Analysis of Cannabinoids and Their Metabolites in Human Urine

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Analysis of Cannabinoids and Their Metabolites in Human Urine Binnian Wei, Lanqing Wang, and Ben C Blount Anal. Chem., Just Accepted Manuscript • Publication Date (Web): 27 Sep 2015 Downloaded from http://pubs.acs.org on October 4, 2015

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Analytical Chemistry

1

Analysis of Cannabinoids and Their Metabolites in Human Urine

2

Binnian Wei1, Lanqing Wang1, Benjamin C. Blount1

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1

Division of Laboratory Sciences, National Center for Environmental Health,

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Centers for Disease Control and Prevention, Atlanta, Georgia, USA

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Send Correspondence to: Dr. B. Wei. E-mail: [email protected]; Telephone: +1 770-488-4564.

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Division of Laboratory Sciences, Centers for Disease Control and Prevention.

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4770 Buford Hwy, NE Rd, Mail stop: F44. Atlanta, GA, US, 30041.

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Conflict of Interest: The authors declare no competing financial interest.

10 11 12 13 14 15

Disclaimer: The findings and conclusions in this study are those of the authors and do not necessarily represent the views of the U.S. Department of Health and Human Services, or the U.S. Centers for Disease Control and Prevention. Use of trade names and commercial sources is for identification only and does not constitute endorsement by the U.S. Department of Health and Human Services, or the U.S. Centers for Disease Control and Prevention.

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Abstract:

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Biologically monitoring marijuana exposure from active and passive use requires

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both a wide linear range and sensitive detection.

19

validated

20

chromatography coupled with tandem mass spectrometry (UHPLC–MS/MS) for

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analysis of urinary ∆9-tetrahydrocannabinol (THC), cannabidiol and cannabinol,

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and two major metabolites of THC, 11-nor-9-carboxy-THC and 11-hydroxy-THC,

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in active users and particularly in people exposed to secondhand marijuana

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smoke (SHMS). The method used positive electrospray ionization (ESI) mode to

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reach the sensitivity needed to detect trace SHMS exposure with limits of

26

detection (LOD) ranging from 0.002 to 0.008 nanograms per milliliter (ng/mL),

27

and 0.005 to 0.017 ng/mL for “free” (unconjugated forms) and “total”

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(unconjugated plus conjugated forms) measurements, respectively. These LODs

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were approximately 10–100 times more sensitive than those reported in the

30

literature. To reduce or avoid time-consuming repetitive sample preparation and

31

analysis, the method simultaneously monitored multiple reaction monitoring

32

transitions in negative ESI mode to quantify high analyte levels typically found in

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the urine of active marijuana users (linear dynamic range of 12.5 – 800 ng/mL).

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The validation results indicated this method was accurate (average inter-/intra-

35

day bias: < 10%), precise (inter-/intra-day imprecision: < 10%) and fast (6

36

minutes run time). In addition, sample preparation throughput was greatly

37

improved using an automation liquid-handling system, meeting the needs for

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potential large-scale population studies.

a

multifunctional

method

using

We have developed and

ultra-high

39


performance

liquid

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Main Text

40

41

1. Introduction

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Increasing use of marijuana both medicinally and recreationally1, 2 may lead to

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increased health risks resulting from exposure to both cannabinoids and the

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toxic chemicals found in marijuana smoke.3-5 Traditionally, cannabinoids and

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their metabolites, i.e. ∆9–tetrahydrocannabinol (THC), cannabidiol (CBD) and

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cannabinol (CBN), and two major metabolites of THC, 11–nor–9–carboxy–THC

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(COOH-THC) and 11–hydroxy–THC (OH-THC) (Figure S-1), are measured in the

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urine of people to assess their exposure to marijuana products and smoke.

