The two most commonly used potency measurement methods in cannabis analysis are infrared spectroscopy and chromatography. We have spent many months in this column on how to properly use infrared spectroscopy to determine analyte concentrations in cannabis samples. So, now we will discuss chromatography. In this first of a series of columns, I will begin a discussion of the theory of chromatographic separations. Our goal will be to derive equations that will help you better perform experiments to optimize your separations.

 is Greek for color writing (1). It was discovered by Russian chemist Michael Tswett around 1900. The name refers to the fact that he used the technique to separate plant pigments, and in the glass columns he was using he could clearly see colored bands separating from each other. The importance of chromatography is underscored by the fact that two of its early practitioners, who laid much of the groundwork in the field, Archer J.P Martin and Richard L.M. Synge, were awarded the Nobel Prize in Chemistry in 1952 for their work (2). In an absurd byproduct of my research into the history of chromatography, I found there exists a record album called 

 by a band called Second Person, one of whose members is Julia Johnson, sister of former United Kingdom (UK) Prime Minister Boris Johnson (3). Small world, isn’t it?

Chromatography purifies mixtures by separating them into their components. An instrument that uses chromatography to separate a mixture is called a 

. Many different types of chromatography are used depending upon the chemical species that need to be separated. A chromatograph makes use of chemical differences between analytes such that they spend different amounts of time inside the instrument. As a result, the different components come out of the instrument at different points in time. The separated chemical species can be identified or quantified using a detector. The detector used can vary from visual identification using the color of sample molecules when working with glass columns, to mass spectrometers (4). The output of a chromatograph is called a 

, which is typically a plot of detector response on the 

Note that this chromatogram is a plot of detector response versus time in minutes, and that there are seven peaks present. The position of each peak in a chromatogram is denoted by its 

, a measure of how long an analyte spent inside the chromatograph. Note that the first peak in Figure 1 has a retention time of 9 min, and that the other peaks came out at later retention times. The size of the peak in a chromatogram can be used to measure the concentration of the chemical species in the original sample that gave rise to that peak (5).

The presence of seven peaks in Figure 1 means the detector saw at least seven different components from the sample exit the instrument. Assuming all components in the sample were completely separated, this means there were seven different components present in the sample. However, chromatography does not always completely separate all the components in a mixture. It is possible for different chemical species to have the same retention time. For example, there may be two or more chemical species responsible for the peak with a retention time of 9 min in Figure 1. Although laboratories sometimes use retention times to identify chemical species this should never be done since a retention time is not unique to a chemical species. Therefore, molecular identification techniques such as infrared spectroscopy (6,7) and mass spectrometry (5) should be used.

 as it’s called in chromatography, of length (L), diameter (D), and radius (r) as shown in 

The volume of this column is given by the equation for the volume of a cylinder which is seen in 

where V is the column volume, r is the radius of column, and L is the length of column.

Now imagine that we pack our column with perfectly spherical particles all the same size (an oversimplification but needed here to keep things simple). This column packing is called a 

 because it does not move and is seen in Figure 2. An example of stationary phase is silica. Once the column is packed, the stationary phase will occupy a volume denoted by V

, and the total empty space in the column, made up of the spaces and interstices between stationary phase particles, is called the 

. Because the total contents of the column are simply the sum of the volumes of the stationary phase and the void volume, we can write an equation for the total column volume (V

 is a fluid that passes through the column. When our chromatographic column is filled with this fluid it will fill all the voids, thus the volume of the mobile phase or V

In gas chromatography (GC) the mobile phase is a gas such as hydrogen, nitrogen, or helium (5). GC has been used to measure cannabinoid concentrations (8) and terpene concentrations (9) in cannabis samples. In liquid phase chromatography the mobile phase is a liquid (5). An example of a liquid mobile phase that has been used to measure cannabinoids is a mixture of water and acetonitrile (9,10). We define the 

 in a chromatographic experiment as the volume per unit time of mobile phase that exits the column. For example, if 1 mL of mobile phase exits the column per minute then the flow rate is 1 mL/min.

Imagine injecting a sample into the head of the column that has zero affinity for the stationary phase, that is, it will not absorb on or “stick to” the particles the column is packed with. As we will see, it is the fact that different molecules have different affinities for a given column packing that is behind the separation mechanism of chromatography (5,8–10). Since our molecule has zero affinity for the column packing, it will only have access to the voids and interstices in the column, that is, the void volume. Let’s assume our void volume is 1 mL. If the flow rate is 1 mL/min, all the mobile phase currently in the column will be flushed out of the column in 1 min. Along with it will come any molecules with zero affinity for the column. In this case, the chromatogram would have a peak 1 min after injection corresponding to the exiting of our zero affinity molecule from the column. In chromatography, the amount of time it takes a molecule with zero affinity for the stationary phase to exit the column is called the 

Imagine we mix up some tetrahydrocannabinol (THC) in ethanol and inject it at the head of our chromatographic column as seen in 

, and that we pump a water mobile phase through a column packed with spherical silica particles where the surface hydroxyl (OH) groups have been derivatized with alkyl chains with 18 carbons in them. In the parlance of chromatography this is a “C18” column. The purpose of the C18 chains is to convert the polar silica surface into one that is non-polar to provide a column with affinity for non-polar molecules. The C18 chain consists of methyl and methylene groups only so it is non-polar, and the bonding scheme of the chains to the silica surface looks something like this: CH

. This setup, where the mobile phase is polar and the stationary phase is non-polar is referred to as 

 chromatography typically involves a polar surface such as naked silica covered with OH groups and a non-polar solvent such as hexane. Normal phase chromatography is used to separate polar molecules since they have an affinity for the column (5). Reversed-phase chromatography typically involves a non-polar stationary phase, such as a C18 column, and a polar mobile phase such as water (5). Reversed-phase chromatography is used to separate non-polar molecules since they have an affinity for the column (5). Reversed-phase liquid chromatography is frequently used to measure THC, cannabidiol (CBD), and other cannabinoids in cannabis samples (9,10). The type of chromatography to use on your sample can be guided by the classic chemical idea of “likes dissolve likes.” In the case of chromatography, the appropriate adage is “likes adsorb likes.” We will discuss the different types of separation mechanisms in more detail in later columns.

