Comprehensive Profiling of Phytocannabinoids and Semisynthetic Cannabinoids in Seized Materials from Northern Italy

Authors: Sara Casati ⚬, Roberta F. Bergamaschi, Lea Sicuro, Alessandro Ravelli, Sofia Vanerio, Valentina Orsi, Camilla Ronco, Enrica Torretta, Alberto Rimoldi, Gabriella Roda, Alessio Battistini, Luca Mollica ⚬, Paola Rota ⚬

Affiliations: † Laboratorio di Tossicologia Forense, Dipartimento di Scienze Biomediche, Chirurgiche e Odontoiatriche, 9304Università degli Studi di Milano, via Luigi Mangiagalli 37, 20133 Milan, Italy; ‡ 189821Fondazione IRCCS Ca’ Granda Ospedale Maggiore Policlinico, via Francesco Sforza 35, Milan 20135, Italy; § Dipartimento di Biotecnologie Mediche e Medicina Traslazionale, 70301Università degli Studi di Milano, via Fratelli Cervi 93, Segrate 20054, Italy; ∥ Dipartimento di Scienze Farmaceutiche, 9304Università degli Studi di Milano, via Trentacoste 2, Milan 20134 Italy; ⊥ Dipartimento di Scienze Biomediche, Chirurgiche e Odontoiatriche, 9304Università degli Studi di Milano, via Della Commenda 10, Milan 20121, Italy; # Institute for Molecular and Translational Cardiology (IMTC), San Donato Milanese, Milan 20097, Italy

License: © 2026 The Authors. Published by American Chemical Society CC BY 4.0 This article is licensed under CC-BY 4.0

Article links: DOI: 10.1021/acsomega.6c00941  |  PMC: PMC13261594

Relevance: Relevant: mentioned in keywords or abstract

Full text: PDF (2.1 MB)

Introduction

Cannabis sativa L. produces a chemically diverse array of phytocannabinoids, with more than 150 compounds identified to date.ref. ref1 The term phytocannabinoids refers to cannabinoids naturally occurring inCannabis sativa, whereas semisynthetic cannabinoids (SSCs) denote compounds produced through chemical modification of these natural constituents (e.g., hydrogenation, acetylation, or structural derivatization). The major acidic phytocannabinoids, Δ⁹-tetrahydrocannabinolic acid (Δ⁹-THCA) and cannabidiolic acid (CBDA), undergo nonenzymatic decarboxylation during drying or heating, yielding Δ⁹-tetrahydrocannabinol (Δ⁹-THC) and cannabidiol (CBD), respectively (Figure ). While Δ⁹-THC, the principal psychoactive constituent ofCannabis, largely defines the legal status ofCannabis-derived products, CBD is nonpsychoactive and has been extensively investigated for its potential therapeutic properties.ref. ref2

In recent years, CBD has also emerged as a key chemical precursor for the synthesis of SSCs. More generally, SSCs obtained through the chemical modification of CBD and/or Δ⁹-THC have increasingly been used as substitutes for Δ⁹-THC in commercial and illicit products, partly to circumvent regulatory controls focused on Δ⁹-THC.

Among these, Δ⁸-tetrahydrocannabinol (Δ⁸-THC) was one of the first compounds to gain widespread attention.ref. ref3 Although Δ⁸-THC occurs naturally only in trace amounts in agedCannabis,ref. ref4 it can be readily produced from CBD via acid-catalyzed cyclization reactions, yielding mixtures of Δ⁸– and Δ⁹-THC.ref5,ref6

Since 2019, a rapidly expanding range of SSCs has been detected in treated herbal material, resins, vape liquids, and edibles.

HHC is most commonly encountered as a synthetic product,ref. ref7 although trace amounts have been detected inCannabis plants,ref8−ref9ref10 where it is thought to arise as a degradation product of Δ⁹-THC,ref. ref9 and it typically occurs as a mixture of 9R-HHC and 9S-HHC epimers,ref7,ref11 with epimeric ratios depending on the cannabinoid precursor used in its synthesis.

More recently, the heptyl homologues of HHC, collectively referred to as hexahydrocannabiphorol (HHCP), have emerged on the market.ref. ref12 Structurally, HHCP shares features with both Δ⁸– or Δ⁹– THCP and HHC, combining an extended alkyl side chain with a hydrogenated cannabinoid scaffold.

A further emerging trend involves the marketing of acetylated derivatives of previously known cannabinoids (reported in blue in Figure ). In particular, THCOAc analogues have been reported for Δ⁸– and Δ⁹-THC, as well as an acetylated derivative of CBD, commonly referred to as cannabidiol diacetate (CBD2OAc) and for HHC and HHCP, known as HHCOAc and HHCPOAc, respectively. These compounds are generally obtained via acetylation of the corresponding parent cannabinoids using acetic anhydride, acetyl chloride, or other conventional acetylating reagents.ref13,ref14

To the best of our knowledge, fully detailed and reproducible synthetic routes for HHCOAc and HHCPOAc have not been reported, although early literature describes related acetylation procedures without complete experimental details;ref. ref10 however, these compounds are presumed to be prepared by acetylation of their corresponding nonacetylated precursors using approaches analogous to those employed for the preparation of THC acetates.

