I am honored to contribute to this inaugural edition of 

! As the organizer of many cannabis science events (including the Cannabis Science Conference, Canna Boot Camps and Analytical Cannabis Symposia at Pittcon), I have worked to bridge gaps between the cannabis industry, analytical science, and medicine. This magazine represents a major leap forward in bridging these gaps, and the timing could not be more perfect.

It is my great pleasure to feature an interview-style article in each issue of 

 where I will introduce you to cannabis science mavens and cover a diverse range of important topics. For this first installment, I focused on a cornerstone of cannabis science-cannabis quality control (QC) testing laboratories, and their evolution during the past decade.

Cannabis laboratory testing has evolved from a garage “pot science” to a multimillion-dollar industry in a very short time span. There are many companies that have played important roles in catalyzing cannabis science. Laboratories like Steep Hill, Trace Analytics, Nordic Labs, and EVIO Labs come to mind as leaders instilling quality in laboratory testing. Recently, I sat down with Lori Glauser, Co-Founder, COO, and Director of EVIO Labs, and Chris Martinez, President of EVIO Labs Florida, to discuss EVIO as well as what lies ahead for cannabis quality control testing.

Could you please introduce EVIO Labs to our readers? How many laboratory locations does EVIO have now?
	Lori Glauser:

 EVIO Labs has quickly become a national provider of accredited cannabis testing, providing high quality analytical and consulting services for the cannabis industry. We currently have eight operating laboratories in five states (see Figure 1, right). We have several additional laboratories in various stages of development, and are working toward our goal of opening 18 locations by the end of 2018.

What are the challenges of working in different states? How do you handle those challenges (different strategies)?
	Glauser:

 Each cannabis-legal state has unique and evolving regulations. As a multistate operator, we work to standardize our methods across labs and across states, and building our business to efficiently meet the needs of all the states rules can be challenging. So far, no two states’ testing requirements are identical. It is imperative for us to stay abreast of the regulatory changes in each state we work in, and also keep an eye on regulatory changes in states where we don't yet have labs to establish where we want to go next. We anticipate that testing regulations will converge in the future, and we will be prepared to meet a future national standard.

When I think about the evolution of cannabis science, EVIO Labs comes to mind. EVIO has expanded operations and invested in credentialed, knowledgeable staff, and world-class analytical instrumentation. How is EVIO evolving cannabis testing?
	Chris Martinez:

 EVIO Labs Florida (see photos in Figure 2, right) is evolving the cannabis testing landscape by bringing together numerous laboratory partners dedicated to providing clients with consistent high-quality cannabis analytical services backed by quality assurance and consulting services supported by the expertise of an extensive network of industry professionals. Our clients gain access to a unique network of published research scientists, content experts, consulting partners, and business leaders to accomplish the needs of even the most complex projects.

How challenging is it to operate high-quality laboratories in an industry where customers don’t always understand why quality is so important or expensive?
	Glauser

: Customer education is key. Some of our customers who have been in the industry for a long time launched their businesses in an unregulated or lightly regulated environment. They are challenged with transforming their businesses to meet a whole new regulatory paradigm-not just for testing, but for security, inventory tracking, packaging, and more. Many of these customers are going through a learning process, and we help provide them with that education. For instance, we explain to customers that cannabis testing costs have increased largely because of the increased quality requirements. In the past, a $50 potency test run on an instrument that may not have been validated or calibrated was sufficient. Now requirements for quality testing panels that include pesticides, residual solvent, and microbiological contaminants using expensive instruments run by experienced scientists have increased to a $400 or more compliance test in regulated states. With the new quality requirements come far more accurate, thorough, and dependable results.  The requirements are not unlike tests performed for food, nutraceutical, and pharmaceutical products where the cost of testing can reach $1000 or more and take weeks to complete. We estimate that full compliance testing in states like Nevada, Oregon, and California range from under 1% to 5% of the of the wholesale value of the batch, depending on batch sizes.

During the early months of the transition of Oregon's cannabis rules last fall, we did receive a lot of calls from customers who were dismayed by the new rules and frustrated with the increased prices. We counsel each customer and explain to them the process, including the additional instrument, expertise, and time requirements involved in each test. Another challenge customers face is that the turnaround time increases when the quality standards are increased. What used to take one day can take three or more days because of the level of rigor and quality checks required for each test. For growers who have already waited months for them to grow, harvest, and cure product, the extra few days waiting for test results can be stressful, especially for those who need to sell their product to generate revenue.

What types of certifications and accreditations does EVIO Labs have?
	Glauser

: All of our laboratories are either currently accredited or are seeking accreditation in their respective states. Our Oregon locations are accredited by the Oregon Environmental Laboratory Accreditation Program (ORELAP). Our laboratories in Colorado and Florida have been accredited to the ISO 17025 standard, and we are currently going through the accreditation process in our California and Massachusetts locations. Our Berkeley, California, location currently has a temporary license by the Bureau of Cannabis Control, and we are building out our Orange County location now, with expected licensing this spring.

How time-consuming or expensive is it to obtain ISO 17025 accreditation?
	Glauser

: The first time EVIO worked toward accreditation, which was based on the The NELAC Institute (TNI) standards, was in 2016, and the process took approximately six months. During that time, we refined and developed more than 120 standard operating procedures, policies, methods, and forms; trained our staff to meet the quality requirements; validated our instrumentation and methods; and performed "proficiency tests," which are tests we perform that are verified by third parties for accuracy. The TNI standard is a more rigorous set of nearly 900 requirements that includes all of the approximately 300 ISO 17025 standards. Once our first four laboratories were accredited, each laboratory thereafter took less time because we use the same core quality management system for all of our laboratories. EVIO Labs Florida used EVIO's core quality system and was accredited within three months. We anticipate each new laboratory will take approximately two months to go through the process. Any laboratory that is getting accredited for the first time to ISO standards should plan a 4–6 month timeline, depending on how well designed their quality management system is. The expense of engaging with the accreditation agency, generally under $10,000, is nominal compared to the massive effort required by our technical staff to prepare for accreditation-which could be upwards of $250,000 in labor plus the opportunity cost associated with our technical staff working on building the quality management system rather than serving customers.

How different are the accreditation processes from state to state? Is there one state that has a particularly good system?
	Glauser:

 States are quite different, but several are converging on the ISO 17025 standard, and third-party verification of laboratory proficiency. Nevada, for instance, doesn't have a third-party accreditation requirement; however, the state's health department has taken a keen interest in regulating the laboratories directly. They have gone so far as shutting down laboratories temporarily for identifying quality lapses. Oregon's more stringent TNI requirements are enforced by the Oregon Health Authority's ORELAP department. We believe this is a particularly good system because the guidelines are very clearly laid out, and it provides far more detailed guidance than the ISO 17025 standards alone. Representatives in other states including New York have indicated that they look to Oregon's system as a model. California is implementing an ISO 17025 program, which I predict will become the minimum standard nationwide. I also believe that states that have a mandatory proficiency test, such as required in Oregon and newly required in California, is important. This means laboratory test results are periodically validated by a third-party to ensure accuracy.

Improved quality management systems (QMS) play a critical role in the cannabis testing evolution. Can you comment on other programs that are critical to professional integrity?
	Glauser:

 All EVIO Labs operate under a common and rigorous QMS to ensure that our results are accurate and reliable. We also implement a continuous improvement program that brings together best practices, as well as customer, regulator, and assessor feedback from across our growing network of laboratories.

Our four Oregon locations are NELAP accredited by ORELAP and licensed by the Oregon Liquor Control Commission (OLCC) to test cannabis. This is an accreditation standard that meets and exceeds the ISO 17025 requirements, and is the quality standard for all of our laboratories. Our strategy for quality management is continuous improvement of our standards, as we bring together best practices from all the laboratories we acquire or build. Our qualified lab directors from across the nation meet at least weekly to share best practices, and continuously improve our core quality management system. For example, whenever one laboratory goes through the accreditation process, we gather feedback from the accreditation agencies, and apply any improvements to our laboratories nationwide.

Can you please tell us how EVIO Labs Florida maintains a competitive edge, and what do you envision for the future of EVIO Labs and the cannabis testing industry?
	Martinez:

 As a veteran-owned enterprise we fulfill the state mandated diversity plan requirement for current license holder renewals and new cultivator applications. EVIO is the first to market in Florida with a world-class ISO-accredited lab that has support of EVIO’s laboratories across the United States. Our customer service is what we pride ourselves on, making ourselves available 24/7 to discuss results and support clients in reaching only the highest quality cannabis for Florida patients. The future for EVIO is bright. We are currently in expansion mode with our second laboratory in Gainesville, Florida, and are in the process of identifying a third location in Lakeland, Florida. The recent expansion into multiple cities is patient driven. EVIO offers clients the latest technology available for testing cannabis at the most competitive rates in the country. As EVIO raises the bar in cannabis testing nationwide we anticipate future laboratories will take note of the immense responsibility they have to support clean and safe cannabis through the latest analytical technology available and passionate staff who constantly put patients first. 

