Danielle Mackowsky

Research scientists in the cannabis field are tasked with validating robust methods that can be seamlessly transitioned into production laboratories. Unlike typical disciplines where controls are easily (and legally) obtained through known manufacturers, analytical chemists working for both consumable vendors as well as cannabis laboratories must do their best to develop methods often without such resources at their disposal. As the industry matures and additional regulations are adopted, the evolution of the pesticide testing subsection continues to be vastly different depending on the jurisdiction one does business in. This creates an interesting challenge for commercial scientists tasked with developing methods that will appeal to a majority of their consumers, while also generating unexpected hurdles to said laboratories once the methods are placed into production. This article provides an overview of best practices and method development techniques for pesticide testing in cannabis from a technical perspective.

	Unlike other analytical fields, the cannabis laboratory testing industry has yet to reach its maturity. Without cohesive testing requirements among states where the plant is legal, laboratories often struggle to develop coordinated methodologies for the analysis of pesticides and mycotoxins. Although both of these classes of compounds have been widely studied in other matrices, the complexity of the cannabis plant presents additional analytical challenges that do not need to be accounted for in other agricultural products. Up to one-third of the overall mass of the cannabis seed, half of the usable flower, and nearly all of the extracts can be contributed to essential oils such as terpenes, flavonoids, and actual cannabinoid content (1). The biodiversity of this plant is exhibited in the more than 2000 unique strains that have been identified, each with its own pigmentation, cannabinoid profile, and overall suggested medicinal use (2). Although early studies suggested the use of hops as a “matrix-matched” substitution for laboratories beginning method development, experienced technicians quickly realized the two matrices could not be treated as equals.

	The struggle to acquire blank, matrix-matched material for reference standards is a problem unique to cannabis laboratories. Clinical and food safety scientists are accustomed to purchasing blank, certified negative biological matrices for all of their testing panels and are even required to pass blind proficiency tests using these matrices. Though studies have been completed on best practices for laboratories analyzing for cannabinoid content in baked goods, cannabis laboratories looking for pesticides and mycotoxins do not have the luxury of simply purchasing (or growing) reference material (3). This hurdle then complicates the establishment of a relevant proficiency testing program. Without having a source for a pesticide-free, legally obtained cannabis flower, most proficiency testing programs are reduced to being essentially a system suitability check with a solvent standard containing the compounds in question (4).

	In addition to the complications surrounding the acquisition of a blank matrix, cannabis laboratories also find challenges in sourcing certain pesticide standards. Even after a supplier is identified, the cost of some compounds can prove to be prohibitive to laboratories looking to start up or expand a pesticide panel. Because state law makers have produced banned pesticide lists without consulting the proper scientists, growers, and cannabis experts, many laboratories find themselves attempting to obtain the active ingredients in pesticides that have been banned in the United States for decades. Additionally, some compounds are so unstable in typical extraction conditions that separate methods may have to be developed by the laboratory to comply with state regulations. Even if a reasonable list of pesticides to test for is established within a state, there is no guarantee that the limits of detection (LOD) and the limits of quantitation (LOQ) that need to be demonstrated by each laboratory are meaningful or significant. As shown in Table I, the detection level established in the state of Nevada for abamectin, bifenthrin, captan, cypermethrin, daminozide, fludioxonil, imidacloprid, paclobutrazol, and thiamethoxam are in the parts-per-billion (ppb) range. Such low LOD’s can be extremely difficult, if not impossible, to establish if laboratories do not have extremely sensitive and robust instrumentation. This problem is further complicated by the small batch production common in cannabis, and Nevada does not specify a minimum sample mass to be collected for testing.

	The development of an extraction method that can be used in a wide variety of laboratory settings is critical to the emerging field of medicinal marijuana testing. Within environmental and food testing laboratories, the QuEChERS (quick, easy, cheap, effective, rugged, and safe) technique has been widely used for the past 13 years. In 2003, Anastassiades and Lehotay published the first QuEChERS application, which discussed the determination of pesticide residues in produce (5). Since then, QuEChERS has become the analytical gold standard for the testing and analysis of a wide variety of difficult edible matrices, including oil, egg, meat, fish, wine, and beverage samples (5–9). Using disposable consumables, hundreds of pesticides can be analyzed in a single extraction with the QuEChERS approach. In addition to pesticide residues, other chemical classes such as antibiotics, veterinary drugs, mycotoxins, polyaromatic hydrocarbons, bisphenol A, and phthalates are routinely monitored using this technique (6–11). This approach generates much less solvent waste than what is typically associated with complex organic extractions and is a relatively easy analytical method to train technicians on. This advantage allows for laboratories to effortlessly adapt this system of sample preparation with minimal cross-training or cost to the customer and patient. 

