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Journal of Analytical Toxicology logoLink to Journal of Analytical Toxicology
. 2024 Mar 29;48(5):350–358. doi: 10.1093/jat/bkae022

Insights into the human metabolism of hexahydrocannabinol by non-targeted liquid chromatography–high-resolution tandem mass spectrometry

Florian Pitterl 1, Marion Pavlic 2, Jianmei Liu 3, Herbert Oberacher 4,5,*
PMCID: PMC11165647  PMID: 38687640

Abstract

Hexahydrocannabinol (HHC), 6,6,9-trimethyl-3-pentyl-6a,7,8,9,10,10a-hexahydrobenzo[c]chromen-1-ol, is a semi-synthetic cannabinoid that has presented challenges to analytical laboratories due to its emergence and spread in the drug market. The lack of information on human pharmacokinetics hinders the development and application of presumptive and confirmatory tests for reliably detecting HHC consumption. To address this knowledge gap, we report the analytical results obtained from systematic forensic toxicological analysis of body-fluid samples collected from three individuals suspected of drug-impaired driving after HHC consumption. Urine and plasma samples were analyzed using non-targeted liquid chromatography–high-resolution tandem mass spectrometry. The results provided evidence that HHC undergoes biotransformation reactions similar to other well-characterized cannabinoids, such as ∆9-tetrahydrocannabinol or cannabidiol. Notably, HHC itself was only detectable in plasma samples, not in urine samples. The observed Phase I reactions involved oxidation of C11 and the pentyl side chain, leading to corresponding hydroxylated and carboxylic acid species. Additionally, extensive glucuronidation of HHC and its Phase I metabolites was evident.

Introduction

Hexahydrocannabinol (HHC), 6,6,9-trimethyl-3-pentyl-6a,7,8,9,10,10a-hexahydrobenzo[c]chromen-1-ol, is a semi-synthetic cannabinoid first described in 1940 (1). Recently, HHC has gained broader public attention as an alternative to ∆9-tetrahydrocannabinol (THC), initially in the USA and soon after in Europe.

Notably, in May 2022, it was identified in Denmark in a food product marketed as a sleep aid (2). Large-scale production of HHC involves using cannabidiol (CBD) as a precursor. In the first step, CBD undergoes acid-catalyzed intramolecular cyclization, primarily yielding delta8-tetrahydrocannabinol (∆8-THC) as the main product (3). Subsequently, ∆8-THC is converted to HHC by catalytic hydrogenation, leading to mixtures of (9S)- and (9R)-HHC epimers (4, 5). As of March 2023, HHC has been reported in 22 European countries (2). Despite initially being sold openly, several European countries, including Austria, have taken actions to control HHC.

The emergence and spread of HHC on the drug market have presented challenges to analytical laboratories, particularly due to the lack of information on its human pharmacokinetics, hindering the development and application of reliable tests for detecting HHC consumption. Given its chemical structure, it is reasonable to expect that HHC undergoes biotransformation reactions similar to other well-characterized cannabinoids such as THC, CBD and ∆8-THC. Figure 1a illustrates potential reactive sites of HHC. The primary metabolic route demonstrated by THC and other cannabinoids involves allylic hydroxylation at the position 11, followed by oxidation to the carboxylic acid as the final oxidized metabolite (6). Another observed metabolism route for CBD includes oxidation of the pentyl side chain (7, 8). Phase II metabolism primarily involves glucuronidation, with parent drugs and their oxidized forms as precursors (9).

Figure 1.

Alt text: Molecular structure of HHC. (a) Possible sites of metabolic transformation and (b) likely occurring fragmentation reactions in MS-MS experiments are highlighted. sites of metabolic transformation and (b) likely occurring fragmentation reactions in MS-MS experiments are highlighted.

Molecular structure of HHC. (a) Possible sites of metabolic transformation and (b) likely occurring fragmentation reactions in MS-MS experiments are highlighted.

While some information on HHC metabolism in animals is available from in vitro experiments, the identification of nine monohydroxylated metabolites of the (9 R)-HHC epimer was reported in microsomal preparations obtained from five animal species (10). However, no other Phase I or Phase II metabolites were mentioned. Additionally, analysis of livers from mice treated with ∆11-THC indirectly suggested a possible occurrence of HHC hydroxylation at the position 11, followed by oxidation to the carboxylic acid (11).