As we work together to move cannabis science forward, it is important to honor the pioneering individuals that not only pave the way for others, but also bring quality and professionalism to the industry. Join me every quarter for interviews with key opinion leaders in many areas of cannabis science, and please continue to support and grow 

 is the columnist and editor of “Cannabis Crossroads” and a contributing editor to 

magazine. Crossney is also the president and CEO of CSC Events. Direct correspondence to: 

Articles

The Evolution of Cannabis Quality Control Testing

The discovery of nonregistered pesticides contamination in cannabis plant material in Colorado prompted the need to set mandatory reporting limits (MRLs) for these pesticides in the state to ensure public safety. Since then, the Colorado Department of Public Health and Environment (CDPHE) undertook the responsibility for setting such MRLs. Colorado’s novel approach to determine MRLs for each pesticide involved forming a working group with its in-state laboratories and expert volunteers to agree on analytical procedures to determine method detection limits (MDLs) using an industry-standard approach. The authors describe how this approach formed a collaborative public-private partnership where state government agencies worked side by side with laboratories and growers toward a common goal. The multilaboratory study was completed yielding MDLs for 13 pesticides in cannabis flowers using a QuEChERS (quick, easy, cheap, effective, rugged, and safe) extraction with analysis by liquid chromatography-tandem mass spectrometry (LC-MS/MS). The authors share the procedures used for the study, an overview of the data generated by the participating laboratories, and the new Colorado MRLs, which became effective on January 1, 2018.

The regulation of pesticide use on cannabis is a complex issue for states with legalized marijuana. Pesticide use is regulated at the federal level; however, cannabis is not a specifically listed crop on any registered pesticide label, and the United States Environmental Protection Agency (US EPA) does not consider cannabis to fit into any general crop groups, such as herbs or spices (1). In addition, no risk assessments have been conducted specifically for pesticide use on cannabis and tolerance limits have not been defined. Pesticide use in cannabis was recognized as a public health and safety concern, but the information deficit and limited available resources left the state of Colorado grappling with how to handle this complex issue when the Denver Department of Environmental Health placed the first holds on 100,000 marijuana plants for improper use of banned pesticides in March 2015.

In the following months, many additional quarantine and recall events occurred. In November 2015, Colorado issued a statewide pesticide policy and executive order (2) addressing the use of pesticides in marijuana. These documents mandated state agencies to consider cannabis contaminated with banned pesticides a threat to public safety, until such a time that scientific assessment establishes which additional pesticides can be safely applied to marijuana.

After issuance of the executive order and statewide policy, the next question was how to get private laboratories certified to perform pesticide testing so the state could mandate this testing for cannabis. Two primary pieces of information were needed: what pesticides are laboratories expected to test for and what are the associated limits? The executive order has frequently been called a zero tolerance policy; while it does establish zero tolerance for the use of a banned pesticide, analytically it is not possible to test to a concentration of zero. Without defining minimum testing requirements, different results would be reported across laboratories. To address these issues, the state initiated a stakeholder process to determine what pesticides should be tested for and what the associated mandatory reporting limits (MRLs) should be.

The Colorado Department of Public Health and Environment (CDPHE) took the lead on oversight of this stakeholder process, which engaged marijuana testing facilities, cultivators, product manufacturers, scientific experts, and other interested parties. The working group first tackled the list of required analytes. The final list selected included 13 pesticides that were considered the most important for testing in Colorado because they are banned for use and have been previously detected in marijuana in the state.

The working group then determined that in the absence of established pesticide tolerance limits, the regulatory limits should be based upon laboratory detection capability. At that time, to obtain this information, a unique public-private collaboration was initiated to design and execute a multilaboratory method detection limit (MDL) study engaging the resources of Colorado’s private marijuana testing facilities and the state’s Department of Agriculture (CDA) laboratory. The results of the study would be used to inform the regulatory limits for the list of 13 pesticides in marijuana. A technical subcommittee of scientists was formed from the stakeholder working group to assist CDPHE in designing the study and reviewing the results.

Following the decision to design a multilaboratory study (MLS), Restek volunteered to help CDPHE form a study design and oversight (SDO) team to ensure the multilaboratory study’s success. Since no study of this kind had ever been attempted in the past and there was no standardized analytical method protocol for participant laboratories to follow, the SDO team’s goal was to minimize variables by applying study guidelines based on industry-standard practices. Also, since the marijuana flower buds are an expensive commodity and time was of the essence given the governor’s directive, careful planning and execution were required to produce statistically valid data that the state could use for decision-making upon completion of the first round.

The team consisted of two CDPHE personnel, one representative from Neptune and Company, one from Lowry Consulting-all based in Colorado-and three from Restek Corporation located in Bellefonte, Pennsylvania. The team began to meet regularly to design guidance documents by which labs would be asked to conduct their testing.

The overall study was designed to progress through three sequential phases one built upon another, culminating with participating laboratories completing all of the steps toward reporting MDLs for pesticides in marijuana buds. These steps included determining an instrument detection limit (IDL), matrix quality control check (MQC), and a formal MDL. Three documents were constructed by the SDO team to guide laboratories through the MLS process followed by the technical subcommittee review, which included the volunteer labs. The technical subcommittee decided that liquid chromatography-tandem mass spectrometry (LC-MS/MS) would be the required analytical technique for the analysis to produce the best MDL data.

The first document described the steps to be taken by each laboratory to establish IDLs for each compound on their LC-MS/MS instrument. While some laboratories had already established such limits, this procedure was important as a starting point since some laboratories had either just purchased their instrument, or had not already completed this step in their laboratory. It was also necessary to establish whether the participant laboratories would be able to detect all 13 compounds at similar levels before spiking into actual matrix. 

The second document, an MQC procedure, described steps to be taken by each laboratory to spike all 13 pesticides into cannabis flower extracts at varying concentration levels, then analyze eight replicates at each level to observe detectable levels they could achieve given acceptance criteria before executing the MDL study. The third document, the MDL procedure, described steps laboratories would follow to spike a dried flower matrix, verified to be free of the analytes of interest, then take it all the way through extraction, cleanup, and analysis. It is important to note that all three documents incorporated guidelines established by the SANTE 2015 Guidance document on analytical quality control and method validation procedures for pesticides residues analysis in food and feed (3). Such guidelines included transition ion ratio and calibration acceptance criteria. The MDL document incorporated the procedure from the 

Other steps taken by the SDO team and stakeholder working group in Colorado included further minimizing variables, requiring all participating laboratories to use the same reference standards from a single source and a composite sample consisting of cannabis buds donated by a grower in Colorado. Restek donated two reference standards mixes and OrganaLabs donated the Cannabis buds. The Colorado Department of Agriculture (CDA) prepared extracts and dried plant composite materials to the study laboratories by grinding, compositing, and extracting throughout the various phases of the study. 

The SDO team decided that at each of the three phases of the study laboratories would submit their results in spreadsheets provided along with each study document. The results were accompanied by raw data components as outlined in each of the three documents. All data submitted were reviewed by the SDO team to maintain confidentiality of the study. It was also decided, after completion of the IDL phase, that a nondisclosure agreement would be signed by the SDO team members to protect any proprietary information laboratories might share during the course of the study. This step contributed to a two- to three-month delay in the process before launching into phase two.

Before launching the MLS, training was offered to laboratory personnel and the CDPHE to ensure that participants were aligned on a general procedure in the absence of a published test method. The training was coordinated by Restek and Shimadzu personnel, who designed a one-day hands-on pesticides sample preparation and analysis workshop. CDPHE hosted the event, providing laboratory space and safety equipment for participants. Three suppliers contributed equipment, consumables, instrumentation, and personnel free of charge to the study group and CDPHE. Spex Sample Prep loaned its grinder for sample processing, Shimadzu Scientific Instruments loaned its LC-MS/MS instrument and personnel for extract analyses, and Restek donated reference standards, sample preparation consumables, and personnel time. Restek and Shimadzu instructors led 19 participants through the steps of a hands-on QuEChERS (quick, easy, cheap, effective, rugged, and safe) pesticides extraction with dispersive solid-phase extraction (dSPE) clean-up and filtration, followed by instrument setup and analysis of extracts by LC-MS/MS. Each participant prepared his or her own sample extract from donated dried 

buds, spiking 11 pesticides at 100 ppb and two pesticides at 1000 ppb into 1.5-g samples. The combined results of the 19 extracts are shown in Table I. (See upper right for Table I, click to enlarge.)

Before the study was launched, it was decided that the participant laboratories would follow their own internal method procedures for the extraction and analysis of pesticides in a cannabis bud matrix, and incorporate what they had learned from the training as necessary. A 3.0-g sample would be utilized by CDA to create the composite extracts for the MQC study phase. Participant laboratories used between 1.5 g to 2.0 g of sample for the MDL phase because the actual amount varied from one laboratory to another based on their internal procedures.

The initial stakeholder groups were formed in January 2016 to address the Executive Order 2017-015 (1) and implementation of pesticide testing. The group spent several months gathering information concerning the determination of pesticide limits from other state programs, comparing regulatory limits, and evaluating the best scientific approach before the decision was made to work with the laboratory community as a whole on establishing pesticide MRLs.

The study design, organization, and initial recruitment of laboratories to participate was completed by July 2016. Although nine laboratories volunteered to participate in the study, MDL data from eight laboratories would eventually be received by the close of the study. The IDL study was initiated in mid-July 2016 with data delivery reported in August; however, the data submittal timeline was extended to laboratories that required additional time. The data for the IDL studies were compiled and released in November 2016 followed by the MQC phase, which began with the release of study samples in January 2017. After the MQC study data were gathered and processed, it was released in April 2017. The final step was for the laboratories to generate their MDLs using homogenized dried bud samples, which were provided to the laboratories in June 2017 with a study close date of July 2017. The data submitted by eight volunteer laboratories were statistically evaluated and reviewed, then released to the participating laboratories, CDPHE, and CDA in September 2017. From start to finish, the entire study took less than 20 months to complete.

