Chemicals and reagents
Cyclopropylfentanyl hydrochloride for animal experiments was acquired from Cayman Chemical (Ann Arbor, MI, USA). A stock solution of 1 mg/mL cyclopropylfentanyl in sterile saline was prepared, and 300 µL aliquots were stored at –20 °C. Cyclopropylfentanyl hydrochloride and furanylfentanyl-d5 for analytical standards were acquired from Cayman Chemical. Cyclopropylnorfentanyl and norfentanyl-d5 were acquired from Chiron AS (Trondheim, Norway). Chromasolv methanol (MeOH) of LC–MS grade was acquired from Honeywell Riedel-de Haën (Seelze, Germany). Ammonium formate and formic acid (98%) were acquired from VWR International AS (Oslo, Norway). Ethyl acetate, n-heptane, nitric acid, and sodium hydroxide were acquired from Merck (Darmstadt, Germany). Disodium tetraborate decahydrate was acquired from Chemi-Teknik AS (Oslo, Norway). Saline was acquired from Hospira (Lake Forest, IL, USA), whereas sodium heparin was acquired from Thomas Scientific (Thomas Scientific, Swedesboro, NJ, USA). Type 1 water (18.2 MΩ) purified with a Synthesis A 10 milli-Q system from Millipore (Billerica, MA, USA) was employed.
Animals and surgery
Male Sprague–Dawley rats (300–400 g, Envigo, Frederick, MD, USA) were double-housed under conditions of controlled temperature, humidity, and light period (22 ± 2 °C, 45 ± 5% humidity, light period 7 AM–7 PM), with ad libitum access to food and water. The animal experiments were approved by the Institutional Animal Care and Use Committee of the NIDA Intramural Research Program, and all procedures were performed in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. Vivarium facilities were fully accredited by the Association for Assessment and Accreditation of Laboratory Animal Care. Animal experiments were designed to minimize the number of rats included in the study.
Rats were anesthetized with intraperitoneal ketamine (75 mg/kg) and xylazine (5 mg/kg), and indwelling catheters constructed of Silastic (Dow Corning, Midland, MI, USA) and vinyl tubing were implanted into the right jugular vein, as described previously (Concheiro et al. 2014). In short, the proximal Silastic end of the catheter was advanced to the atrium, whereas the distal vinyl end was exteriorized on the nape of the neck and plugged with a metal stylet. Directly after catheter implantation, while still under anesthesia, the rats received surgically implanted temperature transponders (14 × 2 mm, model IPTT-300; Bio Medic Data Systems, Seaford, DE, USA). The transponders were implanted s.c. via a prepackaged sterile guide needle delivery system, along the midline of the back, posterior to the shoulder blades. The radio frequency signals emitted from the temperature transponders were received by a compatible handheld reader system (DAS-7006/7r; Bio Medic Data Systems) to allow noninvasive measurement of body temperature (Elmore and Baumann 2018). Postoperatively, rats were single housed and given one week to recover prior to the experiments. A total of 24 rats were used for these studies, and each subject was used for only one experiment.
Animal experiments
One week after surgery, rats were brought into the laboratory in their home cages and allowed 1 h to acclimate to the surroundings. Experiments were carried out between 9 AM and 6 PM. Polyethylene extension tubes were attached to 1- mL tuberculin syringes, filled with sterile saline solution, and connected to the vinyl end of the catheters. By threading the extension tubes outside the cage, blood sampling was facilitated by an investigator remote from the rat. Catheters were flushed with 0.3 mL of 48 IU/mL heparin saline to facilitate blood withdrawal.
To prepare cyclopropylfentanyl for injection, a 300 μL aliquot of 1 mg/mL cyclopropylfentanyl was thawed, before being serially diluted in sterile saline solution to yield concentrations of 30, 100, and 300 μg/mL. Groups of rats received s.c. injections of saline vehicle (control) or 30, 100, or 300 μg/kg cyclopropylfentanyl on the lower back between the hips (N = 6 rats per dose condition). The drug doses were selected based on preliminary in vivo experiments and our published in vitro binding data (Baumann et al. 2018), which show that cyclopropylfentanyl and fentanyl display nearly equivalent affinity for µ-opioid receptors. Animals were randomly assigned to each dose group. Blood samples (300 μL) were withdrawn via catheters directly before injection and at 15, 30, 60, 120, 240, and 480 min after injection. Samples were collected into 1 mL tuberculin syringes before being transferred to 1.5 mL plastic tubes containing 5 μL of 250 mM sodium metabisulfite as a preservative and 5 μL of 1000 IU/mL heparin as an anticoagulant. Blood samples were centrifuged for 10 min at 1000 g at 4 °C. Plasma was decanted into cryovials and stored at − 80 °C until analysis. The rats received an equal volume of saline solution (300 µL) via the intravenous catheter after each blood withdrawal to maintain volume and osmotic homeostasis.
