In this study, we evaluated the stability of carboxylated phytocannabinoids analyzed with positive and negative electrospray (ESI+ and ESI-, respectively) liquid chromatography–mass spectrometry (LC–MS) as a function of drying gas temperature. We used both chromatographic and flow injection analysis (FIA) conditions. Post-chromatographic (in-source) acid decarboxylation was not observed over a drying gas temperature range of 75 °C through 300 °C in ESI+ mode under chromatographic conditions. FIA experiments further demonstrated no correlation between in-source acid decarboxylation in ESI+ mode over a drying gas temperature range of 100 °C through 350 °C. Conversely, chromatographic experiments in ESI- mode did reveal a decrease in tetrahydrocannabinolic acid (THCA) and cannabidiolic acid (CBDA) relative abundances and concomitant increases in the relative abundances of tetrahydrocannabinol (THC) and cannabidiol (CBD) as a function of drying gas temperatures. These data highlight a potential concern for the quantitation of acid phytocannabinoids using ESI- mode LC–MS technologies 

 genome encodes synthase enzymes (1) for the synthesis of carboxylated phytocannabinoids such as Î

-tetrahydrocannabinolic acid (THCA), cannabidiolic acid (CBDA), and cannabigerolic acid (CBGA). In the living plant, psychoactive Î

-tetrahydrocannabinol (THC) does not exist at significant concentrations nor does cannabidiol (CBD)-another cannabinoid that has garnered interest in recent years. THC and CBD are formed through decarboxylation of THCA and CBDA, respectively, through exposure to light, heat, and curing processes (2). Figure 1 illustrates the acid to neutral decarboxylation of THCA and CBDA. The reaction order, k

, and the activation energy (EA) of acid -> neutral decarboxylation has been determined over specific temperatures and times in 

 flowering buds (3). Chemical testing can also influence cannabinoid acid decarboxylation if the testing system exposes the sample to heat (4) as in the injection port of a gas chromatography (GC) system. Liquid chromatography–mass spectrometry (LC–MS) systems also expose the acid phytocannabinoids to heat in the electrospray ionization (ESI) source region. To our knowledge, the effect on acid decarboxylation as a function of temperature in the ESI source region has not been reported. In fact, a PubMed query of “phytocannabinoid decarboxylation in the ESI source” and related queries yielded zero results. One study evaluated the changes in chemical profiles of extracts at ambient temperature versus decarboxylation at a series of temperature ranges (5). Our study used ESI LC–MS in both positive and negative modes to determine the effect of electrospray source temperature on the stability of acid phytocannabinoids in neat solutions. (See upper right for Figure 1, click to enlarge.)

Experimental
	We make the following assumption: under the conditions of a high performance liquid chromatography (HPLC) method, decarboxylation is mitigated and acid phytocannabinoids are stable under the time frame of the analysis as they transport across the column stationary phase. The null hypothesis states the acid phytocannabinoids are equally stable and remain relatively unchanged in the ESI source. To reject the null hypothesis, we must observe a decrease in the relative abundance of the carboxylated species with a commensurate increase in the relative abundance of the decarboxylated analogues. To make this determination, we used electrospray ionization in both positive and negative modes under chromatographic and flow injection analysis (FIA) conditions. In the various experiments, the drying gas temperature was varied from 75 °C through 350 °C and we monitored (M+H)

 adducts specific to the carboxylated and decarboxylated chemical species.
	
	


	THCA, THC, CBDA, and CBD were obtained from Cerilliant. All cannabinoid reference standards were obtained in U.S. Drug Enforcement Administration (DEA) exempt format at 1.0 mg/mL in organic solvent. LC–MS grade methanol, acetonitrile, and formic acid were obtained from Agilent Technologies. Millipore de-ionized (DI) water was used for the aqueous mobile phase.


	An LC–MS system (Agilent Technologies) equipped with a diode array detector (DAD) and a nebulizer ESI source was used. The mass spectrometer was operated in ESI+ and ESI- modes over the mass range of 100–500 

. For each experiment, the drying gas temperatures were 100 °C, 150 °C, 200 °C, 250 °C, and 300 °C. The LC–MS conditions are shown below.


	Column: Agilent InfinityLab Poroshell 120 EC-C18, 3.0 × 50 mm, 2.7 Î¼m
	Mobile Phase: A) 0.1% (V/V) formic acid aqueous phase; B) 0.05% (V/V) formic acid organic phase
	Flow rate: 1.0 mL/min
	Column temperature: 50 °C
	Injection volume: 5 µL
	Diode array detector: 230 nm and 270 nm


	Drying gas flow: 13 L/min
	Nebulizer: 55 psig
	Vcap: 3500


	Time (min)                    %B
	0                                   60
	1.0                                60
	7.0                                70
	8.2                                95

High Resolution Accurate Mass (HRAM) FIA Experiments


	A quadrupole time-of-flight (QTOF) mass spectrometer (Agilent Technologies) equipped with a dual ESI source was used. The QTOF was operated in ESI+ mode over the mass range of 100–500 

. Samples were directly infused into the electrospray source with a syringe pump at a rate of 200 µL/h. The drying gas temperature was incremented stepwise at 50 °C intervals from 100 °C through 350 °C. The QTOF parameters and drying gas temperature program are shown below. 


	Drying gas flow: 12 L/min
	Nebulizer:30 psig
	Vcap: 4000
	Fragmentor: 150
	Skimmer: 65
	Acquisition rate: 1 Hz


	0.0 to 1.5 min, Hold 100 °C
	1.5 to 2.5 min, Gradient to 150 °C
	2.5 to 3.5 min, Hold 150 °C
	3.5 to 4.5 min, Gradient to 200 °C
	4.5 to 5.5 min, Hold 200 °C
	5.5 to 6.5 min, Gradient to 250 °C
	6.5 to 7.5 min, Hold 250 °C
	7.5 to 8.5 min, Gradient to 300 °C
	8.5 to 9.5 min, Hold 300 °C
	9.5 to 10.5 min, Gradient to 350 °C
	10.5 to 11.5 min, Hold 350 °C

HRAM Chromatographic Confirmation Experiments


	An ultrahigh-pressure liquid chromatography (UHPLC) system and time-of-flight (LC/TOF) mass spectrometer (Agilent Technologies) equipped with a dual ESI source was used. The column was an Agilent Poroshell 120 EC-18 3.0 mm x 50.0 mm, 2.7 µm particle size (PN 699975-302). The flow rate was 0.5 mL/min. The injection volume was 5 µL. The TOF mass spectrometer was operated in ESI- mode over the mass range of 100–1000 

 with reference mass on. For each experiment, the drying gas temperatures were 75 °C, 100 °C, 175 °C, 200 °C, and 250 °C. The LC/TOF conditions are given in Tables I and II. (See upper right for Table I and Table II, click to enlarge.)