The reported data collected across all of the analytes consisted of a small number of results for each analyte. The removal of any outliers would have had a large impact on the data. As a consequence, only one data point was removed as an outlier for the analyte permethrin. The data point was removed because it was an order of magnitude greater than any of the other data reported for permethrin.

Basic robust statistics were calculated: the mean, median, variance, standard deviation, percent relative standard deviation, and the upper limits 95% and 99% confidence intervals.

Figure 1 displays how the data compare across the analyte list for average MDL, minimum and maximum values, and the confidence intervals. (See upper right for Figure 1, click to enlarge.) The data for spiromesifen and malathion demonstrate the greatest amount of variability for the analytes in the study. Though not surprising, the extent of the variability was not expected. Removing spiromesifen, the remaining analytes fall into two categories, analytes below 30 ppb and 15 ppb, which is similar to MDL values found in other complex plant matrices.

Figure 2 shows relative standard deviations (%RSDs) for the eight volunteer laboratories for all analytes and represents several key factors that are being addressed by the industry: The lack of reference methods, inexperience with this complex analysis, and an extremely complex interfering matrix. (See upper right for Figure 2, click to enlarge.) The SDO team recognized these factors as key influencers affecting the higher %RSD values.

The SDO team recommendation was for the state to adopt the 99% confidence interval as the initial MRL for each analyte. This recommendation would incorporate the capabilities of the current laboratory community into the MRL determination, ensuring that the laboratories would at least be able to detect the analytes of interest.

The CDPHE was tasked with determining the MRLs for the pesticides based on the data and recommendation of the SDO. CDPHE staff including toxicologists, statisticians, marijuana laboratory certification staff, and analytical scientists were gathered. This group reviewed known toxicological data and other state pesticide MRL information to set the recommended MRL. They developed recommended MRLs for each pesticide and these were presented to the stakeholder group. After receiving public comment and feedback, the information was collected and the MRLs at the 99% confidence interval were incorporated into draft rules for consideration in official rule-making procedures.

The state of Colorado announced the rule-making process was completed and Rule 712 was adopted on November 17, 2017 and became effective January 1, 2018. The new rule, in addition to other contaminants, includes the 13 pesticides now required for testing and the MRLs are based on the MDLs from this multilaboratory study (4). See Table II for a summary of the MRLs. (See upper right for Table II, click to enlarge.) The approach of incorporating MDLs to set MRLs was challenged by members of the analytical community in Colorado who cited that they were not risk-based. However, regulators in Colorado persisted, explaining that the approach was sound given that the MRLs are not meant to be risk-based limits, but are instead set to ensure laboratories can detect banned pesticides that are not permitted for use in Colorado with acceptable precision, accuracy, and sensitivity from a cannabis bud matrix.

A number of noteworthy achievements have resulted from this effort. It is important to recognize this is the first and only study of its kind in the cannabis industry. This effort demonstrated that public-private partnerships can successfully resolve challenges while saving time and money through volunteer effort. It also demonstrated that when such efforts are undertaken with appropriate leadership, experience, and planning, the timeline for completion is shortened. Taking less than 20 months to complete a study of this magnitude is generally considered astoundingly fast in the science community. In addition, using a science-based approach to setting regulatory levels for contaminants in cannabis buds to protect public health is also a novel and uncommon approach among rule-making practices. Its successful application in this case demonstrates a viable approach that might be applied to similar challenges in other states as cannabis regulations emerge and develop. Finally, observing government and industry working together as a stakeholder community, volunteering time, equipment, materials, and space to solve a problem is not only encouraging, but shows that when given proper definition of a problem accompanied by a solution the community will rise to the challenge and deliver beyond expectations.

The study represents a public-private partnership among state and local government agencies with the marijuana industry, testing laboratories, cultivators, extraction laboratories, and infused product manufacturers. The goal was to provide a scientific, fact-based recommendation for the determination of pesticides in marijuana MRLs. As we noted with regard to the large variability of the %RSDs, the industry will benefit from standardized test methods, and greater experience. The process should be reviewed in the future once laboratories have developed improved methods and gain additional experience. The MDL procedures for each laboratory should be completed annually. A review of these collected MDL studies would yield a good potential metric to repeat the study. This will allow the MRLs to be adjusted to the current laboratory competencies.

The authors wish to thank the following individuals, government agencies, and companies for their significant contributions to the success of this collaborative effort: Eric Petty, CDPHE; Julie Kowalski, Trace Analytics; Jeff Lowry, Lowry Consulting; Jack Cochran, VUV Analytics; Jeff Dahl, Shimadzu Scientific Instruments. The Colorado volunteer cannabis testing laboratories: Gobi Labs, TEQ Analytical Laboratories, RM3 Labs, Nordic Analytical Laboratories, Agricor Laboratories, Aurum Labs, and CDA’s Laboratory. The Colorado Department of Revenue, The Colorado Department of Agriculture, The Colorado Department of Public Health & Environment, Neptune and Company, Organa Labs, Spex Sample Prep, Shimadzu Scientific Instruments, and Restek Corporation.

 is with Restek Corporation in Bellefonte, Pennsylvania. 

 is with the Colorado Department of Public Health & Environment (CDPHE) in Denver, Colorado. 

 is with Neptune and Company, Inc., in Lakewood, Colorado. Direct correspondence to: 

Factual and Policy Issues Related to the Use of Pesticides on Cannabis

, (paragraph 4), CDA link: https://drive.google.com/file/d/1V_zVKP9R8qpdgYJ2r OrJR39b6MA1M9AH/view.

Office of the Governor, Denver, Colorado, D2015-015 

Executive Order Directing State Agencies to Address Threats to Public Safety Posed by Marijuana Contaminated by Pesticide

European Commission Directorate-General For Health and Food Safety, SANTE 2015 Guidance document on analytical quality control and method validation procedures for pesticides residues analysis in food and feed (SANTE/11945/2015 30 November-1 December 2015 rev. 0).

, 40 Appendix B to Part 136 -Definition and Procedure for the Determination of the Method Detection Limit-Revision 1.11. (J U.S. Government Printing Office, Washington, D.C., 2011).

Colorado Retail Marijuana Code Adopted Rules, CCR 1 212-2, Rule 712, Section R 712 - 

Retail Marijuana Testing Facilities: Sampling and Testing Program, E. Permissible Levels of Contaminants

, (Adopted November 17,2017; effective January 1, 2018), pp. 124-125.

Using Method Detection Limits to Set Mandatory Reporting Limits for Banned Pesticides: How a Public-Private Partnership Improved Cannabis Regulation in Colorado

Evolution in the legislation regarding the production and consumption of cannabis and related products for medicinal and recreational uses has led to emerging regulations regarding the potency, terpene profiles, and impurity content of such products. Among the impurities, or contaminants, that require testing are metals. Because of their toxicity and carcinogenicity, most jurisdictions are, at a minimum, requiring testing for arsenic, lead, cadmium, and mercury. Metals accumulate in plant material through normal metabolic processes. Furthermore, some plants are capable of hyperaccumulating metals to concentrations well above those in the soils and waters that nourish them. Some such metals are beneficial or nutritional in nature, whereas others show varying levels of toxicity. As such, testing for metals, particularly ones that are toxic, are important for products destined for human consumption. Quantitation of metals within cannabis materials can be accomplished through a variety of analytical techniques. Such amenable technologies include atomic absorption (AA) spectroscopy, inductively coupled plasma-optical emission spectroscopy (ICP-OES), and inductively coupled plasma-mass spectrometry (ICP-MS). As with all aspects of analytical instrumentation, each method has its advantages and disadvantages. Here, we present an overview of analytical methodologies, challenges facing the analyst, and notes on regulatory stipulations.

Recent shifts in legislation efforts in the United States and other countries have led to the use of cannabis and related products (for example, cannabis-based extracts, tinctures, and “edibles”) for medicinal or recreational purposes. In the United States, legalization efforts have been concentrated at the state level, with nine states and the District of Columbia allowing the full use of cannabis for medicinal and recreational purposes, whereas 29 states as well as Guam and Puerto Rico allow various levels of medicinal use of cannabis. In addition, 13 other states have decriminalized cannabis, which means criminal penalties from simple possession are removed. However, there are still three states that completely prohibit any possession or use of cannabis. As of the start of 2018 with legalization taking effect in California followed shortly thereafter by Vermont, approximately 1 in 5 United States residents (~21%) live in states with completely legalized cannabis. Abroad, several countries offer various levels of legality, including full legality (such as in Uruguay), partial legality (in the Netherlands), or have decriminalization statutes that remove criminal penalties for cannabis possession and use.

Because the final products are destined for human consumption through inhalation, ingestion, or topical application, jurisdictions in which cannabis is legal have recognized the need for control of various chemical aspects of the plant and its derivative products. Such regulations focus on potency, pesticide contamination, residual solvent content, moisture content, and metal content, and they necessitate the use of a variety of analytical techniques including high performance liquid chromatography (HPLC), liquid chromatography-mass spectrometry (LC-MS), gas chromatography (GC), GC-MS, and elemental spectroscopy methods (1). These regulations have been recognized as particularly important for medicinal cannabis because consumers may have weakened metabolic systems or compromised immune responses, limiting the effectiveness of their bodies to tolerate or process extraneous chemicals, toxins, metals, or biological material (2). For the purpose of this article, we explore the nature of, regulations on, and analytical methods for determining the metal content of cannabis and associated materials.