This report aims to expand our understanding of human HHC metabolism by summarizing the analytical results from systematic toxicological analysis of samples collected from individuals suspected of drug-impaired driving. Human body fluids, including plasma and urine, were obtained from three subjects under suspicion of HHC consumption. While urine samples tested positive for 11-nor-9-carboxy-∆9-tetrahydrocannabinol (THC-COOH) using an immunochemical assay, recent THC consumption was ruled out since all samples tested negative for THC and 11-hydroxy-∆9-tetrahydrocannabinol (11-OH-THC) via gas chromatography–mass spectrometry (GC–MS). The indeterminate GC–MS results for THC-COOH hinted at potential consumption of a THC-related compound. Through non-target analysis using liquid chromatography–high-resolution tandem mass spectrometry (LC–MS-MS), we identified HHC as the consumed cannabinoid. Furthermore, the available mass spectrometric data allowed us to elucidate significant biotransformation pathways of HHC in humans.

Materials and methods

Chemicals and reagents

Water, methanol (MeOH), acetonitrile (ACN) and acetone (all high-performance liquid chromatography grade) were purchased from Honeywell (Seelze, Germany). Acetic acid (purissima) was obtained from Sigma-Aldrich (St. Louis, MO, USA). Reference standards of ∆9-THC, 11-OH-THC, THC-COOH and cannabinol (CBN) were obtained from Lipomed (Arlesheim, Switzerland). A reference standard of (9 R)-HHC was obtained from Cayman Chemical Company (Ann Arbor, MI, USA).

Casework samples

The findings reported in this study are based on the analysis of human body fluids collected from three individuals suspected of drug-impaired driving. These samples were collected by physicians during clinical examinations of the drivers’ fitness. For two cases, both blood and urine samples were collected, while one case involved only blood samples. After collection, the samples were transported to our laboratory without cooling. Plasma samples were obtained by centrifugation of blood samples at 4000× g for 5 min at ambient temperature. Following this initial processing, samples were stored at −20°C until analysis.

Immunoassay

Immunological testing of the urine samples was accomplished on an Indiko Plus Clinical Chemistry Analyzer (Thermo Scientific, Waltham, MA, USA) applying the CEDIA THC assay (Thermo Scientific) according to the manufacturer’s protocol. The cut-off value for THC-COOH was 25 µg/L.

Quantitation of THC, 11-OH-THC and THC-COOH by GC–MS

A comprehensive description of the method used for quantitative GC–MS analysis of THC, 11-OH-THC and THC-COOH is available in the electronic supplementary material.

Non-targeted LC–MS-MS

The LC–MS-MS system consisted of a Waters ACQUITY UPLC (Waters, Manchester, UK) coupled to a TripleTOF 5600+ mass spectrometer (Sciex, Toronto, Canada). The chromatographic separation was accomplished on a Kinetex biphenyl column (2.6 µm, 100 Å, 100 × 2.1 mm; Phenomenex, Aschaffenburg, Germany) using a 15 min linear gradient of 2–98% ACN in an aqueous 0.5% acetic acid solution. The column temperature was 50°C. The flow rate was 200 µL/min. The “partial loop overfill” mode was used to inject 7.5 µL of the sample. The mass spectrometer was operated in positive electrospray ionization mode using a DuoSpray ion source. The spray voltage was set to 5.5 kV. Gas flows were 30 arbitrary units for the nebulizer gas and 50 arbitrary units for the turbo gas. The temperature of the turbo gas was adjusted to 400°C. The instrument was operated at a mass resolution of ∼30,000 for MS and ∼15,000 for MS-MS (tested with reserpine, m/z = 609.8507). To ensure a mass accuracy of better than 15 ppm, the mass spectrometer was automatically recalibrated every 10 sample injections using a calibration solution delivered via the calibration delivery system (Sciex). The scan range was m/z 100–700 for MS and m/z 50–700 for MS-MS. A duty cycle in the data-dependent acquisition mode included a single MS scan (accumulation time 100 ms) followed by eight dependent MS-MS scans (accumulation time 100 ms each) in the high-sensitivity mode with dynamic background subtraction. The intensity threshold for triggering MS-MS experiments was set to 100 counts. MS-MS spectra were acquired at 35 eV with a collision energy spread of 10 eV. Former target ions were excluded for 30 s after two occurrences.

