Rapid Determination of 24 Synthetic and Natural Cannabinoids for LC–MS-MS Screening in Natural Products and Drug Inspection Applications

Feb 26, 2018
Figure 1: Chemical structures and accurate masses of the 24 cannabinoids
Click to enlarge, Figure 1: Chemical structures and accurate masses of the 24 synthetic and natural cannabinoids investigated.
Table I: Spectra with associated structures searched with NIST search software
Click to enlarge, Table I: Spectra with associated structures searched with NIST search software
Abstract / Synopsis: 

Marijuana, the common or slang term for cannabis in its herbal form, is one of the most widely used illicit drugs in the world. Synthetic cannabinoids have similar psychotropic effects to the natural ones and are rapidly being integrated by the illicit market. To deal with their expanding number and diversity, a targeted and untargeted liquid chromatography–tandem mass spectrometry (LC–MS-MS) screening method was developed, allowing for the simultaneous analysis of 24 synthetic and natural cannabinoids in 8 min for a wide variety of samples such as herbal smoking mixtures, incense sticks, serums, and cannabis plant material. The particular advantage of this LC–MS-MS method is that the full scan event in the MS acquisition provides accurate masses for all detected species and thus allows postanalysis identification of initially untargeted compounds.

The primary psychoactive compound in cannabis is Δ9-tetrahydrocannabinol (THC), which exerts its activity by interacting with the CB1 and CB2 cannabinoid receptors in the brain. Synthetic analogs of THC, when smoked or orally ingested, can mimic the psychotropic effects of cannabis by binding the same receptors (1). Cannabimimetic compounds can be found in herbal mixtures (incense stick, cigarette, Spice, K2), serums, and other matrices, often with no indication of their presence. The European monitoring center for drugs and drug addiction reported in 2009 (2) that spice products (3) were generally used by teenagers and young adults because it enabled them to pass drug-screening tests. Many governments have taken legal action to control specific synthetic cannabinoids. In the United States, they have been listed as Schedule I drugs on the Controlled Substances Act, and in Canada as Schedule II drugs. However, small structural modifications from the controlled substance result in new analogs being legal. Therefore, in May 2012 the United States amended the Act and proposed a bill to place all cannabimimetic agents as Schedule I drugs (4,5).

Given the structural similarities between THC and its synthetic analogs, identification of the compounds in seized samples is a continuous challenge for government agencies. As quickly as analogs are added to controlled substance lists, new ones are synthesized, making their monitoring a moving target (6–9). These new generation analogs are obviously not present in mass spectrometry (MS) or UV spectra libraries; therefore, routine methods for screening cannabimimetics in both products and biofluids rapidly become less effective.

The most common methods of cannabinoid analysis use gas chromatography (GC)–MS (10,11) and liquid chromatography (LC)–MS (9,12–14). GC–MS approaches use MS spectra of known cannabinoids for identification, and most LC–MS approaches are used in multiple reaction monitoring (MRM) mode. Both are targeted methods that are limited to screening known species and are therefore always one step behind in the monitoring of this dynamic designer drug market. To address this shortcoming, we report a targeted–untargeted high-resolution MS approach to screen a variety of samples for cannabinoid-like compounds. The current method includes a high-resolution, nontarget scan that allows identification of all species if present in the sample and targets a selection of 20 synthetic and four natural cannabinoids (Figure 1). (See upper right for Figure 1, click to enlarge. Caption: Figure 1: Chemical structures and accurate masses of the 24 synthetic and natural cannabinoids investigated.) 


Reagents and Standards

LC–MS-grade methanol and acetonitrile were from J.T. Baker (TekniScience) and LC–MS-grade formic acid (98%) was from Fluka (Sigma-Aldrich). High performance liquid chromatography (HPLC)-grade water from a Milli-Q Reference A+ system (Fisher Scientific) was used to prepare all aqueous solutions and mobile phases.

A total of 24 standards were investigated. JWH-018 and JWH-210 were obtained from Toronto Research Chemicals Inc. AKB48 N-(5-fluoropentyl) analog, AM-694, AM-694 3-iodo isomer, AM-1220, AM-2201, AM-2201 2'-naphthyl isomer, AM-2201 N-(4-fluoropentyl) isomer, CP 47,497, (±)-CP 47,497-C8-homolog, (±)-epi CP 47,497-C8-homolog, (±) 3-epi CP 47,497-C8-homolog, JWH-019, JWH-073, JWH-081, JWH-122, JWH-200, JWH-250, and STS-135 were obtained from Cayman Chemical. Δ9-Tetrahydrocannabinol and cannabinol were purchased from Alltech/Grace. Cannabidiol was purchased from Lipomed and cannabigerol was purchased from THC PHARM.