This rapid diversification of SSCs highlights the urgent need for selective analytical methods capable of distinguishing closely related isomers, epimers, and homologues in seized materials, as also evidenced by the recent works of Hundertmark et al.,ref. ref15 Tanaka et al.,ref. ref14 and Holt et al.ref. ref16 Such structural modifications are pharmacologically relevant, as structure–activity relationship studies indicate that relatively small changes, including alkyl side-chain extension or acetylation, can markedly influence cannabinoid receptor interactions.ref7,ref17,ref18 Cannabinoid receptor type 1 (CB1) is the primary receptor responsible for the psychoactive effects of THC and cannabinoid receptor type 2 (CB2) is associated with an immunomodulatory role.ref. ref19 In particular, extension of the alkyl side chain generally increases CB1 receptor potency,ref19−ref20ref21 while the 9R epimers of hydrogenated cannabinoids exhibit higher affinity and cannabimimetic activity than their 9S counterparts.ref. ref7 Despite anecdotal early reports on certain acetylated cannabinoids,ref. ref22 reliable contemporary pharmacological and toxicological data for many SSCs remain scarce.

In response to these challenges, in the present study a liquid chromatography–tandem mass spectrometry (LC-MS/MS) method was developed and validated for the identification and quantification of 31 SSCs and major phytocannabinoids inCannabis –derived products. The initial identification of Δ⁸-THCOAc, Δ⁹-THCOAc, 9R,S-HHCOAc, and cannabinol acetate (CBNOAc) was performed by using in-house synthesized reference material. The validated method was successfully applied to 151Cannabis-derived products seized in Northern Italy between 2024 and 2025. In addition, molecular docking simulations were performed on cannabinoid receptors CB1 and CB2 to provide a structure-based framework for interpreting how acetylation and side-chain modification influence ligand–receptor interactions. Docking results support an increased CB1 binding propensity for acetylated and side-chain-extended derivatives in agreement with the previously documented pharmacological activity of the investigated compounds.

Materials and Methods

Chemicals and Reagents

Ultrapure water (H₂O), acetonitrile (ACN), ethanol (EtOH), and methanol (MeOH) were of analytical grade and purchased from Carlo Erba (Milan, Italy). Formic acid (98%–100%) was purchased from Merck (Darmstadt, Germany). Δ⁸-THCOAc, Δ⁹-THCOAc, 9R– and 9S-HHCOAc, and CBNOAc were initially custom-synthesized by our laboratory as described in paragraph 2.2. The reference materials cannabichromene (CBC) (1 mg/mL in MeOH), cannabidivarin (CBDV) (1 mg/mL in MeOH), cannabigerol (CBG) (1 mg/mL in MeOH), cannabinol (CBN) (1 mg/mL in MeOH), Δ⁸-THC (1 mg/mL in MeOH), Δ⁸-THCP (1 mg/mL in MeOH), and Δ⁹-THC (1 mg/mL in MeOH) were purchased from Merck (Darmstadt, Germany). CBD (10 mg/mL in MeOH) was purchased from PhytoLab GmbH & Co. KG (Vestenbergsgreuth, Germany). CBD2OAc (5 mg/mL in ACN), Δ⁹-THCV (1 mg/mL in MeOH), Δ⁸-THCOAc (1 mg/mL in ACN), Δ⁸-THCA (1 mg/mL in ACN), Δ⁸-Tetrahydrocannabiphorol acetate (Δ⁸-THCPOAc) (10 mg/mL in ACN), CBNOAc (1 mg/mL in ACN), Δ⁹-THCP (1 mg/mL in ACN) were purchased from Cayman (Ann Arbor, Michigan, USA). Cannabinoids Acids 7 Standard (cannabichromenic acid (CBCA), CBDA, cannabidivarinic acid (CBDVA), cannabigerolic acid (CBGA), cannabinolic acid (CBNA), Δ⁹-THCA, and Δ⁹-tetrahydrocannabivarinic acid (Δ⁹-THCVA) at the concentration of 1 mg/mL in ACN) was purchased from Restek (Bellefonte, Pennsylvania, USA). 9R-HHC (0.1 mg/mL in MeOH), 9R-HHCOAc (0.1 mg/mL in MeOH), 9R-HHCP (0.1 mg/mL in MeOH), 9R-HHCPOAc (0.1 mg/mL in MeOH), 9S-HHC (0.1 mg/mL in MeOH), 9S-HHCOAc (0.1 mg/mL in MeOH), 9S-HHCP (0.1 mg/mL in MeOH), and 9S-HHCPOAc (0.1 mg/mL in MeOH) were obtained from Comedical (Trento, Italy).

The internal standards (IS) CBD-d ₃ (0.1 mg/mL in MeOH), Δ⁹-THC-d ₃ (0.1 mg/mL in MeOH), and Δ⁹-THCA-d ₃ (0.1 mg/mL in ACN) were purchased from Merck (Darmstadt, Germany).

Synthesis of Cannabinoid Acetates

Chemistry

All chemicals and solvents used were of analytical grade and purchased from Sigma-Aldrich (St. Louis, MO). Solvents were dried using standard methods and distilled before use. The progress of all reactions was monitored by thin-layer chromatography (TLC) carried out on 0.25 mm Sigma-Aldrich silica gel plates (60 F254) using UV light, anisaldehyde/H₂SO₄/EtOH solution. Flash chromatography was performed with normal-phase silica gel (Sigma-Aldrich 230–400 mesh silica gel). Nuclear magnetic resonance spectra were recorded at 298 K on a Bruker AM-500 spectrometer equipped with a 5 mm inverse-geometry broadband probe and operating at 500.13 MHz for ¹H and 125.76 MHz for ¹³C. The ¹H and ¹³C resonances were assigned by ¹H–¹H (COSY) and ¹H–¹³C (HSQC and HMBC) correlation of two-dimensional (2D) experiments. Chemical shifts are reported in parts per million and are referenced for ¹H spectra to a solvent residue proton signal (δ = 7.26 ppm for CDCl₃) and for ¹³C spectra, to the solvent carbon signal (central line at δ = 77.00 ppm, for CDCl₃). The ¹H NMR data are tabulated in the following order: multiplicity (s = singlet, d = doublet, t = triplet, br = broad, m = multiplet, app = apparent), coupling constants are given in Hz, number of protons, and assignment of proton(s). In this context, the symbols α and β refer exclusively to substituents resonating at higher and lower chemical shift values (ppm), respectively, and do not indicate stereochemical configuration.