	Traditional solid-phase extraction (SPE) columns and liquid–liquid extraction techniques cannot successfully provide laboratories with the reproducible, cost effective, and fast results needed for cannabis analysis. Unlike several commonly extracted biological matrices, plant-based materials do not easily flow through the porous frits and sorbent of an SPE column. In addition, they do not contain the same endogenous matrix interferences found in biological samples that ultimately need to be removed for accurate quantitation. Liquid-liquid extraction often requires large amounts of undesirable, costly, and toxic solvents to be used, and its overall schematic makes batch processing extremely difficult. The above limiting factors allow for QuEChERS to make a desirable transition to the cannabis laboratory community for pesticide and mycotoxin analysis.

	High performance liquid chromatography (HPLC)-grade acetonitrile, HPLC-grade methanol, and American Chemical Society (ACS)-grade formic acid were purchased from Spectrum. Neat pesticide and mycotoxin standards were purchased from Sigma-Aldrich, Chem Service, and Ultra Scientific.

	Dried cannabis samples were thoroughly blended into a fine powder using a Robot-Coupe food processor and dry ice. Figures 1a and 1b illustrate a cannabis sample before and after treatment. Homogenized 1-g cannabis samples were weighed into 50-mL polypropylene centrifuge tubes. To each of these samples, 10 mL of ultrapure water was added. Following a brief vortex mixing step, samples were allowed to hydrate for 15 min to improve extraction efficiency. Acetonitrile (10 mL) containing 2% formic acid was then added to each tube. The contents of a Mylar pouch containing an unbuffered QuEChERS blend (4 g of magnesium sulfate and 1 g of sodium chloride) were added, followed by a 5-min shake on a Spex 2010 Geno/Grinder homogenizer at 1500 rpm. All samples were then centrifuged for 5 min at ≥3000 rcf. Dispersive solid-phase extraction (dSPE) was then used for further sample cleanup, including concentration of the pesticides and mycotoxins in a 1-mL aliquot of the resulting organic layer. A blend of magnesium sulfate (150 mg), primary secondary amine (PSA, 50 mg) and ChloroFiltr (UCT Inc.) polymeric-based sorbent (50 mg) was packed into a 4-mL SpinFiltr tube. For comparison purposes, a second blend containing magnesium sulfate (150 mg), PSA (50 mg), and 7.5 mg of graphitized carbon black (GCB) was also used on an additional set of QuEChERS aliquots. PSA assists in the removal of pigmentation and organic acids, whereas magnesium sulfate further removes any remaining water from the extract. ChloroFiltr, which was designed for the selective removal of chlorophyll, was selected because of its effectiveness in removing pigmentation without sacrificing the recovery of planar analytes. GCB is commonly used in agricultural applications for the removal of pigmentation. A SpinFiltr tube (UCT Inc.) was selected for the sorbent blend to be packed into because of its reduction in sorbent carry over and increased sample purification abilities from the inclusion of a 0.2-µm PTFE frit. After transfer to this tube, all samples were vortexed for 30 s followed by centrifugation at ≥3000 rcf for 5 min. The resulting sample was then transferred to an autosampler vial for analysis.

	A Thermo Scientific Dionex UltiMate 3000 LC system coupled to a Thermo Scientific TSQ Vantage tandem mass spectrometer was used for all pesticide and mycotoxin analysis. EZ method (scheduled SRM) was used as the acquisition method with Xcalibur (version 2.2) software (Thermo Fisher Scientific) for data processing. A 100 mm x 2.1 mm, 3-µm Selectra Aqueous C18 HPLC column and a 10 mm x 2.1 mm, 3-µm Selectra Aqueous C18 guard column supplied by UCT were used for analyte retention and separation. Although other HPLC packing phases could be used, the polar modification of the aforementioned C18 material proved to be necessary for the retention of some of the extremely polar analytes included in the method. Mass spectrometry (MS) parameters and retention times for all pesticides and mycotoxins are shown in Table II.