Once the THC molecules are injected into the column, assume they will adsorb on the C18 stationary phase since it has an affinity for them. Imagine that the THC partitions in such a way that ½ the molecules are adsorbed and ½ are in solution at any given point in time. There are only a finite number of adsorption sites on our column packing, so the THC molecules adsorb along a finite length of column, as illustrated by the red block in Figure 3. This slug of material will contain ethanol solvent and THC molecules in solution in contact with stationary phase particles with adsorbed THC. The front of this slug of material is called the “solvent front” as seen in Figure 3 because it will be the first part of our sample to exit the column. We now have our sample injected into our column and are ready to begin the separation process. We can think of a chromatographic separation as a race since different molecules pass through the column at different speeds. Thus, the situation we see in Figure 3 is the beginning of the race. However, we will have to wait to see who wins this race until next column, as a discussion of the details of how a separation happens is too long for this column.

Chromatography is an analytical chemical technique used to separate mixtures into their component chemical species. This is typically done by packing a column with a stationary phase, injecting the sample at the head of the column, and then passing a mobile phase of fluid through the column. Based on their affinities for the column packings different chemical species will leave the column at different points in time. As the different chemical species exit the column they can be quantitated and identified using different detectors. We defined the column volume, void volume, stationary volume, void time, and retention time, concepts we will need going forward to understand chromatographic separation mechanisms.

https://en.wikipedia.org/wiki/Chromatography

https://en.wikipedia.org/wiki/List_of_Nobel_laureates_in_Chemistry

https://en.wikipedia.org/wiki/Julia_Johnson

https://en.wikipedia.org/wiki/Gas_chromatography–mass_spectrometry

, 5th Edition, (Marcel Dekker, New York, 1995, Boca Raton, Florida) 2011.

 Fundamentals of Fourier Transform Infrared Spectroscopy,

 2nd Edition (CRC Press, Boca Raton, Florida, 2011).

Infrared Spectral Interpretation: A Systematic Approach 

T. Ruppel, M. Kuffel, “Cannabis Analysis: Potency Testing Identification and Quantification of THC and CBD by GC/FID and GC/MS,” PerkinElmer Application Note, 2013.

M.W. Giese, M.A. Lewis, L. Giese, and K.M. Smith, 

A. Hazekamp, A. Peltenburg, R. Verpoorte, and C. Giroud, 

, PhD, is Founder, CEO, and Chief Technical Officer of Big Sur Scientific. He is the inventor of the BSS series of patented mid-infrared based cannabis analyzers. Dr. Smith has done pioneering research and published numerous peer-reviewed papers on the application of mid-infrared spectroscopy to cannabis analysis, and sits on the editorial board of 

. He has worked as a laboratory director for a cannabis extractor, as an analytical chemist for Waters Associates and PerkinElmer, and as an analytical instrument salesperson. He has more than 30 years of experience in chemical analysis and has written three books on the subject. Dr. Smith earned his PhD on physical chemistry from Dartmouth College.
Direct correspondence to: 

Articles

Cannabis Analysis 

In part I in a new series of columns, we begin a discussion of the theory of chromatographic separations. Our goal is to derive equations that will help you better perform experiments to optimize your separations.

Principles of Chromatography, Part I: Theory

The lack of federal oversight with regard to heavy metals in consumer cannabis products in the US has left individual states to regulate its use. Cannabis is legal in 37 states, while 18 states and Washington, D.C. allow its use for adult recreational consumption (1). However, the cannabis plant is known to be a hyper-accumulator of heavy metals in the soil, so it is critical to monitor levels of elemental contaminants to ensure cannabis products are safe to use. Unfortunately, there are many inconsistencies with heavy metal limits in different states where cannabis is legal. The vast majority of these states define the “big four” heavy metals as lead (Pb), arsenic (As), cadmium (Cd), and mercury (Hg). New York State also requires the testing for chromium (Cr), nickel (Ni), copper (Cu), antimony (Sb), and zinc (Zn), while Michigan requires inorganic As (not total) and also adds Cr, Ni, and Cu to the big four. Maryland and a few other states also include Cr as well as the big four

 Some states base their limits directly in the cannabis, while others are related to human consumption per day. Others take into consideration the body weight of the consumer, while some states do not even have heavy metal limits. To complicate the situation, certain states only require heavy metals in the cannabis plant or flower, while some give different limits for the delivery method such as oral, inhalation, or transdermal (2).