General Acetylation Procedure

Starting from 1 mg of pure or purified starting material, the sample was diluted in CH₂Cl₂ (0.2 mL) and then treated with Et₃N (0.05 mL) and Ac₂O (0.025 mL). The reaction was stirred at 50 °C and monitored by TLC (Hexane/AcOEt 9:1, v:v). After 1 h, the reaction was quenched by the addition of MeOH (0.2 mL) and the solvent was removed under vacuum. The crude product was purified by flash chromatography (Hexane/AcOEt 95:5, v:v).

Acetylation of Diastereomeric Mixture of 9R-HHC and 9S-HHC

¹H-NMR assignments for the diastereomeric mixture of 9R-HHC and 9S-HHC acetates refer to the structure numbers in Figure :

Starting from 1 mg of a purified diastereomeric mixture (7:3 ratio, 9R:9S) of 9R-HHC and 9S-HHC, and applying the general acetylation procedure, the desired acetylated mixture of 9R-HHCOAc and 9S-HHCOAc was achieved in high yield (1 mg, 89% and 7:3 diastereomeric ratio). ¹H NMR analysis of the 9R-HHCOAc and 9S-HHCOAc mixture showed: ¹H NMR (500 MHz, CDCl₃) δ 6.53–6.51 (br s app, overlapping, H-4), 6.37–6.35 (br s app, overlapping, H-2), 2.60–2.29 (overlapping, Hα-10, H-10a, and 2H at C1′), 2.28 and 2.27 (2 × OCOCH₃), 1.88–0.98 (overlapping, 2H at C7, 2H at C8, H-9, 2H at C2′, 2H at C3′, 2H at C4′, Hβ-10, and CH₃ at C9), 1.45 (overlapping, 6a), 1.36 (overlapping, CH₃α at C6), 1.06 (overlapping, CH₃β at C6), 0.94 (d, J = 6.6 Hz, CH₃ at C9), 0.88 (overlapping, t app, J = 6.7 Hz, H-5′), 0.80 (q app, J = 12.9 Hz, Hβ-10).

Acetylation of Δ⁸-THC

¹H and ¹³C assignments for Δ⁸-THCOAc refer to the structure numbers in Figure :

Starting from 1 mg of Δ⁸-THC and applying the general acetylation procedure, the desired Δ⁸-THCOAc was achieved in high yield (1 mg, 88%). ¹H NMR and ¹³C NMR analysis of Δ⁸-THCOAc showed: ¹H NMR (500 MHz, CDCl₃) δ 6.56 (br d, J = 1.4 Hz, 1H, H-4), 6.40 (br d, J = 1.4 Hz, 1H, H-2), 5.42 (s app, 1H, H-8), 2.72 (dd, 1H, J = 4.4, J = 17.1 Hz, Hα-10), 2.59 (ddd, J = 4.4, J = 11.3, J = 12.9 Hz, 1H, H-10a), 2.50 (m, 2H, 2H at C1′), 2.28 (s, 3H, OCOCH₃), 2.12 (m, 1H, Hα-7), 1.91 (m, 1H, Hβ-10), 1.84–1.72 (overlapping, 2H, Hβ-7, and H-6a), 1.68 (s, 3H, CH₃ at C9), 1.62–1.50 (overlapping to water signal, 2H, 2H at C2′), 1.37 (s, 3H, CH₃α at C6), 1.35–1.22 (overlapping, 4H, 2H at C3′ and 2H at C4′), 1.09 (s, 3H, CH₃β at C6), 0.87 (t, J = 6.9 Hz, 3H, H-5′). ¹³C NMR (125 MHz, CDCl₃) δ 169.0 (OCOCH₃), 154.5 (C5), 142.8 (C1), 133.9 (C3), 122.7 (C9), 119.7 (C8), 116.0 (C11), 115.3 (C4), 114.3 (C2), 77.7 (overlapping to solvent signal, C6), 44.6 (C6a), 36.1 (C10), 35.3 (C1′), 31.7, 31.5 (C2′ and C3′), 30.5 (C10a), 27.7 (C7), 27.4 (CH₃α at C6), 23.6 (CH₃ at C9), 22.5 (C4′), 21.3 (OCOCH₃), 18.5 (CH₃β at C6), 14.0 (C5′).

Acetylation of Δ⁹-THC

¹H and ¹³C assignments for Δ⁹-THCOAc refer to the structure numbers in Figure :