	The column oven was maintained at 40 °C, the autosampler temperature was 10 °C, and the injection volume of all samples was 5 µL. Mobile-phase A was 5 mM ammonium formate in purified water (Milli-Q, EMD Millipore) with 0.1% formic acid, and mobile-phase B was 5 mM ammonium formate in methanol with 0.1% formic acid. The mobile-phase flow rate was 300 µL/min. The gradient for all sample analysis was as follows: 0 min, 0% B; 2–5 min, 50% B; 5.5–9 min, 60% B; 12–15 min, 100% B; and 15.1–20 min, 0% B. Wash solvent utilized was 1:1 (v/v) methanol–water.

	MS/MS was performed with heated electrospray ionization in negative mode (ESI-). The MS/MS system conditions were set as follows: discharge current at 4500 V (ESI

); sheath gas of nitrogen at 55 arbitrary units; auxiliary gas of nitrogen at 40 arbitrary units; ion sweep gas at 0 arbitrary units; vaporizer temperature at 400 °C; ion transfer capillary temperature at 350 °C; collision gas of argon at 1.5 mTorr; Q1 peak width of 0.7 Da full width half maximum (FWHM); and Q3 peak width of 0.7 Da FWHM.

	Quantitation was performed against a six-point matrix-matched calibration curve. With the exception of thiabendazole, which had an isotopically labeled internal standard, no internal standards were used for quantitation. All other compounds were quantified using absolute analyte peak areas. Using four replicates, absolute recovery was in the range of 70–100% for most compounds, with reproducibility less than 10% (Table III). The use of GCB was explored as an alternative to the chosen dSPE sorbent blend. Ultimately, better recoveries were obtained using the polymeric sorbent designed for chlorophyll removal (Table IV), with significant differences in recovery highlighted in red. If GCB must be used during the dSPE step, a very small quantity, between 2.5 and 7.5 mg is recommended. Sample cleanliness was similar for both dSPE sorbent blends, as shown in Figure 2. A traditional dSPE vessel, with selected sorbent blend packed into a 2-mL or 15-mL centrifugation tube, was not used to save overall sample extraction time. The use of a larger centrifugation tube with a filter instead allowed for a larger sample to be recovered without a tedious pipetting step. 

	Acetonitrile containing 2% formic acid and unbuffered QuEChERS salts were selected in the extraction step instead of other salt and acetonitrile combinations to combat the acidic ochratoxin A from being retained on the PSA sorbent during dSPE cleanup. This methodology contributed to the reduced recovery of pymetrozine, a basic analyte, as a result. The acidic pH resulted in insufficient partitioning of pymetrozine into the organic solvent layer.

	In spite of this compromise, some ochratoxin A was still retained on the PSA sorbent. Further acidification of the extraction solvent, such as increasing overall formic acid concentration to 5%, may improve the recovery of ochratoxin A. This modification may be explored if this mycotoxin is the most critical compound to the overall success of the extraction. Low recovery and high variability were obtained for ochratoxin A at the lowest fortification level because of the lower signal intensity observed at this concentration. Better results were obtained when evaluating the more highly concentrated samples. This lower signal intensity issue was also observed with acephate, aldicarb sulfoxide, chlorpyrifos, pyrethrin I, spiromesifen, and triethylphosphorothioate at lower concentration levels. Additional liquid chromatography–tandem mass spectrometry (LC–MS/MS) optimization and the inclusion of suitable isotopically labeled internal standards would further improve the overall performance of this method.

	Abamectin is prone to sodium (and potassium) adduct formation. The use of an ammonium buffer (acetate or formate) in the organic mobile phase can help to reduce sodium adducting by forming the [M+NH

 adduct. Despite this mobile phase selection, the MS signal for abamectin was still quite low and recovery data could only be generated for the highest spiked samples (500 ng/g). Spiromesifen is also an insensitive compound that exhibited a very weak signal for the protonated molecular ion (

 371.4). In spite of this, an intense peak at 

 273.1 was observed, possibly because of in-source fragmentation, and was successfully used for quantitative purposes.