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In the last decades, a number of analytical methods have been applied to

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analyze THC, OH-THC, COOH-THC, CBD and CBN in urine samples, including

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methods employing liquid or gas chromatography (LC or GC) coupled with either

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a single quadrupole mass spectrometry (MS) or tandem mass spectrometry

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(MS/MS).6-10 Reported limits of detection (LOD) for these analytes ranged from

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0.2 to 5.0 nanograms per milliliter (ng/mL). Given the relatively high exposure

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levels resulted from active marijuana smoking, these LODs meet the needs

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accordingly for desired detection rates. Nevertheless, persons smoking

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marijuana could also cause passive exposure of other people through inhalation

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of secondhand marijuana smoke (SHMS), which are often characterized by low

59

biomarker

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environmental circumstances, smoker density and exposure duration. While most

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studies have focused on active use of marijuana, information on SHMS, including

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exposure characteristics and adverse health consequences, is still limited in the

levels

although

collectively

depending


on

factors

including


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open literature. A sensitive analytical method to quantify trace biomarker levels is

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therefore indispensable for effectively monitoring and assessing SHMS exposure.

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The objective of this study was to develop and analytically validate a

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sensitive ultra-high pressure LC (UHPLC) – electrospray ionization (ESI)

67

combined with MS/MS method for quantifying THC, COOH-THC, OH-THC, CBD

68

and CBN in urine from people exposed to marijuana smoke, particularly to SHMS.

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To the best of our knowledge, the analyte-specific LODs achieved in this method

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for urine samples were approximately 10 – 100 times more sensitive than

71

previous studies. In addition, time-consuming repetitive preparation and analysis

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of unknown urine samples often become necessary when analyte levels fall out

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of a method’s linear dynamic ranges. We simultaneously monitored multiple

74

reaction monitoring (MRM) transitions of those five analytes under both positive

75

and detuned negative ESI modes for low-concentration (LODs – 50 ng/mL) and

76

high-concentration (up to 800 ng/mL) samples, respectively. This combined

77

analysis greatly increased the applicable quantitation ranges and facilitated the

78

data acquisition by reducing or avoiding potential repetitive sample preparation

79

and analysis. Finally, we increased sample preparation throughput using an

80

automation liquid-handling system to meet the needs for potential large–scale

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population studies.

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2. Experimental Section

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Details on chemicals and materials are provided in the Supporting Information.

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Standard Solution Preparation 4 ACS Paragon Plus Environment

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Blank urine pools, used as matrix for calibration standard (calibrators) and quality

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control (QC), were prepared using human urine anonymously collected in

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compliance with Institutional Review Board protocol. The CDC Human Subjects

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Review Board determined this activity was not human subject research. Blank

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urine samples were combined to form a matrix urine pool, which was kept at 4 oC

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and stirred overnight to ensure thorough mixing. This urine pool was

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subsequently spiked with target analytes to form calibrators and QCs.

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We prepared working solutions for calibrators and QCs from serial

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dilutions of primary stock solutions with methanol and water (v/v: 60:40), and

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stored them in Teflon–capped amber glass vials at –24 °C. 50 µL of each

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working solution was automatically added to 500 µL of blank urine (see Sample

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preparation section), creating calibrators at 0.001 to 800 ng/mL, and QC samples

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at 0.05, 25 and 500 ng/mL. The internal standard spiking solution had

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concentrations of 0.06 ng/µL for CBD-d3, CBN-d3 and 0.1 ng/µL for OH-THC-d3,

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COOH-THC-d3 and THC-d3. Details regarding the standard solution preparation

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are given in the supporting information.

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Sample preparation

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We prepared calibrators, QCs, blanks and unknown urine samples following

103

same procedures, as depicted in Figure 1. Briefly, urine samples stored at

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temperatures ≤ –65 °C were thawed and vortex-mixed for 10 minutes at room

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temperature. To measure “total” concentrations (unconjugated and conjugated

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forms), 500 µL of each urine sample (blank and unknown) was transferred to

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each well in the 96-well plate, followed by adding 50 µL of calibrator and QC 5 ACS Paragon Plus Environment


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working solutions to each blank urine sample. Working solutions were replaced

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with water for unknown samples. Then, 50 µL of internal standard solution and

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50 µL of enzyme solution (Escherichia coli, type IX-A, 20 units/µL, 0.5 M