Pharmacodynamic endpoints were determined at each blood withdrawal by an experienced rater. Catalepsy score and body temperature were obtained just prior to each blood withdrawal, whereas hot plate latency was measured thereafter. On each test day, one investigator prepared solutions of cyclopropylfentanyl and administered the drug to rats, whereas another investigator performed the behavioral scoring with no knowledge about which dose had been administered. Over the course of the 1-min observation period, catalepsy behaviors were scored based on the following three overt symptoms: immobility, flattened body posture, and splayed limbs, as previously described (Elmore and Baumann 2018). Each symptom was scored as 1 = absent or 2 = present at each time point. For each rat, catalepsy scores at each time point were summed, giving a minimum possible score of 3 and a maximum possible score of 6. Subsequently, body temperature was rapidly measured using a handheld reader sensitive to signals emitted by the surgically implanted transponder. Immediately after each blood withdrawal, rats were placed on a hot plate analgesia meter (IITC Life Sciences, Woodland Hills, CA, USA) set at 52 °C. Rats were removed from the hot plate when they exhibited jumping, flinching, or paw licking as a response to the heat stimulus and were then returned to their home cages. A timer triggered by a foot pedal was used to record the time spent on the hot plate. To prevent tissue damage, a 45 s cut-off was employed.
Analytical methods
Determination of cyclopropylfentanyl and cyclopropylnorfentanyl in rat plasma samples was performed using a previously described method (Bergh et al. 2018) with minor modifications. Stock solutions of cyclopropylfentanyl, cyclopropylnorfentanyl, and their internal standards (ISs) were prepared in MeOH and stored at − 20 °C. We used furanylfentanyl-d5 and norfentanyl-d5 as ISs for cyclopropylfentanyl and cyclopropylnorfentanyl, respectively. The optimal choice would have been to use isotopic labeled cyclopropylfentanyl and cyclopropylnorfentanyl, but at the time our method was developed and validated, such compounds were not commercially available. The ISs were therefore chosen based on our previously published and fully validated method developed for determination of cyclopropylfentanyl and 25 other fentanyl analogs (Bergh et al. 2018). Here, furanylfentanyl-d5 was used as IS for cyclopropylfentanyl, whereas norfentanyl-d5 was used as IS for all N-dealkylated metabolites, with acceptable results. Working solutions for 7 calibrators and 5 quality control (QC) samples were prepared separately by diluting stock solutions in MeOH. Calibrators (10–5000 pg/mL) and QC samples (7.5–4000 pg/mL) were prepared by diluting 25 µL of the working solutions with 100 µL commercial pooled blank Sprague Dawley rat plasma containing K2EDTA (Tebu-Bio, Roskilde, Denmark).
Rat plasma samples (100 μL) were transferred to plastic tubes, and IS and MeOH were added prior to vortexing. Borate buffer (pH 11) was added, and the tubes briefly vortexed before addition of an ethyl acetate/heptane mixture, followed by 1 min of vortexing. The tubes were centrifuged, and the supernatants were transferred to glass tubes containing nitric acid in MeOH and evaporated to dryness under a stream of N2. The samples were reconstituted in mobile phase and centrifuged prior to transfer of the supernatants to autosampler vials for UHPLC-MS/MS analysis.
Rat plasma samples were analyzed using an Acquity UHPLC™ system (Waters, Milford, MA, USA) coupled to a Xevo-TQS triple quadrupole MS with an electrospray ionization interface (Waters). The analytes were chromatographically separated on a Kinetex biphenyl column (Phenomenex, Verløse, Denmark) kept at 60 °C with a mobile phase consisting of 10 mM ammonium formate pH 3.1 (solvent A) and MeOH (solvent B) delivered at a flow rate of 0.5 mL/min. The separation was performed using a 9-min gradient profile. Data were obtained and processed using Masslynx™ 4.1 software (Waters). For more detailed information about the UHPLC-MS/MS analysis, see Bergh et al. (2018). The MS/MS parameters used, and the retention times of the analytes and ISs, are presented in Table 1.
Table 1 MRM transitions, cone voltages, collision energies, and retention times Method validation
The UHPLC-MS/MS method was validated in accordance with the Scientific Working Group for Forensic Toxicology guidelines (AAFS 2019), with minor modifications. The validation was performed by examining the following parameters: linearity, intermediate precision and bias, limit of detection (LOD), limit of quantification (LOQ), recovery, matrix effects (ME), matrix interferences, stability, and carry-over.
The linearity was determined based on 8 assays of 7 calibrators prepared in pooled rat plasma from drug-naïve control rats with one replicate per concentration level. Weighted calibration curves (1/x), excluding the origin, were constructed by plotting calibrator concentration against analyte/IS peak height ratio. The calibration curves were considered acceptable with a correlation coefficient (R2) ≥ 0.99 and residuals ≤ ± 20%.