	Acequinocyl is an insecticide that is included in the Nevada monitoring list (Table I). Although this compound could be readily detected by LC–MS/MS as a [M+H]

 parent ion when analyzing neat solvent standards, poor results were obtained when conducting spiking experiments and from the matrix-matched calibration curve. Thus, it was omitted from the method. To explain these results, its overall stability was examined. According to Ying and colleagues (12) it undergoes rapid phototransformation in an aquatic environment and rapid hydrolysis under neutral and alkaline conditions. The resulting major transformation product is hydroxyacequinocyl. In the previously mentioned study, it was observed that the highest intensity peak occurred at 

 343.3, because of the loss of ethenone (CH

=C=O). They also determined that positive atmospheric pressure chemical ionization (APCI

) provided vastly superior signal response over ESI

. For the problematic compounds included in this method, different LC–MS/MS parameters, including interface settings, should be further evaluated to obtain better signal intensity and lower detection limits.

	The method outlined above allows for the simultaneous analysis of 47 pesticides and five mycotoxins in cannabis in one simple QuEChERS procedure. By allowing for batch processing, this methodology allows for cannabis laboratories to save time and minimize solvent usage, leading to decreased costs. Analysis of the samples was performed by LC–MS/MS using an aqueous C18 HPLC column, which allowed for improved retention of the more polar pesticides included in the method. The method was evaluated by fortifying cannabis samples with each compound at four levels. The average recovery obtained was generally in the range of 70–100%, and the reproducibility was ≤10%.

	With the legalization of cannabis becoming widespread, this simple method is designed for implementation in either start up or established cannabis laboratories that are looking to streamline their sample preparation process, decrease their solvent usage, and provide their clients with accurate and fast results. By using an established extraction technique, such as QuEChERS, laboratories do not need to waste valuable research and development time on perfecting the best sorbent blends and consumables to use. Overall, this method is amendable to various pesticide monitoring lists for states that have already legalized cannabis or for those jurisdictions planning to do so.

		“Recommended Methods for the Identification and Analysis of Cannabis and Cannabis Products,” United Nations Office of Drugs and Crime (2009).

		C.E. Wolf, J.L. Poklis, and A. Poklis, 

		“Proficiency Testing for Medical Marijuana Independent Laboratories,” Nevada Division of Public and Behavioral Health (2015).

		M. Anastassiades, S.J. Lehotay, D. Stajnbaher, and F.J. Schenck, 

		S.C. Cunha, S.J. Lehotay, K. Mastovska, J.O. Fernandes, M. Beatriz, and P.P Oliveira, 

		M. Villar-Pulido, B. Gilbert-Lopez, J.F. Garcia-Reyes, N.R. Martos, and A. Molina-Diaz, 

		X. Ying, H. Cheng, X. Hao, H. Gao, M. Zhang, L. Wu, and H. Wang, 

Brian Kinsella, Tina Fanning, Danielle Mackowsky

 are with UCT, LLC, in Bristol, Pennsylvania. 

 is with Ace Analytical Laboratory in Las Vegas, Nevada. Direct correspondence to: 

Articles

The method described here allows for the simultaneous analysis of 47 pesticides and five mycotoxins in cannabis in one simple QuEChERS
procedure. This simple method is designed for implementation in start-up laboratories and in established laboratories that wish to streamline
their sample preparation process, decrease solvent usage, and obtain accurate and fast results.

Pesticide and Mycotoxin Analysis: Mastering the Complexity of the Cannabis Matrix

As a result of the rapid growth of the cannabis industry, many testing laboratories are looking for efficient, reliable, and cost-effective analytical methods to analyze chemical residues, such as pesticides, mycotoxins, solvent residues, terpenes, and heavy metals, as well as cannabinoid concentration in cannabis-infused edibles and beverages. In this article, QuEChERS (quick, easy, cheap, effective, rugged, and safe), a sample preparation technique widely adopted in the food testing industry, is introduced to the discipline of forensic testing as a viable method for the extraction of pesticides and cannabinoids in various complex sample matrices. The claimed amounts of cannabinoids versus the actual amounts are compared, as well as the pesticide residue levels in edible and beverage samples.