Metallic elements are known for their complex biochemistries and variety of both beneficial and deleterious health effects within the human body. Although many metals serve nutritional and metabolic purposes within the human body, others can act as toxins or carcinogens (3). For example, iron consumed in food is used by the body to transport oxygen within blood and is also used in cellular respiration, whereas consumption of mercury or lead can cause damage to the nervous system (4). Because of these effects, both positive and negative, the content of metals in products destined for human consumption (for example, food and pharmaceutical products) is often regulated by a local or federal oversight agency (such as the US Food and Drug Administration [FDA] or the US Environmental Protection Agency [EPA]).

Like all plants, cannabis uptakes metals from its environment as a result of normal plant metabolic functions. Some of these metals are naturally occurring, and leach into groundwater that the plant uptakes from the soils and minerals in which it grows. Other metals precipitate in rainwater as metal-laden atmospheric aerosols that are dissolved into precipitation. Most of these atmospheric metals arise from anthropogenic sources, such as smelting, refining, and other industrial processes. Finally, metals may be introduced into the plant’s environment as constituents of fertilizers, pesticides, herbicides, and fungicides used to increase crop yield. Regardless of their provenance, when metabolized, metals are absorbed and transported through the plant roots and into plant tissue, where they often reside as metal complexes within proteins and extracellular fluids (5).

Cannabis is so effective at uptaking metals from its environment that hemp (a nonpsychoactive variety of cannabis) has been used as a tool in the bioremediation of metal-contaminated sites. For example, hemp was used to leach radioactive strontium and cesium from the soils surrounding Chernobyl (6). In fact, cannabis has been called a 

of various trace metals, including lead, cadmium, magnesium, copper, chromium, and cobalt (7), which leads to concern that these elements may occur in high concentrations in cannabis plants cultivated for human consumption.

One of the primary challenges facing cannabis analysis is non-uniform regulations. Because cannabis remains illegal at the federal level, oversight on regulations is left to the state’s discretion. As such, limits on metal concentrations, as well as which metals require testing, vary from state to state. For example, most states require testing for the “big four” heavy metals: arsenic, cadmium, lead, and mercury. Certain states go further, such as Maryland, requiring testing for barium, chromium, selenium, and silver in addition to the big four. Adding further complication, the concentration limits for various products can also vary state by state, with some jurisdictions taking a stricter approach and others more relaxed. There is also variation by method of ingestion (for example, inhalational versus oral administration) with tolerances of metals for inhalational products being stricter than for those that are orally administered (that is, eaten or ingested).

Metals analysis is carried out on aqueous samples in which the material of interest is dissolved. The dissolution process can be carried out by a few different methods, the most simple of which is conventional hot-plate or hot-block digestion. Samples are placed into a vessel with solvents, typically strong acids such as nitric acid (HNO

) or hydrochloric acid (HCl). For organic matrices such as cannabis, hydrogen peroxide (H

) is often used to increase the oxidation potential of the solvents. The vessels are capped and refluxed at a moderate temperature, often for ≥8 h, to ensure complete digestion.

A more common method for sample preparation for elemental spectroscopy analysis is microwave digestion. A small amount of solid sample is placed into a digestion vessel along with solvents, similar to a hot-plate digestion procedure. The vessels are sealed and placed into the microwave, which allows the solvents within the vessels to reach very high temperatures and pressures, increasing the efficiency of dissolution. A typical microwave digestion of cannabis products is complete within ~45 min.

In the case of either digestion method, the molecular constituents of the digested material are destroyed during dissolution, but the constituent elements remain in solution. After dissolution, the resulting solution is typically diluted with ultrapure water to an appropriate matrix acid concentration and is ready for analysis.

Critical to carrying out digestions for metals analysis is the use of clean labware, “trace metals grade” reagents, and ultrapure (typically >18 MÎ©•cm) deionized water. As we explore in the sections below, common elemental spectroscopy methods are sensitive to metals ranging in concentration from parts per billion down to parts per trillion. Thus, reagents, such as acids, containing 1 ppm of various heavy metals may contain 50 ppb of said metals after they are diluted to the appropriate concentration, which is much higher than detection limits and, often, regulatory limits.

Long a staple of analysis labs, atomic absorption (AA) spectroscopy, is perhaps the most common analytical technique for metals analysis. Atomic absorption operates under the principle of the Beer-Lambert law, with gaseous free atoms absorbing light of a characteristic wavelength. The ratio of the intensity of the light shone through a blank sample to the intensity of light shone through one containing the analyte is the absorbance, and corresponds to the concentration of the element within the sample.

Atomic absorption spectroscopy is often attractive because of its relatively low cost compared to other elemental techniques. The instruments themselves as well as cost of operations (that is, consumables) tend to be far less that inductively coupled plasma (ICP) techniques. Additionally, with the use of a graphite furnace, detection limits can be in the low parts-per-billion to high parts-per-trillion range; however, with a graphite furnace comes increased costs.

Despite the low cost and potential for low detection limits, AA is notably limited in terms of sample throughput; these limitations arise from a few fundamental aspects of the technique. First, each element analyzed requires the use of a hollow cathode lamp, many of which are “single element” lamps, though “multielement lamps” do exist. As such, the number of elements per sample is limited by the number of lamps that the instrument can hold. Second, each element quantitated requires its own analysis. For example, during a typical analysis, the instrument will perform an injection-aspiration of the sample while transilluminating with the first hollow cathode lamp. Then, the instrument will switch lamps and repeat the injection-aspiration for any subsequent elements.

As a final limitation to the technique, elements such as As, Bi, Sb, Se, and Bi are simply not very sensitive by flame AA and benefit from the use of a hydride vapor generator to form volatile hydrides before sample injection. This approach necessitates the use of an additional piece of laboratory equipment as well as reagents. Additionally, it may require separate sample preparations because, often, potassium iodide (KI) is used to reduce As in solution from As(V) to As(III) for vapor generation; however, the use of KI may induce the precipitation of mercury as potassium tetraiodomercurate(II) from the sample solution. Thus, separate sample preparations for As and Hg when using a hydride vapor generator for analysis may be required.

Laboratories on a tight budget, testing only a few elements, or those that are not as concerned with sample throughput may benefit from the use of AA with a hydride vapor generator as their analytical technique for metals.

Inductively Coupled Plasma-Optical Emission Spectrometry 


Inductively coupled plasma-optical emission spectrometry (ICP-OES) represents the “middle-of-the-road” for elemental analysis with a price-point and performance intermediate to AA and ICP-mass spectrometry (MS). ICP-OES operates under the principle that adding energy to an element will cause an electron to attain a higher energy level. When that electron falls back to its initial level, the element emits a quantum of light of known wavelength. The wavelengths of light correspond to an individual element whereas the intensity, or brightness, of the light corresponds to concentration. Compared to AA, ICP-OES offers several advantages when it comes to cannabis testing, though with increased performance comes some potential drawbacks.

First, it is a true, simultaneous, multielement technique. Characteristic wavelengths of emission lines for multiple elements are diffracted and detected simultaneously, precluding the need for running sequential analyses for each element. Additionally, with many instruments, an entire spectrum is collected at once, allowing the user to retroactively analyze a dataset for elements, even if they were not included in the initial method.

Secondly, detection limits are improved compared to flame AA, with sensitivity for most elements in the range of low parts per billion. Detection limits can be improved for most elements by approximately one order of magnitude through the use of an ultrasonic nebulizer to increase nebulization efficiency. However, even when using an ultrasonic nebulizer, some cannabis laboratories have found that after the samples have been prepared and diluted, the required sensitivity is pushing the limit of the instrument’s fundamental capabilities.

Lastly, ICP-OES is more amenable to complex or dirty matrices than ICP-MS, allowing for easier analysis of brines, oils, and samples with high levels of total dissolved solids. Despite similar sample introduction systems, ICP-OES benefits in this regard from not physically pulling ions into the instrument, like ICP-MS. In the latter technique, high salts and other complex matrices can build up in the system interface region, high concentrations of easily ionized elements (such as Na, K) contribute to the space-charge effect, and high concentrations of matrix elements can contribute to polyatomic and isobaric interferences.

In contrast to AA, ICP-OES systems are more expensive to purchase and operate. They require the use of argon gas at a rate of ≥10 L/min to maintain the plasma as well as specialized and consumable glassware for sample introduction and analysis, such as nebulizers, spray chambers, and torches. Some ICP-OES instruments maintain a flow of argon even when the instrument is in standby mode to purge the optical bench.


The emerging leader in metals analysis in cannabis products is ICP-MS because of its high sensitivity and rapid sample throughput resulting from near-simultaneous multielement data acquisition. ICP-MS benefits from using the highly energetic ICP as an ion source coupled with the mass specificity and rapid switching of a quadrupole for mass filtering.

An ICP-MS system uses an ICP source that is similar to that used in ICP-OES, but in this case, it is using the argon plasma to form cations rather than to generate light from atoms in the sample. The cations are then pulled into the mass analyzer portion of the instrument by differential vacuum, where they are mass-filtered using a quadrupole, allowing for rapid switching between masses transmitted to the detector and high mass specificity.

The high sensitivity of ICP-MS instrumentation, with detection limits often in the range of tens- to single-digit parts per trillion range, can be attributed to the efficiency with which an ICP generates cations for almost all elements on the periodic table. Such sensitivity allows for detection of trace elements in a variety of matrices, even ones that are very dilute. This high sensitivity enables analysts to reduce interfering compounds from the matrix through dilution and to analyze a variety of sample types.