Low Resolution LC–MS Confirmatory Experiments


	For the ESI+ mode experiments, solvent standards containing CBDA, CBD, TCHA, and THCA were prepared at 250 µg/mL in acetonitrile. For the ESI- mode experiments, a solvent standard containing THCA was prepared at 500 µg/mL in LC–MS grade acetonitrile.


	A 100 µg/mL solution of THCA was prepared in LC–MS grade acetonitrile.


	A solvent standard containing CBDA and THCA was prepared at 500 µg/mL in LC–MS grade acetonitrile.


	In ESI+ mode, CBDA, CBD, THC, and THCA are observed at the proper retention times and the DAD signal remains relatively constant over the five injections of increasing drying gas temperature. No CBD or THC are observed under the CBDA or THCA peaks at any drying gas temperature. Figures 2 and 3 illustrate the ESI+ mode LC–MS and DAD chromatograms at 100 °C and 300 °C, respectively. (See upper right for Figures 2 and 3, click to enlarge.)

In ESI- mode, there is an inverse correlation between the relative abundance of THCA and drying gas temperature while the relative THC abundance is positively correlated with increasing drying gas temperature. Figures 4 and 5 illustrate the 359.2 

 ESI-extracted-ion chromatograms (EICs) for THCA and THC at 100 °C and 300 °C, respectively. (See upper right for Figures 4 and 5, click to enlarge.) Table III demonstrates the absolute area counts and relative ratios for the loss of THCA and the increase of THC as the temperature of the drying gas is increased. (See upper right for Table III, click to enlarge.)


	The FIA temperature program was bracketed into three regions:
	1. Low temperature range: 0.0 min to 3.5 min, 100–150 °C
	2. Mid temperature range: 3.5 min to 7.5 min, 150–250 °C
	3. High temperature range: 7.5 min to 11.5 min, 250–350 °C

FIA high resolution accurate mass EIC chromatograms specific for THCA and THC were extracted within each region. In this study, the THCA (M+H)

 pseudomolecular ion and (M+Na)+ adduct were observed with exact masses of 359.2217 

, respectively. THC was observed as the (M+H)

. The mass accuracy, molecular formula generation (MFG) from the empirical spectra, and the isotopic fidelity were measured for each chemical species. These data are shown in Figures 6 and 7 for the mid-temperature range of 150–250 °C. (See upper right for Figure 6 and Figure 7, click to enlarge.)

Table IV illustrates the relative HRAM EIC ion abundance for each chemical species representing THCA and THC and their respective ion ratios. In FIA ESI+, the decreasing THC/THCA ion ratios reveal the fact that little to no acid phytocannabinoid decarboxylation is occurring under these conditions. This suggests the ESI+ pseudomolecular ion of THCA is relatively stable and does not undergo significant decarboxylation over the temperature range of 100 °C through 350 °C. (See upper right for Table IV, click to enlarge.)


	In ESI negative mode, THCA is observed at the proper retention time and THC is observed under the THCA peak at all drying gas temperatures. The relative abundance of the acid is reduced with commensurate increase in the relative abundance of THC as the drying gas temperature is increased from 75 °C through 250 °C with complete loss of the acid compounds at 200 °C and 250 °C for CBDA and THCA, respectively. Figure 8 is the series of HRAM EICs representing CBDA and CBA over the drying gas temperature range of 75°C through 250°C. Figure 9 is the series of HRAM EICs representing THCA and THC over the drying gas temperature range of 75 °C through 250 °C. In both cases, the relative abundance of the acid phytocannabinoids is reduced while the relative abundance of the decarboxylated analogues CBD and THC are increased as the drying gas temperature is increased. This is evidence of in-source decarboxylation of the acid phytocannabinoid. (See upper right for Figures 8 and 9, click to enlarge.)

In this study, we used TOF, QTOF, and single quadrupole LC–MS systems in ESI+ and ESI- modes to evaluate the effect of temperature in the source region on the stability of acid phytocannabinoids. We initially used a chromatographic approach with a single quadrupole LC–MS coupled with a diode array detector. Under these conditions, we putatively determined that no decarboxylation of the acid phytocannabinoids THCA and CBDA occurs over the drying gas temperature range of 100 °C through 300 °C in ESI+ mode. However, under the same chromatographic conditions and temperature range, the in-source loss of the relative abundance of THCA with commensurate increase of the relative abundance of THC was observed. The increase in the percent area ratio of THC/THCA taken from Table III is exponential.

We next used FIA on a LC-QTOF system in ESI+ mode to remove any effects of chromatography and confirm the absence of THCA decarboxylation under these conditions. Lastly, using a LC-TOF system with chromatography in ESI- mode we injected a mixture of CBDA and THCA over the temperature range 75 °C through 250 °C. Through this experiment, we confirmed post-chromatographic, in-source decarboxylation of the acid cannabinoids as evidenced by the observation of CBD and THC under the CBDA and THCA chromatographic peaks. Under these conditions, complete loss of the acid cannabinoids is observed at 200 °C and 250 °C for CBDA and THCA, respectively. This observation agrees with data from reference 3.

Given these observations, we propose the (M+H)

 adducts of acid phytocannabinoids are stable in ESI+ mode and the energetics of decarboxylation are not favorable under these conditions. Figure 10 illustrates the putative (M+H)

 pseudomolecular ion for THC. Conversely in ESI- mode, the (M-H)- species readily decarboxylates as a function of drying gas temperature in the source through the mechanism shown in Figure 11. (See upper right for Figures 10 and 11, click to enlarge.)

With chromatography, the mass spectral detection of CBD or THC under the chromatographic CBDA and THCA peaks, respectively, confirms post-chromatographic decarboxylation in the source region. When using ESI+ with chromatographic or FIA conditions, decarboxylation of the acids are not observed. However, ESI- mass spectra data reveal the acid phytocannabinoid decarboxylation exponentially increases as the drying gas temperature of the ESI source is increased. We therefore reject the null hypothesis for ESI- LC–MS analyses and posit acid phytocannabinoids fully decarboxylate in ESI- mode with source temperatures ≥ 200 °C. These data highlight a potential concern for the quantitation of acid phytocannabinoids using ESI- mode LC–MS technologies.

Agilent products and solutions are intended to be used for cannabis quality control and safety testing in laboratories where such use is permitted under state or country law.