	To date, 25 states and the District of Columbia in the United States have legalized the medical use of marijuana, while four states and the District of Columbia have also legalized the recreational use of marijuana (1). Although the federal government still classifies any use or possession of the drug as illegal, all 50 states are starting to see an increase in the number of edible marijuana samples within their borders. As a result, many testing laboratories are looking for fast, reliable, and cost-effective methods to determine cannabis potency and chemical residues in cannabis edibles and beverages. The pros and cons of legalization are still heavily debated throughout the country, but all scientists agree that uniform testing policies and procedures need to be established as soon as possible and that overall sample cleanup is the main issue within these analyses. Many states legalized the use of recreational and medicinal marijuana without establishing any analytical protocols that are commonplace and routine in other scientific industries. Without any sort of regulatory control, laboratories in these states can analyze samples and report results without any fear of repercussion.

	For states where any consumption or possession of marijuana is still illegal, forensic laboratories are saddled with the task of analyzing this contraband. Even though it is extremely common for drug chemistry laboratories to directly test plant material and even oil-based products for the presence of tetrahydrocannabinol (THC) and related compounds, the use of edibles and beverages recreationally has brought along a new set of analytical challenges. Drug chemistry departments are not necessarily equipped to handle this type of analysis. In spite of forensic toxicology laboratories being capable of handling extractions from various biological matrices, baked goods and hard candies have few similarities to their everyday case load.

	One concept that does appear to be readily universal in the edible marijuana community is the establishment that 10 mg of THC is equal to one dose for this drug (2). While this standard works in theory, many edible manufacturers produce products that range from multiple doses in one package to just containing one individual dose. If proper interpretation of dosing instructions is not noted, some consumers may be put at risk for either undesirable effects from consuming too much in one sitting or no relief of symptoms at all for medicinal users. An imperative concept that novice edible users need to keep in mind is that digesting marijuana is not the same as smoking it when discussing the plant’s pharmacology. Those who have prior experience with smoking marijuana may assume that one brownie or one package of gummy bears equals one marijuana cigarette, though that is far from the case. Employees at marijuana shops may not be educated enough on this issue or be aware that edibles do not necessarily contain the THC amount as advertised on their packaging. This fact alone makes it even more critical that effective testing measures are put in place to ultimately protect the end users.

	The development of an extraction method that can be used in a wide variety of laboratory settings is critical to the emerging fields of recreational and medicinal marijuana testing. Within environmental and food testing laboratories, the QuEChERS (quick, easy, cheap, effective, rugged, and safe) sample preparation method has been widely used for the past 13 years. In 2003, Anastassiades and Lehotay published the first QuEChERS application, which discussed the determination of pesticide residues in produce (3). Since then, QuEChERS has become the analytical gold standard for the testing and analysis of a wide variety of edible matrices, including oil, egg, meat, fish, wine, and beverage samples (4–9). Using disposable consumables, hundreds of pesticides can be analyzed in a single extraction with the QuEChERS approach. In addition to pesticide residues, other chemical residues such as antibiotics, veterinary drugs, mycotoxins, polycyclic aromatic hydrocarbons (PAHs), bisphenol A, and phthalates are routinely monitored using this technique (10–14). The solvent waste generated is much less than what is typically associated with complex organic extractions. This technique is a relatively easy analytical method for technicians to learn, allowing for laboratories to effortlessly adopt this system of sample preparation.

	Neither traditional solid-phase extraction (SPE) columns nor liquid–liquid extraction techniques can successfully provide laboratories with the reproducible and fast results needed for cannabis food analysis. Unlike biological matrices, edible products do not easily pass through the porous frits and sorbent of an SPE column. In addition, they do not contain the same endogenous matrix interferences found in biological samples that ultimately need to be removed for accurate quantitation. Lastly, when analyzing the cannabinoid content in edible samples, the final extract often needs to be diluted rather than concentrated before instrumental analysis. This is in stark contrast to forensic samples, which often require concentration of target analytes at trace levels. Liquid–liquid extraction often requires large amounts of undesirable and toxic solvents to be used. The above limiting factors allow for QuEChERS to make a desirable transition to the forensic community.