These inconsistencies and the fractured nature of state-based limits would make it extremely complicated to implement at the federal level unless there was a completely fresh assessment of the regulations. For example, why does New York State currently have action limits for nine elemental contaminants whereas the vast majority of the other states only require the big four. In other words, what do New York regulators know about the toxicological impact of the five additional elements, compared to say California that only regulates four. Or why does Michigan require inorganic arsenic (iAs) while every other state just lists total arsenic. Furthermore, how can some states justify no heavy metal action limits at all? It might all become a moot point when federal regulators have oversight of the industry, but what will that regulatory panel look like? The disparity in state-based limits can be a good indicator, but there might also be clues in the way federally approved cannabidiol (CBD) drug formulations have historically been regulated, together with an understanding of how national standards and testing organizations have approached writing standardized methods and the development of reference materials.

To highlight the disparity between state-based limits and federal guidelines for pharmaceutical formulations, the Food and Drug Administration’s (FDA) Botanical Review Team in the Office of New Drug Products recently publisheda study entitled “Quality Standards in State Programs Permitting Cannabis for Medical Uses,” which compared the maximum allowable limits of the big four heavy metals for states that had medical cannabis programs (MCP) with 

 Chapter <232> permitted daily exposure (PDE) concentrations in inhalation and oral drug formulations (3). Although the actual states are not mentioned, it can be seen from 

 that in the majority of cases, the eight medical programs reported are very different to the USP limits. In fact, it can be seen by the data in red that they are significantly higher than the federal limits.

There is no obvious reason as to why the state programs action limits are so different, but the researchers’ overall conclusion was three-fold:

Testing for the four heavy metals in inhalation and oral products was lacking in the majority of the state medical programs.

Even when present, it was inconsistent across the different medical programs.

It did not always align with USP recommendations.

Clearly, there is a need for more consistency across state lines, particularly as the industry inevitably moves in the direction of federal oversight. This is further compounded by the fact that there is compelling evidence in the public domain that only monitoring the big four heavy metals is not enough to ensure consumer safety. But how many metals should there be in an expanded panel, particularly as there is no comprehensive understanding of the sources of elemental contaminants in the cannabinoid production processes. Moreover, besides the big four, there is no consensus on the toxicity impact of other heavy metals in cannabis and hemp, as there have been no risk studies carried out regarding heavy metal contaminants and for that reason, consumer health is likely being compromised.

At a recent American Society for Testing and Materials (ASTM) workshop dedicated to the measurement of heavy metals in cannabis and hemp consumer products, compelling evidence was presented by several researchers that suggested 10–15 elemental contaminants are worthy of consideration (4). Moreover, the National Institute of Standards and Technology (NIST) is developing a 13-toxic element hemp certified reference material (CRM) through its Cannabis Quality Assurance Program (CannaQAP) to include Pb, Cd, As, Hg, beryllium (Be), cobalt (Co), Cr, manganese (Mn), molybdenum (Mo), Ni, selenium (Se), uranium (U), and vanadium (V) (5). In addition, ASTM International’s D37 Cannabis Committee has recently developed and approved a standardized inductively coupled plasma-mass spectrometry (ICP-MS) method for up to 24 different elemental contaminants in cannabis. As a result, it is going to offer testing laboratories the flexibility to select the heavy metals that are important in their jurisdiction and, in particular, to encourage state regulators to expand the panel beyond the big four to those additional metals that are worthy of regulatory scrutiny.

The only solid evidence we have at this current time for what could be a federally regulated panel is with the FDA approved CBD-based drug Epidiolex, which is available in the US to treat childhood seizures. Manufactured by UK-based GW Pharmaceuticals, it went through the regulatory process four years ago to get it approved in the US and had to show compliance by meeting PDE limits for up to 24 elemental impurities as defined in USP Chapter 232 (6) and 

International Council for Harmonisation of Technical Requirements for Pharmaceuticals for Human Use (ICH)

Moreover, the USP recently published a draft monograph for CBD as an active pharmaceutical ingredient (API) for a federally approved drug formulations which stated that (8): “Elemental impurities in official drug products are controlled according to the principles defined and requirements specified in Elemental Impurities—Limits 232, as presented in the General Notices 5.60.30.”

In the long term, this could possibly indicate that the FDA will regulate CBD products for up to 24 elemental contaminants when it eventually has oversight of the cannabis industry. But more importantly, in the short term it implies that CBD being manufactured in the US for recreational or medicinal purposes does not meet the purity requirements for federally-approved drugs, because currently it only has to comply with the state’s maximum limits for heavy metal contaminants, which in most US states is typically only Pb, Cd, As, and Hg.

However, it’s important to stress that a panel generated by pharmaceutical regulators isn’t necessarily one that should be used by the cannabis industry, as the process for manufacturing cannabinoids is very different to drug products. At some point, the cannabis industry needs to carry out a comprehensive risk assessment study to provide evidence as to what metals are worthy of consideration similar to what the pharmaceutical regulators did to characterize elemental impurities in drug products and formulations (9). To better understand this risk assessment process and how it might have a bearing on a cannabis panel of heavy metal contaminants, let’s take a closer look at how pharmaceutical regulations for elemental impurities came into existence.

Federal Regulations for Drug Substances and Products

The pharmaceutical industry began the process to overhaul regulations and methodology for elemental impurities more than 20 years ago when it updated its 100-year-old semi-quantitative sulfide precipitation colorimetric test for a small suite of heavy metals to eventually arrive at a method to monitor 24 elemental impurities in drug products using plasma spectrochemistry. Moreover, they completely reassessed the toxicological impact of these elemental contaminants based on well-established animal models and defined them by PDE limits according to the mode of administration (oral, parenteral, inhalation, and transdermal) and classified them by toxicity and the probability of finding them in the drug manufacturing process.