Peter J.W. Stone, Sue D’Antonio, Nikolas C. Lau, and Wendi A. Hale

 are with Agilent Technologies in Santa Clara, California. 

 is with Agilent Technologies and The Johns Hopkins School of Medicine in Baltimore, Maryland. Direct correspondence to: 

P.J.W. Stone, S. D’Antonio, N.C. Lau, W.A. Hale, and A. Macherone, 

Articles

This study highlights a potential concern for the quantitation of acid phytocannabinoids.

The Stability of Acid Phytocannabinoids Using Electrospray Ionization LC–MS in Positive and Negative Modes

This study considers the use of ethanol as a co-solvent in a carbon dioxide extraction of cannabinoids and its effects on extraction time and overall raw yield. A control test was initially conducted to calculate the volumetric flow of liquid and supercritical CO

 and to record the amount of material collected. Additional runs were then performed with the injection of a 5% by volume stream of ethanol, first with hops and then confirmed on cannabis. Upon separation of the oil from the ethanol it was determined that the oil was extracted up to 300% faster and with roughly 2% greater overall yield.

There is much discussion around increasing solubility power of a solution with the assistance of a co-solvent, but few, if any, studies consider the increase in solubility of CO

 with the presence of ethanol as a co-solvent. Often, various solvents are mixed together to obtain a larger amount of solute before it’s saturated. However, unlike many experiments involving co-solvents, CO

 as a solvent cannot exist at room temperature and pressure. That means, to determine if the solubility of the solution has increased, a dynamic research model is needed instead of the typical static models that can be used with other solvents. An increase in the solubility of the solution is just one concern. The ability of CO

 to select different cannabinoid fractions at specific times must still be retained, proving the introduction of ethanol can improve extraction times without detracting from the flexibility offered by CO

 as a solvent. Otherwise, a straight ethanol extraction might be chosen over a carbon dioxide and ethanol co-solvent extraction. Throughout this study, two key factors were considered: increase solubility as measured by an increase in yield, and extraction selectivity as measured by color and viscosity. The oils that are lighter in color and less viscous offer a decrease in post-processing time, making them more desirable to many end users. These more volatile components will separate out sooner than the waxes, fats, and chlorophylls that contribute to a darker, thicker end product. 

This experiment was completed with a beta co-solvent module that consisted primarily of an ethanol tank and pump with some instrumentation and a check valve to ensure unidirectional flow of the ethanol into the system (see Figure 1). The ethanol tank supplied the ethanol pump with room pressure and room temperature ethanol. The pump would then increase the pressure of the ethanol from atmospheric to the operating pressure of the carbon dioxide system. A variable frequency drive would control the speed of the positive-displacement pump, and therefore the flow rate of the ethanol, to allow for a specific injection quantity. Volumetric percentages were chosen based on the standard for measuring co-solvent efficiencies. Finally, the necessary pressure valves and reliefs were added to connect this module to a CO

 extraction system. The piping and instrumentation diagram of the co-solvent test system is shown in Figure 1. A CO

 flow meter, placed in line between the separators and the pump on the closed loop CO

 extraction system, was used to monitor the flow of CO2. This returned a standard cubic feet per minute (SCFM) for the CO

, used to calculate a L/min flow rate based on liquid or supercritical CO

 at either 1200 PSI and 75 °F or 1800 PSI and 105 °F. The closed loop system allowed for the conversion of SCFM (mass flow) to L/min (volumetric). (See upper right for Figure 1, click to enlarge.)

The connection point between the co-solvent module and the CO

 extraction system was located just before a temperature control heat exchanger (red arrow in), which is located immediately before the inlet tubing for the extractor, where the raw material is located. (See upper right for Figure 2, click to enlarge.) This structure would allow the ethanol to mix with the CO

 stream, thus avoiding a temperature drop and ensuring the mixture was homogenized before going into the extractor. The CO

 and co-solvent mixture would then travel through the raw material and collect the oil before traveling to the separator. There the saturated solution would be depressurized, allowing the mixture of oil and ethanol to fall out of the solution and allowing the gaseous CO

 to return to be recompressed by the diaphragm compressor. 

This experiment was performed on both hops and cannabis, at both subcritical and supercritical parameters. For the subcritical test, parameters of 1200 PSI and 75 °F were used. For the supercritical test, parameters of 1800 PSI and 105 °F were selected.

This was done to match the standard operating parameters and capabilities of the CO

 system. Any subcritical and supercritical parameters could have been used. Due to the increased pressure requirements and the associated losses in volumetric efficiency in the CO

 compressor, the main difference between the subcritical and supercritical runs was the CO

 flow rate. In turn, the ethanol flow rates differed. But since both subcritical and supercritical were compared to their control tests, and not to each other, this did not factor into the findings. The original experiment at our facility in Ohio was done using hops because of regulations in Ohio prohibiting testing with cannabis. The hops used were Columbus hops. In each test, hop pellets were ground, their input weights recorded, and then loaded into the extractor to be used as the raw material. A control test was done on both sets of data points before the co-solvent test was run. During this test, data samples were taken every 10 min and weights were recorded to get a yield curve in grams of oil versus time for a CO

 only extraction. The sample time started only after the system was up to pressure and flow had begun between the extractor and the separator. A representative portion of every sample was kept to evaluate the control extraction relative to the co-solvent extraction in terms of both color and consistency. After the control tests were completed, the same tests were replicated, with the only difference being the addition of a 5% by volumetric flow ethanol stream. This ethanol stream was started only after the system was up to pressure and flow had begun. Samples were again taken every 10 min, with each sample containing both botanical oil and ethanol. The samples were weighed and kept apart from one another until separation of the oil from the ethanol. Each individual ethanol and oil sample was then put through a rotary evaporator to separate the ethanol from the extracted oil. Both the weight of the oil and the volume of the ethanol were recorded. The ethanol volume was recorded to verify the flow rate of the ethanol pump since there was no ethanol flow meter present during the test runs. In the absence of an ethanol flow meter for this experiment, measurements of ethanol used were taken throughout the process to verify that the amount used was consistent with the pump motor speed. Once the control and experimental hops tests were completed, subsequent control and experimental tests were run in Pueblo, Colorado to verify the results on cannabis. The cannabis strain used was a high tetrahydrocannabinol (THC) strain that had been decarboxylated prior to the extraction test. Test cannabis had also been ground to approximately 100 µm for the test runs. The CO

 extraction system used in conjunction with the cannabis test had a higher CO

 flow rate than the one used with the hops tests, but the ethanol flow rate was increased to keep the volumetric percentages the same.

The hops test matrix can be seen in Tables I and II. (See upper right for Tables I and II, click to enlarge.)