	High performance liquid chromatography (HPLC)-grade acetonitrile, HPLC-grade methanol, and American Chemical Society (ACS)-grade acetic acid were purchased from Spectrum. Also, 35 neat pesticide standards were purchased from Sigma-Aldrich, Chem Service, or Ultra Scientific. A 2-ppm working solution containing 35 pesticides was prepared in acetonitrile. Three cannabinoids including THC, cannabidiol (CBD), and cannabinol (CBN) were purchased from Cerilliant in 1-mg/mL solutions. A 10-ppm mixture of the three cannabinoids was prepared in acetonitrile. Neat triphenyl phosphate was purchased from Cerilliant and diluted to 10 ppm in acetonitrile.

	Baked goods, chocolate bars, and hard candies were ground into a fine powder using a SPEX 6770 freezer mill before extraction. A hard candy sample before and after grinding is shown in Figures 1a and 1b. Gummy-based candies can be ground into powder with the presence of liquid nitrogen; however, the powder will return to its elastic gel state when the temperature rises. Thus, gummy-based samples were cut into fine pieces instead of using a freezer mill. Carbonated beverages, such as sodas, were degassed for 30 min before analysis, while oil samples were extracted without any sample pretreatment.

	Homogenized 1-g samples (baked goods, chocolate bars, hard candies, gummy bears, or oil samples) were weighed into a 50-mL centrifuge tube. To each of these samples, 10 mL of reagent water was added. Samples were hydrated for 1 h using a horizontal shaker. For beverage samples, 10 mL of degassed sample was added to 50-mL tubes without the water addition and the 1-h hydration step. Internal standard and 10 mL of acetonitrile containing 1% acetic acid were added to all samples, which were then shaken for 1 min using the SPEX Geno/Grinder homogenizer. A proprietary blend of QuEChERS extraction salts supplied by UCT was added to each tube, and the tubes were shaken vigorously to break up any salt agglomerates. The extraction salts help to facilitate phase separation and partition target analytes from the aqueous layer into the acetonitrile layer. After shaking, the samples were centrifuged for 5 min at 3000 rcf. Three distinct layers are formed after centrifugation as demonstrated in Figure 1c. The top layer is the organic phase (acetonitrile) containing pesticide residues, cannabinoids, and the organic-soluble matrix coextractives; the middle layer is the insoluble matrix components and water containing the water-soluble matrix components, such as sugars; and the bottom layer is the undissolved excess extraction salts.

Photographs of a hard candy (a) before and (b) after freezer mill grinding; (c) after QuEChERS extraction; and (d) the extracts before (left) and after (right) dSPE cleanup.

	For pesticide residue analysis, 1 mL of the supernatant was transferred to a 2-mL dispersive solid-phase extraction (dSPE) tube containing a proprietary blend of sorbents (UCT), and shaken for 1 min using the Geno/Grinder, then centrifuged for 5 min at 3000 rcf. This process removes chlorophyll, sugars, organic acids, and fatty compounds from the sample extracts by retaining them onto the sorbents. The resulting clean extract, illustrated in Figure 1d, is then diluted 2x with reagent water and analyzed by LC–MS/MS.

	For cannabinoid content analysis, dSPE was not necessary because of the high cannabinoid concentration in the acetonitrile extract. Instead, serial dilutions ranging from 200x to 20,000x were carried out to obtain a concentration (a few hundred parts per billion) that is suitable for LC–MS/MS analysis.

	A Thermo Scientific Dionex UltiMate 3000 LC system coupled to a Thermo Scientific TSQ Vantage tandem mass spectrometer was used for pesticide and cannabinoid analysis. Xcalibur (Version: 2.2) software was used for data acquisition and processing. A 100 mm x 2.1 mm, 3-µm 

 Selectra Aqueous C18 HPLC column and 10 mm x 2.1 mm, 3-µm 

 guard column supplied by UCT were used for analyte retention and separation. The aqueous C18 HPLC column was selected because of the high polarity of several pesticides covered in this study, such as methamidophos and acephate. Chromatograms of 35 pesticides and triphenyl phosphate (IS) are shown in Figure 2, with the retention order listed in Table I. The same HPLC column and mobile phases were also used for cannabinoid analysis to provide ease without the need of switching the HPLC column and mobile phases when analyzing the pesticide and cannabinoid samples in succession. The aqueous C18 HPLC column showed great separation of CBD and THC, two compounds that have the same multiple reaction monitoring (MRM) transitions. An example chromatogram of a diluted mint milk chocolate sample spiked with 70 ppb of three cannabinoids using this aqueous C18 column is shown in Figure 3.

