Standards and chemicals
Certified reference materials of amitriptyline, bupropion, citalopram, desipramine, desmethylcitalopram, duloxetine, fluoxetine, imipramine, mirtazapine, nortriptyline, paroxetine, sertraline and trazodone were purchased from LGC Standards (Teddington, London, UK); certified reference materials of clomipramine, desmethylvenlafaxine, doxepin, hydroxybupropion, norfluoxetine, norsertraline, trimipramine, venlafaxine, 7-aminoclonazepam, 7-aminoflunitrazepam, alprazolam, bromazepam, clonazepam, diazepam, flunitrazepam, lorazepam, midazolam, nitrazepam, nordiazepam, oxazepam, temazepam, zolpidem, 6-monoacetylmorphine, N-desmethyltramadol, buprenorphine, codeine, fentanyl, hydrocodone, hydromorphone, meperidine, methadone, morphine, naloxone, naltrexone, oxycodone, oxymorphone, tramadol, bupropion-d9, citalopram-d6, clonazepam-d4, codeine-d3, diazepam-d5, duloxetine-d3 and morphine-d3 from Cerilliant (Round Rock, TX, USA); methanol, acetonitrile, ammonium formate and formic acid from Merck (Darmstadt, Germany); ultrapure deionized water was purified by Milli-Q from Millipore (Billerica, MA, USA). All solvents used in the extraction procedure were HPLC grade. Quantisal™ oral fluid collection devices and elution buffer were purchased from Immunalysis (Pomona, CA, USA).
Calibrators, quality control, and internal standards
The stock solutions of the substances were prepared by dilution of the reference certified material in methanol. Dilutions of the stock solution in methanol were made to create calibrators at 2.5, 5, 25, 50, 100, 150, 250 ng/mL for amitriptyline, bupropion, hydroxybupropion, citalopram, desmethylcitalopram, desipramine, venlafaxine, desmethylvenlafaxine, doxepin, fluoxetine, imipramine, mirtazapine, nortriptyline, sertraline, trazodone, trimipramine, clomipramine, duloxetine, norfluoxetine, norsertraline and paroxetine (antidepressants); at 2.5, 5, 25, 50, 75 and 125 ng/mL for 7-aminoclonazepam, 7-aminoflunitrazepam, alprazolam, bromazepam, clonazepam, diazepam, flunitrazepam, lorazepam, midazolam, nitrazepam, nordiazepam, oxazepam, temazepam and zolpidem (benzodiazepines); and at 5, 25, 50, 100, 150 and 250 ng/mL for morphine, codeine, 6-monoacetylmorphine, buprenorphine, fentanyl, hydrocodone, hydromorphone, meperidine, methadone, naloxone, naltrexone, N-desmethyltramadol, oxycodone, oxymorphone and tramadol (opioids). In this work, zolpidem was reported in benzodiazepine’s group, to embrace all method substances.
Quality control (QC) working solutions were prepared by another analyst (different from the individual preparing the calibrators). The low-quality control (LQC) solutions were prepared in methanol at concentrations of 15 ng/mL for opioids; 7.5 ng/mL for benzodiazepines; 7.5 ng/mL for amitriptyline, bupropion, hydroxybupropion, citalopram, desmethylcitalopram, desipramine, venlafaxine, desmethylvenlafaxine, doxepin, fluoxetine, imipramine, mirtazapine, nortriptyline, sertraline, trazodone and trimipramine and 15 ng/mL for clomipramine, norfluoxetine, paroxetine, duloxetine and norsertraline. Medium-quality controls (MQC) solutions were prepared in methanol at 125 ng/mL for antidepressants and for opioids and at 40 ng/mL for benzodiazepines. High-quality control (HQC) solutions were prepared in methanol at 200 ng/mL for antidepressants and for opioids and at 100 ng/mL for benzodiazepines. More information about QC working solutions are summarized in Table 1.
Internal standard (IS) solutions were made from dilutions of the stock solutions of certified reference materials, to produce a single IS mixture working solution at the concentration of 5 ng/mL for codeine-d3, morphine-d3, clonazepam-d4 and diazepam-d5, and 125 ng/mL for bupropion-d9, citalopram-d6 and duloxetin-d3. All solutions were prepared in methanol and stored in amber glass vials at − 20 °C.