Starting from 1 mg of Δ⁹-THC and applying the general acetylation procedure, the desired acetylated Δ⁹-THCOAc was achieved in high yield (1 mg, 88%). ¹H NMR and ¹³C NMR of Δ⁹-THCOAc analysis showed: ¹H NMR (500 MHz, CDCl₃) δ 6.54 (br d, J = 1.4 Hz, 1H, H-4), 6.4 (br d, J = 1.4 Hz, 1H, H-2), 5.97 (s app, 1H, H-10), 3.06 (br d, J = 10.1 Hz, 1H, H-10a), 2.52–2.45 (overlapping, 2H, 2H at C1′), 2.28 (s, 3H, OCOCH₃), 2.16–2.09 (overlapping, 2H, 2H at C8), 1.89 (m, 1H, Hα at C7), 1.69–1.62 (overlapping, 4H, CH₃ at C9 and H-6a), 1.61–1.52 (overlapping to water signal, 2H, 2H at C2′), 1.40 (s, 3H, CH₃α at C6), 1.39–1.20 (overlapping, 5H, 2H at C3′, 2H at C4′ and Hβ at C7), 1.08 (s, 3H, CH₃β at C6), 0.87 (t, J = 6.9 Hz, 3H, 3H at C5′).¹³C NMR (125 MHz, CDCl₃) δ 168.9 (OCOCH₃), 154.5 (C5), 149.3 (C1), 142.8 (C3), 134.9 (C9), 123.1 (C10), 115.3 (C4), 115.1 (C11), 114.0 (C2), 77.5 (overlapping to solvent signal C6), 45.6 (C6a), 35.4 (C1′), 34.1 (C10a), 31.5 (C3′), 31.1 (C8), 30.6 (C2′), 27.5 (CH₃α at C6), 24.9 (C7), 23.5 (CH₃ at C9), 22.6 (C4′), 21.3 (OCOCH₃), 19.4 (CH₃β at C6), 14.0 (C5′).

Acetylation of CBN

¹H and ¹³C assignments for CBNOAc refer to the structure numbers in Figure :

Starting from 1 mg of CBN and applying the general acetylation procedure, the desired CBNOAc was achieved in high yield (1 mg, 88%). ¹H NMR and ¹³C NMR of CBNOAc: ¹H NMR (500 MHz, CDCl₃) δ 7.80 (s, 1H, H-10), 7.14 (br d, J = 7.9 Hz, 1H, H-7), 7.83 (br dd, J = 1.1 Hz, J = 7.9 Hz, 1H, H-8), 6.72 (br d, J = 1.7 Hz, 1H, H-4), 6.40 (br d, J = 1.7 Hz, 1H, H-2), 2.59–2.53, 2.36 (overlapping, 2H, 2H at C1′), 2.36 (s, 3H, CH₃ at C9), 2.32 (s, 3H, OCOCH₃), 1.66–1.57 (overlapping, 8H, 2H at C2′, CH₃α at C6 and CH₃β at C6), 1.37–1.28 (overlapping, 4H, 2H at C3′ and 2H at C4′), 0.89 (t, J = 6.9 Hz, 3H, H-5′). ¹³C NMR (125 MHz, CDCl₃) δ 169.1 (OCOCH₃), 154.3 (C5), 147.3 (C1), 144.5 (C3), 137.5 (C10a), 136.8 (C6a), 128.3 (C8), 126.8 (C9), 125.6 (C10), 122.8 (C7), 116.3 (C2), 115.8 (C4), 114.2 (C11), 77.7 (overlapping to solvent signal, C6), 35.5 (C1′), 31.4 (C3′), 30.4 (C2′), 29.7 (CH₃α at C6), 26.9 (CH₃β at C6), 22.5 (CH₂ at C4′), 21.5 (CH₃ at C9), 21.4 (COCH₃), 14.0 (C5′).

Standard Solutions, Calibrators, and Quality Control Samples

All the solutions were stored in the dark at −20 °C. Working solutions were prepared from stock solutions (0.1 or 0.01 mg/mL) by appropriate dilution in MeOH and used for the preparation of calibration curves and quality control (QC) samples at 1 μg/mL and 100 ng/mL for all compounds, and at 1 μg/mL for ISs (CBD-d ₃, Δ⁹-THC-d ₃, and Δ⁹-THCA-d ₃).

Calibration standards containing:were freshly prepared for each analytical batch by adding suitable volumes of working solutions to 1 mL of MeOH.

• 0, 0.5, 1, 2.5, 5, 10, 25 ng/mL (corresponding to 0, 1, 2.5, 5, 10, 25, 50% since 50 ng/mL was set as 100%) for Δ⁸-THC, Δ⁹-THC, Δ⁸-THCA, Δ⁹-THCA;
• 0, 1, 2.5, 5, 10, 25 ng/mL (corresponding to 0, 0.1, 0.25, 0.5, 1, 2.5% since 1 μg/mL was set as 100%) for CBC, CBCA, CBD, CBDA, CBDV, CBDVA, CBG, CBGA, CBN, CBNA, Δ⁹-THCV, Δ⁹-THCVA, Δ⁹-THCOAc, 9R– and 9S-HHC;
• 0, 1, 2.5, 5, 10, 25, 50 ng/mL (corresponding to 0, 0.001, 0.0025, 0.005, 0.01, 0.025, 0.05% since 100 μg/mL was set as 100%) for CBD2OAc, CBNOAc, Δ⁸-THCOAc, Δ⁸-THCP, Δ⁸-THCPOAc, Δ⁹-THCP, 9R-HHCOAc, 9R-HHCP, 9R-HHCPOAc, 9S-HHCOAc, 9S-HHCP, and 9S-HHCPOAc;
• 10 ng/mL for CBD-d ₃, Δ⁹-THC-d ₃, and Δ⁹-THCA-d ₃;

Collection and Preparation of Cannabis and Cannabis-Derived Products Samples

An aliquot of 25 mg from each flower (n = 69), resin (n = 69), wax (n = 3), vape-liquid (n = 9), and powder (n = 1) sample was extracted with 2.5 mL of cyclohexane and mixed for 15 min at room temperature. 10 μL aliquot of the initial solution (S1) was added to 990 μL of MeOH to obtain a second diluted solution (S2). A third solution (S3) was prepared by diluting a 10 μL aliquot of the S2 in 990 μL of MeOH.