Intermediate precision and accuracy were determined based on 8 assays of 4 different QC sample concentrations prepared in both pooled plasma from drug-naïve control rats and in commercial pooled rat plasma with one parallel pr. matrix source. Precision was determined as the coefficient of variation (% CV). Accuracy, given as bias, was calculated as the percent deviation between the measured mean of the different QC samples and the respective nominal concentration. Intermediate precision and accuracy were determined for all assays collectively and deemed acceptable at a % CV and bias ≤ ± 20%.
LOD was determined based on three assays of five different QC sample concentrations prepared in both pooled rat plasma from drug-naïve control rats and commercial pooled rat plasma with one parallel pr. matrix source. LOD was defined as the lowest concentration which produced a signal-to-noise ratio (S/N) ≥ 3. LOQ was determined based on 8 assays of 4 different QC sample concentrations prepared in both pooled plasma from drug-naïve control rats and commercial pooled rat plasma with one parallel pr. matrix source. LOQ was defined as the lowest QC sample concentration where the S/N ≥ 10 and the intermediate precision and accuracy were ≤ ± 20% for the transition of quantification.
Recovery and ME were determined by analyzing three sets of samples fortified with analytes at two concentration levels (25 and 4000 pg/mL). Ten blank plasma samples from two different sources (4 samples of pooled plasma from drug-naïve control rats and 6 samples of commercial pooled rat plasma) were fortified with analyte pre-extraction (set 1) or post-extraction (set 2). Additionally, five replicates of reconstitution solution were fortified with analyte (set 3). IS was added to all samples post-extraction. Recovery was determined as the ratio between the peak heights of analyte added pre-extraction (set 1) and the peak height of analyte added post-extraction (set 2). ME was determined as the ratio between the peak height of analyte added post-extraction (set 2) and the peak height of analyte added to reconstitution solution (set 3), as described by Matuszewski et al. (2003). The ME was deemed acceptable within the range of 80–120%.
Matrix interferences were assessed in the plasma samples taken from the rats prior to cyclopropylfentanyl exposure (N = 17). Because neither of the ISs used were isotope labeled analogs of the analytes, which can contain traces of the unlabeled analyte, interferences from the ISs were not evaluated.
The stability of cyclopropylfentanyl and cyclopropylnorfentanyl in fortified commercial pooled rat plasma and extracted rat plasma samples fortified pre-extraction were evaluated in triplicates at two concentration levels (25 and 4000 pg/mL). The stability of fortified plasma samples was investigated after two freeze/thaw cycles and after storage for up to 2 months at − 80 °C. The stability of extracted samples was evaluated for up to 2 days in the autosampler at 10 °C. Samples were deemed stable if the deviation from the initial concentration was ≤ ± 20%.
Carry-over was assessed by injecting an extracted rat plasma sample fortified with the highest calibrator (5000 pg/mL) succeeded by three samples of blank extracted matrix. Carry-over was considered present if the blank samples displayed a peak height > 10% of the peak height at LOQ.
Data analysis and statistics
All pharmacodynamic and pharmacokinetic data were statistically evaluated employing GraphPad Prism version 8.0 (GraphPad Software, La Jolla, CA, USA). ED50 values were calculated in GraphPad Prism by nonlinear regression analyses of mean catalepsy scores and hot plate responses over the first 2 h after cyclopropylfentanyl injection. Time course data for body temperature, catalepsy scores, and hot plate latency were evaluated using two-way analysis of variance (dose × time) followed by Tukey’s multiple comparison tests. Time-concentration profiles for cyclopropylfentanyl and cyclopropylnorfentanyl were subjected to two-way analysis of variance (dose × time) followed by Tukey’s multiple comparison tests. Thermo Kinetica version 5.1 (Thermo Fisher Scientific, Philadelphia, PA, USA) was used to determine pharmacokinetic constants for cyclopropylfentanyl and its metabolite cyclopropylnorfentanyl, including concentration maximum (Cmax), time for concentration maximum (Tmax), area-under-the-curve (AUC), elimination constant (Ke), and plasma half-life (T1/2). To determine differences between dose groups, pharmacokinetic constants for each analyte were compared by one-way analysis of variance (dose) followed by Tukey’s post-hoc test. The observed AUC values, from 0 to 8 h post-injection, were compared to predicted AUC values which were calculated for the 100 and 300 μg/kg cyclopropylfentanyl doses by multiplying the observed value at 30 μg/kg by 3.33 and 10, respectively. Predicted and observed values for cyclopropylfentanyl and cyclopropylnorfentanyl AUC were analyzed by two-way ANOVA (dose × condition). Relationships between plasma concentrations of analytes and body temperature, catalepsy score, or hot plate latency were evaluated using a Pearson’s correlation analysis. Specifically, for each subject, the AUC value calculated for each analyte was plotted with respect to the mean temperature, summed catalepsy score, or mean hot plate latency values across all times points. p < 0.05 was employed as the minimum threshold for statistical significance for all comparisons performed.