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ammonium acetate, pH 6.8) were added into all samples. After gentle mixing

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(“Pre-Mix”), the 96-well plate was incubated at 37 °C for 2 h. Then, 50 µL of 10N

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NaOH was added to each well, and incubated at 70 °C for 20 minutes. After

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cooling the plate to room temperature, 400 µL of formic acid, water and methanol

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(v/v/v: 12.5:12.5:75) was added to each well. To measure “free” unconjugated

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concentrations, enzyme and NaOH solutions were replaced with 450 µL of formic

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acid, water and methanol (v/v/v: 5.5:27.8:66.7), and no incubation was performed.

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After mixing for 5 minutes, the plate was centrifuged for 30 minutes at –5 °C, and

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then 0.96 mL mixtures from each well was transferred onto the 96-well SPE plate,

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pre-equilibrated with 1.0 mL of methanol and 1.0 mL of buffer (5mM ammonium

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formate, 0.05% formic acid). After soaking for 10 minutes, the mixtures were

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pushed through the SPE under approximately 1.0 psi positive pressure. Then,

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the samples were washed with 1.0 mL of water and 1.0 mL of methanol and

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water (v/v: 60:40). After drying for 15 minutes with nitrogen (25 psi), the samples

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were collected in a second 96-well plate by elution with 1.0 mL of methanol, and

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then evaporated to dryness using a TurboVap evaporator (Biotage, Charlotte, NC,

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USA) at room temperature. The residuals were reconstituted with 50 µL of

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method and water (v/v: 50:50). After 5 minutes of vortex, 10 µL of each sample

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was injected into the LC system.

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Instrumentation and Operation 6 ACS Paragon Plus Environment

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The “Pre-Mix” procedure (Figure 1), also the critical one impacting the accuracy

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and precision of analytical results, was performed on a Hamilton automated

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liquid–handling system – Microlab Star (Reno, NV, USA), aimed at improving

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data reproducibility and sample preparation throughput. On an as-needed basis

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in future, the throughput can be further expanded using a fully automated system

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illustrated in our previous study.11 Additional information can be obtained upon

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request.

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Chromatographic separation was achieved using a Kinetex reversed

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phase column (100 mm × 2.1 mm, particle size 2.6 µm, C18) (Phenomenex,

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Torrance, CA, USA) on a Shimadzu UHPLC system (Columbia, MD, USA) at a

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flow rate of 0.4 mL/min. Column temperature was maintained at 40 °C during the

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entire analysis. The gradient program contained 5.0 mM of ammonium formate

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with 0.05% formic acid (solvent A), and acetonitrile (solvent B). After injection, LC

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flow during the first 2.5 minutes and the last 1.5 minutes was directed to a waste

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container by the switching valve. Only the flow occurring between 2.5 and 4.5

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minutes was directed to the MS. Detailed gradient conditions are provided in

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Table – S1. Figures–2 (a, c, e, g, i) and –2 (b, d, f, h, j) depict representative

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chromatograms for urine samples processed with the “free” and “total” methods,

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respectively.

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MS/MS analysis was performed on a Sciex triple quadrupole 6500 with a

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TurboIonSpray source (Foster City, CA, USA). ESI+ and ESI- modes were used

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to acquire scheduled MRM transition data. Optimum MS source parameters were

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as follows: source temperature: 600 °C; ionspray voltage (ESI+/ESI-): 5500/-4500



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V; ion source gas-1: 80 psi and gas-2: 90 psi; curtain gas: 35 psi; target scan

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time (ESI+/ESI-): 0.18/0.041 seconds. Two MRM precursor/product transitions for

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each native analyte and one transition for the isotope labeled internal standard

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were monitored. The collision offset energy (CE) under ESI- mode were “de-

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tuned” to higher value so as to yield a lower response of the corresponding MRM

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transition to avoid detector saturation when analzying high concentrations

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typically found in active smokers’ urine. Detailed MRM transitions and voltage

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settings are given in Table S–2, and the instrument method for data acquisition is

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also described in the Supporting Information.