As an analytical technique, ICP-MS has historically suffered from the stigma of being a complicated, esoteric technique. However, most modern instruments are coupled with software that provides for a comparable ease of use, with automated instrument tuning, method setup, and post-run analysis. Furthermore, many instruments also use collision or collision-reaction cell technology to reduce the effect of polyatomic interferences on analyte ions.

The two primary downsides to ICP-MS, like ICP-OES, are relatively high cost of ownership and susceptibility to interferences when analyzing complex matrices. Like an ICP-OES system, an ICP-MS system will consume argon at a rate of >10 L/min, as well as employing the use of consumable glassware and interface cones. Furthermore, in high-salt or otherwise complex matrices, ICP-MS can suffer from degradation of signal, interferences from matrix elements, or the 

, in which easily ionized and abundant elements defocus the ion beam of low-concentration analyte elements. However, because of the high sensitivity of ICP-MS, dilution of samples can help resolve matrix-related problems while still preserving analyte concentrations in levels that are quantifiable.

The rapid changes in the legal status of cannabis and related products have necessitated regulations on their chemistry, including potency, residual solvents, pesticide contamination, and metals content. Because of metals’ potentially harmful biological effects, metals analysis of cannabis and its related products is critical to minimizing risk to the consumer. Although such regulations vary between states and jurisdictions, the majority require testing for arsenic, cadmium, lead, and mercury (at a minimum) because of their particularly hazardous properties.

Metals analysis can be conducted using a variety of instrumentation and methodologies-including AA, ICP-OES, and ICP-MS-each with their individual strengths and weaknesses. Atomic absorption spectroscopy benefits from low costs, but suffers from slower sample throughput; ICP-OES offers similar detection limits and more rapid sample processing than AA at a slightly higher cost; and ICP-MS offers exceptional sensitivity and throughput, but at the highest initial and operating costs.

Despite the higher costs and analytical complications associated with the method, most laboratories are pursuing ICP-MS as their technique of choice because of the high sensitivity, high sample throughput, and adaptability to other methods and industries. Because the technique has long been used in testing of food, water, and pharmaceutical products, contract laboratories with existing ICP-MS systems are viewing cannabis testing as a supplemental revenue stream. Alternatively, as cannabis testing evolves, laboratories may use their ICP-MS instruments for detection of metals in cannabis growth media, soils, fertilizers, water for hydroponic growth, and so on. Furthermore, as regulations evolve and are codified, it is likely that the number of metals requiring testing will increase and acceptable concentrations of said metals will decrease, necessitating the use of fast and sensitive analytical techniques, such as ICP-MS.

Andrew P. Fornadel, PhD, Daniel L. Davis, Robert H. Clifford, PhD, 

are with Shimadzu Scientific Instruments in Columbia, Maryland. Direct correspondence to: 

S. Kuzdzal, R. Clifford, P. Winkler, and W, Bankert, “A Closer Look at Cannabis Testing,” Shimadzu Scientific Instruments Whitepaper (2017). Retrieved 16 January 2018: 

https://www.ssi.shimadzu.com/industry/literature/Shimadzu_Whitepaper_Emerging_Cannabis_Industry.pdf

E. Charkowski, “Hemp ‘Eats’ Chernobyl Waste, Offers Hope for Hanford,” in 

http://rediscoverhemp.com/hemp-eats-chernobyl-waste-offers-hope-for-hanford/

M. Girdhar, N.R. Sharma, H. Rehman, A. Kumar, and A. Mohan, 

A.P. Fornadel, D.L. Davis, R.H. Clifford, and S.A. Kuzdzal, 

Metals in Cannabis and Related Substances-Regulations and Analytical Methodologies

The overproliferation of cannabis strains following the establishment of cannabis-complicit states in the United States has led to confusion and resulted in a lack of transparency for the consumer at
the dispensary.

As various states in the United States are starting up cannabis programs one by one with little cooperation or standardization, entire countries are doing so with efficiency at the same pace. Globally, about 2.25% of the population consumes cannabis. Both medical and recreational cannabis make up a multibillion-dollar global industry. Lawful cannabis programs have already been implemented in Canada, Mexico, the United Kingdom, the Netherlands, Australia, Germany, Italy, Israel, Poland, the Czech Republic, Uruguay, Peru, South Africa, and other countries, regardless of global drug policy. Most of these countries regulate cannabis as a medicine. Israel in particular is at least 10 years ahead of the United States in instituting standardization in the production and extraction of cannabis for clinical trials leading to patented drug products in line to gain approval from the US Food and Drug Administration (FDA) and entry into the lucrative prescription drug market in the United States. Cannabis companies stateside need to play catch up to gain market share, starting with distinguishing the staggering number of cannabis strains being grown.

The overproliferation of cannabis strains following the establishment of cannabis-complicit states in the United States has led to confusion and resulted in a lack of transparency for the consumer at the dispensary. In Washington state alone, more than 39,000 cannabis strains are being grown. Further confounding the situation for the recreational cannabis consumer and patient is the sole reliance placed on tetrahydrocannabinol (THC) content to establish the inherent value of flower rather than taking the entirety of the pharmacologically active chemoprofile of the plant into account. Both state regulators and cannabis testing laboratories can help relieve the growing uncertainty. Regulators should institute broader chemical profiling, genotyping, and mandatory cannabis cultivar registration with specific criteria required before ever growing the cultivar. And those cannabis testing laboratories that go beyond simply quantifying cannabinoid potency and quality assurance are in a unique position to demystify cannabis strains through expanded chemometrics, genotyping, and consumer education. Both approaches are aimed to give cannabis consumers more confidence in what they are purchasing. On the upside, findings indicate that cannabis consumers are willing to pay a premium for genotyped, authenticated flower.

Because the vast majority of cannabis being grown in the United States is a Type I drug-type defined as having a cannabidiolic acid-tetrahydrocannabinolic acid (CBDA:THCA) ratio of <0.5, various efforts have been underway to make sense of cannabis strain names through the use of data analytics on broader cannabis flower chemoprofiles. It turns out that cannabis terpenoids not only imbue pharmacologically important attributes, but uniquely provide a basis for a secondary nomenclature after established cannabinoid content. We, and others, have shown that while unique terpenoid chemoprofile patterns exist and can be assigned to a cannabis cultivar, the absolute amounts of any given terpenoid can be influenced by genetic, epigenetics, environmental, and cultivation factors.

The self-inflicted confusion within the United States cannabis industry is an opportunity to demonstrate ingenuity against the rapidly maturing global cannabis industry, where outdoor growing costs are a fraction of the indoor energy intensive grows here. Throwing out unnecessary impediments is a priority, starting with nonstandard strain naming in favor of the agronomic and horticultural practice of registered “cultivar” names based on full genome sequencing and broad chemometrics to allow for consistent, reproducibly effective comparisons within the United States cannabis industry and enable trademarking or patenting of specific cultivars. As the industry rapidly evolves toward big agriculture versus boutique cannabis growers, cannabis cultivar authentication will be a key to success and keeping market share in the United States.

 is the Chief Science Officer with Digipath Labs in Las Vegas, Nevada. Direct correspondence to: 

Empowering the Cannabis Consumer in a Rapidly Expanding Global Market

Welcome to the inaugural issue of our new publication, 

. We are thrilled that our group is a part of the cutting edge in cannabis science and will help unveil what the future holds for this industry. In alliance with the Cannabis Science Conference, this new professional journal and website will focus on educating the legal cannabis industry about the science and technology of analytical testing and quality control, including topics such as

Sample preparation techniques for analytical testing

Development of analytical methods for potency testing

Identifying contaminants such as pesticides, residual solvents, heavy metals, and mycotoxins

Quality control for cannabis, extraction, and processing

Laboratory best practices and certification

Analytical methods for the characterization of cannabinoids and terpenes

With our portfolio of leading science and pharmaceutical brands, including 

LCGC, Spectroscopy, Nutritional Outlook, PharmTech

, we have extensive experience keeping readers in fields such as analytical chemistry, pharmaceutical development and manufacturing, and nutritional supplements informed about the latest techniques, best practices, and regulatory information. Given the knowledge and experience we have from producing those thought-provoking and industry-relevant brands, launching 

 is a perfect next step into the future of analytical science. This new publication will mirror the quality of content and circulation that makes our other publications so successful.

The recent acceleration in the acceptance of cannabis as a valuable medical treatment places the industry at a critical juncture. With the launch of 

, we are taking an important step to connect the scientific community we serve with the fast-growing cannabis industry.

 will be published quarterly in 2018 and will appear at all the major trade shows this year. Be sure to pick up a copy of our latest issue at the next show near you, or visit our website, www.cannabissciencetech.com, for the latest information on subscriptions, newsletters, articles, and more.

The New Frontier in  Analytical Science: Cannabis

! I am beyond excited to be the editor-in-chief of this new publication, which will focus on educating and sharing knowledge with the analytical science community about all scientific aspects of cannabis. As the push continues to legalize cannabis in some form in more and more states, there is an important need to address the analytical challenges that go hand in hand with cannabis testing, quality control, and sample preparation. Questions ranging from which technique to use to how to handle the varying state regulations, and concerns about consumer safety, dosing, and legal matters are all topics we plan to address in this publication. Many refer to the cannabis industry as the new “wild west,” but it is my belief that analytical science will help bring order to (or tame) the “wild west” and help raise cannabis to a position of legitimacy in the science, medical, and legal communities.