 232 - Elemental Impurities (6) together with 

 Q3D Guidelines (7). While the measurement procedures were defined in 

 Chapter 233 which describes the plasma spectrochemistry methodology (inductively coupled plasma-optical emission spectrometry [ICP-OES] or ICP-MS), a microwave digestion procedure, and a full set of validation protocols (10). 

 represents the full list of USP and ICH elemental PDE limits in microgram per day (µg/day) per delivery method and toxicological classification, which are explained in greater detail in 

It’s also important to emphasize that the data in Table II are maximum limits per day. So, for a suggested daily dosage of 10 g, these limits should be divided by 10 to calculate the maximum allowable limits in the drug products in microgram per gram (µg/g). However, even though 10 g is a typical maximum dosage for drugs, we have no way of knowing in what quantities consumers use cannabis products. So, if larger quantities are being consumed these PDE limits will be even lower. Furthermore, the mode of administration will also impact the regulated limit, so inhalation PDEs in most cases are significantly lower than the oral ones. In addition, the classification number will impact the frequency of testing with the Class 1 and 2A metals of higher priority that the Class 2B and 3 metals. In fact, the classification can offer clues as to what elements from this list would be worthy of consideration for an expanded panel to regulate cannabis and hemp consumer products. For example, Class 1 and 2A would warrant inclusion in any regulated panel, whereas Class 2B metals would likely not be required at all, because they are not used in the manufacturing process. On the other hand, Class 3 metals may not be required for oral products but would be required for inhalation products such as vaping devices.

Currently in the life cycle of the cannabis industry, it is not obvious where the heavy metal regulatory landscape will end up. Clearly, the big four are not enough, but what is a realistic panel that reflects the real world of cannabinoid production? New York and Michigan states are leading the charge with an additional five and four metals respectively, with a few other states like Maryland and Missouri adding chromium to the big four. But it’s clear from the recent study carried out by the FDA’s Botanical Review Team that they view the efforts of the states to regulate heavy metals as being inadequate. However, recent activity from the USP has sent signals to the industry that if they want to produce CBD as an API for use as a federally approved drug formulation, they are going to have to monitor up to 24 elemental contaminants to meet the quality attributes. And it’s encouraging that standards organizations are developing standardized methods to meet these demands with NIST developing a 13-toxic element hemp CRM and ASTM approving an ICP-MS method for up to 24 different elemental contaminants in a wide variety of cannabis and hemp related samples. The evidence is clearly going in that direction, and it should be only a matter of time before the industry moves beyond the big four and has an expanded regulated panel that is more in line with the pharmaceutical industry, and better reflects consumer safety concerns.

R. Thomas, "The importance of measuring heavy metal contaminants in cannabis and Hemp," available at: 

https://cdn.technologynetworks.com/ac/Resources/pdf/the-importance-of-measuring-heavy-metal-contaminants-in-cannabis-and-hemp-312957.pdf

 (2022) “Quality standards in state programs permitting cannabis for medical uses,” 

R. Thomas, "A recap of ASTM’s workshop on measuring elemental contaminants in cannabis and hemp consumer products,"

https://www.analyticalcannabis.com/articles/a-recap-of-astms-workshop-on-measuring-elemental-contaminants-in-cannabis-and-hemp-consumer-313229

https://www.nist.gov/programs-projects/nist-tools-cannabis-laboratory-quality-assurance

https://www.usp.org/chemical-medicines/elemental-impurities-updates

https://www.ema.europa.eu/en/documents/scientific-guideline/international-conference-harmonisation-technical-requirements-registration-pharmaceuticals-human-use_en-32.pdf

https://www.gmp-compliance.org/gmp-news/usp-draft-monograph-for-cbd

R. Thomas and A. DeStefano, "Understanding sources of heavy metals in cannabis and hemp: Benefits of a risk assessment strategy – part 1," 

https://www.analyticalcannabis.com/articles/how-to-better-understand-sources-of-heavy-metal-contaminants-in-cannabis-part-1-what-can-we-learn-313862

 is the principal of Scientific Solutions, an educational consulting company that serves the needs of the trace element user community. He has worked in the field of atomic and mass spectroscopy for almost 50 years, including 24 years for a manufacturer of atomic spectroscopic instrumentation. He has served on the American Chemical Society (ACS) Committee on Analytical Reagents (CAR) for the past 20 years as leader of the plasma spectrochemistry, heavy metals task force, where he has worked very closely with the United States Pharmacopeia (USP) to align ACS heavy metal testing procedures with pharmaceutical guidelines. Rob has written more than 100 technical publications, including a 15-part tutorial series on ICP-MS. He is also the editor and frequent contributor of the "Atomic Perspectives" column in 

 magazine, as well as serving on the editorial advisory board of 

. In addition, Rob has authored five textbooks on the fundamental principles and applications of ICP-MS. His most recent book is a new paperback version of 

Measuring Heavy Metal Contaminants in Cannabis and Hemp

 published in December, 2021. Rob has an advanced degree in analytical chemistry from the University of Wales, UK, and is also a Fellow of the Royal Society of Chemistry (FRSC) and a Chartered Chemist (CChem).

Navigating the Lab

Currently in the life cycle of the cannabis industry, it is not obvious where the heavy metal regulatory landscape will end up. Clearly, the big four are not enough, but what is a realistic panel that reflects the real world of cannabinoid production?

Expanding the Panel of Heavy Metal Contaminants in Cannabis and Hemp Beyond the Big Four:  What Is the Regulatory Evidence Telling Us?