Sample preparation for non-target analysis

0.25 mL of the sample was mixed with 10 µL internal standard solution (1.0 µg CBN per mL ACN). For protein precipitation, 0.75 mL ACN was added. The mixture was vortexed for 20 s and centrifuged for 5 min at 4000 × g. The supernatant was separated, diluted with 2 mL of a 0.1 M aqueous acetic acid solution, and submitted to solid-phase extraction employing SPE-ED Scan ABN columns (200 mg/3 mL, Applied Separations, PA, USA). The cartridges were equilibrated with 2 mL methanol and 2 mL water. After sample application, the cartridges were washed twice with 3 mL water. To remove water, cartridges were centrifuged for 5 min at 4000 × g, and dried with N2 for 20 min. Elution was accomplished with 2 mL acetone. The eluate was evaporated to dryness and reconstituted in 50 µL of ACN in water (50/50, v/v).

Quality control in non-target analysis

Several quality control samples were processed and/or analyzed together with each batch of casework samples. These included system suitability test samples, blank samples, zero samples and spiked samples. The spiked samples contained THC, 11-OH-THC, THC-COOH, HHC and CBN (50 ng/mL each).

For assessing the limits of identification of the applied experimental setup, urine and plasma samples were spiked at various concentrations with the available cannabinoid reference standards and analyzed. The observed limits of identification were in the low nanograms per milliliter range.

Tandem mass spectral library search

Compound identification was accomplished with a previously published workflow that involves conversion of tandem mass spectral data in an open format, automated library search and expert reviewing (12, 13). Raw data files in wiff-format were converted into the “mascot generic format” (mgf) using MSConvert from ProteoWizard (14). The MS-MS spectra part of the mgf file was extracted with a program written in ActivePerl 5.6.1 (Active State Corporation, Vancouver, Canada). Thus, all MS-MS spectra were available as plain text files containing peak list information, and they were used as input for database search. Library search was accomplished with “MSforID Search” (15) written in Pascal using Delphi 6 for Windows (Borland Software Corporation, Scotts Valley, CA, USA; now Embarcadero Technologies, Inc., San Francisco, CA, USA). The acquired tandem mass spectra were matched to an extended version of the “Wiley Registry of Tandem Mass Spectral Data” using the following settings: “m/z tolerance” ±0.01 and “intensity threshold value” 0.01. The library contained spectra of the following cannabinoids: THC, 11-OH-THC, THC-COOH, CBD, ∆8-THC, cannabigerol and CBN. A library search annotation was considered as putatively correct if the precursor ion mass error was within 20 ppm of the predicted ion mass and the average match probability value was >5.0. Finally, the correctness of tentative identifications was checked by expert reviewing. For this purpose, the corresponding tandem mass spectra were written to a single file and imported into the MS Search program (version 2.0 g, National Institute of Standards and Technology, Gaithersburg, MA, USA).

Annotation of HHC metabolites

The raw LC–MS-MS data were processed with PeakView software (version 2.0, Sciex). To identify potentially occurring metabolites, m/z values of protonated molecular ions and corresponding fragment ions were calculated using the integrated mass calculator. Extracted ion chromatograms (XICs) of the protonated molecular ions were generated with an m/z width of 20 ppm. To verify the chemical (sub-)structures of putatively detected metabolites, fragment ion m/z values were compared to those observed in reference spectra of HHC and other cannabinoids. Noteworthy fragmentation reactions observed for HHC are depicted in Figure 1b.

Results and discussion

Urine and plasma samples collected from three individuals suspected of HHC consumption were submitted to systematic forensic toxicological analysis that involved (a) immunological testing of urine samples , (b) GC–MS analysis of plasma samples targeting THC, 11-OH-THC and THC-COOH and (c) non-targeted LC–MS-MS analysis of urine as well as plasma samples. The obtained analytical results provided evidence that all three persons had consumed HHC.