 Chromatograms of 35 pesticides and triphenyl phosphate (IS) retained on the UCT Selectra Aqueous C18 HPLC column.

 Chromatogram of a diluted (2000x) mint milk chocolate sample spiked with 70 ppb of three cannabinoids.

	The column oven was maintained at 40 °C, and the samples in the autosampler were kept at 10 °C. The injection volumes were 2 and 5 μL for pesticide and cannabinoid analysis, respectively. Mobile-phase A was 10 mM ammonium acetate in Milli-Q water, and mobile-phase B was methanol with 0.1% formic acid. The mobile-phase flow rate was 300 μL/min. The gradient for pesticides was programmed as follows: 0 min, 0% B; 1–3.5 min, 50% B; 6–9 min, 95% B; and 9.1–14 min, 0% B. For cannabinoid analysis, the gradient program was as follows: 0–0.5 min, 60% B; 3–7 min, 95% B; and 7.1–10 min, 60% B.

	Tandem MS was operated with heated electrospray ionization (HESI) in positive mode. The MS/MS conditions were set as follows: spray voltage at 3500 V; sheath gas of nitrogen at 50 arbitrary units; auxiliary gas of nitrogen at 40 arbitrary units; vaporizer temperature at 450 °C; ion transfer capillary temperature at 350 °C; collision gas of argon at 1.5 mTorr; Q1 peak width of 0.4 Da full width half maximum (FWHM); and Q3 peak width of 0.7 Da FWHM. The optimization of the MS/MS transitions was performed individually for each analyte by infusing 1 μg/mL of standard in acetonitrile at 10 µL/min in a 50:50 A–B mobile phase at a flow rate of 300 µL/min. Scheduled selected reaction monitoring (SRM) with a cycle time of 0.5 s was set for data acquisition. The precursor and product ions, collision energies, and S-Lens RF values of the 35 pesticides, IS, and three cannabinoids are listed in Table I.

Semiquantitative Determination of Pesticide Residues

	Pesticide residues were analyzed semiquantitatively because of the lack of negative control samples for each sample matrix, as well as the large number of pesticides at different concentration levels making the standard addition method impractical. Matrix-matched calibration curves were generated using the post-spiked blank extracts of green tea sample to semiquantify the pesticide residues in edibles and beverages. Appropriate amounts of the pesticide spiking solution were added into the tea extracts to generate six-point matrix-matched calibration curves with concentrations at 5, 10, 25, 50, 100, and 250 ng/mL. The responses were found to be linear (

 > 0.99) over the entire concentration range. Accuracy and precision results at 10-ng/g and 50-ng/g spiking levels are shown in Table II.

	A variety of edibles, including chocolate bars, brownies, hard and gummy candies, CBD oil, and beverage samples, were analyzed for pesticide residues using the QuEChERS extraction and dSPE cleanup method described above, and the results are summarized in Table III. Four products were found to contain bifenazate at concentrations ranging from 10 to 1221 ng/g, although some of them claimed to be “organically grown” products. The highest concentration was found in a CBD oil sample, where the pesticide might have been enriched when manufacturing the concentrated oil product from the raw plant materials. Bifenazate is commonly used to control mites on agricultural products. Though it is not terribly harmful to humans in comparison to other pesticides, there is a lack of research regarding the health risk of constant consumption of this pesticide. Most pesticide research does not take into account the oral dosing over a long period of time, especially in people with compromised immune systems.

