At the end of the last column (which you can find online at 

www.cannabissciencetech.com/view/principles-of-chromatography-part-i-theory

), we had injected a sample at the head of a chromatography column and were waiting to see what happened next. In this second installment in our discussion on chromatography theory, we discuss via the mechanisms of equilibrium, adsorption, and de-adsorption how molecules move through a chromatography column and how separations occur. A critical variable in determining the retention time of molecules is the fraction of time they spend adsorbed to the stationary phase. Simply put, if you can arrange your chromatography experiment such that the analyte molecules spend different amounts of time adsorbed, a separation is possible.

For more background on the discussion contained here, I refer you to the references (1,2) and my recent column on chromatographic theory (3). Using a horse race analogy to describe a chromatographic separation may seem silly, but in the end, it works well. At injection time all molecules are “in the starting gate” as illustrated in 

 (note that I left the stationary phase out of the figures below for ease of viewing, but we will assume it is there and doing its job)

The purple block at the head of the column in Figure 1 is our sample immediately after injection.

Imagine the sample contains tetrahydrocannabinol (THC) dissolved in ethanol, and that the mobile phase is composed of water and acetonitrile. Thus, this example discusses liquid chromatography (LC), but the principles developed here apply to both gas chromatography (GC) and LC. Both LC and GC are capable of measuring cannabinoid concentrations in samples (4,5). The water–acetonitrile mobile phase mimics the conditions of my favorite LC cannabis potency method (5).

Upon injection some of the THC molecules will adsorb on the stationary phase, indicated in red in Figure 1, and some will stay dissolved in the mobile phase, indicated in blue in Figure 1. The purple rectangle at the head of the column represents what I call a slug of sample, but a scientifically better term is 

. The band is purple because the red stationary phase molecules and the blue mobile phase analyte molecules occupy the same length of column, and when you combine red and blue you get purple. Recall that the front edge of the band of sample is known as the 

Above the head of the column in Figure 1 you can see the designation in red of C

, which denotes the concentration of analyte adsorbed on the stationary phase, and the height of the red rectangle gives the size of C

. The designation below the column in blue, C

, is the concentration of analyte in the mobile phase. The height of the blue rectangle below the column gives the magnitude of C

. The red and blue lines to the right of the red and blue rectangles in Figure 1 show that there is no analyte in the portion of the column just beyond the solvent front.

Now, at this point in the injection we have some THC molecules adsorbed and some in solution, which occupy the same length of column. However, because of the dynamic nature of molecules this situation is not static, and at any given point in time some molecules are in the process of adsorbing on the stationary phase and others are de-adsorbing and entering the mobile phase. The term “equilibrium” at the top of Figure 1 means we are assuming that the rates of adsorption and de-adsorption are the same. We will also assume for the sake of simplicity that THC has an affinity for our stationary phase, and that at equilibrium half the molecules are adsorbed on the stationary phase and half are dissolved in the mobile phase. This is why the red and blue rectangles in Figure 1 are the same size. We will also assume that the ethanol molecules that the THC was dissolved in have no affinity for the stationary phase and spend all their time in the mobile phase. The arrow of time points to the right in Figure 1 denoting the direction of mobile phase flow.

 illustrates the situation in our chromatography column, 10 s after injection.

 is the same as before because the molecules that were adsorbed are still adsorbed and are indicated by the pink band at the head of the column. The blue rectangle denoting C

 has moved down the column because these molecules are dissolved in the mobile phase. As a result, C

 at the column head is now zero. In Figure 2 we call this situation disequilibrium because the stationary phase and mobile phase analyte molecules are no longer in equilibrium because they are in different sections of the column and no longer in contact with each other.

 represents the situation inside our chromatographic column 20 s after injection.

Note that the pink slug of analyte molecules that was at the head of the column in Figure 2 is now gone. These adsorbed molecules had fresh mobile phase above them containing no analyte molecules, and so given the opportunity de-adsorbed and dissolved in the mobile phase. Therefore, C

 has gotten bigger to the left in the column compared to Figure 2.