The findings of this study are from the research initiative submitted to the Pennsylvania State Department of Health in accordance with House Bill 1024. The research study involved evaluating the effectiveness of solvent-based extraction methods for eliminating or substantially diminishing microbial contamination from feed materials (for example, Cannabis sativa) during the extract manufacturing process. This study investigated both supercritical carbon dioxide and hydrocarbon extraction technologies. Organic Remedies Inc.’s unidirectional aseptic manufacturing infrastructure and processes were used starting with contaminated plant material to prepare primary extracts followed by essential, decarboxylated, and distilled oils. Testing of extracts for microbial contamination according to Pennsylvania State regulations was carried out by a state certified laboratory. Results of testing associated with starting plant material revealed the presence of 28,000 cfu/g total aerobic bacteria, 17,975 cfu/g enterobacteriaceae, and “too numerous to count” cfu/g total yeast and mold. E. coli, salmonella spp., and mycotoxins were not detected. Results of testing all extracted materials showed that they were free of the tested microbial contaminants and fungal toxins. Data presented here show the results of testing for the hydrocarbon extraction pathway and the supercritical carbon dioxide pathway. In addition to testing for microbial contamination, the data show results for content of delta-9 tetrahydrocannabinol (THC), cannabidiol (CBD), total cannabinoids, and total terpenes, all of which were used to evaluate extraction efficiency.

Organic Remedies Inc., is a vertically integrated grower and processor located in Carlisle, Pennsylvania. As a licensed Clinical Registrant (CR), Organic Remedies Inc., participates with the Philadelphia College of Osteopathic Medicine, an Academic Clinical Research Center (ACRC), in research studies that explore the medicinal qualities of 

 and its related formulated products. With the recent refinement of the Pennsylvania Medical Marijuana Regulations (1), ACRCs and CRs were allowed to undertake studies exploring whether the extraction process and associated refinement protocols are capable of producing materials that were within Department of Health quality standards when contaminated plants were utilized as feed material.

The current microbial contaminant limits for 

 feed material intended for use in extracted product manufacturing are non-detect, non-detect, <10,000 cfu/g, <10,000 cfu/g, and <1,000 cfu/g for 

total aerobic bacteria, total yeast and mold, and bile tolerant gram-negative (

) bacteria, respectively (1). Pennsylvania microbial limits (1) are relatively strict in comparison to other states’ standards; the rationale for such strict limits is a result of cannabis’ definition as a medicine in the current regulatory framework of the state. Moreover, no form of post-harvest sterilization is permitted; therefore, if a batch of flower or trim fails microbial testing it must be destroyed.In an efficiently operating indoor grow the cost of dried or cured plant material to the producer can be as much as $2.00/g; in a greenhouse setting, that cost can be as low as $0.75/g. As a result, plant material not conforming to PA microbial criteria has substantial negative impacts on business revenue, employment opportunities, and the cost of medical cannabis products.

Sterilization of cash crops is not uncommon in the food or nutraceutical industries; however, those products are very different from cannabis because they are usually fruits and herbs that are intended for oral consumption by the consumer. Furthermore, products such as fruits, vegetables, and herbs have very different morphological characteristics compared to 

 inflorescence intended for vaporization or the manufacturing of products created via extraction. For the food and nutraceutical industries, numerous sterilization techniques are outlined in the 

 (USP) (2) and permitted by the US Food and Drug Administration (FDA) and—when coupled with final product testing—can reliably ensure consumer safety. Some examples of terminal sterilization technologies that are permitted by the FDA include steam-sterilization, dry-heat sterilization, gas sterilization, ionizing radiation, and sterilizing filtration (2). Furthermore, the USP and FDA describe a process-based microbial risk mitigation technique for manufacturing called “Unidirectional Aseptic Processing” (2). That process, which does not diminish the importance of terminal sterilization and final product testing, incorporates numerous terminal sterilization techniques in the manufacturing process, appropriate air handling infrastructure, and intensive training of personnel (2).

 differs from other cash crops regarding the intended use of its manufactured products. In fact, the entire manufacturing process—including the variety of final product formats—differs from other agricultural industries insofar as it is intended to produce botanical, extracted raw materials, and formulated products. Further complicating the manufacturing of 

 products is that there is no single set of regulations governing the quality of raw material or final products; this lack of central governance is due to the classification of

 -THC)—as a Schedule I controlled substance (3).Therefore, regulatory frameworks are created on a state-by-state basis and recommendations set forth by the FDA or their associated advisory bodies may not be wholly adopted. For example, Pennsylvania does not allow for the use of federally accepted microbial remediation technology to sterilize contaminated plant material prior to market distribution. Additionally, the state does not allow 

growers or processors to utilize plant material contaminated with bacteria or yeast and mold in the manufacturing of raw materials or formulated products via solvent extraction. The logic behind these types of regulations is sound in that they are set up to be risk averse and to ensure that patients in the medical cannabis market of Pennsylvania have access to safe therapies; however, the current (restrictive) framework overlooks the capabilities of the extraction process including the associated infrastructure to deactivate or remove microbial contaminants.

In cooperation with the Philadelphia College of Osteopathic Medicine and with the approval of the Pennsylvania Department of Health, Organic Remedies Inc. designed and executed the proceeding study to investigate the potential of a well-designed solvent extraction and raw-material manufacturing process to deactivate or remove microbial contaminants from compromised feed material during the production of materials used to create consumer products.