For LC-MS/MS analysis, each sample was prepared as follows:

– 10 μL aliquot of S2 was added to 10 μL of IS and 980 μL of MeOH (final dilution 1:10000);

– 50 μL aliquot of S3 was added to 10 μL of IS and 940 μL of MeOH (final dilution 1:200000);

– 5 μL aliquot of S3 was added to 10 μL of IS and 985 μL of MeOH (final dilution 1:2000000).

LC-MS/MS Analysis

Analyses were performed on a 1290 Infinity UHPLC system (Agilent Technologies, Palo Alto, CA, USA) coupled to a QTrap 5500 triple quadrupole linear ion trap mass spectrometer (Sciex, Darmstadt, Germany). Compounds were separated on a Kinetex HPLC XB-C18 column (100 mm × 2.1 mm i.d., 2.6 μm) (Phenomenex, CA, USA) using 0.1% formic acid in water (mobile phase A) and 0.1% formic acid in acetonitrile (mobile phase B).

The 17 min linear gradient was the following: from 0.00 to 4.80 min solvent B increased from 70% to 75% and held 75% from 4.80 to 7.00 min, then solvent B increased to 82% from 7.00 to 12.00 min, and to 90% from 12.00 to 12.10 min, then held at 90% to 15.00 min, decreased to 70% from 15.00 to 15.10 min, and finally held at 70% from 15.10 to 17.00 min for re-equilibration.

The flow rate was 0.30 mL/min and the injection volume was 3 μL. The column thermostatic oven was kept at 40 °C. The working conditions and parameters of the MS were optimized as follows: the ion source was ESI, operating in positive mode; the resolution of the precursor ion selector (Q-1) and product ion selector (Q-3) was 0.7 ± 0.1 amu; the curtain gas, ion source gas 1, and ion source gas 2 were set at 25, 45, and 10 psi, respectively; the source temperature was 550 °C; the ionization voltage was 5500 eV; and the entrance potential was 10 eV. MS acquisition was performed by Multiple Reaction Monitoring (MRM) mode. The MRM conditions and parameters including ion transitions, declustering potential (DP), and relative collision energy (CE) are provided in Table . The data acquisition and processing were performed using Analyst1.6.2 and MultiQuant2.1.1 software (Sciex, Darmstadt, Germany), respectively.

Validation Procedure

The method was fully validated according to the international recommendations for the validation of a new analytical method in forensic toxicology.ref. ref23 Selectivity was evaluated using blank matrices, when available, and in the absence of a suitable blank matrix, by a standard-addition procedure at low, medium, and high concentration levels. Six-point calibration curves for each compound, with eight replicates per level, were generated from the peak area ratios of the analytes to the IS versus nominal analyte concentration, including zero as a calibration level without forcing the intercept through the origin. Linearity was assessed by ordinary least-squares regression and verified by lack-of-fit analysis of variance (ANOVA) using replicated calibration levels. Sensitivity was expressed in terms of LOD (Limit of Detection) and LOQ (Limit of Quantification). The LOQ was determined as the lowest concentration with values for precision and accuracy within ±20% and a signal-to-noise (S/N) ratio of the peak areas ≥10, whereas the LOD was defined as the lowest concentration with a S/N of the peak areas ≥3. Precision and accuracy of the method were determined through the analysis of duplicate Quality Control (QC) samples at low, medium, and high concentration levels in eight different days for each compound according to their calibration ranges. Precision and accuracy were determined by calculating the coefficient of variation (CV%) and the Bias% of the QC samples and they were considered satisfactory if Bias was ± 15% of the nominal value (20% near LOQ); precision was within 15% RSD (20% near LOQ). The stability of the analytes at low and high concentration levels was assessed by comparing the mass spectrometric responses obtained after repeated injections of calibrators stored under different conditions: at room temperature for 24 h, at 4 °C for 48 h, and at −20 °C for 48 h, 15 days, and 30 days. The matrix effect was evaluated by fortifying each matrix with standard analyte solutions at the LOQ and at three additional concentration levels (low, medium, and high), and comparing the results with those obtained in spiked solvent after subtraction of the endogenous contribution of each analyte. The matrix effect was considered satisfactory if it was <20%, calculated as the percentage difference between the signal in the spiked matrix (blank-corrected) and the signal in the spiked solvent. Extraction recovery was indirectly assessed through extraction repeatability of Δ⁹-THC, CBD, and Δ⁹-THCA by performing six independent extractions (CV% < 20%).

Molecular Structures

The structures of cannabinoid receptors CB1 and CB2 were obtained from the Protein Data Bank (https://www.rcsb.org/) in complex with agonists (PDB: 5XRA ref. ref24 and 5ZTY,ref. ref25 respectively). The structures of Δ⁹-THC and 9S-HHC were obtained from the NIH PubChem repository (https://pubchem.ncbi.nlm.nih.gov/), while the structures of Δ⁸-THC, Δ⁸-THCOAc, Δ⁸-THCP, Δ⁸-THCPOAc, Δ⁹-THCOAc, Δ⁹-THCP, Δ⁹-THCPOAc, 9S-HHCOAc, 9S-HHCP, 9S-HHCPOAc, 9R-HHC, 9R-HHCOAc, 9R-HHCP, and 9R-HHCPOAc were manually edited starting from the downloaded structures using PyMOL.ref. ref26

Molecular Protein–Ligand Docking

Docking simulations were conducted using AutoDock 4.2.6, a software suite designed to predict interactions between macromolecules and small ligands.ref. ref27 The atomic charges, torsional flexibility, and protonation states of the enzyme and of the ligands were set up using, respectively, the AutoDock Tools graphical interface and Meeko (https://meeko.readthedocs.io). The ligand binding site for performing the docking calculation was identified on the basis of the ligand’s position in the original PDB structure, i.e., its geometric center was used to center the docking box for all the systems considered in the present work. The docking grid was set to 50 Å × 50 Å × 50 Å along the x, y, and z axes, with a grid spacing of 0.403 Å. Docking simulations employed a genetic algorithm, with each configuration file corresponding to a single docking run, generating up to 100 possible binding poses. These poses were then statistically analyzed based on their predicted dissociation constants (K _i) expressed in terms of ligand concentration.