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3. Results and Discussion

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We first evaluated different 96-well plates containing various materials, including

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C18, mixed-mode cation/anion-exchange polymer, strongly hydrophilic water

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wettable polymer, and inert diatomaceous earth. Overall, C18 SPE provided the

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highest sample extraction recoveries. The high lipophilicity of these analytes

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enabled us to apply high percentages of methanol in washing solvent (50–70%)

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to purify the urine samples on C18 SPE plates. We found washing solvent with

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methanol and water (60:40) had a balanced performance between extraction

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recoveries and matrix effects.

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Because ESI+ analysis yielded higher sensitivity for all analytes, we

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monitored their MRM transitions using ESI+ for low-concentration samples (LOD

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– 50 ng/mL). To avoid variation caused by MS detector saturation using ESI+

175

analysis, we simultaneously monitored those five analytes under ESI- mode to


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quantify any unknown samples whose concentrations were in the range of 12.5 –

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800 ng/mL. Under both ionization modes, we observed excellent calibration

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linearity with average determination coefficients (R2) ≥ 0.995. We also assessed

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data consistency between two ionization analyses by comparing fortified samples

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whose concentrations fell into the overlapping linear dynamic ranges of two

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modes, and we observed excellent agreement in analytical results (Tables 2 and

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S4). Expectedly, this combined analysis would greatly facilitate data acquisition

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by increasing the applicable dynamic ranges and reducing the repeating rates of

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sample preparation and analysis.

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One of the challenges in measuring urinary cannabinoids and their

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metabolites is that the high lipophilicity of these compounds can result in

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substantial adsorption to materials used for sample preparation, i.e. tips and 96-

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well plates. To minimize chemical loss during sample preparation, we added 0.3

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mL methanol into all samples before SPE clean-up. After injection, the analytes

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could also absorb to other instrumental surfaces, i.e. tubing, column, LC injection

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loop, needle and port etc. Over time, amounts accumulated on LC parts can

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potentially cause carry-over contamination. To avoid this issue, we added the

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following LC autosampler washing program: before each injection, the auto-

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sampler measuring line was washed with 600 µL of acetonitrile and water (v/v:

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55/45), and both internal and external surfaces of the sampling needle were

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rinsed with 300 µL of acetonitrile, 2-propanol and water (v/v/v: 45/45/10). We

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evaluated the effectiveness of this washing program by measuring the carry-over

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concentrations in the subsequent blank urine samples following the highest 9 ACS Paragon Plus Environment


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calibrator (800 ng/mL). The results indicated that no targeted peaks at the

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retention times of all analytes were detected.

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Thermal instability of these analytes in urine was another issue that

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requires extra caution in the entire processes of sample collection, transport,

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storage and analysis. Our stability tests with fortified urine at room temperature

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(~25 oC) indicated >80% of total CBD, CBN and THC were lost after 24 hours

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(Figure – 3: b, d and f). The losses of total OHTHC and COOH-THC at 25 oC

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were approximately 5 and 12% after 24 h, respectively. At 4 oC, approximately

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10 – 30% of total CBD, THC and CBN decomposed after 3 days, and no obvious

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losses of total OHTHC and COOH-THC were observed (Figure – 3: a, c and e).

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We did not observe significant decrease in total analyte concentrations≤ –20 oC

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during the same study period. These results are consistent with those reported in

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a recent study by Desrosiers, et al.

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temperatures ≤ –20 oC and the stability to indoor light are in progress. Available

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evidences indicated that urine samples should always be kept frozen during

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storing and shipping. For analysis, samples need to be prepared in a timely

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manner and analyzed within one day. The high-throughput liquid-handling

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automation

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plates/instrument/day) presented in this study would better meet such needs so

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as to maintain the quality of analytical results.

system

and

fast

12

Studies regarding long-term stability at

UHPLC-MS/MS

analysis

(~two

96-well

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One of the main advantages of this method, aside from the broad

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applicable quantitation ranges, high-throughput preparation and fast analysis, is

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the marked sensitivity that is essential for effectively monitoring and assessing 10 ACS Paragon Plus Environment