On the other hand, the molecules that in Figure 2 were in the mobile phase now have a piece of column where no molecules were adsorbed and have taken the opportunity to adsorb. This is why C

 is higher to the right in Figure 3 compared to Figure 2, and the net effect is that the adsorbed molecules have shifted to the right. Now molecules are adsorbing and de-adsorbing so the phrase “adsorption/de-adsorption” appears in Figure 3.

 illustrates the situation in our chromatography column 30 s after injection.

 is the same as in Figure 3 because the adsorbed molecules have stayed adsorbed, hence the red rectangles in Figures 3 and 4 are in the same place. However, the molecules that were in the mobile phase in Figure 3 have moved to the right due to the flow of the mobile phase and are now above a section of column with adsorbed molecules. Hence the blue rectangle representing C

 has moved to the right in Figure 4 compared to Figure 3. Now that we have mobile phase containing analyte molecules above stationary phase with absorbed analyte molecules, equilibrium is re-established, and we find ourselves in a situation similar to Figure 1, which is why our sample band is purple again. However, compared to Figure 1 the net change is that our band of analyte molecules has moved to the right. Thus, figures 1 through 4 illustrate the mechanism via which an analyte moves down a column.

 shows the situation in our chromatography column after 40 s.

Now, we begin the whole process over again. Molecules in the mobile phase have moved to the right in the figure hence the blue rectangle representing C

 has moved compared to Figure 4, whereas the adsorbed molecules have stayed put, giving the pink rectangle in the column in the figure and the fact that the red column representing C

 has not moved. Thus, in this fashion adsorbed and mobile phase molecules “leapfrog” each other down the column until they reach the end.

Our discussion so far explains how a band of analyte molecules moves down a chromatography column, but how does chromatography achieve a separation when there are multiple analytes present?

The total time it takes for any molecule to exit the column, the elution time, is the sum of the time it spends adsorbed on the stationary phase and the time it spends moving with the mobile phase, as given by 

 is the time spent in mobile phase, and t

 is the time spent in the stationary phase.

Remember that our sample is a mixture of ethanol and THC, and we assumed ethanol had no affinity for the stationary phase. This means for ethanol t

 is zero, and it will move through the column at the flow rate (3) of the mobile phase. The time it takes a nonadsorbing molecule to exit a column is t

 or the void time, and is given by the flow rate and void volume (the amount of space in the column not occupied by stationary phase), as seen in 

 is the void volume in mL, and f is the flow rate in mL/min.

Assuming for our THC–ethanol separation the void volume is 2 mL, and the flow rate is 1 mL/min, the ethanol molecules will elute at (2 mL/1 mL/min) or 2 min, which is t

 shows a chromatogram with a peak with a t

 of 2 min that we will assume is from unabsorbed ethanol molecules.

 for all molecules is the same because when they are in the mobile phase they all flow at the same speed, the flow rate. What needs to vary then for molecules to elute from a column at different times is for them to have different values of time spent in the stationary phase, t

. To use our horse race analogy, imagine one horse runs a straight, unobstructed path to the finish line while another has fences, trees, and barriers to go over or around. Naturally, the horse with the unobstructed path finishes first, or in our case the molecule that spends the least amount of time adsorbed on the stationary phase elutes first.

? In a nutshell, the affinity of the molecule for the stationary phase. This then is the necessary condition that needs to be met for a chromatographic separation to take place. A large part of the art and science of chromatographic method development is making sure all the analytes in a mixture have different elution times. More on that later.

All this means there hopefully is a mathematical relationship between elution time, the quantity we care about in a separation, and t

. To derive this equation, we first note that the fraction of time analyte molecules spend in the mobile phase and the fraction of time spent adsorbed on the stationary phase equals 1 as follows:

 is the fraction of time spent in mobile phase and F

 is the fraction of time spent in the stationary phase.

 is true because an analyte molecule can partition between two phases, stationary and mobile, so the total time spent in the column is the sum of the time spent in each phase, or 1 in this case.

The fractions of time spent mobile and stationary can be obtained from a chromatogram as follows:

where all these variables are defined above, and realizing that for a given separation t

 and substitute it into Equation 3 we obtain

Note that the denominator in equation 6 contains the quantity (1-F

). As the fraction of time an analyte spends stationary F

) goes down and hence te goes up. This is the mathematical relationship between F

 and retention time we have been seeking. Given our example for THC, F

 is 2 min. This means in this case THC will elute at (2 min/0.5) or 4 min, which is the second peak seen in Figure 6.