Results and Discussion

Cannabinoid Acetates Synthesis

At the time this study began, authentic reference standards of major cannabinoid acetates, including acetylated Δ⁸-THC, Δ⁹-THC, and HHC epimers, were not commercially available. Therefore, these compounds were synthesized in accordance with the procedures described in the Materials and Methods section (Section sec2.2 ). Unlike earlier studies,ref15,ref16 where acetylation of cannabinoids was primarily used as a simple derivatization step without purification, the methodology adopted in this work included isolation of the acetylated products via flash column chromatography, followed by full structural characterization by NMR spectroscopy. This purification step was critical to remove residual reagents and byproducts, ensuring sufficient purity for comprehensive spectroscopic analysis.

Acetylation was carried out using acetic anhydride and triethylamine in dichloromethane, starting from milligram-scale quantities of pure precursors. TLC monitoring indicated complete conversion within 1 h at 50 °C, with no significant degradation or side reactions observed. Purification by silica gel flash chromatography using a solvent mixture of hexane/ethyl acetate yielded clean fractions of the acetylated products. Notably, the acetylation of the diastereomeric mixture of 9R– and 9S-HHC, affording the corresponding acetates, proceeded without epimerization at C-9, as confirmed by the retention of the original diastereomeric ratio.

The successful formation of the acetate esters was clearly supported by diagnostic NMR signals in CDCl₃ (Supporting Information S1–S4). In the ¹H NMR spectra, the appearance of a characteristic singlet corresponding to the methyl group of the acetate moiety (δ ≈ 2.28–2.32 ppm, 3H) confirmed acetylation of the phenolic hydroxyl group. Moreover, the disappearance of the phenolic OH signal observed in the starting materials further supported complete esterification. In the ¹³C NMR spectra, the presence of the ester carbonyl resonance (δ ≈ 169 ppm) together with the acetate methyl carbon signal (δ ≈ 20–21 ppm) provided additional confirmation of acetate formation.

Comprehensive structural characterization was achieved using both one- and two-dimensional NMR experiments (COSY, HSQC, and HMBC), allowing unambiguous assignment of all proton and carbon resonances. Overall, the synthetic and analytical strategy described herein provides purified and fully characterized cannabinoid acetate standards, addressing the limitations of reports that relied on unpurified derivatization products or incomplete spectroscopic data.

Method Validation

The LC-MS/MS method was developed and fully validated for the identification and quantitation of phytocannabinoids and SSCs as follows.

Chromatographic Separation

In recent years, several techniques for the analysis ofCannabis-derived products have increasingly been tested. The two most common analytical methods are based on gasref11,ref28−ref29ref30 and liquidref31,ref32 chromatography (GC and LC); however, high temperatures in the GC injection port turn carboxylate forms and, partially, acetylated forms (as shown in Supporting Information Figure S5) into their respective nonacidic or nonester forms, thus in the analytical result the specific contribution of each of them remains unknown. As a result, the detection and the quantitation of Δ⁹-THC do not necessarily imply that it was already present in the original sample but rather derived from the conversion of the carboxylate precursor Δ⁹-THCA and/or Δ⁹-THCOAc. To determine Δ⁹-THC and Δ⁹-THCA separately, a derivatization step prior to GC analysis is needed. Moreover, Cheng and Kerrigan suggested that GC liner and inlet conditions can influence the in situ formation of Δ⁹-THC from CBD cyclization.ref. ref33 Contrarily, LC is the most effective and versatile technique for the analysis of complex mixtures of cannabinoids without derivatization, since it prevents decarboxylation of acidic forms, and it allows a separate identification and quantification of the neutral forms and their acidic or ester precursors. However, we found that a minor quantity of the acetylated form was de-esterified in the ion source of the LC-MS/MS system, when its temperature was set around 500 °C. We demonstrated that the acetylated forms that were converted in nonester forms were directly proportional to their initial concentration (Supporting Information Figure S6). Moreover, since the conversion occurs in the ion source, after the chromatographic separation, contrary to GC analysis, the contribution of the Δ⁹-THCOAc-derived Δ⁹-THC shows a different RT rather than the original Δ⁹-THC. Numerous studies, including those reviewed by La Maida et al.,ref. ref34 have explored the chromatographic analysis of Δ⁹-THC, Δ⁸-THC, and other THC isomers across diverse matrices, noting their highly similar behavior. GC-MS effectively differentiates various THC isomers, while 9R– and 9S-HHC epimers separate readily via TLC, GC (native or TMS-derivatized), or LCref13,ref35,ref36 . Holt et al.ref. ref16 quantified Δ⁸-, Δ⁹-, and Δ¹⁰-THCOAc in cannabis products using GC-MS, and Tanaka et al.ref. ref14 achieved partial Δ⁸– and Δ⁹-THCOAc separation by LC-PDA-MS. To our knowledge, this study introduces the first validated LC-MS/MS method that satisfactorily resolves, identifies, and quantifies multiple epimeric and isomeric SSC formsincluding Δ⁸– and Δ⁹-THCOAc, Δ⁸– and Δ⁹-THCPOAc, 9R– and 9S-HHCOAc, and 9R– and 9S-HHCPOAc. Chromatographic separation is illustrated in Figure , achieved on a C18 column with an ACN-based mobile phase. A 17 min step gradient ensured baseline resolution of early-eluting cannabinoid acids, neutral cannabinoids, and late-eluting acetylated derivatives, with no methanol-related interferences observed.