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SHMS exposure. We determined the LODs and limits of quantitation (LOQ) by

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preparing and analyzing four low-concentration urine pools (0.005, 0.010, 0.025

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and 0.050 ng/mL) over 3-month period. We first determined the standard

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deviation (SD) of each pool’s concentration quantified using first MRM transition

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with confirmation by the second transition (Table S-2) (More details regarding

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analytical specificity is described in the supporting information), and then we

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plotted the SD of each pool against the concentration, and finally obtained the S0

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given as the Y-intercepts

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and 10S0, respectively. We observed the method for unconjugated form had

231

more sensitive LODs (0.002 – 0.008 ng/mL, or equivalently 0.064 – 0.242 fmol

232

on-column) than those for “total” form (0.005 – 0.017 ng/mL, or equivalently

233

0.159 – 0.514 fmol on-column) (Table S-3). This was most likely the result of the

234

tandem-hydrolysis procedure (enzymatic-alkaline hydrolysis). During hydrolysis

235

incubation at higher temperature, analytes can partially degrade due to their

236

thermal instability; meanwhile, they can also partially bind to added enzyme (E.

237

Coli.) because of the high lipophilicity and then precipitate by centrifugation.

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Despite the mass loss during sample preparation, the detection sensitivity

239

achieved in this method for urine samples were 10–100 times the values (0.2 –

240

5.0 ng/mL) reported in the literature.6-9

13-15

. LODs and LOQs (Table 1) were calculated as 3S0

241

We used three sets of samples at low, medium and high concentrations

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(0.025, 25 and 500 ng/mL, respectively) to determine the optimized extraction

243

recoveries and matrix effects (details in supporting materials).11 Average

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extraction recoveries for all analytes ranged from 54 to 93% and 18 to 75% for



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“free” and “total” measurements, respectively (Table 1). Ion suppression due to

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matrix effect varied from – 9 to 32, and – 41 to 32 for “free” and “total”

247

measurements, respectively.

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Calibrators and QCs were always freshly produced using the liquid-

249

handling automation system, and then prepared in the same manner along with

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unknown samples, and laboratory blanks. We observed excellent inter-day and

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intra-day accuracy (90–110%), and imprecision was less than 10% (Table 2 and

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Table S–4) based on replicate analyses of a serial of fortified urine samples

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prepared by spiking known amounts of analytes over a 3 consecutive months.

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Acknowledgement

255

We thank Ernest McGahee and Jason Terranova for maintaining the

256

performance of the automated liquid-handling system. The manuscript has

257

certainly benefited from those constructive suggestions from the reviewers.

258

Supporting Information

259

The supporting information consists of experimental details, LC gradient program,

260

MRM ion transitions, LODs in terms of fmol on-column, accuracy and precision

261

for “total” method, molecular structures of five analytes, as mentioned in the text.

262

A complete instrument method file for data acquisition was also uploaded

263

separately.

