SD was assumed to be 20 cm H2O/mL for all groups. These effect size estimates yielded >99% power to detect a difference in lung elastance as a function of varying inhaled remifentanil doses for both the acute and the repeated exposure experiments. For all comparisons, P < 0.001 was considered to be statistically significant. All statistical comparisons were 2-sided. R 3.1.1 (R Foundation for Statistical Computing, Vienna, Austria) and GraphPad Prism (La Jolla, CA) were used to perform the power calculations and statistical analyses.
Rats exposed to increasing concentrations of remifentanil aerosol exhibited increasing depth of analgesia, as indicated by increased time to tail flick with significant analgesia measured at 5 minutes with doses ≥750 μg/mL compared with the pretest baseline (naive) (P < 0.0001) and inhaled saline (P = 0.0002) groups. The maximal measurable level of analgesia was achieved after 5 minutes using a 1000 μg/mL solution of remifentanil, as measured by time to tail flick (P < 0.0001), and no further increase in analgesia was detectable using 2000 μg/mL (Fig. 1). At greater doses (1000 and 2000 μg/mL), rats appeared sedated; the rats were not moving or grooming for 2 to 3 minutes after removal from the restrainer. However, there was no loss of consciousness (loss of righting reflex) or adverse events noted regardless of dose. Even the rats exposed to the highest concentration were visually normal/fully recovered within 2 to 3 minutes of cessation of remifentanil delivery.
Onset of Action and Recovery
Studies evaluating the time of onset of analgesia were performed using the 1000 μg/mL solution concentration. The onset of action for aerosolized remifentanil was rapid, reaching maximal measurable effect approximately 2 minutes after the initiation of exposure. This maximal measurable analgesic effect was maintained until cessation of administration at 5 minutes. After the 5-minute exposure to inhaled remifentanil, recovery was rapid and complete. Recovery was visually apparent at 3-minute postadministration, with animals moving around and engaging in grooming behaviors. In addition, baseline sensitivity to a painful stimulus was observed within 3 minutes of cessation of remifentanil delivery. Analgesic effect was significantly different than baseline (time 0) at time points 2 to 7 minutes (P < 0.0001). Time points 1 and 8 to 10 were not significantly different than baseline (Fig. 2).
Remifentanil in Blood
Remifentanil was measured in blood, and its concentration increased over time with continuous inhalation exposure. At 5 minutes, exposure was discontinued and blood concentrations of remifentanil (Fig. 3) rapidly decreased to baseline levels within 3 minutes, essentially mirroring the analgesic effects (Fig. 2).
Acute intratracheal remifentanil exposure up to 200 μg/mL in mice did not significantly alter any of the pulmonary mechanics measurements compared with saline exposure. Lung parameter response to methacholine challenge in acute remifentanil-exposed mice was also no different than saline-exposed mice (lung elastance and other parameters shown, Supplemental Digital Content 2, Supplemental Figure 4A, http://links.lww.com/AA/B383). Animals subacutely exposed to inhaled remifentanil did not show any alterations in pulmonary mechanics. However, after exposure to inhaled remifentanil, mice subacutely exposed to a methacholine challenge showed a significantly diminished change in lung resistance (P < 0.0001), airway resistance (P = 0.0001), tissue resistance (P < 0.0001), and lung elastance (P = 0.0013) compared with animals subacutely exposed to inhaled saline, while lung compliance and tissue elastance were unchanged (lung elastance, Fig. 4; Supplemental Digital Content 2, Supplemental Figure 4B, http://links.lww.com/AA/B383). On the basis of these data, it was concluded that remifentanil aerosols did not cause lung irritation, bronchospasm, or other adverse pulmonary effects. Furthermore, a potential benefit of decreased irritant-induced bronchoconstriction was observed with repeated administration of inhaled remifentanil, evidenced by a decrease in methacholine-induced changes in lung resistance, lung elastance, airway resistance, and tissue damping.
Toxicity and Histopathology
Rats subacutely exposed to inhaled remifentanil had no obvious difference in behavior, eating habits, or weight gain compared with unexposed rats. Histopathologic examinations of rat lung and nasal turbinates by a veterinary pathologist revealed no evidence of inflammatory changes or tissue damage with subacute exposure to remifentanil aerosol. (Fig. 5A–F shows representative images in 4×–40×. More images are available.)