Both urine samples tested positive using the CEDIA THC immunoassay, which primarily targets THC-COOH. Another plausible explanation for the observed results is the cross-reactivity of HHC metabolites. This hypothesis finds support in studies conducted in the 1980s, which highlighted that oxidized forms of HHC might lead to false-positive test results in immunoassays developed for detecting THC-COOH (16).

The applied GC–MS method utilized selected ion monitoring to target specific fragments of THC, 11-OH-THC and THC-COOH. The analysis of the three plasma samples yielded negative results for THC and 11-OH-THC. However, the results for THC-COOH were indeterminate. While a detectable peak was observed for each ion within the specified retention time window, the ion ratios deviated from the acceptable ranges. The occurrence of interfering ions related to one or more THC-related compounds (i.e., HHC metabolites) likely contributed to these deviations, though their exact origin remains speculative.

For non-target analysis, LC–MS-MS with data-dependent acquisition control was employed (13). Compound annotation was accomplished through tandem mass spectral library search. All samples tested positive for THC-COOH. This observation was indicative for a recent THC consumption. However, negative tests for THC, 11-OH-THC and other common psychoactive substances suggested that the observed impairment was linked to another compound that was not covered by the applied annotation workflow. To track down the identity of this substance, tentative matches to cannabigerol that were observed in the plasma samples proved useful. Cannabigerol is typically a minor constituent of cannabis, making it challenging to detect in plasma samples. To confirm the compound’s identity, a cannabigerol reference standard was analyzed under identical experimental conditions. The obtained mass spectrometric information, including the m/z of the precursor ion and the corresponding fragment ion spectrum, supported a putative match of the compound to cannabigerol (Figure 2a and b). However, due to the observed non-conformance in retention times, identity was excluded.

Figure 2.

Alt text: Fragmentation spectra of (a) an unknown compound detected in one of the analyzed plasma samples, (b) cannabigerol in a reference sample and (c) HHC in a reference sample.

Fragmentation spectra of (a) an unknown compound detected in one of the analyzed plasma samples, (b) cannabigerol in a reference sample and (c) HHC in a reference sample.

To find an alternative candidate, a PubChem search was conducted using the molecular formula of cannabigerol (C21H32O2). Among the 3,779 results, one was identified as HHC. By analyzing an HHC reference standard, the identity of the unknown compound was verified as HHC. In this case, the mass spectrometric data (Figure 2c) as well as the chromatographic information perfectly matched.

HHC and cannabigerol are structurally related compounds, with cannabigerol being considered the open-chain form and HHC the related cyclic form. Due to ring-opening reactions involved in the tandem mass spectral fragmentation of cyclic cannabinoids, the fragment ion mass spectra of these compounds often exhibit a high degree of similarity to the spectra of their open forms (e.g., THC vs. CBD). This similarity also applies to HHC and cannabigerol, resulting in fragmentation patterns that are difficult to differentiate from each other (Figure 2). As a consequence, if a software-supported tandem mass spectral library search is utilized, HHC spectra may be matched to cannabigerol if the library contains cannabigerol-specific spectra but lacks HHC-specific spectra.

As previously mentioned HHC was detected only in the analyzed plasma samples (Figure 3a), suggesting that it may have undergone metabolic modifications before being excreted in urine. Drawing parallels with the human metabolism of THC and CBD, it was anticipated that metabolic transformation reactions would involve Position 11 as well as the pentyl side chain (6–8). Additionally, it was expected that both the parent drug and its oxidized forms might undergo glucuronidation reactions (9). Leveraging the available LC–MS-MS data, these hypotheses were verified, enabling insights into the biotransformation pathways of HHC in humans.

Figure 3.

Alt text: Representative XICs of HHC and proposed human transformation products: (a) HHC, (b) HHC glucuronide, (c) hydroxylated HHC, (d) glucuronide of hydroxylated HHC, (e) HHC oxidized at C11 to the carboxylic acid, (f) glucuronide of HHC oxidized at C11 to the carboxylic acid, (g) HHC oxidized at C3′ to the carboxylic acid, and (h) glucuronide of HHC oxidized at C3′ to the carboxylic acid.