Blank oral fluid samples were mixed with Quantisal™ elution buffer according to the manufacturer’s dilution (1:3, v/v), fortified with the working standard solutions and used for method development and validation.
To demonstrate that the analytical method was fit for purpose, oral fluid samples collected from volunteers participating in parties and electronic music festivals were analyzed (n = 38). The inclusion criteria were age greater than 18 years old and self-report use of the synthetic drug in the last 24 h. The sample collection was performed anonymously, and procedures performed in this study involving oral fluid samples from human volunteers were in accordance with the ethical standards of the University of Campinas committee (Comitê de Ética em Pesquisa da UNICAMP—CEP, CAAE 88770318.0.0000.5404), and with the ethical standards as laid down in the 1964 Declaration of Helsinki and its later amendments or comparable ethical standards.
To perform the liquid-liquid extraction (LLE), 500 µL of sample collected with Quantisal™ oral fluid device was transferred to a 5 mL polypropylene tube, followed by 25 µL of IS solution, 500 µL saturated solution of sodium tetraborate and 1 mL of methyl tert-butyl ether (MTBE). The mixture was vortexed using BenchMixer™ XL multi-tube vortexer (Benchmark Scientific, Sayreville, NJ, USA) for 2 min at 2500 rpm. After that, the samples were centrifuged at 987g for 5 min and the organic layer (700 µL) was transferred to a new 2 mL polypropylene tube and dried under nitrogen stream (10 psi/40 °C) using a TurboVap evaporation system (Biotage, Uppsala, Sweden). The samples were resuspended with 100 µL of a mixture solution (mixture of water and methanol 80:20, v/v, containing 0.1% formic acid and 2 mmol/L ammonium formate) and 1 μL was injected into LC–MS/MS system.
The analysis was performed on a Nexera X2 ultra-high-performance liquid chromatography system coupled to an LCMS8060 triple quadrupole mass spectrometer (Shimadzu, Kyoto, Japan). The chromatographic separation was performed on a biphenyl column (Raptor, 100 × 2.1 mm, 2.7 μm; Restek, Bellefonte, PA, USA), maintained at 40 °C. The mobile phase consisted of ultrapure water containing formic acid (0.1%, v/v) and ammonium formate (2 mmol/L) (A) and acetonitrile (B). The flow rate was 0.4 mL/min, and the elution gradient initialized with 5% B maintained for 0.5 min, followed by a linear increase to 55% B in 5.5 min, and another linear increase to 100% B in 0.5 min, holding at 100% B for 1.5 min and returning to initial conditions over 0.2 min. The system was reequilibrated for 1.3 min before the next injection, with a total chromatographic run of 9.5 min.
The mass spectrometer was equipped with an electrospray ionization source operating in positive mode. The mass spectrometer conditions were: interface temperature at 400 °C, desolvation temperature at 350 °C, heat block temperature at 400 °C, drying gas (N2) flow at 5 L/min, heating gas flow (air) at 15 L/min, nebulizing gas (N2) flow at 3 L/min and collision-induced dissociation gas pressure (Ar) at 270 kPa. The analyses were performed in multiple reaction monitoring (MRM) mode. For each compound, two MRM transitions were selected, one as quantifier and one qualifier for confirmative identification, except for tramadol (only one transition was chosen). Individual chromatographic retention times and MRM information were presented in Table 2. Data were acquired and processed using LabSolutions 5.97 software (Shimadzu).
Method validation was performed based on the Scientific Working Group for Forensic Toxicology (SWGTOX) guidelines . The parameters evaluated were limit of quantification (LOQ), linearity, interference studies, bias, imprecision, matrix effect, carryover, stability, dilution integrity and recovery.
Analytes identification criteria considered (1) a symmetrical chromatographic peak with retention time within ± 2% of the average calibrator retention time, (2) signal/noise ratio higher than 3 for both qualifier and quantifier ions and (3) the ratios of the two transitions within a maximum of ± 30% of those established by the calibrators, varying more for those with low intensity for the major transition .
Limit of quantification
The LOQ was defined as the lowest concentration of the standard calibration curve that fulfilled identification criteria, with a signal-to-noise ratio of at least 10, acceptable bias and imprecision. The LOQ for all analytes was evaluated using three replicates per run, over 3 days with three different sources of the blank matrix.
