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

February 28, 2018
Table I: Pesticide and mycotoxin detection levels for the state of Nevada
Click to enlarge, Table I: Pesticide and mycotoxin detection levels for the state of Nevada
Figure 1: Cannabis sample
Click to enlarge, Figure 1: Cannabis sample before (left) and after (right) homogenization with dry ice.
Abstract / Synopsis: 

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. (See upper right for Table I, click to enlarge.) 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.

Experimental

Reagents and Standards
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.

Sample Preparation
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. (See upper right for Figure 1, click to enlarge, caption: Figure 1: Cannabis sample before (left) and after (right) homogenization with dry ice.) 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.

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