Representative XICs of HHC and proposed human transformation products: (a) HHC, (b) HHC glucuronide, (c) hydroxylated HHC, (d) glucuronide of hydroxylated HHC, (e) HHC oxidized at C11 to the carboxylic acid, (f) glucuronide of HHC oxidized at C11 to the carboxylic acid, (g) HHC oxidized at C3′ to the carboxylic acid, and (h) glucuronide of HHC oxidized at C3′ to the carboxylic acid. The color code refers to the possible sites of metabolic transformation of HHC as depicted in Figure 1.

In a first attempt to identify the expected HHC metabolites, we calculated the m/z values of protonated molecular ions and constructed corresponding XICs (Figure 3). By utilizing accurate mass information, we were able to provide evidence for the occurrence of oxidation reactions. These reactions included (i) hydroxylations (+O; Δm/z = 15.9949; Figure 3c), (ii) methyl oxidation to carboxylic acid (+O2, −H2; Δm/z = 29.9741; Figure 3e), and (iii) pentyl side chain oxidation to carboxylic acid (−(CH2)2, +O2, (−H2; Δm/z = 1.9429; Figure 3g). Furthermore, we observed signs of glucuronidation for both HHC and its Phase I metabolites (+C6H8O6; Δm/z = 176.0321; Figure 3b, d, f and h).

Remarkably, two forms of putative HHC glucuronide were detected (M1.1 and M1.2; Figure 3b). Glucuronidation of HHC is limited to the hydroxyl group located at C1. As semi-synthetic HHC consists of a mixture of (9S)- and (9R)-HHC epimers, the glucuronidation process yields diastereomers that can be chromatographically separated (Figure 3b).

Furthermore, multiple peaks were observed in the XIC traces specific for hydroxylated HHC forms and the corresponding glucuronides (Figure 4c and d). The presence of different sites of modification and the formation of diastereomers offered plausible explanations for the occurrence of multiple peaks in the glucuronide chromatograms.

Crucially, some of the detected hydroxylated forms (M3.1, M3.2, M4.1 and M4.2; Figure 3c) were considered to be interferences only. These interferences were attributed to in-source fragmentation of the corresponding glucuronides (Figure 3d). In-source fragmentation of glucuronides generates ions identical to those of the precursors (17), which are then detected in MS mode and contribute to signals in the precursors’ XIC traces at the retention times of the glucuronides. As a result, among the five hydroxylated HHC species detected, only one was considered to be present in the unconjugated form (M2; Figure 3c), while the other four species were identified as glucuronides (M3.1, M3.2, M4.1 and M4.2, Figure 3d).

In the XIC traces of the carboxylic acids as well as of the corresponding glucuronides, only one prominent peak was observed (M5, M6, M7 and M8; Figure 3e–h). This suggests that either no diastereomers were formed in these cases, or if they were formed, the chromatographic method used was not capable of separating the diastereomers. Minor peaks in the XIC traces of the carboxylic acids at the retention times of the carboxylic acid glucuronides indicated in-source fragmentation of the glucuronides (Figure 3e–h).

To provide additional evidence for the proposed structures of HHC metabolites, we examined the available tandem mass spectral data. Spectra representing the individual metabolites are provided in msp-format in the electronic supplementary material.

Figure 5 displays the tandem mass spectra of several glucuronides, specifically selected as representative examples due to their ability to provide structural information on both the conjugated and corresponding unconjugated forms (18). The dissociation of the glucuronides involved neutral loss of the glucuronic acid (Δm/z = 176.0321) and the generation of fragment ions identical to those of the precursor ions of the original forms (Figure 5). Certain secondary fragment ions were crucial for identifying the site of modification.

Figure 4.

Alt text: Representative fragmentation spectra of the proposed HHC Phase II metabolites. (a) M1.1: HHC glucuronide 1, (b) M3.2: HHC-OH (6-9) glucuronide 2, (c) M4.1: HHC-OH (1′–5′) glucuronide 1, (d) M6: HHC-COOH glucuronide and (e) M8: a putative metabolite obtained by oxidation of C3′ to the carboxylic acid and subsequent glucuronidation (HHC-COOH (3′) glucuronide).