264

References


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1. Murray, R. M.; Morrison, P. D.; Henquet, C.; Di Forti, M., Cannabis, the mind and society: the hash realities. Nature Reviews Neuroscience 2007, 8, (11), 885-895. 2. UNODC, United Nations Office on Drugs and Crime. World Drug Report. 2014. 3. Hall, W.; Degenhardt, L., Adverse health effects of non-medical cannabis use. The Lancet 2009, 374, (9698), 1383-1391. 4. Moir, D.; Rickert, W. S.; Levasseur, G.; Larose, Y.; Maertens, R.; White, P.; Desjardins, S., A comparison of mainstream and sidestream marijuana and tobacco cigarette smoke produced under two machine smoking conditions. Chem Res Toxicol 2008, 21, (2), 494-502. 5. Volkow, N. D.; Baler, R. D.; Compton, W. M.; Weiss, S. R., Adverse health effects of marijuana use. New Engl J Med 2014, 370, (23), 2219-2227. 6. Kemp, P. M.; Abukhalaf, I. K.; Manno, J. E.; Manno, B. R.; Alford, D. D.; Abusada, G. A., Cannabinoids in humans. I. Analysis of Δ9-tetrahydrocannabinol and six metabolites in plasma and urine using GC-MS. J Anal Toxicol 1995, 19, (5), 285-291. 7. Grauwiler, S. B.; Scholer, A.; Drewe, J., Development of a LC/MS/MS method for the analysis of cannabinoids in human EDTA-plasma and urine after small doses of Cannabis sativa extracts. Journal of Chromatography B 2007, 850, (1), 515-522. 8. Fernández, M. d. M. R.; Wille, S. M.; Samyn, N.; Wood, M.; López-Rivadulla, M.; De Boeck, G., On-line solid-phase extraction combined with liquid chromatography– tandem mass spectrometry for high throughput analysis of 11-nor-Δ 9tetrahydrocannabinol-9-carboxylic acid in urine. Journal of Chromatography B 2009, 877, (22), 2153-2157. 9. Scheidweiler, K. B.; Desrosiers, N. A.; Huestis, M. A., Simultaneous quantification of free and glucuronidated cannabinoids in human urine by liquid chromatography tandem mass spectrometry. Clin Chim Acta 2012, 413, (23), 1839-1847. 10. Skopp, G.; Pötsch, L., Stability of 11-nor-Δ9-carboxy-tetrahydrocannabinol glucuronide in plasma and urine assessed by liquid chromatography-tandem mass spectrometry. Clin Chem 2002, 48, (2), 301-306. 11. Wei, B.; Feng, J.; Rehmani, I. J.; Miller, S.; McGuffey, J. E.; Blount, B. C.; Wang, L., A high-throughput robotic sample preparation system and HPLC-MS/MS for measuring urinary anatabine, anabasine, nicotine and major nicotine metabolites. Clin Chim Acta 2014, 436, 290-297. 12. Desrosiers, N. A.; Lee, D.; Scheidweiler, K. B.; Concheiro-Guisan, M.; Gorelick, D. A.; Huestis, M. A., In vitro stability of free and glucuronidated cannabinoids in urine following controlled smoked cannabis. Anal Bioanal Chem 2014, 406, (3), 785-792. 13. DNR. Analytical Detection Limit Guidance & Laboratory Guide for Determining Method Detection Limits. http://dnr.wi.gov/regulations/labcert/documents/guidance/LODguide.pdf [Accessed date: Jan. 22, 2014] 14. ICH, ICH Harmonised Tripartite Guideline. Validation of Analytical Procedures: Text and Methodology. 1994. 15. Taylor, J. K., Quality assurance of chemical measurements. CRC Press: 1987.

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307 308

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Table 1 – Limit of detection (LOD), limit of quantitation (LOQ), average extraction efficiency of sample preparation and average matrix effect.

309

310 311

Extraction efficiency (“free”/”total”), %

Matrix effect (“free”/”total”), %

LOD, ng/mL

LOQ, ng/mL

Analyte

“free”/“total” 1

“free”/“total”

Low2

Mid

High

Low

Mid

High

OHTHC

0.008/0.017

0.028/0.048

93/69

81/67

82/75

1.0/13

-5/1

-9/-32

COOH-THC

0.005/0.015

0.018/0.045

83/58

81/60

83/59

5/9

-4/-6

-2/-41

CBD

0.004/0.009

0.012/0.030

88/22

70/26

71/28

8/23

-9/-4

-5/-9

CBN

0.002/0.007

0.008/0.023

76/20

64/18

71/21

2/-6

-8/-16

-4/-12

THC 0.002/0.005 0.008/0.015 74/18 59/19 54/22 6/32 -7/-5 -7/-12 “Free” and “total” refer to unconjugated and the sum of conjugated and unconjugated forms, respectively. 2 Low, mid and high levels refer to 0.025, 25 and 500 ng/mL, respectively.