Sample 11 on runs 1 and 2 were both taken after an additional hour of running to determine the maximum amount of oil that could be obtained. The benefits of the co-solvent were delayed because the ethanol pump did not start until the system was up to pressure and CO

 flow had started. This was due to the time it took for the ethanol to travel through the heat exchanger and into the raw material, before exiting the raw material to the separator. This travel time was determined by the CO

 flow rate and the length of tubing through which the ethanol would travel before mixing in solution with the carbon dioxide and botanical material. This delay inspired an option in later versions to allow a terpene extraction to occur before the co-solvent portion of the extraction takes place. Many end users perform a CO

-only fractional terpene extraction before the cannabinoid extraction, so this method is representative of one common practice. The delay in co-solvent assistance is evident in the supercritical run as the flow rate was much slower than the subcritical run. This can be noted in the amount of ethanol in the run as well. Comparing the subcritical runs and supercritical runs independently, we can see there is an increase in speed and, in the case of the subcritical run, an increase in the maximum amount of material extracted. A graphical representation of these runs can be seen in Figures 3 and 4. (See upper right for Figures 3 and 4, click to enlarge.)

During the testing process it was clear that additional benefits include both the ease in collecting and cleaning the separator when using a co-solvent. The resulting oil–ethanol mixture was nearly homogenized and could be easily transitioned into a winterization process without the necessary hotplate and stirrer. This benefit manifests as a reduction in energy consumption and time required for post-extraction processing activities. After these tests, it was determined that the presence of ethanol increases the solubility power of the CO

 is still selectable even with the addition of the co-solvent. This was clear from the results of the supercritical test, as seen in the slower speed shown in Figure 5. (See upper right for Figure 5, click to enlarge.)

The cannabis test matrix can be seen in Table III and IV. (See upper right for Tables III and IV, click to enlarge.)

With the cannabis test runs, we were able to increase the flow rates of the CO

 to achieve a better representation of flow rates currently available on the market. We were also able to increase the raw material bed to allow for a longer extraction curve for the increased flow. Extraction curves for the above data can be seen in Figures 6 and 7. (See upper right for Figures 6 and 7, click to enlarge.)

During these runs, the delay in starting the ethanol pump was reduced to achieve a better overall increased extraction curve. In this test, the co-solvent pump was started as soon as liquid or supercritical CO

 was flowing in the lines. This was determined by the pressure approaching 1000 PSI for each run. Midway through the fifth pull, the ethanol pump was stopped to replicate previous experiments by extracting all the oils around this time. The extraction curve for both super and subcritical CO

 were similar. The extraction on the subcritical was roughly 175% faster, and extraction on the supercritical was 300% faster. Furthermore, the mixture darkened in the subsequent samples and held a similar pattern to that seen in hops.

While more testing is needed to consider additional factors about ethanol co-solvent runs, these tests proved the effectiveness of the method at these parameters. An injection of a 5% co-solvent stream can increase efficiency by more than 100%, while still maintaining some selectivity and adding the benefit of cutting out homogenization from the post-processing winterization process. Ethanol being a polar solvent has its pros and cons, but using it in solution with CO

 helps to reduce the removal of the unwanted chlorophyll. With the addition of ethanol to the solution, dialing into a set of extraction parameters to extract desirable cannabinoids and terpenes while leaving the unwanted compounds in the raw material can be accomplished not only with pressure and temperature adjustments of the CO

, but can now also be adjusted by raising or lowering the concentration of ethanol in the stream. This method of co-solvent injection is also beneficial in that all the ethanol pumped into the CO

 system is retained with no loss. This is because of the saturation into the plant material, as is the case with typical ethanol extractions, which can leave 10% or more of the ethanol behind in the plant material. This occurs because ethanol is soluble in liquid of supercritical CO

, allowing for natural movement of any remaining ethanol from the extractor to the separator during recovery.

 is a senior mechanical engineer with Apeks Supercritical, a leading CO

 extraction system manufacturer. Sam has been with the company for almost six years, after graduating from The Ohio State University, where he got a BSME. He leads the team innovating the co-solvent technology at Apeks and has spearheaded several other innovative technologies over the years. Direct correspondence to: SamS@apekssupercritical.com.

This study considers the use of ethanol as a co-solvent in a carbon dioxide extraction of cannabinoids and its effects on extraction time and overall raw yield.

Co-Solvent Ethanol Injection Helping CO2 Extraction

Cannabis is known for being a complex plant with thousands of chemical compounds. The most economically important compound for many years was the cannabinoids, particularly tetrahydrocannabinol (THC). In the past decade we have begun to understand the importance of other groups of natural cannabis compounds such as terpenes and now flavonoids. Flavonoids are naturally occurring secondary metabolic products that can have important functions within the cannabis plant and benefit consumers by their health and healing properties. In this column, we once again go beyond cannabinoids and look at the importance and chemistry of flavonoids in cannabis.

There are thousands of compounds in botanicals. Cannabis is rich in many vitamins, nutrients, and beneficial compounds. Historically, cannabis has been harvested for cannabinoids, but over the past decade there has been increased research and interest in the other active and beneficial compounds such as fatty acids, terpenes, antioxidants, and flavonoids. Many of these beneficial compounds are metabolite compounds produced as an end product of metabolism. Metabolites are small molecules that have many functions including defense, pigments, pheromones, odorants, and catalysts for many biological processes. 

Primary metabolites are involved in growth, development, and reproduction. If a primary metabolite is missing or disrupted, the organism can degrade or die. Examples of primary metabolites are amino acids, antioxidants, and vitamins. Secondary metabolites are not directly involved in critical processes, but have secondary functions. An absence of a secondary metabolite does not necessarily mean the organism’s death. Examples of these secondary metabolites include compounds such as terpenes and flavonoids. 

Flavonoids are secondary plant, algae, or fungus metabolites composed of polyphenolic compounds.  Phenyl groups are cyclic molecules (C

) closely related to benzene (Figure 1). Flavonoids generally have a 15-carbon structure with two phenyl rings and one heterocyclic pyran ring (Figure 2).

In 1664, Robert Boyle first described flavonoid experiments with plants and colors in his book 

Experiments and Considerations Touching Colours

 (1,2). The first flavonoid was identified by a Hungarian biochemist and Nobel Prize winner, Albert Szent-GyÃ¶rgyi in 1930. The name for flavonoids has it root in the Latin 

 meaning yellow and describes the function as pigments in plants. Flavonoid pigments are found to produce several ranges of colors including yellows, blues, and reds with all the mix of colors in between. Color is perceived by optical system as wavelengths of light absorbed and reflected by a substance. Certain chemical compounds such as flavonoids and elements can reflect or transmit energy in visible light bands, which then are seen as particular colors (see Table I). Visible light is in the range of ~400–700 nm. It is within this range of light that humans perceive color. Outside that range of light we find other types of light and energy such as X-ray, ultraviolet (UV), and infrared (Figure 3).