Before I delve into the great content in this issue, I’d like to tell you a little bit about myself and how this magazine came to be. I worked on the analytical chemistry publications 

 for the past 10 years, spending the last seven as the managing editor. During that time, I worked very closely with Laura Bush, the editorial director of those brands. She has been a very supportive colleague and mentor to me over the years. Without Laura this first issue would not be possible, and I would like to thank her tremendously for taking the editorial lead in creating this start-up and organizing the content for our first issue before I was promoted to the editor-in-chief position.

I would also like to extend a heartfelt thank you to Josh Crossney, of JCanna, Inc. and the Cannabis Science Conference, who partnered with us on this magazine. Through this collaboration, we have built the foundation for what we hope will be a long-standing publication. Josh is also a contributing editor and will have a feature in each issue called “Cannabis Crossroads.” Here in our first issue, he interviews the founders of EVIO Labs and discusses the specific challenges faced by accredited cannabis testing laboratories. 

Edibles are a complex matrix to analyze because of the various forms they can come in and the other components involved in the samples. In this issue, Kevin A. Schug, of the University of Texas at Arlington, and his colleagues take a look at different edibles based on their composition, and discuss various sample preparation strategies that may be appropriate for analyzing the different products. Metals testing is another hot-button topic in the cannabis industry that Andrew P. Fornadel and his colleagues from Shimadzu Scientific Instruments tackle in this issue. Fornadel’s article presents an overview of analytical methodologies that can be used for metals testing as well as the challenges facing the analyst and notes on regulatory stipulations. 

Another area that causes concern in the cannabis industry is pesticide testing. Many states have different requirements and testing levels for various pesticides, which leads to problems when laboratories are tasked with testing cannabis samples without specific guidelines to follow. Here, Joseph Konschnik, of Restek Corporation, and his coauthors, describe a collaborative public-private partnership where state government agencies worked side by side with laboratories and growers toward a common goal of agreeing to analytical procedures to determine method detection limits using an industry-standard approach. In another article, Katherine K. Stenerson and Gary Oden of MilliporeSigma discuss several steps that were developed to improve the work flow in the analysis of pesticide residues from cannabis plant material by liquid chromatography-tandem mass spectrometry and gas chromatography-tandem mass spectrometry. The goals of the study were to optimize the extraction and analysis process to reduce sample background, improve pesticide recoveries, and allow direct injection of QuEChERS (quick, easy, cheap, effective, rugged, and safe) extracts, without dilution, for high performance liquid chromatography analysis.

In addition, we have a short tutorial on the QuEChERS method of dispersive solid-phase extraction in which Douglas E. Raynie, of South Dakota State University, explains how the method and salting out are applied to the analysis of cannabinoids of interest. Christian Sweeney, of Cannabistry Labs, describes a goal-oriented approach to extraction processes. Sweeney explains that the wide range of options available has left processors questioning the best method for their extractions, but a goal-oriented approach that focuses on finding the best fit between process and physicochemical product attributes is critical.

We also have two great opinion pieces, one from Susan Audino of S.A. Audino & Associates, LLC, and the other from Cindy Orser of Digipath Labs. Audino discusses the basic principles of quality assurance and laboratory accreditation as well as the current status of voluntary methods in development. Orser addresses the overproliferation of cannabis strains and the need for cannabis cultivar authentication. Enjoy our first issue and be sure to come back for more in June!

Cannabis Science: Taming the Wild West

Many processes have been adopted for extraction and refinement of cannabis. Much like traditional botanical extractions, the aim of these techniques is to provide the desirable qualities of cannabis in a more readily usable form for the delivery method of choice. The wide range of options available has left processors questioning the best method for their extractions. A goal-oriented approach, focused on finding the best fit between process and physicochemical product attributes is critical. Pairing this goal-oriented process development with analytically guided optimization will allow for the development of an ideal process to deliver the best end product.

One of the key challenges when designing a cannabis processing plant is the overwhelming number of options available. Within a cannabis processing plant, decisions about unit operations will need to be made around preprocessing, extraction, and purification. Even after landing on an extraction process between light hydrocarbon, ethanol, solventless, or supercritical fluid options, there are myriad equipment vendors offering the latest and greatest technologies for each process. This plethora of possibilities leads to the unfortunate, but unavoidable question: What’s the best? While this question addresses some basic design options for the process, the far more critical question to ask is, what’s the best for my products?

Answering that question demands a holistic view of the process, and not simply siloed unit operations. This question drives us toward goal-oriented process development, which invariably results in higher efficiencies and higher end-product quality. Regardless of whether the end product is as basic as a simple extract, a high-end consumable, or a pharmaceutical-grade formulation, understanding the physicochemical attributes that drive that product’s quality will assist in making decisions about how to design a cannabis process. These attributes may be things like cannabinoid potency ranges, viscosity, color, clarity, or aroma. Each of these desired properties can be carefully crafted by choosing and arranging the optimal unit operations into a complete process.

While the common parlance in the industry has become “extraction lab,” when designing a cannabis process there are three key stages: preprocessing, extraction, and purification. Each of these stages is filled with options that will affect the properties of the resulting cannabis extract. Making decisions about those stages becomes much more straightforward when you are armed with the requirements of an end product.

Preprocessing may begin as early as making decisions about how to dry and store cannabis raw material. If, for example, the end product requires fresh aroma notes like a live resin, then this unit operation may demand no drying and frozen storage. If, on the other hand, the goal is to deliver the traditional cured flavor of cannabis, then identifying the optimal temperature, humidity, and moisture content will drive the development of that process step. The key is identifying metrics by which to grade our process-in this case, perhaps moisture and terpene content-and having a way to measure it.

A recurring theme in process development is the criticality of analysis. Analytical methods provide the feedback required for design. In the above example, a drying process is proposed for delivering a targeted terpene profile. That process could then be carried out, and the resulting dry cannabis raw material could be analyzed via gas chromatography (GC) to determine if the targeted terpene profile was achieved. Because the target is rarely achieved on the first attempt, the drying process conditions (temperature, humidity) should be altered, the experiment repeated, and more terpene data collected. This iterative process design will result in an optimized approach while keeping the goals of the end product in mind, all because of the feedback provided by analysis.

Another important preprocessing step is raw material size reduction. This step is often skipped, but in many cases, it can enhance the efficiency and quality of a cannabis extraction process. The goal of this process step is to decrease the particle size of the material to be extracted, resulting in an increased surface area available for solvent extraction. Again, taking a holistic view, we might recognize that this would benefit solvent-based extraction processes such as those using butane, ethanol, or a supercritical fluid, but perhaps would be detrimental to some of the solventless extraction processes.

Next, we look to the goal of our process. Let’s assume we are interested in delivering the bioactive components of the cannabis raw material as close to their natural state as possible in our finished extract and optimizing the efficiency of our extraction process. We can then define metrics that tell us if we have accomplished those goals. For example, I may hypothesize that by creating a ground raw material with a consistent and small particle size I will increase the efficiency of my extraction. Again, we turn to our iterative process design mindset and test several size-reduction variables (that is, milling versus grinding, room temperature versus chilled, low-speed grinding versus high-speed grinding), and using an instrument such as a sieve shaker or particle size analyzer I can determine which process variable gives me the smallest and most consistent ground particle size. To achieve the goal of delivering “nature identical” bioactives, I can analyze for target compounds such as the cannabinoids and terpenes before and after the size reduction process to ensure that degradation resulting from thermal or oxidative damage is minimized.

The heart of any cannabis processing plant is the extraction process. This unit operation will have a major impact on every physicochemical attribute of the cannabis extract being processed and will likely represent a significant portion of the capital outlay associated with a processing plant.

Light hydrocarbons, specifically butane and propane, have been used for cannabis extraction for some time (1). This solvent system provides fast extractions and is fairly selective for cannabinoids and terpenes, meaning it is possible to avoid coextracting other potentially unwanted compounds from the cannabis raw material. Light hydrocarbon extraction systems tend to be middle of the road in terms of capital cost, but depending on local safety requirements it may demand a serious infrastructure investment to create a safe working space.

From the lens of goal-oriented process development, light hydrocarbon extractions have two limitations. The first involves tunability. Compared to other extraction processes like supercritical fluid extraction, there are a smaller number of variables that can be adjusted when producing a light hydrocarbon extract, which means the processor has less impact on the properties of the extract produced. The upshot of this lack of tunability is an ease of use and shallower learning curve for operators. Secondly, while butane and propane are highly volatile compounds, a post-processing step, usually referred to as 

, is required to remove the solvent from the extract. This purge step is often carried out in vacuum ovens and involves heating the extract under deep vacuum conditions to drive off the remaining solvent. Through regulations, each state will dictate the degree to which these solvents must be removed. Colorado, for example, has just reduced its limit from <5000 ppm butane to <1000 ppm (2).

Again, analysis becomes a critical step for driving the development of an optimized process. If simple efficiency and throughput is the aim of our process optimization, it is critical to determine which vacuum pressure and temperature combination results in the fastest removal of butane from our extract to achieve a product that meets the regulatory specifications. To determine this, a residual solvent analysis (RSA) must be performed on samples from the purge process to determine how much butane remains at various times under a given set of purging conditions. If, for example, the RSA result on a sample comes back above the regulatory specification, the process developer knows they must either increase the purge time, increase the temperature of the purge, or pull a deeper vacuum.