Two batches of contaminated cannabis inflorescence (Triangle Kush x Fruity Pebbles, TKFP211007H/TKFP211007H1) were combined into a single batch of plant material large enough to support two experiments. Sub-batches were derived from the homogenized parent batch and were treated differently pre-extraction. For hydrocarbon extraction, the plant material's native bud structure was retained and batched in equal-mass replicates. Prior to carbon dioxide extraction, all parent plant material was ground using a Robocoupe Blixer 15 then batched into equal mass replicates.

The total amount of plant material allocated to hydrocarbon and supercritical fluid extraction was 9.67 kg and 12.98 kg, respectively. The mass of plant material in each hydrocarbon replicate (n=5) was 1.93 kg and the mass of each supercritical carbon dioxide replicate (n=5) was 2.60 kg.

Carbon Dioxide and Hydrocarbon Extraction

Hydrocarbon extraction was performed by cooling the extraction solvent to -20 

C before injecting the solvent into the closed-loop extraction system. The extraction process was conducted under dynamic flow (1.0 L/min.) conditions at 1 Bar. Each extraction lasted 15 min with temperature held constant at -20 

C by heat exchangers located before and after the extraction columns.

Supercritical carbon dioxide extraction was conducted under dynamic flow with the solvent pressurized to 285 Bar and heated to 55 

C; the flow rate of the carbon dioxide solvent was 275 g/min over 8 h resulting in a solvent to feed ratio of 51:1. The temperature and pressure conditions in the retrograde phases were 55 

C/46 Bar, in the first, second, and third separators, respectively. The process conditions outlined above resulted in a carbon dioxide extraction output of two different fractions; those fractions were derived from separator 1 and separator 2. No material was obtained from separator 3. Concordantly, each extraction replicate yielded two fractions (denoted A and B) that were tested independently and later combined for downstream processing.

Winterization of the primary hydrocarbon extract was performed in the extraction solvent by maintaining a temperature of -80 

C for 12 h; the raw extract from each replicate was transferred to a vessel suitable to long term storage at -80 

C.Following the extended period of exposure to -80 

C, the raw extract was pumped at a rate of 3 L/min through the CUNO BHN2 bag (25 µm) filtration system at 1 Bar.

Combined extracts (such as the A and B fractions) from each carbon dioxide extraction replicate were dissolved in absolute ethanol at 40 

C with the assistance of a sonic bath. The volume of ethanol used during this process was 10x the mass of the combined extracts. To ensure that the raw extracts were fully dissolved, a hand blender was used. The ethanol and raw extract mixture was then put into a freezer at -40 

C for 12 h; following, the fats and waxes were removed using a 300 mm stainless steel Buchner funnel, Whatman 1 (11 µm) filter paper, and a vacuum pump and flask assembly. The residual fats on the filter were rinsed with clean, cold ethanol (-40 

To remove chlorophyll from the winterized hydrocarbon extracts, 650 g of pharmaceutical grade magnesium silicate was packed into a specialized column that was attached to the closed-loop extraction system. With that column in place, the winterized and filtered extract was pumped down through MgSiO

 at a rate of 1 L/min, at a pressure of 1.5 Bar, and 20 

C until all material was fed in through the system.

Clarification of the winterized and filtered supercritical carbon dioxide extract was conducted by direct addition of peat-derived activated carbon. Next, 35 g of activated carbon was added directly to the winterized extract that was dissolved in ethanol. The mixture was then heated to 50 

C and agitated with an immersion circulator for 30 min. Subsequently, the activated carbon was removed from the ethanolic mixture using a 300 mm Buchner funnel with Whatman 1 filter paper (11 µm) attached to a vacuum pump and flask system operating at 0.1 Bar.

In both extraction paradigms, sterilizing filtration was executed in a similar manner except for a single difference; that difference was the temperature that was maintained during the process. In the case of hydrocarbon extracts, the sterilizing filtration occurred at room temperature. For supercritical carbon dioxide extracts, the winterized and clarified mixture was cooled to -40 

C for 12 h before sterilizing filtration to ensure that all fats and waxes were captured. Both hydrocarbon and supercritical carbon dioxide extracts were pumped into the 3M cartridge filtration system (<0.2 µm) at a pressure of 1 bar until all extract moved through the system.

For hydrocarbon extracts, the solution was transferred to a 10 L rotary evaporator. The conditions for solvent recovery of the hydrocarbon solvent were 36 

C, 0.5 Bar, and 100 RPM. The solvent recovery system was continuously fed until all material was transferred and the essential oil of cannabis was the consistency of maple syrup.

For supercritical carbon dioxide extracts, a 10 L rotary evaporator was used and the conditions were 55 

C, 0.15 Bar, and 100 RPM. The solvent mixture was continuously fed into the system until all material was transferred and the essential oil was the consistency of maple syrup.

The purging process is unique to hydrocarbon manufacturing and is the terminal process for creating raw materials that are used in hydrocarbon-extract product manufacturing. The purging process was executed in a radiant heat vacuum oven coupled with a scroll-pump and cold trap assembly. The extract was purged at 30 

C and 0.07 Bar in a thin layer for 22 h.

Decarboxylation of the refined carbon dioxide cannabis extract was conducted in a boiling flask and stirring-mantle assembly coupled with a vacuum pump and cold trap. The decarboxylation was conducted over 2 h at a temperature of 165 

C and a vacuum level of 0.001 Bar with constant agitation.