Stock solutions of individual standards were prepared separately in 10-mL volumetric flasks at an approximate concentration of 100 μg/mL in methanol. Diluted stock solutions (100 ng/mL to 1 μg/mL) were directly infused into the mass spectrometer for adjustment of the experimental parameters for each analyte. A standard mixture of the 24 components was also prepared and injected to adjust the chromatographic separation of all analogs.

Sample Preparation

To determine the effectiveness and robustness of the LC–MS-MS method, 10 seized samples and a cannabis sample were analyzed. The samples were present in two forms: tablets and herbal products (incense stick, cigarette, and a cannabis plant). They were finely ground, then aliquots of 5–10 mg of the resulting powder were transferred to 10-mL volumetric flasks and dissolved in 70:20:10 methanol–water–acetonitrile containing 1% formic acid. Following this, the solutions were vortexed for 2 min, sonicated for 10 min, and vortexed again for 3 min. The supernatant was filtered through a 0.45-μm pore polytetrafluoroethylene (PTFE) syringe filter (Phenomenex). Herbal samples required an additional centrifugation step at 3500 rpm for 10 min to avoid mass overloading of the syringe filter. Filtrates were diluted 10–100-fold in 80:20 water–acetonitrile, the initial mobile phase, before injection.

LC–MS-MS Operating Conditions

Data were acquired on an LTQ Orbitrap XL mass spectrometer coupled to an Acella HPLC system (Thermo Scientific). Xcalibur 2.1 and Thermo LTQ Tune Plus 2.5.5 software (Thermo Scientific) were used to control the system and process the data. External mass calibration was used throughout the project. Four analytical columns were initially tested for their chromatographic performance: 100 mm × 2.1 mm, 3.5-μm dp XTerra C18 and 100 mm × 2.1 mm, 1.7-μm dp Acquity BEH C18 columns, both from Waters; a 75 mm × 2.1 mm, 2.6-μm dp Kinetex C18 column from Phenomenex; and a 100 mm × 2.1 mm, 2.6-μm dp Accucore aQ C18 column from Thermo Scientific. Two eluent systems were tested during method development, water–methanol and water–acetonitrile, both containing 0.1% formic acid, under generic gradient conditions (5–95% organic).

Optimized separations were carried out using the Accucore aQ column coupled to a 4 mm × 2.0 mm Phenomenex C18 guard column, both maintained at 40 °C, and the water–acetonitrile gradient. The autosampler temperature was set at 10 °C to avoid sample degradation. Eluents consisted of 0.1% formic acid in water (eluent A) and 0.1% formic acid in acetonitrile (eluent B), and the initial mobile phase contained 20% B. The following gradient elution was applied at a flow rate of 350 μL/min: 20–58% B over 1 min, held at 58% B for 1 min, increased to 85% B over 1 min, then held at 85% B for 2 min. Eluent B was then returned to 20% B over 0.2 min. The system was allowed to reequilibrate for 2.8 min, giving a total cycle time of 8.0 min. The injection volume was 3–5 μL. A needle wash step using 70:20:10 methanol–water–acetonitrile was included in the method. A 5-μL blank, consisting of the initial mobile phase, was injected after each sample to monitor and reduce any potential carryover.

The electrospray interface was operated in positive ion mode. Nitrogen was used as both sheath gas and auxiliary gas while helium was used as collision gas. Using direct infusion, instrumental parameters were adjusted semiautomatically for every analyte using the tune tool in the LTQ Tune Plus software. The parent ions of all analytes showed similar behavior due to their similar structures. After screening every compound individually, the experimental parameters of the full scan event that were found to be suitable for all analytes were set to the following values: sheath and auxiliary gas at flow rates of 44 and 17 (instrument units), respectively; spray voltage, +3500 V; capillary temperature, 310 °C; capillary voltage, 28 V; tube lens, 101. The MS2 and MS3 transitions for every compound were also determined using the tune tool by varying the normalized collision energy. They ranged between 25% and 33% after they were optimized. Therefore, a three-step collision energy function set at 25%, 30%, and 35% was used to perform average fragmentation on every compound. The ion transitions MS2 and MS3 for each standard are shown in Table I, where the MS3 transitions arise from the MS2 value shown in boldface type. (See upper right for Table I, click to enlarge.) Mass spectra were acquired from m/z 50 to 1000 using two scan events: the first was a Fourier transform (FT)-MS full scan for accurate mass detection and the second was a data dependent step with MS-MS acquired only for precursors from the parent mass list with a dynamic exclusion of 10 s. Every standard was then injected onto the column individually to determine its retention time and confirm the parent ion accurate mass, MS2 and MS3. To determine if there were any interactions between the compounds, a mixture of the 24 standards was injected.