1

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Table 2 – Accuracy and precision of replicate analysis of five compounds in fortified urine samples by “free” method (without enzymatic-alkaline hydrolysis). Within-day

315

Between-day

ESI Target, Error RSD Error RSD measured measured mode ng/mL % % % % OHTHC + 0.025 0.0244 -2.3 9.2 0.0245 -1.9 5.7 0.100 0.103 3.0 5.0 0.105 5.5 3.5 1.25 1.24 -0.8 2.2 1.28 2.7 3.0 25.0 26.3 5.1 5.3 26.9 7.6 2.1 – 25.0 26.4 5.4 8.1 24.6 -1.6 7.6 100 107 7.0 4.9 107 7.0 1.0 500 480 -4.1 2.2 475 -5.1 2.9 COOH-THC + 0.025 0.0245 -2.2 6.5 0.0247 -1.2 6.6 0.100 0.105 5.4 5.7 0.104 4.0 4.2 1.25 1.19 -4.7 9.9 1.26 1.1 0.5 25.0 26.5 6.0 3.6 26.8 7.3 1.8 – 25.0 26.3 5.2 4.6 26.5 5.8 2.4 100 105 4.9 8.3 107 7.3 4.6 500 473 -5.4 3.3 457 -8.6 1.8 CBD + 0.025 0.0246 -1.5 6.6 0.0244 -2.6 6.5 0.100 0.099 -0.6 4.4 0.110 9.6 8.3 1.25 1.15 -8.4 1.5 1.26 1.1 9.8 25.0 25.9 3.4 2.9 25.7 2.6 2.4 – 25.0 27.2 8.8 2.0 24.8 -1.0 4.8 100 107 6.7 4.9 105 4.8 8.2 500 453 -9.5 1.4 453 -9.4 3.3 CBN + 0.025 0.0264 5.6 9.0 0.0253 1.3 5.7 0.100 0.109 8.7 5.6 0.108 7.6 4.7 1.25 1.23 -1.3 3.9 1.23 -1.9 4.0 25.0 25.8 3.1 6.1 26.8 7.0 3.8 – 25.0 25.4 1.6 9.7 25.7 2.9 3.4 100 98.3 -1.7 7.5 104 3.7 3.2 500 469 -6.2 3.0 464 -7.3 3.1 THC + 0.025 0.0249 -0.5 9.9 0.0274 9.7 3.1 0.100 0.108 8.0 5.1 0.106 5.5 7.4 1.25 1.18 -5.9 4.4 1.23 -1.3 5.1 25.0 26.7 6.7 7.7 27.3 9.1 2.7 – 25.0 25.3 1.3 7.8 25.8 3.2 5.2 100 107 7.2 4.7 106 6.0 3.6 500 476 -4.8 2.6 473 -5.4 4.7 Abbreviation: ESI = electrospray ionization; RSD = relative standard deviation

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Figure Legends

318

(All figures are in single-column format)

319

Figure 1 – Sampling preparation scheme. Abbreviations: ISTD – internal standard spiking solution; Conc. – concentration. “Pre-Mix” included adding ISTD and enzymatic solution (if mearing “total” concentration) to urine samples followed by gentle mixing. “Hydrolysis” included enzymatic and alkaline procedures in order, and was omitted to measure “free” (“unconjugated”) concentrations. “SPE Extraction” included SPE pre-equilibrium, sample clean-up and elution. “Condensation” included sample evaporation and residual reconstitution.

320 321 322 323 324 325 326 327 328 329 330 331 332 333

Figure 2 – Chromatograms of fortified urine samples analyzed on the Sciex triple quadrupole 6500. Chromatograms illustrated in left column (0.010 ng/mL) and right column (0.025 ng/mL) were for “free” (“unconjugated”) and “total” (“conjugated” + “unconjugated”) concentrations, respectively. Figure 3– Thermal stability of OHTHC, COOH-THC, CBD, CBN and THC in fortified urine pools. Figures (a), (c) and (e) show the results for samples stored at 4 ⁰C and (b), (d) and (f) display the results for samples stored at 25 ⁰C.

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Figure – 1

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Figure – 2

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Figure –3:

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“For TOC only”

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Page 20 of 20

Analysis of Cannabinoids and Their Metabolites in Human Urine

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