Chemicals that allow for the absorption and reflection of light are said to have chromophores. In these molecules the energy difference between molecular orbitals is in the energy range of visible light. Visible light hits the electrons in the orbital and excites them moving the electron from a ground state to an excited state. This excitation produces a change in the shape of a molecule called a conformational change. 

 are specific structural forms or interactions that allow for visible light to be transmitted. One of the most common chromophores is a conjugated Ï-bond. Pi-bonds are double bonds within the p-orbital of a molecule. Conjugated Ï-bonds occur when three or more p-orbitals within a molecule form alternating double and single bonds within a molecule as seen in the alternating double and single bonds within the flavonoid phenyl rings (Figure 1).

Additional structures within a molecule can also be important in the perception of color by altering or increasing the transmission of visible light wavelengths. An

 is a functional group in the molecule; attached to the chromophore. These functional groups (such as alcohols, carboxylic acids, and nitrogen containing compounds) change the ability of the chromophore to absorb light. This change can alter the wavelength or intensity of the absorption and therefore change the visible color observed. It is the ability to absorb and reflect light that allows flavonoids to function to capture wavelengths of light, protect plants from UV radiation, attract pollinators, and defend against predators (3).

These phytochemical properties make flavonoids powerful phytonutrients and antioxidants, which make them in demand for human nutrition and health. Antioxidants have been a buzz-word in human health for several decades. These compounds inhibit oxidation, which is the loss of electrons that creates free radicals with unpaired valence electrons. These radical ions, atoms, or molecules are highly reactive and are important in many biological processes, but due to their highly reactive  nature they can also cause unplanned changes or damage to other molecules, cells, and tissues causing disruptions to health and disease. Often these radicals can cause chain reactions that damage cells. Antioxidants can stop these chain reactions. Antioxidants can be compounds that are intentionally produced to preserve products and foods from oxidation such as preservatives and antirust compounds or antioxidants can be naturally occurring compounds that inhibit oxidative free radicals from undergoing chain reactions and causing cell damage and disease. Studies have shown that flavonoids can produce positive results in combatting heart disease, diabetes, and some cancers (4–6).

Flavonoids can be categorized into three general classes by backbone structure: bioflavonoids, isoflavonoids, and neoflavanoids. 

 (or flavonoids) are designated as compounds where the phenyl ring is attached to the C2 carbon of the benzopyran ring. 

 have their attachment to the C3 carbon and 

 have the attachment of the phenyl ring at the C4 carbon of the benzopyran ring (Figure 4).

There are dozens of subgroups for all of the structural flavonoid groups. Table II shows important subgroups and functional groups of flavonoids. These groups produce the antioxidant effects and color pigmentations in many plants. Reds, blues, and purples are produced by anthocyanins. Yellows are produced by many flavonoids including anthoxanthins, chalcones, and aurones. Proanthocyanids produce black, brown, tan, and red pigments. (See upper right for Table II, click to enlarge.)

Cannabis is rich in many beneficial compounds including flavonoids. It is estimated that up to 10% of the cannabis plant compounds are flavonoids with up to almost 3% by weight in dried leaves and buds. The list of flavonoids in cannabis depends on which strains are being examined and the color of the plants. There are a dozen basic cannabis flavonoids found to some degree in the majority of the strains. Cannabis has also been found to have its own flavonoids exclusive to cannabis species (cannaflavins A, B, and C). Cannaflavins were first discovered by a researcher in London in 1986. Marilyn Barrett at the University of London identified cannaflavin A and cannaflavin B (Figure 5).

The majority of cannabis flavonoids are either flavones or flavonols, which are anthoxanthins. These compounds are primary responsible for yellow colors (Table III). 

Cannabis is known for variations in colors and patterns.  As plants grow they are primarily green and yellow. As the plant matures, many color variations can occur in the leaves and buds. The primary color of cannabis is green from the presence of chlorophyll. During the changes in temperature, however, such as during changes of season the production of chlorophyll stops and other colors can present as other compounds masked by chlorophyll become apparent. 

Some of the more spectacular colors of cannabis are blues, purples, reds, and pinks due to flavonoids known as 

. These compounds are often dependent upon pH. These compounds are antioxidants and give the red and blue hues of grapes, berries, and cannabis especially strains such as Purple Haze, Blueberry, and Blue Dream. Red and pink colors tend to be rare, but include strains like Pink Panther. Yellow and oranges can be seen in strains with high carotenoids (terpenoids) or flavonoids such as Lemon Kush and Orange Bud.

Flavonoids in cannabis are antioxidants, colorants, and plant protectants. These compounds are not psychoactive but can play a role in suggested entourage effects where the flavonoids present in cannabis act in a synergistic action with other compounds such as the cannabinoids and terpenes to produce antioxidant, antidepressant, anti-inflammatory, and disease fighting properties. Just as in the case of the importance of terpenes in cannabis, the importance of flavonoids will need a lot more investigation and testing to truly elucidate the beneficial effects of these compounds for health.

“The Project Gutenberg EBook of Experiments and Considerations Touching Colours, by Robert Boyle.” n.d. Accessed March 23, 2020. 

http://www.gutenberg.org/files/14504/14504-h/14504-h.htm

M. Sandu, L.M. BÃ®rsa, and L.G. Bahrin,  

F. Ferreyra, M.L., S.P. Rius, and P. Casati, 

V. Ponzo, I. Goitre, M. Fadda, R. Gambino, A. De Francesco, L. Soldati, L. Gentile, P. Magistroni, M. Cassader, and S. Bo, 

https://doi.org/10.1186/s12967-015-0573-2

https://doi.org/10.1097/MD.0000000000010686

M.K. Chahar, N. Sharma, M.P. Dobhal, and Y.C. Joshi, 

F. Ververidis, E. Trantas, C. Douglas, G. Vollmer, G. Kretzschmar, and N. Panopoulos,  

A.J. Herrington, n.d. “What Do The Colors of Marijuana Mean?” 

https://hightimes.com/guides/colors-marijuana-mean/

is a Senior Applications Scientist with SPEX CertiPrep and a member of both the AOAC and ASTM committees for cannabis.

We once again go beyond cannabi¬noids and look at the importance and chemistry of flavonoids in cannabis.

Beyond Potency: Flavonoids-the Purples, Reds, and Blues 

The selection and use of the right growth substrate for your cultivation is essential for healthy high-producing cannabis plants. There is no “one-size fits-all” when it comes to deciding on the growth substrate that will support and provide the nutritional environment across the life cycle of your plants. In this column, we explore some options for growth substrates that are essential to the health of your plants.