Results and Discussion

Method Development and Validation

As seen in Figure 1, many of the 24 compounds have similar structures, which makes the chromatographic separation challenging. Between the mobile phases tested, the acetonitrile gradient gave better selectivity. Between the columns tested, the Accucore aQ gave the best selectivity and retention of early eluted analytes.

  1. L. Console-Bram, J. Marcu, and M.E. Abood, Prog. Neuro-Psychopharmacol. Biol. Psychiatry 38(1), 4–15 (2012).
  2. Understanding the 'Spice' phenomenon, Lisbon, 2009, http://www.emcdda.europa.eu/publications/thematic-papers/spice, consulted 09/22/ 2014.
  3. K.A. Seely, J. Lapoint, J.H. Moran, and L. Fattore, Prog. Neuro-Psychopharmacol. Biol. Psychiatry 39(2), 234–243 (2012).
  4. Synthetic Drug Abuse Prevention Act of 2012, https://www.govtrack.us/congress/bills/112/s3190/text, consulted 09/22/2014.
  5. L.N. Sacco and K. Finklea, "Synthetic Drugs: Overview and Issues for Congress," 2014, https://www.hsdl.org/?view&did=757033, consulted 09/22/2014.
  6. S. Dresen, N. Ferreirós, M. Pütz, F. Westphal, R. Zimmermann, and V. Auwärter, J. Mass Spectrom. 45(10), 1186–1194 (2010).
  7. A.D. de Jager, J.V. Warner, M. Henman, W. Ferguson, and A. Hall, J. Chromatogr. B: Anal. Technol. Biomed. Life Sci. 897, 22–31 (2012).
  8. K.G. Shanks, G.S. Behonick, T. Dahn, and A. Terrell, J. Anal. Toxicol. 37(8), 517–525 (2013).
  9. N. Uchiyama, M. Kawamura, R. Kikura-Hanajiri, and Y. Goda, Forensic Sci. Int. 227(1–3), 21–32 (2013).
  10. T. Sobolevsky, I. Prasolov, and G. Rodchenkov, Forensic Sci. Int. 200(1–3), 141–147 (2010).
  11. A. Grigoryev, S. Saychuk, A. Melnik, N. Moskaleva, J. Dzhurko, M. Ershov, A. Nosyrev, A. Vedenin, B. Izotov, I. Zabirova, and V. Rozhanets, J. Chromatogr. B: Anal. Technol. Biomed. Life Sci. 879(15–16), 1126–1136 (2011).
  12. K.B. Scheidweiler and M.A. Huestis, J. Chromatogr. A 1327(0), 105–117 (2014).
  13. A.A.M. Stolker, J. van Schoonhoven, A.J. de Vries, I. Bobeldijk-Pastorova, W.H.J. Vaes, and R. van den Berg, J. Chromatogr. A 1058(1–2), 143–151 (2004).
  14. S. Kneisel, M. Speck, B. Moosmann, T.M. Corneillie, N.G. Butlin, and V. Auwarter, Anal. Bioanal. Chem. 405(14), 4691–4706 (2013).
  15. K. Scheidweiler, M.Y. Jarvis, and M. Huestis, Anal. Bioanal. Chem. Published on-line: September 16, 2014, 1–15 (2014).
  16. J.P. Danaceau, E.E. Chambers, and K.J. Fountain, "Analysis of Synthetic Cannabinoids from Urine for Forensic Toxicology using Oasis HLB μElution plates and CORTECS UPLC Columns," Waters Corporation, 2013, http://www.waters.com/waters/library.htm?lid=134763848&lset=1&locale=en_CA, consulted 09/22/2014.
  • Twitter
  • Facebook
  • Linkedin
  • Rss