Linearity was evaluated with calibration range from 0.5 to 50.0 ng/mL for antidepressants (except clomipramine, duloxetine, norfluoxetine, norsertraline, and paroxetine from 1.0 to 50.0 ng/mL), from 0.5 to 25.0 ng/mL for benzodiazepines and from 1.0 to 50.0 ng/mL for opioids. Linearity was evaluated with six-point calibration curves over 5 days, by linear least squares regression (1/x2 weighting) for all analytes. Calibrators were required to quantify within ± 20% of each target concentration, with correlation coefficient (r) greater than 0.99.
Oral fluid samples were fortified with common pharmaceuticals and drugs of abuse/metabolites at 200 ng/mL, extracted and injected into the LC−MS/MS. No peaks were visualized in each analyte’s detection window that satisfied identification criteria. Supplementary Table 1 includes all pharmaceuticals evaluated as potential interferents (selectivity). Ten blank samples from different sources were extracted and analyzed to evaluate possible endogenous interferences. In addition, the potential contribution of native ions present in commercial deuterated ISs was evaluated comparing the blank oral fluid pool with and without IS additions. No interfering peaks should be visualized that satisfied identification criteria.
Bias was evaluated in the triplicate analysis of fortified matrix samples, at three different concentrations (low, medium, and high) over 5 days. It was calculated considering the percentages of nominal deviation from the target concentration. The highest average acceptable bias from the target concentration was ± 20%. Results are presented in percentages.
The imprecision was evaluated in the triplicate analysis of fortified matrix samples, at three different concentrations (low, medium, and high) over 5 days. Both within-run and between-run imprecisions were calculated using the one-way ANOVA (p < 0.05) approach with the varied factor (run number) as the grouping variable . Using this approach, imprecision is considered as relative standard deviation percentage (%RSD) within the triplicate analysis in one day (n = 3) and for 5 days (n = 15) for each concentration. Imprecision values with %RSD less than 20% were considered acceptable.
Matrix effects were evaluated by comparison of target peak areas in six blank samples from different sources fortified with analytes after extraction (at low and high QC levels) with the average target peak areas of a set of neat standards. Results were expressed as percentages considering a negative result indicative of matrix suppression, and a positive result of matrix enhancement.
Carryover was assessed analyzing blank samples immediately after the highest point of the calibration curve was analyzed. It was considered absent if all analyte’s peak were below LOQ values.
All the stability studies were conducted at low and high QC concentrations (n = 6) in triplicate. On day zero, they were aliquoted in 5 mL polypropylene tubes and stored at 25 °C (room temperature), 4 °C (refrigerator) and − 20 °C (freezer). After 3, 7, 15, 30 and 60 days, aliquots of each QC were fortified with IS and quantified using freshly prepared calibration curves. These drug concentrations were compared to those of the initial QC samples.
Sample stability after three freeze–thaw cycles at − 20 °C was evaluated in triplicate on day zero and after quantifying each concentration, the other triplicates were stored at − 20 °C. After three freeze–thaw cycles (one cycle = 24 h), triplicates samples were quantified against a newly prepared calibration curve.
For evaluation of processed samples stability when storage in autosampler, low and high QCs and calibrator samples were extracted and analyzed immediately. These extracts were stored on the autosampler at 10 °C and re-injected after 12, 18 and 24 h. The peak areas of these stored QCs were compared to those obtained immediately.
In all stability studies, analytes were considered stable if the concentration was within ± 20% of the initial concentration.
For dilution integrity studies, a triplicate of blank oral fluid samples was fortified with 500 ng/mL and diluted 20-fold in a blank oral fluid-Quantisal™ buffer mixture. If the measured concentration times the dilution factor is within ± 20% of the target concentration, the integrity of the dilution is established.
Recovery (extraction efficiency) was performed in two batches: the first using six replicates of blank samples fortified with analytes at the low and high concentrations, extracted with the proposed procedure and injected into the LC–MS/MS; the second, using six replicates of blank samples extracted by the proposed procedure and, the final extract was fortified with the analytes at low and high QC concentrations and injected into the LC–MS/MS. The average peak area of the samples fortified prior to extraction divided by the average peak area of the samples fortified after extraction is multiplied by 100 to give the percent extraction efficiency.