We often are asked the question, “What is the best growth substrate for my cannabis?” We wish there was a simple, single answer, but the answer is always, “It depends.” It is important to start to think of cannabis cultivation as a horticultural system that can be optimized and not simply an agricultural one-size-fits-all crop. When deciding on the growth substrate, also called the 

, you will need to consider all of the variables that will affect the taste, yield, uniformity, and quality, “organic properties,” whether you are growing indoors or will cultivate sun-grown plants, and the potential benefits of the complex soil food web versus possible plant pathogens that could damage your buds, especially if you cultivate outdoors.

This column focuses on hydroponic substrates to be used in controlled environment agriculture (CEA) settings, such as greenhouses and warehouses. If you are cultivating outdoors, we assume you will be planting in soil and we will discuss this option only briefly in this column.

Let’s take a moment to discuss the differences between soil and a hydroponic substrate. In general, Earth’s soil is rich with organic material, such as bacteria, fungi, insects, and decaying leaves that provide part of the complex nutrient system that will feed the plants. We might think of it as a “living soil” because it is filled with microscopic life that assists with nutrient development and absorption. In contrast, hydroponic substrates are a set of “root-supporting” granular or fibrous materials that have little or no components able to provide nutrients to the plants. All nutrients for hydroponic systems must come from the water-based nutrient solutions that you provide through fertigation. Hydroponic growth substrate materials are best considered as an architectural support structure that ensure the roots have a strong and solid base to support the heavy plant structures as shown in Figure 1, such as the stems, leaves, and flowers. The hydroponic growth substrate is porous, meaning that it is filled with tiny holes and complex pathways for the flow of water and nutrients around the growing root system. The solids in a hydroponic growth substrate are inert support materials, while the hollow pore space allows room for the roots to grow and fertigation liquids to flow. The pore space allows for the flow of water, nutrients, and air (oxygen) to the roots for immediate absorption into the plant.

Hydroponic substrates perform several functions to enable plant growth. First, and most obvious, is that the substrates provide an anchor that enables the cannabis plant to grow and remain upright. It does so by providing a complex three-dimensional (3D) matrix for the root system to infiltrate the pore space and support the plant. However, this is just the beginning of the benefits of the hydroponic substrate: it provides a habitat for the overwhelming number of beneficial and benign microorganisms to live. This high concentration and close proximity make them readily available to colonize the growing roots and microscopic root hairs. Colonizing roots with beneficial and benign organisms limits or prevents the attachment of harmful pathogenic fungi and bacteria.

Hydroponic substrates, such as coco coir, have the capacity to bind  nutrients. This is particularly true for micronutrients such as calcium and magnesium. Regardless of the substrate choice, the surface of the substrate itself will become colonized with microbes that create an enhanced complex surface structure, called a 

, that can assist with nutrient absorption. This biofilm develops with time and becomes a local source of micronutrients as the microbes increase in number and build the more complex surface structure.

Finally, the hydroponic substrate ensures the maintenance of a proper balance between air (oxygen) and the nutrient solution for the root system. The selection of the appropriate substrate will guarantee that these essential components of plant health are available at the right time and the right place to optimize plant growth and maintenance from seedling to harvest.

Your choice of hydroponic substrate depends on the type of cultivation facilities you are growing in and the goals of your cultivation.

Coco coir seems improbable since it is made from the rough fibrous husk of coconuts, but it is an excellent organic product as a hydroponic substrate (Figure 3). It has been used for thousands of years as a strong material in a wide variety of products from door mats to plant growing medium. It is highly inert and holds water very well. It is a sustainable resource that provides an Earth-friendly approach to your operations.

Rockwool is made from the mineral basalt and contains no organic material (Figure 4). It is a sterile, porous, easy-to-work-with lightweight hydroponic substrate. It is not yet recyclable, but can be repurposed into an outdoor soil amendment. The key factor for selection of rockwool is its sterility, cleanliness, and ability to maintain a proper water to air mixture at the root level with the nutrient solution.

There are four key parameters that affect the performance of the hydroponic substrate. Comparing and contrasting the performance of these parameters in your choice of substrate will ensure that you can make a logical and fully-vetted decision on which to use. The key parameters are:

We prefer top drip irrigation with frequent “watering cycles.” Infrequent heavy watering creates runoff flow channels through the substrate. The result is fewer roots at the top of the substrate matrix and lowered nutrient uptake for the plants. Some runoff, about 20%, is needed to flush salt build-up for a drain to waste system.

The best pH for a hydroponic system ranges between 5.5 and 6.5, slightly acidic. pH is important to maintain optimal nutrient availability for the plant, but is not as important as most people think. If you are running a drain to waste drip irrigation system where nutrients are being replenished with each irrigation cycle, it is much less important. If your system is a closed-loop recycled system more care must be taken to maintain proper pH.

Because of the minimal ability of rockwool to retain nutrients, the hydroponic solution must be supplied at a higher level and be more nutrient rich than for coco coir. The coco coir itself provides some slow-release nutrient value by virtue of its organic composition, but it is minimal.

Rockwool can retain 90–94% of its volume with water. While coco coir retention varies with the source; it is typically in the mid-60%. Both are satisfactory in this key parameter. Irrigation should be short and frequent to enable full saturation of the matrix and drying to properly occur in both substrates.

For those of you interested in producing an “organic” product the choice is clear: select coco coir as your hydroponic substrate. Once it is properly prepared, it provides an easy-to-work-with medium that fosters a strong growth environment to support the plant and for bio-film formation for the root-based beneficial microbes supporting nutrient absorption. However, be aware that the system is messy to work with and brings with it additional labor costs for preparation and clean-up. It is also not sterile because that is not part of the vendors’ process; I have seen fungal infections that were ultimately attributed to the coco coir source. So, coco coir provides an excellent hydroponic substrate, but you must be willing to work within the constraints of preparation and possible fungal infections. With diligence and care, it can be your go-to substrate in your greenhouse or warehouse.

In addition, we recommend rockwool for hydroponic use. Rockwool is an inorganic porous material formed at extremely high temperatures through a manufacturing process. On the plus side, it arrives at your cultivation site as a sterile product. It will not introduce pathogens to your facility. If your focus is on minimizing contamination, it is the clear choice. It’s easy to start clean and stay clean with rockwool. The use of inorganic mineral salt fertilizers makes the system more suitable for short and repeated irrigation as it reduces likelihood of plugging and clogging of fertigation lines. This means it is much more compatible for automated irrigation systems because there is reduced contact between workers and the plants, thus lowering the opportunities for the transfer of plant and human pathogens. If your top priority is to start clean and stay clean, we recommend implementing a rockwool substrate across your cultivation facility. You will thank yourself with healthy plants over their full life cycle.