Representative fragmentation spectra of the proposed HHC Phase II metabolites. (a) M1.1: HHC glucuronide 1, (b) M3.2: HHC-OH (6–9) glucuronide 2, (c) M4.1: HHC-OH (1′–5′) glucuronide 1, (d) M6: HHC-COOH glucuronide and (e) M8: a putative metabolite obtained by oxidation of C3′ to the carboxylic acid and subsequent glucuronidation (HHC-COOH (3′) glucuronide).

An important fragmentation reaction shared by HHC and other cannabinoids involves cleavage of bonds between Positions 10 and 10a, 5 and 6 as well as 6a and 10a (Figure 1b). The resulting (2,6-dihydroxy-4-pentylphenyl) methylium ion retains the pentyl side chain while losing the Positions 8–11. In the case of HHC, this specific fragment ion exhibits an m/z value of 193.1223 (Figure 2c), which was indeed observed in the fragment ion mass spectra of the putative HHC glucuronides (M1.1 and M1.2; Figure 4a). This fragmentation reaction proved valuable for distinguishing hydroxylation at Positions 8–11 from hydroxylation at the pentyl side chain (Figure 1a). For the former species (M3.1 and M3.2), the fragment ion at m/z 193.1223 was detected (Figure 4b). For the latter species (M4.1 and M4.2), two fragment ions with specific m/z shifts to 209.1159 (+O; Δm/z = 15.9918) and 191.1082 (+O, −H2O; Δm/z = −2.0152) were observed (Figure 4c).

Carboxylic acids, including THC-COOH, are known to exhibit characteristic neutral losses of −H2O (Δm/z = 18.0106) and subsequent −CO (Δm/z = 46.0055) during collision-induced dissociation (19). The accurate mass information provided evidence of two HHC-derived carboxylic acid species as well as their corresponding glucuronides, with oxidation reactions occurring at C11 (M5 and M6) and C3′ (M7 and M8). In collision-induced dissociation, cleavage of water and CO was observed for both putative HHC metabolites (Figure 4d and e). Nevertheless, with the available fragmentation information, including reference data from THC-COOH, unequivocal verification of the chemical structures was only achieved for 11-nor-9-carboxy-hexahydrocannabinol (HHC-COOH, M5) and its glucuronide (M6). For the other species, doubts about the correctness of the proposed structures still exist. Further analyses, involving specific in vitro assays and chemical synthesis (5, 7, 20), would be required to increase identification confidence and to verify the preference for C3′ over other positions of the pentyl side chain for oxidation.

In an attempt to validate accomplished annotations, we compared the tandem mass spectral information of selected metabolites with data obtained from analyzing reference standards. The reference standards analyzed included 8-hydroxy-HHC, 9-hydroxy-HHC, 10-hydroxy-HHC, 11-hydroxy-HHC as well as 11-nor-9-carboxy-HHC. The results are provided in the electronic supplementary material Figures S1 and S2. The different hydroxylated reference compounds exhibited very similar fragmentation patterns, and the observed fragment ions corresponded well with those observed for related compounds in the human samples (M2, M3.1 and M3.2). Additionally, the fragment ion mass spectrum of the 11-nor-9-carboxy-HHC standard aligned with the corresponding spectra annotated in the human samples (M5 and M6).

Based on the LC–MS-MS data, the study was able to identify 11 HHC metabolites, and an overview of the compounds detected in the analyzed plasma and urine samples is provided in Table I. The proposed metabolic transformation reactions are summarized in Figure 5. Transformation of HHC involves the formation of HHC glucuronide (M1), HHC-COOH (M5) and HHC-COOH glucuronide (M7). Hydroxylation appears to occur at either Positions C8 to C11 or the pentyl side chain. Given the observation of the corresponding carboxylic acid, it is highly likely that C11 is the primary site of oxidation (M2), followed by glucuronidation (M3.1 and M3.2). While confirmation is still pending, there is some evidence supporting oxidation of C3′, resulting in hydroxylated and carboxylic acid species (M7), along with their corresponding glucuronides (M4.1, M4.2 and M8).