Selectivity, Linearity, LOD, and LOQ

Across all analytes and matrices, no additional coeluting peaks were observed. When a blank matrix was unavailable, the chromatographic peaks of endogenous and spiked analytes showed comparable retention times and ion ratios. For each analyte, the ordinary linear model showed no statistically significant lack of fit over the investigated range (p > 0.05). The mean %CV, LOD, and LOQ are listed in Table .

Inter-Day Precision and Accuracy

The method showed good performance, with CV values ≤ 15% (≤20% at the LOQ) for precision and bias ≤ 15.6% for accuracy. Precision and accuracy data for all analytes are reported in Table .

Stability

Calibrators stability was assessed by monitoring the percentage deviation of each analyte in the vial over time. After 24 h of storage at room temperature, deviations ranged from 1.00% to 15.68%, except for 9R- and 9S-HHCPOAc (<30%). After 48 h, deviations ranged from 2.45% to 15.92% at 4 °C (except for 9R and 9S-HHCPOAc (<40%)) and from 3.13% to 15.54% at −20 °C. Long-term calibrator stability was assessed exclusively at −20 °C obtaining a deviation ranging from 2.43% to 15.57% after 15 days and from 5.16% to 15.78% after 30 days.

Matrix Effect and Extraction Repeatability

Across all analytes and matrices, the matrix effect was within the predefined acceptance limits, ranging from 0–22% at the LOQ and low level, 0–19% at the medium level, and 0–16% at the high level, thus confirming the suitability of solvent-based calibration for quantitative purposes. Regarding extraction repeatability, the coefficients of variation for Δ⁹-THC, CBD, and Δ⁹-THCA obtained from six replicate extractions were consistently below 20% for all matrices, indicating acceptable and reproducible extraction performance. Considering the close structural similarity among the investigated analytes, these percentage values can reasonably be assumed to be representative for all compounds included in the method.

Analysis of SeizedCannabis –Derived Samples

The validated LC-MS/MS method was subsequently applied to characterize phytocannabinoid SSCs in 151 cannabis-derived products seized between 2024 and 2025 in Northern Italy. The samples primarily consisted of cannabis flowers (45.7%, n = 69) and resins (45.7%, n = 69), while vape liquids (6.0%, n = 9), waxes (2.0%, n = 3), and powder (0.7%, n = 1) represented a smaller proportion.

An overview of the frequency of detection of all cannabinoids is reported in Table , which summarizes the complete quantitative data set. Δ⁹-THC and Δ⁹-THCA were detected in almost all samples, while Δ⁹-THCOAc (85%), Δ⁹-THCP (61%), Δ⁸-THC (18%), and Δ⁸-THCOAc (6%) were the most frequently identified SSCs.

Cannabis Flowers

Δ⁹-THCA was detected in 100% of flower samples, while Δ⁹-THC was also present in nearly all samples; concentrations ranged from 0.05% to 37.86% (mean: 12.89%; median: 11.37%) for Δ⁹-THCA and 0.13–14.11% (mean: 3.73%; median: 3.07%) for Δ⁹-THC and overall Δ⁹-THCA exceeded Δ⁹-THC in 90% of samples, confirming that acidic cannabinoids predominate in plant material.

Δ⁹-THCOAc was detected in 84% of samples at trace levels (range: LOQ–0.008%; median: 0.0004%). Δ⁹-THCP was also frequently observed at low concentrations (range: LOQ–0.03%; median: 0.0009%).

Δ⁸-THC was not detected in flower samples; however, Δ⁸-THCOAc was identified in two samples (0.02%), which also contained multiple hydrogenated cannabinoids (HHCs and derivatives), suggesting possible exogenous addition.

Resins

Δ⁹-THCA and Δ⁹-THC were detected in 99% of resin samples with Δ⁹-THCA ranging from 0.09% to 48.14% (mean: 14.27%; median: 10.79%), and Δ⁹-THC from 0.17% to 26.04% (mean: 10.74%; median: 10.17%); Δ⁹-THCA exceeded Δ⁹-THC in 58% of samples, whereas the opposite trend was observed in 42%, indicating partial decarboxylation or processing and Δ⁸-THC was frequently associated with samples where Δ⁹-THC > Δ⁹-THCA, being quantified above the LOQ in 18 samples (mean ∼3%), suggesting potential manipulation or addition of a synthetic product, although trace Δ⁸-THC was also detected in additional samples.

Δ⁹-THCOAc was highly prevalent (range: LOQ–6.99%; median: 0.0008%), supporting the hypothesis of either natural occurrence at trace levels or intentional addition at higher concentrations, and Δ⁹-THCP was also commonly detected at low concentrations (range: LOQ–0.0041%; median: 0.0019%).

Some resin samples exhibited unusually high CBD contents (up to 63.7%), consistent with CBD-rich chemotypes. One such sample also contained significant levels of Δ⁹-THCOAc (7.0%), Δ⁸-THC (2.1%), and Δ⁸-THCOAc (0.6%), while Δ⁹-THC and Δ⁹-THCA were, respectively, 1.3 and 0.1% indicating a complex and possibly altered composition.