When choosing the right hydroponic substrate for your cultivation site, you need to consider your overall goals and the system that you want to optimize for taste, yield, quality, and uniformity in your final products. If your priority is to grow organically, you would select coco coir. On the other hand, if your priority is to grow to minimize plant diseases, you would select rockwool. Regardless of the substrate you choose, you will still have to manage plant pathogens and pests. In all cases, growing and producing healthful and desirable buds is a hard job, but well worth the extra effort when you bring a measurable and scientific approach to cultivation. We have seen both of these particular substrates produce healthy plants and a wonderful final product.

 is a project manager, systems engineer, and scientist focused on today’s commercial cannabis enterprise.  She is the CEO of Agate Biosciences LLC, an innovation consultancy. With her PhD from The University of Texas at Austin, she focuses on investment assessments, due diligence, and project management. She spent 27 years at NASA’s Jet Propulsion Laboratory as a scientist and project manager, leading advanced technology development projects and proposal development in the complex government-regulated environment. Now, as the Chief Executive Officer of Agate Biosciences, she brings the best practices of detailed research, project management, and systems engineering to angel investing, venture capital, and private equity clients.

 is a scientist and technologist who cares deeply about the cultivation and health of plants in the cannabis industry. With his PhD in microbiology from the University of California, Davis, Plant Growth Laboratory, he solves the most challenging problems in hydroponics, from studying the root microbiome to developing nutrients and lighting systems to ensure plant health and a disease-free life cycle. He spent 22 years at NASA’s Jet Propulsion Laboratory as a scientist, technologist, and research leader before becoming the President of Agate Biosciences, a consulting firm for project management, systems engineering, and science in CEA for the past eight years. He leads developments to optimize sustainability, consistency, quality, and yield without compromising plant health. Direct correspondence to: 

Let’s explore some options for growth substrates that are essential to the health of your plants.

The Root of Success:  Using the Right Growth Substrate

In this installment of “Cannabis Crossroads” we examine more than 158,000 peer-reviewed cannabis science articles and report the top authors, most prevalent terms, and emerging trends. Cannabis science is growing rapidly and we used various tools to mine this data to report interesting trends.

What does one do while social distancing during a COVID-19 global pandemic? In this installment of “Cannabis Crossroads” I took on the challenge of exploring more than 158,000 peer-reviewed cannabis articles to determine interesting trends, top authors, and observations. Cannabis science is growing rapidly and we used a powerful analytics platform to take a deep dive into cannabis publications from 1812 to 2020.

In the fast-evolving world of cannabis science, it is important to keep informed of new areas of growth and trends in scientific articles. There are a wide variety of tools available for mining peer-reviewed publications. Simple keyword searches can be performed using Google Scholar. To analyze keyword trends, top authors and explore connections between publication datasets, a platform called Dimensions (Dimensions.ai) is extremely powerful yet simple to use. In addition to publications, Dimensions also brings together grants, datasets, citations, alternative metrics, clinical trials, patents, and policy documents. I used the Dimensions platform to review some interesting cannabis science article trends, authors, and citation information with you. 

Exploration of the keyword “cannabis” in Dimensions generates 158,835 total articles since 1812. The large growth from 1971 to 2019 is shown in Figure 1. (Note: If you include book chapters, proceedings, preprints, and monographs the total number of publications containing “cannabis” increases to 222,896.)

Interestingly, only a small portion of these articles are published in the fields of plant biology (3598 articles) and chemical sciences (5577 articles). A large portion of these articles fall under medical and health sciences (99,932 articles), as shown in Figure 2.

If we look at all fields across all years, there were 17,741 articles that contain the keywords “cannabis” and “chromatography,” and 14,297 articles with the keywords “cannabis” and “mass spectrometry.” Interestingly, there were very large spikes in both datasets in 2010 and 2018. In fact, publications in 2018 outpaced those in 2019 (see Figure 3).

The top authors publishing on “cannabis” and “mass spectrometry” are shown in Table I. Marilyn Ann Huestis at Thomas Jefferson University, and formerly with Chemistry and Drug Metabolism at The National Institute on Drug Abuse (NIDA), National Institutes of Health (NIH), has the largest number of publications containing these two keywords (as well as “cannabis” and “chromatography”).

Dr. Raphael Mechoulam, Hebrew University of Jerusalem, Israel, who isolated numerous cannabinoids, including 

-tetrahydrocannabinol (THC), and elucidated its structure was the ninth ranked author for “cannabis” and “chromatography.” (Note: If we look at just the keyword “cannabis,” Dr, Mechoulam has 229 articles with more than 21,100 citations.)

Top author honors of all things cannabis, however, go to Robin Macgregor Murray of Kings College London, United Kingdom, with 347 articles mentioning “cannabis.” Louisa J. Degenhardt of The University of New South Wales (UNSW), Sydney, Australia placed second with 308 articles and a whopping 43,867 citations!

If we narrow the searches to the confines of the analytical chemistry field, we see far less publications containing “cannabis” (2371 articles in total), but we can also see a noticeable increase in the publication rate after 2017, as highlighted in Figure 4.

What is contributing to the growth in cannabis publications in the analytical chemistry field post-2017? Pesticides became a very hot topic and witnessed a surge in publications in 2018 (1367 publications in all fields, with 71 specifically in analytical chemistry). In addition to contaminant testing, many more recent searches show topics of cannabinoid and terpene identifications.

These analytics show not only growth in cannabis science, but also an evolution. A large portion of older cannabis publications are aimed at studies of addiction, drugs of abuse (DAU), psychology, public health, forensics, and health sciences, whereas the more recent topics appear to be focusing on topics such as quality control, exploration of potential therapeutic value, “whole plant” medicine, nutraceuticals, food safety, genetics, and clinical trials. Regardless, such analytics can help us guide our future endeavors in academia, government organizations, R&D, and more. They can also help us find and connect with other authors publishing on similar topics of interest. Knowledge is power, and these analytics can be used to quickly assess current trends and explore new cannabis science boundaries.

I have shared just a few trends and interesting observations here, and I strongly encourage you to explore your own keyword searches using the Dimensions platform. Perhaps you are more interested in keywords such as “cannabis” and “cancer” or exploring beyond publications into the worlds of the 1490 cannabis-specific grants, 24,122 cannabis patents, and 912 cannabis clinical trials listed from a host of international clinical trial registries.