To our knowledge, this is the first study to evaluate the safety and efficacy of remifentanil inhalation. To that end, this study was designed to be a pilot proof of concept study to show feasibility and safety before progressing to greater fidelity pharmacokinetic/pharmacodynamic studies. Therefore, we used rodent models, because these animals are relatively easy to study but still allow pharmacodynamics to be crudely but effectively elucidated. There also are established models in mice to study pulmonary mechanics after pulmonary drug administration. Our results show that inhaled remifentanil is rapidly absorbed, pharmacologically active, rapidly cleared, and noninjurious.
In this initial study, the chamber design did not mitigate dosing variables, such as quantification of inhaled uptake of drug, rodent positioning in the chamber relative to aerosol introduction, and minute ventilation changes associated with the test procedures or drug effect. Although the actual inhaled dose of the drug by the rats was unknown, however, the dose/response study did show that rats exposed to ≥750 μg/mL concentration of remifentanil aerosol exhibited statistically significant analgesia in response to the tail flick test. Given the aforementioned deficits, it is predicted that the required concentration would be much lower for facemask or direct tracheal delivery because of the large amount of drug that is not inhaled when using a whole-body exposure chamber.
The onset of action from inhaled remifentanil occurred within 2 minutes, and recovery occurred within 3 minutes after cessation of drug administration. In addition, blood concentrations of remifentanil mirrored the analgesic effects, including a rapid spike after pulmonary exposure and a rapid decrease to baseline after cessation. Remifentanil’s rapid onset of action suggests that, inhaled, it could be used to produce rapid analgesia to facilitate short-term clinical procedures, such as establishing IV access in pediatric or mentally delayed patients, whereas rapid drug clearance would limit the potential for airway compromise after drug administration.
Pulmonary mechanics were measured with a FlexiVent to assess acute pulmonary changes in mice. Subacute pathology also was assessed via histologic evaluation of pulmonary tissues in subacutely exposed rats. Significantly lower doses of inhaled remifentanil were used compared with the dose−response study in rats, as mice were used for this experiment, and direct delivery into the trachea via the breathing circuit of the mechanical ventilator rather than a whole-body exposure chamber. It is suspected that doses achieved in mice were comparable with or greater than those achieved in the whole-body exposures using rats. There was no difference between pulmonary mechanics measurements when we compared acute remifentanil with acute saline or when comparing subacute remifentanil and subacute saline. However, although it has not been reported by others using the FlexiVent system, it is possible that the use of ketamine as an anesthetic during these studies may have concealed subtle changes in pulmonary mechanics associated with remifentanil, although significant irritant-like responses occurred after stimulation by a moderate dose of methacholine (25 mg/mL) in mice. In addition, rat tissues were histologically normal.
From these data, we concluded that remifentanil inhalation was nonirritating and noninjurious, even after repeated exposure in both rats and mice. This study agrees with previous studies on inhaled morphine and fentanyl, which also showed no toxicity.5 Regardless, future studies will test inhaled remifentanil in more suitable large animal models, as well as humans, including an assessment of the safety of longer-term and chronically administered inhaled remifentanil.
Repeated administration of inhaled remifentanil may attenuate respiratory hyperresponsiveness. After subacute exposure to inhaled remifentanil, resistance (airway, lung, and tissue) and elastance were less affected by exogenous methacholine challenge compared with subacute saline exposure. Previous studies have investigated the inhaled morphine and fentanyl for relief of dyspnea related to pulmonary disease.5–10 One theory is that opioids mitigate dyspnea by reducing cholinergic responses secondary to the inhibitory action of opioid agonists on the release of acetylcholine in the airway.6,9,10,29 Our findings on remifentanil are consistent with previous reports of a diminished cholinergic response after subacute exposure to inhaled remifentanil. Inhibition of acetylcholine release reduces acetylcholine-induced airway smooth muscle contraction and also reduces acetylcholine-induced increases in mucus secretions. This not only supports our hypothesis that inhaled remifentanil is nonirritating but also suggests that there may be benefit. Again, the safety of chronic dosing will also need to be evaluated.