It is quite unlikely, though, that the only goal of this process is efficiency. Delivering a high level of terpenes is critical both to the bioactive and organoleptic properties of the extract. This challenge adds a new layer of complexity because the methods for improving purge efficiency (increased time and temperature) will also result in greater loss of terpenes. In this case, it is critical to analyze both the residual solvent content and the terpene content throughout the purge process. An example of the type of data that could be generated for this process development is shown in Figure 1. (See upper right for Figure 1, click to enlarge.) During a purge process at a given temperature and pressure, samples can be pulled and analyzed for terpene and residual solvent content. This step not only gives the process developer a sense of how long it will take to produce material within specification, but also the expected loss of terpenes. Similar charts would then be produced at different temperatures and pressures until the developer finds a set of conditions that meets their goal of minimizing purge time and terpene loss.

Ethanol extraction has become increasingly popular because of its fast extraction, low capital costs, and clear designation as a food-grade solvent. Ethanol is a less selective solvent than butane and propane, but depending on the goals of the extraction this characteristic could be either a positive or negative attribute. If the goal is to strip a wide range of compounds from the cannabis raw material beyond just cannabinoids and terpenes, ethanol will certainly accomplish that. Because of this, it is not uncommon for ethanol extraction to be coupled with post-extraction purification processes to remove some unwanted coextractives. Just like with light hydrocarbon extraction, solvent removal becomes the most time-consuming step in the process. For ethanol extracts, it is more common for rotary or falling film evaporation to be used because ethanol requires significantly more energy to evaporate compared to butane. This means that the vast majority of volatile aroma compounds will be stripped from the extract during solvent removal. If those compounds are not important to the end product, then this potential problem is a non-issue, but if your formulation is meant to leverage the entourage effect of cannabinoids and terpenes acting synergistically to drive a condition specific effect, then this approach may not be the ideal process.

) extraction has risen to a place of prominence in the cannabis industry. Its steep learning curve and high capital spend mean it is not the ideal choice for some, but the depth of tunability afforded by its solvent system gives the processor a level of control on the extract produced that no other process can provide. This tunability is due to a unique property of CO

in the supercritical phase. By altering the pressure and temperature conditions of the extraction, the operator can alter the solvent characteristics of the CO

 (4). What this means is that the process developer can design an extraction process to select only the compounds that meet the goals of their end product.

 requires no further work for solvent removal because it simply evaporates from the extract as it is collected from the extractor. This means that after a method is developed for the extraction of cannabis with supercritical fluid CO

, there is no additional solvent removal process where volatile compounds can be lost. This gives the operator the ability to capture truly nature identical aroma profiles from their raw material. Figure 2 illustrates this point by displaying two chromatograms back to back. (See upper right for Figure 2, click to enlarge.) The top chromatogram represents the terpene profile of a cannabis raw material before extraction, while the bottom chromatogram (intentionally flipped for ease of viewing) represents an 18-fold dilution of the terpene extract produced by supercritical fluid CO

 extraction. Although only a few of the notable compounds are labeled specifically, the comprehensive mirroring of the two spectra clearly shows how powerful this extraction technology is at producing truly nature identical extracts.

Post-extraction purification can range from simple filtration, often referred to as 

, to more complex processes, such as distillation or preparative-scale chromatography. Again, the requirements of the end product should drive decisions here. Winterization leverages melting point differences between the various components in an extract to precipitate out unwanted waxes at a reduced temperature. It is then possible to filter away those waxes while retaining most of the other compounds in the extract. This means that while winterization can reduce the overall viscosity of the extract and increase the cannabinoid potency, it will have little effect on the overall color and clarity of the finished extract. If, for example, the end product can use oil of any color or clarity, but the formulation can’t handle some of the plant waxes that may have been coextracted by an ethanol solvent, then winterization may be a perfect choice.

Distillation can purify an extract by selective evaporation. By evaporating, condensing, and collecting the various fractions of an extract that evaporate at different temperatures, an operator can pull a highly pure cannabinoid fraction, often referred to as 

. This distillate will be very high in cannabinoid potency, will be light in color, and will have high clarity, but it will not retain many of the other compounds from the raw material. In fact, because of the high temperatures required for the distillation of cannabinoids, many bioactives in the extract can be degraded in the process. Therefore, if a product demands a fuller spectrum of cannabis bioactives, distillation may not be the ideal technology, but if a high clarity, high cannabinoid potency extract is the goal, it may be a good choice.

Lastly, preparative-scale chromatography can purify the compounds in an extract based on their affinity to the stationary phase within a chromatography column. This process will provide the highest levels of purity for individual cannabinoids (upwards of 95% pure), but will separate out the vast majority of other compounds. This purification process produces an extract that allows a product developer to formulate with highly pure cannabinoids, but it will not deliver the complete efficacy of the raw material.

This small snapshot of the options available shows just how complex the task of process development can be in the cannabis industry. The keys to effective process development are having a complete understanding of the physicochemical attributes the end product demands, and relying on analysis to provide feedback when optimizing processes. By keeping these considerations in mind as guidelines, finding the answer to “what’s the best for my product?” can be a straightforward, though still laborious process.

 is the Director of Science & Technology at Cannabistry Labs in Denver, Colorado. Direct correspondence to: 

(6), 797-803 (2015), doi:10.2131/jts.40.797.

Colorado Department of Revenue, Marijuana Enforcement Division, “Recently Adopted Retail Marijuana Permanent Rules - Effective January 1, 2018.” https://www.colorado.gov/pacific/enforcement/med-rules, p. 123.

C. Sweeney, “Cannabis Extraction and Refinement in Colorado: A 5,280 Ft. View” presented at the Cannabis Science Conference, Portland, Oregon, 2017.

Natural Extracts Using Supercritical Carbon Dioxide

Goal-Oriented Extraction Processes

In this work, several steps were developed to improve the workflow in the analysis of pesticide residues from cannabis plant material by liquid chromatography tandem mass spectrometry (LC–MS/MS) and gas chromatography (GC)–MS/MS. The goals of the study were to optimize the extraction and analysis process to reduce sample background, improve pesticide recoveries, and allow direct injection of QuEChERS (quick, easy, cheap, effective, rugged, and safe) extracts, without dilution, for high performance liquid chromatography (HPLC) analysis. Reduced background and better pesticide recoveries were achieved by the use of a QuEChERS cleanup with zirconia/C18 functionalized silica, primary secondary amine (PSA), and a low-surface-area carbon blend. HPLC analysis was optimized to provide late elution of coextracted cannabinoids relative to the targeted pesticides, thus making it possible for a dump step to be used to prevent them from entering the MS detector. The HPLC column configuration used improved the peak shapes for the earliest eluted hydrophilic pesticides when injecting extracts in 100% acetonitrile. This then allowed a single set of extracts to be analyzed by both GC–MS/MS and LC–MS/MS.

Consumption of cannabis or cannabis-based products is currently legal in some form in 29 states in the United States plus the District of Columbia. Testing of the plant materials and products is required by many of these states; however, the specific test methods and target compound lists are not mandated in all cases. In October 2016, the state of Oregon took a major step forward by requiring that all laboratories testing cannabis be accredited by the Oregon Environmental Laboratory Accreditation Program (ORLEP) and licensed by the Oregon Liquor Control Commission (OLCC) (1). Consequently, Oregon Administrative Rules (OAR) list specific contaminants to be tested in marijuana samples, along with action levels (2). The pesticides on this list include carbamate, organophosphorus, macrocyclic lactone, neonicotinoid, pyrethroid, and triazole fungicides as well as others. Action levels per OAR vary from 0.2 to 1 µg/g, depending on the specific pesticide. In addition, the state of California, which legalized recreational cannabis in 2016, recently released a proposed list of pesticides that includes all but one of those found on the OAR list, plus eight more (3).

Because of its ease of use and applicability to a wide range of pesticides, the QuEChERS (quick, easy, cheap, effective, rugged, and safe) approach has been adopted by many testing laboratories for use on cannabis. After extraction, incorporation of a cleanup step is important for removing pigments as well as other contaminants. A QuEChERS cleanup using a mixture of primary secondary amine (PSA), C18, and graphitized carbon black (GCB) is often chosen for this purpose. PSA removes acidic interferences, C18 removes hydrophobic interferences, and GCB retains some pigments-specifically the green color imparted by chlorophyll. This mixture of sorbents thus retains a wide range of contaminants; however, it also has potential to reduce recoveries of target pesticides that are susceptible to hydrophobic retention on C18, or planar enough in structure to be strongly retained by GCB. In previous work by Stenerson and colleagues with cannabis, an alternative sorbent mix was evaluated for cleanup in the analysis of various pesticides, and found to offer a better balance than a traditional mixture of PSA/C18/GCB with regards to removal of pigmentation and pesticide recovery (4). This sorbent mix contained PSA, a zirconia-coated C18 functionalized silica, and a specialized carbon. The zirconia retains by Lewis acid-base interactions, and has been found to reduce the background of certain fatty compounds as well as some pigments. The carbon is a low surface area graphitized variety that has been shown to have weaker retention of small, planar molecules such as certain pesticides. In this work, the pesticide list tested previously was expanded to include many of those on the OAR list described above. The alternative sorbent mix was compared directly to a conventional blend of PSA/C18/GCB for cleanup and analysis of spiked replicates of cannabis plant material analyzed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) and gas chromatography-tandem mass spectrometry (GC-MS/MS). For the LC-MS/MS portion of the analysis, the column and mobile phase selection was optimized to improve overall workflow, which is described here.