Equation 6 is in my opinion the most important equation in chromatography because it relates elution time, the

-axis quantity we care about that is derived from a chromatogram, to the fraction of time an analyte spends adsorbed, which is determined in turn by the chemical properties of the analyte, stationary phase, and mobile phase. Simply put, if you can get F

 to be different for each analyte in a mixture, you can in theory achieve a separation.

For those of you who are spectroscopically inclined, such as myself, I draw the analogy between Equation 6 and the formula for the peak positions in infrared spectra seen in 

where W is the peak position in wavenumber (cm

), c is the speed of light, k is the force constant for a chemical bond, and µis the reduced mass for a chemical bond.

The force constant, k, is related to chemical bond strength. The reduced mass, µ, is calculated from the mass of the two atoms in a bond. Note that k and µ are chemical properties that are different for different chemical bonds. For example, the values of k and µ for a C-H bond are different than for a C=O bond, which is why they have different peak positions in infrared spectra (6). Equation 7 is in my opinion the most important equation in infrared spectroscopy because it relates peak position, the 

-axis quantity we care about that is derived from a spectrum, to the chemical properties of the analyte. Similarly, equation 6 relates elution time, the 

-axis quantity we care about that is derived from a chromatogram, to the chemical properties of the analyte, stationary phase, and mobile phase. Thus, the two most important equations in spectroscopy and chromatography relate peak position on the 

-axis to molecular properties. Note that equation 6 depends upon the experimental setup, such as the nature of the mobile phase, stationary phase, and analyte, whereas infrared peak positions depend upon the chemical structure of the analyte only. More on this later.

The passage of analyte molecules through a chromatographic column is analogous to a horse race, with different molecules passing through the column at different speeds. Via words, pictures, and equations we found that a key quantity that determines elution time is the fraction of time an analyte spends adsorbed on the stationary phase, and that this is a function of the chemical properties of the analyte, mobile phase, and stationary phase.

Skoog, D.A.; West, D.M.; and Holler, E.J.; 

 6th Edition, Saunders College Publishing: New York, NY, 

Undergraduate Instrumental Analysis 5th Edition

Smith, B.C.; Principles of Chromatography, Part I: Theory, 

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

Ruppel, T.; Kuffel, M.; Cannabis Analysis: Potency Testing Identification and Quantification of THC and CBD by GC/FID and GC/MS, 

Smith, B,C.; Infrared Spectral Interpretation: A Systematic Approach, 

Brian C. Smith, 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: 

Smith, B., Chromatography Theory, Part II: How Separations Occur Illustrated and Quantitated, 

Articles

Cannabis Analysis 

Part II of this series looks at the mechanism via which molecules move through HPLC columns and leads us to a discussion on the equations used to characterize separations.

Chromatography Theory,  Part II: How Separations Occur Illustrated and Quantitated

Every year the cannabis industry advances in market growth, technology, and product innovation.

*Projected* 21% overall year over year revenue growth

The cannabis industry experienced massive growth during the peak of the pandemic in 2020 and 2021. However, 

 demonstrates that a correction is occurring especially in mature markets that experienced the most growth. In 2022, Colorado, Nevada, Oregon, and Washington markets declined to revenues that were seen in 2019.

In 2022, 43% of the market consisted of the sale of products using cannabis extract as the main product or an infusion ingredient. The other 57% of revenue was made up of flower and pre-rolls, as seen in 

. The extraction and processing segment of the industry has the most potential for growth as extracts and infused products have unlimited potential for innovation. Flower and pre-roll products seem to have little room for end product innovation and experience the most price compression. Price compression and the hyper-competitive nature of flower sales have driven cultivators to find ways to convert cannabis biomass into higher value products. The result is an increase in concentrates, other processed products, and higher quality products in order to demand higher prices.