Vape Liquids

Δ⁹-THCA was detected in 44% of vape liquids, while Δ⁹-THC was more consistently present; Δ⁸-THC was identified in all nine samples, with concentrations reaching up to 54%.

Δ⁸-THCOAc was detected in six samples (median: 0.001%), and in three cases co-occurred with Δ⁸-THCP > LOQ, suggesting synthetic formulation.

Δ⁹-THCP was also detected at trace levels (range: 0.0001–0.0012%; median: 0.0008%) in 55% of the samples.

Waxes and Powder

None of the wax samples contained SSCs. However, one sample exhibited the highest Δ⁹-THC concentration observed (70.1%), while the remaining two showed extremely high CBD contents (97.5 and 63.8%).

The single powder sample contained Δ⁹-THCA (88% detection), although no further SSCs were identified.

Structure–Activity Relationship of SSCs

We have also investigated the structure–activity relationship (SAR) of SSCs by means of molecular docking models, which have been computed with the aim of evaluating the entity of their binding to CB1 and CB2. CB1 is the main target of the present study, and the latter serves as a control due to their similar fold but with a different THC orientation in the binding pocket. Binding affinity was evaluated through multiple (100) runs of the docking calculations for a series of ligands among the ones presented above and computing the distribution of the dissociation constants (namely K _i) over all the docking poses. This metric reflects the strength of interaction between an inhibitor and its target: a shift of the distributions toward lower K _i values clearly indicates stronger interaction (i.e., a lower K _i corresponds to a lower free Gibbs energy of binding). The distributions of the computed K _i of the derivatives of Δ⁸-THC, Δ⁹-THC, 9R-HHC, and 9S-HHC (Figure A) revealed in all cases an increase of the affinity toward CB1 as a response to the chemical modifications considered in the present work, i.e., an insertion of the acetate group or the hydrophobic side-chain extension. All the investigated compounds showed a dramatic increase in the binding affinity once the acetate group was introduced in the molecule, ranging from halved average K _i values for both THC forms (Δ⁸-THC and Δ⁹-THC) to a 10-fold decrease for HHC, whereas the elongated side-chain had a strong effect mainly on the binding of both THC derivatives, as previously suggested by Citti et al.ref. ref19 (Δ⁹-THCP_Ki = 1.2 versus Δ⁹-THC_Ki = 40). Moreover, this study shows that 9R epimers of HHC and HHCP had stronger binding affinity than their 9S counterparts based on K _i values (Figure A), confirming that small structural modifications can alter pharmacological profiles.ref7,ref12,ref17,ref18,ref37 The role of the acetate group in increasing the stabilization of receptor–ligand interactions can be explained in our models by the formation of hydrogen bonds with neighboring residues His 178 and Ser 383 (Figure B: relevant information is reported only for Δ⁹-THC, for the other systems they can be found in Supporting Information Figures S7 and S8). Ser 383 stably interacts with all the molecules considered in the present study via its Oγ side chain atom (Figure S7), also supporting recently reported resultsref38,ref39 about the role of these molecules as prodrugs upon acetate hydrolysis, i.e., they can efficiently bind to the receptor even in the absence of the acetate moiety and only due to the ubiquitous presence of O1. A role of further stabilization, with respect to molecules without the acetate functional group, can be envisaged for His 178 (Figure S8), as it stabilizes the complex via Nε2–O hydrogen bonding (roughly in 25% of the poses). Aside from hydrogen bonding, an elongated hydrophobic side-chain can more easily be accommodated by the hydrophobic binding pocket of the receptor, formed by residues Phe 268, Ile 271, Tyr 275, Leu 276, and Trp 279 (Figure C). This is in agreement with recently documented experimental findings about an increased potency (EC50) of compounds as a function of the length of the hydrophobic side-chain, i.e., Δ⁹– and Δ⁸-THC homologues with longer alkyl chains possess increased CB1 activation potential.ref. ref40 The same trend in the energetic/inhibitory profiles has been observed for the receptor CB2 (Supporting Information Figure S9), with a more pronounced synergy between the two modifications: the distributions of K _i are broader, i.e., with a significant part of the K _i values higher than 200 nM, suggesting a less specific interaction with respect to the one with CB1.

Conclusions

The synthetic strategy described herein provides purified and fully characterized cannabinoid acetate standards (Δ⁸-THCOAc, Δ⁹-THCOAc, 9R- and 9S-HHCOAc, and CBNOAc), thereby overcoming limitations of earlier reports that relied on unpurified derivatization products or incomplete spectroscopic characterization. The availability of purified and fully characterized cannabinoid acetate standards is particularly important in light of increasing analytical, forensic, and regulatory interest. The developed LC separation successfully resolved multiple epimeric and isomeric cannabinoid structures, and the validated LC-MS/MS method proved effective for the identification and quantification of both phytocannabinoids (including acidic and neutral forms) and SSCs in Cannabis-derived products. This analytical approach enabled the reliable identification and structural characterization of these compounds, contributing to the expanding knowledge base required for their detection and monitoring. The method presented here could therefore be implemented in routine analytical workflows to enable the detection of SSCs beyond those currently under regulatory control. The present study highlights the high prevalence of SSCs in Cannabis-derived products, posing a potential risk to consumers who are often unaware of the actual composition of the products they consume. Indeed, molecular docking analyses suggested a potentially increased binding propensity toward CB1 for acetylated and side-chain-extended derivatives, although these in silico findings require experimental confirmation. Overall, these findings underline the importance of continued analytical surveillance and further pharmacological investigation of newly emerging cannabinoid derivatives.

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