Quantitative Determination of Cannabinoid Content

	For cannabinoid analysis, content was quantitated by the standard addition method. Diluted QuEChERS extracts were spiked at +50% and +150% of the stated cannabinoid content. Plotting the peak areas of the diluted sample and the two aforementioned spiked samples allowed for the calculation of cannabinoid concentrations in the diluted samples. Excellent linearity (

 > 0.995) was observed for all samples tested. An example of the standard addition calibration curve is demonstrated in Figure 4. Calculation of the cannabinoid contents in the original samples was based on the dilution factors and unit conversion factors. The results (in milligrams per pack) were then compared to the stated content on the packaging of each product, as summarized in Table IV. For all of the cannabis samples tested in this study, only 31% (four out of 13 samples) had accurately labeled the total cannabinoid content within ±20% of the stated values.

 Standard addition calibration curve of THC in a cookie and cream bar sample (500x dilution of the QuEChERS extract, 70 ppb cannabinoids were added for the 50% spiked sample and 210 ppb were added for the 150% spiked sample).

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	Although the US Food and Drug Administration takes care of monitoring the safe production of the majority of edible products in the United States, marijuana-infused foods do not fall under their jurisdiction because the drug is still considered to be illegal in the eyes of the federal government. Recreational users may be primarily focused on getting their money’s worth from these products, but medicinal users have additional, notable concerns with edibles. Many medicinal consumers do not necessarily want to feel the psychoactive effects of marijuana, but rather just want to combat the symptoms of their specific ailment. If a product is inaccurately labeled, medicinal users run the risk of consuming too much marijuana and thus having undesirable side effects. On the contrary, they also could be unnecessarily suffering from side effects of their illnesses because of an edible product containing less cannabinoids than the suggested dosage amount. These concerns are in addition to the possible presence of unwanted chemical residues, such as pesticides, mycotoxins, and heavy metals.

	A fast, reliable, and cost-effective analytical method has been developed for the determination of pesticide residues and cannabinoid contents in cannabis infused edibles and beverages. This method uses the advantages of QuEChERS products to extract 35 pesticides and three cannabinoids (CBD, CBN, and THC) from complex food and beverage matrices. This extraction is then followed by either serial dilution for cannabinoid content analysis, or a dSPE cleanup for the efficient removal of various matrix coextractives, resulting in extracts for trace-level pesticide residue analysis. This hybrid method allows QuEChERS, which is extensively used in the food testing industry, to be used in a forensic or private cannabis laboratory setting.

	The developed method has been successfully applied to the analysis of 13 real cannabis edible and beverage samples. For pesticide analysis, bifenazate was present in four out of the 13 tested samples at varied concentrations ranging from 10 to 1221 ng/g. For cannabinoid content analysis, only 31% (four out of 13) of the samples had accurately labeled the total cannabinoid content within ±20% of the stated value, while the majority of samples (eight out of 13, or 62%) were overlabeled (more than 20% below the labeled amount). One sample was found underlabeled which actually contained 40% more THC than stated.

	Keith Tucker with SPEX SamplePrep LLC is acknowledged for providing the 6770 Freezer mill and 2010 Geno/Grinder for this study. Erik Swiatkowski with UCT is thanked for his help in handling liquid nitrogen when using the SPEX 6770 freezer mill.

		A.A. Monte, R.D. Zane, and K.J. Heard, 

		State Medical Marijuana Laws, National Conference of State Legislators 18 April 2016.

		B. Kinsella, S.J. Lehotay, K. Mastovska, A.R. Lightfield, A. Furey, and M. Danaher, 

		L. Vaclavik, M. Zachariasova, V. Hrbek, and J. Hajslova, 

		T.H. Kao, S. Chen, C.J. Chen, C.W. Huang, and B.H. Chen, 

		S.C. Cunha, C. Cunha, A.R. Ferreira, and J.O. Fernandes, 

Xiaoyan Wang, Danielle Mackowsky, Jody Searfoss

 are with UCT in Bristol, Pennsylvania. Direct correspondence to: 

QuEChERS is introduced to the discipline of forensic testing as a viable method for the extraction of pesticides and cannabinoids in various complex sample matrices.

Danielle Mackowsky

Articles

Pesticide and Mycotoxin Analysis: Mastering the Complexity of the Cannabis Matrix

Determination of Cannabinoid Content and Pesticide Residues in Cannabis Edibles and Beverages