Innovation in the extraction and processing segment is often driven by organizations looking for a competitive edge or ways to reduce operating and production costs. 2021 was the year tetrahydrocannabinol (THC) isomers surfaced on the mainstream hemp market. 2022 was the year of THC isomer regulations, oversupply of hemp and cannabis biomass, and the solventless sweep. In February 2022 we reviewed the foundation of THC isomers (3) due to the rapid rise of accessibility to these isomers, mainly delta-8 THC, with no regulatory agency ensuring these products are safe to consume. Delta-8 THC is intoxicating, but milder than delta-9 THC. Delta-8 THC continued to make national news in 2022 as consumers were purchasing these intoxicating “hemp-derived” products from gas stations across the US. As demonstrated in 

, the process by which delta-8 THC is produced involves converting cannabidiol (CBD) into delta-8 THC using strong acids, harsh solvents, and heat.

In the process, unidentifiable non-THC by-products are showing up on chemical analyses. In addition to this, no one is testing for the solvents and acids that are being used to produce these products. These both are cause for concern among scientists as they could present serious health risks to consumers (4). Each state has reacted differently to the rise of delta-8 and its potential consumer safety risks. In 

 you can see that 21 states have strictly banned or regulated delta-8 THC.

In most adult-use states, delta-8 is regulated, but products are available. Delta-8 THC products are offering hemp operators another revenue stream in a flooded CBD market where many are being forced out of business. It is easy to see how operators would be willing to take the risk associated with producing delta-8 products in the hypercompetitive hemp market with a major oversupply of hemp biomass.

Another result of hemp biomass continuing to be in an oversupply and regulated cannabis market prices in 2022 plummeting was the rise in operators converting CBD into delta-9 THC. Operators in states like Washington and California have begun sourcing hemp biomass, extracting CBD, then converting CBD to delta-9 THC in mass quantities. The cost of hemp and the conversion process is so low, which has resulted in prices for extracts in the adult-use and medical markets to fall to all-time lows.

Counter to the falling prices and market compression, concentrates with clear differentiation rapidly expanded in 2022. Products with minor cannabinoids have increased in revenue by 30–1200% depending on the state (6). With CBD losing its novelty, more impactful minor cannabinoids such as cannabigerol (CBG) and cannabinol (CBN) have captivated consumers. Demand for minor cannabinoids from consumers has driven more companies to offer products with these minor cannabinoids. The demand is there, but more importantly these cannabinoids can be sourced from hemp, which creates an opportunity for organizations to produce in-demand products with higher margins.

Solventless products took the market by storm in 2022, increasing in market share by more than 100% (6). Solventless products are produced without using a solvent-based extraction method. The extract is produced using a rosin press method, ice water hash, or a combination of both methods. Solventless extract has made its way into the market through concentrates, vapes, as well as edibles and are continuing to expand. Solventless concentrates seem to offer a different flavor profile that consumers are enjoying. However, with less THC, more terpenes, and more balanced cannabinoid profiles, these concentrates seem to be offering consumers superior experiences with both flavor and high. Solventless products are demanding much higher prices than their counterparts and consumers are paying it! It seems as though consumers are understanding of the fact that producing solventless concentrates and products costs more and is higher in value than other products. This demonstrates a shift in consumer buying behavior where they are looking for added value beyond price and THC. This is an exciting turn in the market and operators are rapidly pivoting to produce more solventless products.

Among solvent extraction methods, a rise in scalability and streamlining took center stage. More than ever, manufacturers are producing systems that minimize the need for human interaction and maximize automation. Ethanol extraction and solvent recovery has never been faster and more efficient by utilizing automation and continuous feed technology. Supercritical fluid CO

 extraction saw the introduction of dewaxed extract within the CO

 extraction system. This eliminates the need for any post-processing, which dramatically improves extraction efficiency and overall cost of production. It will be interesting to see if these advancements prove the value of their high equipment cost.

The US has experienced a turbulent and unpredictable past few years, however, the future continues to be unpredictable. Nothing seems entirely stable or consistent, which means predictions are even harder to make. While the economy continues to reveal the beginning of a recession, the federal government has recently made some political moves that has the public convinced that federal legalization is around the corner. The recession in combination with these changes in the federal political climate has illuminated a few key predictions for 2023.

Price compression will continue, forcing more companies out of business.