This study does not exactly mimic potential human exposure scenarios that are envisioned for inhaled remifentanil; however, it does provide conclusive evidence that remifentanil can be delivered via inhalation to produce profound analgesia and limited sedation. Limitations to the current study include the use of a whole-body exposure chamber in which large amounts of drug were wasted and in which determining exact doses was not possible. We attempted to overcome this limitation by tracheal administration of aerosolized drug; however, this required delivery of the drug to an anesthetized animal, which prevented the assessment of remifentanil pharmacodynamics. In addition, the translatability of animal pain models and pharmacokinetics are inherently limited because of the relative perception of pain and large interspecies variability in airway and lung structures, aerosol deposition, and blood concentrations after pulmonary exposure. This study also is limited because of physiological volume limitations in drawing serial blood samples for pharmacokinetic analysis in mice and rats. A large animal study or human study would be required for a comprehensive population pharmacokinetic and pharmacodynamic study with clinically generalizable results. In addition, further studies are needed to fully determine the pharmacokinetic/pharmacodynamics relationship in humans because here there was an artificially induced plateau of analgesic effect (20 seconds) at concentrations of remifentanil aerosol of 1000 μg/mL. In general, increasing concentrations of remifentanil in the aerosols increased analgesia (Fig. 1), but because of the safety cutoff in this assay, it is unclear whether the level of analgesia would also continue to increase. We suspect the answer is no, given the profound analgesia and sedation observed under these experimental conditions at the highest doses. Pain testing beyond the 20-second cutoff in our initial studies caused burn injury, suggesting that the pain stimulus at 20 seconds was likely substantial and that the depth of analgesia had reached a near physiological maximum, regardless of the safety cutoff.
Future studies to develop inhaled remifentanil for human use also will need to address the delivery vehicle and its inherent fluid mechanics. First, unlike volatile-inhaled anesthetics, nebulized remifentanil is not an ideal gas. Therefore, gas laws such as partial pressure will not govern its uptake and distribution, as is the case for traditional inhaled anesthetics. Second, issues associated with laminar and turbulent flow may also affect drug uptake. And third, basic respiratory parameters such as spontaneous and controlled ventilation may complicate pharmacokinetic and pharmacodynamic assessments using inhaled remifentanil.
Finally, known side effects of opioid administration in humans also will need to be assessed. Bradycardia, nausea, and chest rigidity are of concern, particularly in patients who do not have IV access. It may also be possible to alter future formulations to include racemic epinephrine or concurrent inhaled benzodiazepine administration to minimize these risks.
The evidence described here supports our hypothesis that inhaled remifentanil would produce rapid onset of analgesia, followed by rapid recovery, while being nonirritating and noninjurious to the airways and lungs of rodents. To further elucidate the translatability of inhaled remifentanil, pharmacokinetic, pharmacodynamic, and safety studies will need to be performed in humans.
Name: Tatjana Bevans, CRNA, MSN.
Contribution: This research is part of this author’s PhD dissertation. This author was involved in study design, data collection, data analysis, and manuscript preparation.
Attestation: Tatjana Bevans is first author and wrote and approves the final manuscript. Tatjana attests to the integrity of the original data and analysis reported in this manuscript. Tatjana is the archival author.
Name: Cassandra Deering-Rice, PhD.
Contribution: This author contributed to data collection, data analysis, and participated in manuscript preparation.
Attestation: Cassandra Deering-Rice approves the final manuscript.
Name: Chris Stockmann, PhD, MSc.
Contribution: This author contributed to data analysis and manuscript preparation.
Attestation: Chris Stockmann approved the final manuscript.
Name: Alan Light, PhD.
Contribution: This author contributed to study design and data analysis.
Attestation: Alan Light approves the final manuscript.
Name: Christopher Reilly, PhD.
Contribution: This author contributed to data collection, data analysis, study design, and manuscript preparation.
Attestation: Christopher Reilly approves the final manuscript. Dr. Reilly attests to the integrity of the original data and analysis reported in this manuscript.
Name: Derek J. Sakata, MD.
Contribution: This author contributed to study design and manuscript preparation.
Attestation: Derek J. Sakata approves the final manuscript.
This manuscript was handled by: Markus W. Hollmann, MD, PhD, DEAA.
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