Cannabinoid distillation was conducted in a wiped film system with a feed rate of 250 g/h at 0.08 mBar; the cannabinoids were condensed at 98 

C, and the terpenes were condensed at -15 

Overview of Material Flow and Testing Points by Manufacturing Line

Material Flow – Batching (parent material) ➔ Extraction ➔ Winterization ➔ Clarification ➔ Sterilizing filtration ➔ Solvent recovery (essential oil) ➔ Purging (purged essential oil)

Testing Points – Parent material, post solvent recovery (essential oil), post purging (purged essential oil)

Material Flow – Grinding (parent material) ➔ Extraction (raw extract) ➔ Winterization ➔ Clarification ➔ Sterilizing filtration ➔ Solvent recovery (essential oil) ➔ Decarboxylation (decarboxylated oil) ➔ Distillation
(distillate oil)

Testing Points – Parent material, post extraction (raw extract), post solvent recovery (essential oil), post decarboxylation (decarboxylated oil), post cannabinoid distillation (distillate oil)

Analytical Testing of Extracted Materials

All testing of samples for cannabinoid potency (%), terpene content (%), microbial contaminants (cfu/g), and mycotoxins (µg/g) was undertaken by Steep Hill Pennsylvania, an International Organization for Standardization (ISO) and International Electrotechnical Commission (IEC) certified laboratory that is approved by the Commonwealth of Pennsylvania for testing of medical marijuana products.

The samples were homogenized and subdivided into the required weight for each analytical test. Each test was completed in replicates of five. Test portions were extracted with an organic solvent and prepared for analysis following Steep Hill PA’s standardized operating procedures. Cannabinoid analysis was conducted using high performance liquid chromatography (HPLC). Limits of quantitation for the cannabinoids CBD, cannabidiolic acid (CBDA), cannabinol (CBN), Δ

 THCA), cannabidivarin (CBDV), tetrahydrocannabivarin (THCV), cannabigerol (CBG), cannabigerolic acid (CBGA), delta-8-tetrahydrocannabinol (Δ

THC), and cannabichromene (CBC) were 0.52, 0.42, 0.28, 0.41, 0.41, 0.39, 0.39, 0.51, 0.50, 0.78, and 0.47 µg/g, respectively. Terpene analyses were conducted using gas chromatography–mass spectrometry (GC–MS) and the limit of quantitation varied between 0.055 and 0.072 µg/g depending on the identity of the analyte. Mycotoxin analyses were conducted using liquid chromatography–mass spectrometry (LC–MS); the limit of quantitation for Aflatoxin B1, Aflatoxin G1, Ochratoxin A, Aflatoxin B2, and Aflatoxin G2 were 1.44, 0.56, 1.44, 1.34, and 1.28 µg/g. Microbial testing was conducted using ready-to-use plates provided by 3M and BioRad. Third-party validations confirm that these platforms are reliable and derived data are repeatable. The manufacturers’ standard operating procedures (SOPs) for sample preparation and analysis were followed. The following ready-to-use agar products were used in this study:

3M Petrifilm Rapid Yeast & Mold Count Plates - 3M ID: 7100039447

3M Petrifilm Aerobic Count Plates - 3M ID: 7100039310

3M Petrifilm Enterobacteriaceae Count Plates - 3M ID: 7100126818

3M Petrifilm E. coli/Coliform Count Plates - 3M ID: 7100126767

Two data matrices were assembled from available data; the hydrocarbon and carbon dioxide matrices were 15x18 and 70x18, respectively. Independent variables in each matrix included the oil identification, replicate, and process-input material mass. Response variables included output mass and the cannabinoids CBD, CBDA, CBN, Δ

 THC, and CBC; additionally, response variables included total yeast and molds, total aerobic bacteria, total enterobacteria, 

The variable “total mycotoxins” was calculated from individual values of the aflatoxins B1, B2, G1, G2 and Ochratoxin A.

Prior to conducting any calculations or analyses, all cannabinoid potency results were mathematically converted to account for the decarboxylation of acid form cannabinoids. This was done by multiplying the acid form percentage of a given cannabinoid by the decarboxylation coefficient (0.87) then adding that to the percentage of the same neutral form cannabinoid. Statistical analysis of each data set was undertaken using Repeated Measures Analysis and Tukey's honestly significant difference (HSD) post-hoc analysis to evaluate the influence of each manufacturing step in removing microbial contaminants. All analyses were executed in R (

The parent material contained 17.27% total cannabinoids (that is, Δ

 THC = 3.94%) and 1.28% total terpenes. Microbial tests indicated that the plant material was contaminated with 28,000 cfu/g total aerobic bacteria, 17,975 cfu/g enterobacteriaceae, and “too numerous to count” cfu/g total yeast and mold. Where “too numerous to count” were found for a specific group of microbial contaminants, a value of 1 x 10

 cfu/g was tabulated in the data matrix for statistical purposes. No 

 or mycotoxins were detected in the parent material.

The most important finding of this study was that both hydrocarbon and supercritical carbon dioxide extraction of contaminated plant material resulted in manufactured raw materials that were free of viable microbial contaminants that were evaluated during this study. No statistical analyses were possible because there was no variance in the dataset following the extraction step; in short, all tested microbial contaminants were not detected after the parent plant material was processed via solvent extraction.