Table I.

Overview of HHC-related compounds detected in the analyzed casework samples

Compound Tentative annotation Possible site(s) of modification Molecular formula m/z Retention time (min) Plasma 1 Urine 1 Plasma 2 Urine 2 Plasma 3
HHC HHC C21H32O2 317.2475 12.4 n.d. n.d.
M1.1 HHC Glucuronide 1 1 C27H40O8 493.2796 9.3 n.d. n.d.
M1.2 HHC Glucuronide 2 1 C27H40O8 493.2796 10.1
M2 HHC-OH (6–9) 8–11 C21H32O3 333.2424 10.6 n.d. n.d.
M3.1 HHC-OH (6–9) Glucuronide 1 8–11 C27H40O9 509.2745 8.9 n.d.
M3.2 HHC-OH (6–9) Glucuronide 2 8–11 C27H40O9 509.2745 9.2
M4.1 HHC-OH (1′-5′) Glucuronide 1 1′–5′ C27H40O9 509.2745 7.7 n.d.
M4.2 HHC-OH (1′-5′) Glucuronide 2 1′–5′ C27H40O9 509.2745 8.1 n.d.
M5 HHC-COOH 11 C21H30O4 347.2217 10.5
M6 HHC-COOH Glucuronide 11 C27H38O10 523.2538 9.2
M7 HHC-COOH (3′) 3′ C19H26O4 319.1903 8.4 n.d.
M8 HHC-COOH (3′) Glucuronide 3′ C25H34O10 495.2225 7.2 n.d.

✓—detected; n.d.—not detected.

Figure 5.

Alt text: Proposed metabolic transformation of HHC in humans based on the screening of plasma and urine samples for HHC metabolites.

Proposed metabolic transformation of HHC in humans based on the screening of plasma and urine samples for HHC metabolites.

Conclusion

As outlined by a recent report of the European Monitoring Center for Drugs and Drug Addiction (2), investigating the pharmacokinetics of HHC should be a priority. To support and advance research in this field, we report important metabolic transformation reactions of HHC in humans. The information was derived from analyzing plasma and urine samples collected from three individuals suspected of HHC consumption. The LC–MS-MS data provided evidence for the occurrence of oxidation reactions involving C11 and the pentyl side chain, leading to the corresponding hydroxylated and carboxylic acid species. Furthermore, signs of extensive glucuronidation of HHC and its Phase I metabolites were observed. The obtained results represent a solid foundation for designing further in vitro and in vivo experiments that enable more specific investigations of the pharmacokinetics, pharmacology and toxicology of HHC as well as the development of reliable analytical tests for the detection of HHC consumption.

Supplementary Material

bkae022_Supp
bkae022_supp.zip (317.7KB, zip)

Contributor Information

Florian Pitterl, Institute of Legal Medicine, Medical University of Innsbruck, Muellerstrasse 44, Innsbruck 6020, Austria.

Marion Pavlic, Institute of Legal Medicine, Medical University of Innsbruck, Muellerstrasse 44, Innsbruck 6020, Austria.

Jianmei Liu, Forensic Chemistry Division, Cayman Chemical Company, 1180 E Ellsworth Rd., Ann Arbor, MI 48108, USA.

Herbert Oberacher, Institute of Legal Medicine, Medical University of Innsbruck, Muellerstrasse 44, Innsbruck 6020, Austria; Core Facility Metabolomics, Medical University of Innsbruck, Muellerstrasse 44, Innsbruck 6020, Austria.

Supplementary data

Supplementary data is available at Journal of Analytical Toxicology online.

Data availability

Data cannot be shared for ethical/privacy reasons.

Author contributions

Florian Pitterl (Conceptualization, Methodology, Formal analysis, Investigation, Writing, Visualization), Marion Pavlic (Resources, Writing), Jianmei Liu (Formal analysis, Resources, Writing, Visualization) and Herbert Oberacher (Conceptualization, Methodology, Investigation, Resources, Writing, Visualization, Supervision).

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

bkae022_Supp
bkae022_supp.zip (317.7KB, zip)

Data Availability Statement

Data cannot be shared for ethical/privacy reasons.


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