As Albert Einstein said, “Any fool can know. The point is to understand.” I sincerely hope that your own, customized keyword searches and analytics will become an important, guiding step in your journey towards understanding your particular cannabis science fields and that you find the Dimensions platform to be useful. Be safe and be vigilant.

About the Dimensions Data Exploration Platform

Dimensions is a linked research knowledge system that reimagines discovery and access to research. Developed by Digital Science in collaboration with more than 100 leading research organizations around the world, Dimensions brings together grants, datasets, publications, citations, alternative metrics, clinical trials, patents, and policy documents to deliver a platform that enables users to find and access the most relevant information faster, analyze the academic and broader outcomes of research, and gather insights to inform future strategy. For more information about the product have a look at their support site or visit 

Joshua Crossney is the columnist and editor of “Cannabis Crossroads” and a contributing editor to Cannabis Science and Technology magazine. Crossney is also the president and CEO of CSC Events. Direct correspondence to: 

The print version of this article contained the incorrect versions of Figures 2 and 4. The correct figures appear here.

A report on the top authors, most prevalent terms, and emerging trends in peer-reviewed cannabis science articles.

Cannabis Science Analytics:  A Look at Top Authors, Keywords and Trends

The insanity of different states using different sampling schemes, analytical methods, different analytes, and different concentrations to regulate hemp growers has been discussed here before. With the promulgation of the U.S. Department of Agriculture’s Interim Final Rule in October 2019, some clarity has been brought to the problem. However, there are still problems with representative sampling, composite samples, and insuring there will be enough testing capacity to meet the demand for testing come harvest season. Here, some thoughts on how to solve this problem are shared.

In a recent column (1), I outlined the problem of different states using different sampling schemes, analytical methods, different analytes, and different concentrations to regulate hemp growers. With the promulgation of the United States Department of Agriculture’s (USDA) Interim Final Rule in October 2019 (2), some clarity has been brought to the problem. However, there are still problems with representative sampling, method differences, and insuring there will be enough testing capacity to meet the demand for testing come this harvest season. Some thoughts on how to solve this problem are discussed below.

Why Is Getting Hemp Testing Right so Important?

The answer: Because cannabidiol (CBD) is medicine and it should be tested like medicine. Hemp is an unusual crop because if it is being grown for CBD, rather than for its traditional uses as food and fiber, farmers are growing medicine. This is in part why the industry is so highly regulated. The absolute limit of 0.3% total tetrahydrocannabinol (THC) means that a farmer’s ability to sell their crop and make a living is affected by changes in total THC content of a few hundredths of a percent. I know of no other crop where such high stakes testing is required by law. Some farmers have literally bet the farm on hemp, weren’t able to sell their crops, and committed suicide as a result (3). Since Federal legalization of hemp (4) thousands of farmers across the US have taken the risk and invested heavily into planting hemp crops. We owe it to them, the backbone of our industry, to give them the best testing data and the fairest chance of being able to sell their crop.

Effects of USDA Interim Final Rule for Hemp Testing

My first article on hemp testing insanity was published in the October 2019 edition of 

 (1). In it I lamented the lack of guidance on hemp testing and how the variation in methodologies across states and third party laboratories produces inconsistencies in test results that are not fair to hemp farmers. As if they heard my call, within days of my article going to press the USDA promulgated an Interim Final Rule (IFR) for the regulation of the hemp industry nationwide (2). As far as hemp testing goes, the interim final rule made the five points below clear.

The Cannabinoid Being Regulated Is Total THC

Total THC is determined using this equation

and the amount of total THC is not allowed to exceed 0.3% by dry weight. This is useful since, as I have pointed out (1), some states were regulating on total THC, others on THC, and others some combination of the two.

Margin of Error Has Been Taken into Account

The IFR defines an “acceptable hemp THC level” by adding the margin of error to 0.3%. For example, if for a given analyzer the error for the determination of total THC in dried hemp is ±0.04 weight percent, the acceptable hemp THC level for samples measured on that analyzer would be 0.34 weight percent total THC or less. This is welcome since all measurements contain error (5) so this must be taken into account when determining hemp legality. Previously, some states were not taking margin of error into account and were insisting on a strict 0.3% total THC limit, whereas others were allowing upwards of 0.1% error and passing samples that were upwards of 0.4% total THC.

A Representative Sample Must Be Collected

The suggestion is to take one cutting per acre, and that cutting should be from the top third of the plant. I will have more to say about this further on.

Proposal to Implement a Nationwide Hemp Testing Laboratory Licensing Program

Please, Dear God, yes do this now please! I have been speaking, researching, and writing upon the inter-laboratory variation problem in this industry for some time (5–9). Briefly, the problem is different laboratories report significantly different results on the same samples. This is caused by human error, a lack of standard analytical methods, and the absence of appropriate standard reference materials. Inter-laboratory variation is a significant problem for our industry, and the only way to solve it is governmental oversight imposing scientifically correct practices and standard methods and samples on the industry. A federal laboratory licensing program is a step in the right direction.

Gas Chromatography, Liquid Chromatography, and Alternative Methods May Be Used for Testing

I bring this up because I have had people tell me that the USDA requires gas or liquid chromatography be used for hemp testing. This is not true as I have confirmed with in-person meetings with the science staff at the USDA. The whole point of the phrase “alternative methods” is that the USDA realizes that nonchromatographic technologies exist or may be developed to test hemp that are just as viable as gas chromatography (GC) or high performance liquid chromatography (HPLC). For example, there exist infrared spectroscopic methods for analyzing hemp that are in use (9,10). To be clear, the USDA does not test hemp, does not have a hemp testing method, and does not approve or disapprove of any given hemp testing method. If people mistakenly force themselves to only use chromatography for hemp testing, they are denying themselves the benefits of alternative technologies.

2020 Growing Season Hemp Testing Challenges

How can we solve hemp testing problems such as representative sampling, composite samples, and insuring there will be enough testing capacity to meet the demand?

April 2020

The Stability of Acid Phytocannabinoids Using Electrospray Ionization LC–MS in Positive and Negative Modes

Co-Solvent Ethanol Injection Helping CO2 Extraction

Beyond Potency: Flavonoids-the Purples, Reds, and Blues

The Root of Success: Using the Right Growth Substrate

Cannabis Science Analytics: A Look at Top Authors, Keywords and Trends

Hemp Testing Insanity II: Impending Hemp Testing Harvest Season Challenges

How to Distinguish Between Hemp and Cannabis Plants Using Raman Spectroscopy

Extraction Facility Design: Important Factors to Consider Before Selecting Extraction Equipment

Cannabis Cultivation is More Math than Science: The Role of Data in A Successful Grow Operation