The essential oil manufacturing process, which included extraction, winterization, clarification, sterilizing filtration, and solvent recovery, extracted 1185.2 g of cannabinoids from the parent material that contained 1670.2 g of cannabinoids. The essential oil was free of all tested microbial contamination and mycotoxins following those manufacturing steps (see 

The purged oil was also found to be free of all tested microbial contaminants and mycotoxins (see Figures 1-3 and Table I); the transition of material from the essential oil phase to this phase was 93.9% efficient where 1112.3 g of cannabinoids were recovered during the purging process from the essential oil that contained 1185.2 g of cannabinoids.

Five extractions, each with 2 fractions, were 84.6% efficient where 1730.8 g were extracted from the plant material that contained 2046.3 g of cannabinoids. Additionally, raw extracts from the supercritical carbon dioxide process were free of all tested microbial contaminants and fungal metabolites (see 

Following winterization, clarification, sterilizing filtration, and solvent recovery, the essential oil was found to be free of all tested microbial contaminants and mycotoxins (see Figures 4-6 and Table II).Overall, those processes were 78.8% efficient for cannabinoid retention where 1363 g of 1730.8 g of cannabinoids were retained through the four steps.

The decarboxylation step was found to be 97.2% efficient for cannabinoid retention and 1325.4 g of 1363 g of cannabinoids were recovered during the process. Additionally, the decarboxylated oil was found to be free of all tested microbial contaminants and fungal metabolites (see Figures 4-6 and Table II).

The cannabinoid distillation step was found to be 96.0% efficient; present in the decarboxylated oil was 1325.4 g of cannabinoids and the distillate oil contained 1272 g of cannabinoids. Finally, the distillate oil was free of all tested microbial contaminants and mycotoxins (see Figures 4-6 and Table II).

Overall, our results suggest that the tested microbial contaminants are not transferred from feed material to the raw materials produced during extraction and refinement and that those raw materials are suitable for use in product manufacturing.

Sterilizing of products during the manufacturing process or prior to distribution is not uncommon in the food, pharmaceutical, and medical device industries. The US FDA has reviewed a variety of sterilization methods for use in the biomedical and food industries such as irradiation, gas sterilization, steam or dry heat sterilization, sterilizing filtration, and liquid dips (4,5). Furthermore, the US FDA presented a challenge in 2019 to encourage companies to develop a new sterilization technique or set of techniques capable of replacing ethylene oxide in the food and medical device manufacturing industries because of its flammability and potential carcinogenicity (5-7). One promising technique for product sterilization is the application of supercritical carbon dioxide including additives such as water (8,9), hydrogen peroxide (10,11), peracetic acid (4,12), and ethanol (13). Additionally, supercritical carbon dioxide without additives has been shown to remove notable organisms such as 

Hydrocarbon extraction utilizes a hydrocarbon solvent and submicron filtration (such as, sieve <0.2 µm) in the manufacturing process to produce products such as sugars, diamonds, liquid live resins, and diamond sauces. The results of our experiment utilizing contaminated plant material suggested that the techniques and infrastructure used here can create raw materials free of viable microbial contaminants and fungal metabolites that were tested here (no data shown for mycotoxins; 0 ppb at all stages for both technologies). Therefore, we assert that the hydrocarbon extraction, processing techniques, and infrastructure utilized in this study are suitable for creating raw material from contaminated feed material that meet or exceed current PA Department of Health regulations for extracted materials (1).

Supercritical carbon dioxide manufacturing also takes advantage of numerous aseptic processing technologies. At the extraction stage, supercritical carbon dioxide was utilized without the introduction of additives. It is unclear whether microbial deactivation or removal was due to supercritical carbon dioxide alone because there are a variety of materials interacting in the supercritical carbon dioxide during extraction (for example, water, cannabinoids, terpenes, and so on). However, it is apparent that the tested microbial contaminants were removed or deactivated during the first stage of our supercritical carbon dioxide manufacturing process. Furthermore, after the extraction process the extract was winterized, clarified, and filtered through a <0.2 µm sieve prior to solvent recovery; our results suggest that microbial deactivation or removal was maintained through those steps. Subsequent supercritical carbon dioxide manufacturing processes leverage heat to decarboxylate the acid-form cannabinoids and distill the neutral-form cannabinoids. Both of those processes were executed at temperatures greater than 100 

C for extended periods of time; our results suggest that, at both of those stages, microbial deactivation or removal is maintained and that those products are free of fungal metabolites. Therefore, we assert that the infrastructure and techniques leveraged during this study can produce raw materials that meet or exceed PA Department of Health’s current regulations (1).

This study, which evaluated the potential of a well-designed unidirectional aseptic processing approach to both hydrocarbon and supercritical carbon dioxide extract manufacturing, suggests that the extraction process is capable of deactivating or removing the tested microbial contaminants when highly contaminated 

inflorescence is used as feed material. Finally, the absence of viable microbial contamination tested herein was maintained throughout all raw material manufacturing steps in both the hydrocarbon and supercritical carbon dioxide schemes.

We would like to thank the Pennsylvania Department of Health for allowing us to undertake this study as well as the Pennsylvania Cannabis Coalition and the Organic Remedies Inc. board for financial support. Additionally, we would like to thank the Philadelphia College of Osteopathic Medicine and Duquesne University for their time and support of this research initiative. Finally, this work could not have been completed without the support of the Specialists and Technicians employed in the Organic Remedies Inc. laboratory.

Peer-Reviewed Article

This study, which evaluated the potential of a well-designed unidirectional aseptic processing approach to both hydrocarbon and supercritical carbon dioxide extract manufacturing, suggests that the extraction process is capable of deactivating or removing the tested microbial contaminants when highly contaminated Cannabis spp. Inflorescence is used as feed material.

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