Dried cannabis was pulverized using an IKA T10 Ultra Turrax mixer. Next, 1.9 g was weighed into a 50-mL centrifuge tube and spiked with pesticides at 50 ng/g. After a 10-min equilibration time, the sample was mixed with 10 mL of deionized water and allowed to sit for 30 min. Then 10 mL of acetonitrile was added, and the sample was shaken at 2500 rpm for 30 min. A mixture of 4 g of magnesium sulfate, 1 g of sodium chloride, 0.5 g of sodium citrate dibasic sesquihydrate, and 1 g of sodium citrate tribasic dihydrate was added, and the sample was shaken for 1 min. The sample was then centrifuged at 5000 rpm for 5 min, and the supernatant was removed for cleanup.

Next, 1 mL of extract was added to a 2-mL tube containing the mixture of cleanup sorbents. Two different sorbent mixtures were used:

(400 mg, 400 mg, 400 mg, and 1200 mg, respectively)

PSA/zirconia-coated C18 functionalized silica/low-surface-area graphitized carbon (400 mg, 480 mg, and  80 mg, respectively)

Samples were shaken for 1 min, and then centrifuged at 5000 rpm for 3 min. The supernatant was removed for analysis.

A single set of extracts was analyzed directly after cleanup, and without dilution, on both an Agilent 6460 LC-MS/MS system and an Agilent 7000C GC-MS/MS system. Pesticides that did not yield response by LC-MS/MS were analyzed by GC-MS/MS. The HPLC column used was a 10 cm x 2.1 mm, 3.0-µm Ascentis RP-Amide column, equipped with a 3 cm x 2.1 mm, 5-µm RP-Amide guard column. A gradient of 5 mM ammonium formate, 0.1% formic acid in 5:95 water-acetonitrile and 5 mM ammonium formate, 0.1% formic acid in 95:5 water-acetonitrile was used at a flow rate of 0.4 mL/min. The gradient started at 10% organic, held for 1 min at 10%, ramped to 100% in 13 min, and finally held at 100% for 6 min. The sample injection volume was 5 µL, and the column temperature was maintained at 30 °C.

GC analysis was performed on a 20 m x 0.18 mm, 0.18-µm film thickness SLB-5ms column, with an oven program starting at 50 °C, with a 2-min hold, and ramping at 8 °C/min to 325 °C with a 10-min hold. Helium carrier gas was used at a constant flow rate of 1.2 mL/min. The injection port temperature was 250 °C, and a 1-µL injection was performed in pulsed splitless mode with a 50-psi pulse held until 0.75 min. The liner used was a 4-mm i.d. tapered FocusLiner containing quartz wool.

Quantitation was done against a five-point matrix-matched calibration curve prepared in unspiked cannabis extract. Separate curves were prepared for each cleanup. No internal standards were used, thus all recovery values reported are absolute.

Typical cannabis samples analyzed by testing laboratories contain high levels of cannabinoids, often in the range of 20-25% by weight. These compounds will coextract with the pesticides during the QuEChERS process, and are only partially removed during cleanup. In the case of LC-MS/MS analysis, these remaining cannabinoids can build up on the detector, resulting in the need for more frequent system maintenance. In this study, column and mobile-phase selection were based on conditions that would force elution of the cannabinoids as late as possible in the run, ideally after the pesticides. Under these conditions, the diverter valve on the LC-MS/MS system could be set to flow to waste after elution of the last pesticide. This setting will then prevent a majority of the cannabinoids from entering the detector.

To facilitate the appropriate high performance liquid chromatography (HPLC) conditions, a screening experiment was designed to study elution of the major cannabinoids relative to the targeted pesticides on several different column chemistries, and using both acetonitrile and methanol based gradients. The column chemistries screened were as follows:

All columns were 10 cm x 2.1 mm columns with 2.7-µm fused-core particle packings. The fused-core particle architecture version of these chemistries was initially chosen for both efficiency and speed. The HPLC conditions were similar to those listed previously in the experimental section, with ultraviolet (UV) used for detection, and ammonium formate omitted from the mobile phase. Samples were injected in 100% acetonitrile to emulate samples resulting from the QuEChERS extraction. As expected, on all five columns this injection solvent resulted in poor peak shapes of the earliest eluted pesticides. Focusing on overall retention profiles, the RP-Amide column yielded the least amount of overlap between the pesticide and cannabinoid elution ranges. Overlap was notably less than obtained using the C18 column (Figure 1), which is a commonly used chemistry for this application. (See upper right for Figure 1, click to enlarge.) In addition, comparing acetonitrile to methanol in the mobile phase, using the former in the gradient eluted the pesticides faster, further decreasing overlap with the cannabinoids.

To simplify the method as much as possible, the same QuEChERS extract was analyzed by both HPLC and GC. However, as indicated previously in the column screening experiment, injection of 100% acetonitrile into the high aqueous starting conditions of the gradient produced poor peak shapes for the early eluted pesticides. To improve the peak shapes of these compounds, a 3-µm fully porous RP-Amide column was substituted for the 2.7-µm fused-core RP-Amide column. The installation of a guard column further improved peak shape (Figure 2), most likely because of increased retention and improved mixing of the sample with the mobile phase. (See upper right for Figure 2, click to enlarge.) In addition, when working with high background samples, the use of a guard column is highly recommended to extend the life of the analytical column.

As expected, the coextracted chlorophyll generated a QuEChERS extract with a deep green color. After cleanup, a majority of the green color was removed by both cleanups, with the final extracts appearing as a light yellow. Analysis by GC-MS in full scan mode (not shown) produced similar peak patterns between both cleanups and the uncleaned extract, but a difference in amplitude of background peaks (which consisted mainly of terpenes and cannabinoids). Overall reduction in background was compared by summation of total peak area for each chromatographic analysis. Compared to no cleanup, the alternative sorbent mix reduced background by 35% and PSA/C18/GCB by 31%.

Using the optimized HPLC conditions described previously for the final LC-MS/MS analysis of the cannabis extracts, minimal overlap was observed between two of the major cannabinoids present in the samples, and the later eluted pesticides. Figure 3 shows an extracted ion chromatogram (EIC) of 

314.5, taken from a full scan LC-MS analysis of a cannabis extract. (See upper right for Figure 3, click to enlarge.) This mass represents the molecular ion of the two major cannabinoids detected in the sample extract; tetrahydrocannabinol (THC) and cannabidiol (CBD). As indicated, the last pesticide analyzed, pyridaben, was eluted just before cannabidiol. The most abundant cannabinoid present, THC, was eluted well after. Column flow could be switched to waste after elution of pyridaben, preventing some of the CBD and all of the THC from entering the MS system. Other cannabinoids; specifically cannabigerol (CBG), cannabinol (CBN), cannabidiolic acid (CBDA), cannabichromene (CBC), cannabigerolic acid (CBGA), and tetrahydrocannabinolic acid (THCAA), are known to be eluted after CBD on the RP-Amide phase. Thus, if present in a cannabis sample, all of these could also be diverted to waste as well.


The pesticides included in this study represented a majority of those on the OAR list. Two pesticides from this list, avermectin B1a and naled were not analyzed because of lack of response. Avermectin is prone to sodium and potassium adduct formation. The presence of ammonium formate in the mobile phase should reduce this occurrence (because it is monitored as an ammonium adduct). However, even with these measures, others have also reported issues with low-level detection of this compound (5,6). Naled is susceptible to adsorption by PSA, and thus did not make it through the cleanup process with either sorbent mix.

Comparing spike data between the two cleanup methods, the alternative mix exhibited better overall performance than PSA/C18/GCB. Looking at data for the individual pesticides (Table I), bifenthrin, chlorantraniliprole, clofentezin, fenproximate, fludioxinil, and hexythiazox showed notably better recoveries using the alternative sorbent mix. (See upper right for Table I, click to enlarge.) Although none of these are completely planar in structure, it is possible that recovery was reduced using PSA/C18/GCB because of the hydrophobic retention on the GCB phase, which has a higher surface area than the carbon in the alternative mix.

Several pesticides exhibited poor recoveries after both cleanup techniques. Bifenazate recovery was around 50%. This compound is susceptible to oxidation to bifenazate-diazine, which may have occurred to some degree during the extraction and cleanup process (7). Daminozide is a carboxylic acid, and thus its recovery was affected in both cleanups by the presence of PSA. Matrix interference prevented the analysis of dichlorvos in the PSA/C18/GCB extracts. In the extracts cleaned with the alternative mix, the peak could be detected, but recoveries were low and variable. This low recovery is most likely because of the retention on the zirconia-based sorbent as a result of Lewis acid-base interaction between the zirconia and the phosphate group present in the structure of dichlorvos. This same behavior has been observed by the author and others for this compound when using zirconia sorbents (8-10). Etofenprox is a very hydrophobic pesticide (log 

 = 7.1) and may exhibit poor extraction efficiency or retention by the C18 and zirconia portions of the cleanup sorbents (although less so on the later). Imazalil is a relatively polar pesticide, which can be retained by PSA. Recovery issues have also been observed by others with this compound when using zirconia containing sorbent mixtures (10,11). Low recoveries of spinosyn A and D, which are large macrocyclic lactones, have been reported when using C18, carbon, and zirconia-containing sorbents (12). The use of citrate buffering in the QuEChERS extraction has been observed to increase recovery, possibly by displacement of the analytes from the zirconia (10). Low spiroxamine recovery could not be attributed solely to cleanup, and may be because of an issue with extraction efficiency.

March/April 2018

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