To combat this, companies will shift to producing more premium products to maximize margins.

This will also lead to more rapid and expansive product innovation to offer a competitive edge.

THC isomers will remain available on the market unless consumers demand regulation.

Hemp will continue to plummet in price and will supply the THC operators with cheap access to CBD for conversion to THC.

With more blood in the water, mergers and acquisitions will continue to rise.

New medical and adult-use states will enter the space creating markets that demand higher prices. This will create more opportunity for companies to enter more states and multi-state operators (MSOs) to capture higher profit margins.

With the federal administration shedding a more positive light on the cannabis industry and the potential for safe banking, more investment money will enter the space.

This will drive growth and innovation in both plant touching and ancillary cannabis companies.

Equipment, real estate, and operating capital loans and financing options will become more readily available.

Extraction methods and equipment will become more efficient and selective, creating more options for product formulation.

US adults on the fence about whether or not to accept cannabis as a medical or recreational product will finally enter the cannabis space as consumers.

Whether or not these predictions come to fruition, the cannabis industry as a whole has continued on an upward and positive trajectory both from a revenue generating and political perspective. This year will be telling as to the cannabis industry’s true ability to be “recession-proof” as well as understanding the timeline for federal legalization.

Mining for Gold: BDSA Market Outlook, Key Insights, and Ways to Win 

https://bdsa.com/wp-content/uploads/2022/10/BDSA-Mining-for-Gold-Sept-2022.pdf

Cannabis Product Sales by Category U.S. 2020-2022 

https://www.statista.com/statistics/1281580/us-cannabis-product-sales-by-category/

Friesen, L. How Are Tetrahydrocannabinol—THC—Isomers Entering the Market? 

https://cen.acs.org/biological-chemistry/natural-products/Delta-8-THC-craze-concerns/99/i31

CBD Oracle Delta-8 is Available in 30 States While Others Try to Ban It 

https://cbdoracle.com/news/policy/delta-8-thc-legal/

CBN and CBG Sales See Rapid Growth as CBD Sales Slow in Cannabis Markets 

https://bdsa.com/cbn-and-cbg-sales-see-rapid-growth/

 is the founder, CEO, and Chief Extractor of Heylo. With a background in chemistry and clinical research, Lo was inspired to explore cannabis as a medicine and to enter the emerging industry. She joined Eden Labs, a leading CO

 extraction equipment manufacturer to support and expand a Research and Development department. There she managed the development of their latest and greatest CO

 extraction system. In 2017, after working with Eden Labs and another cannabis processor, Lo launched Heylo with a mission to help people get more out of life with cannabis.

Friesen, L., Extraction 2022 Highlights and 2023 Predictions, 

Extraction Science

There goes another year in the books. This review of 2022’s extraction greatest hits takes a look at discoveries, advancements in technology, and other highlights from the year. This article also lays out some predictions for 2023, which is set up to be an interesting year with less access to funding that could slow the development of new technology or innovation in the extraction space.

Extraction 2022 Highlights and 2023 Predictions

In this issue, we will continue the discussion of energy and elements used in atomic spectroscopy by looking at one of the most commonly used analytical techniques in the elemental analytical laboratory—inductively coupled plasma-atomic emission spectroscopy (ICP-AES). This technique is used to achieve high levels of accuracy while being able to discern increasingly low levels of detection in comparison with some of the other techniques we have reviewed. We will examine the structure and function of modern ICP systems to understand how to optimize these techniques for different types of samples and analytical targets.

The discussion of atomic spectroscopy started with the basic principles of changes in energy states that result in the absorption, emission, and other changes in energy measured by atomic absorption spectroscopy (AAS) or atomic emission spectroscopy (AES). In the first elemental spectroscopy column (September 2022), we looked at the form and function of many absorption techniques and started to examine the fundamentals of emission spectroscopy leaving off with the fundamental skeleton of inductively coupled plasma (ICP) (1). The basic components of emission spectroscopy techniques are similar, starting with sample induction to a nebulizer then the passage of the sample to an excitation source leading to optics that directs emission to a detector (see 

Most modern laboratories rely heavily on atomic emission spectrometers with a plasma source either ICP-AES or ICP-mass spectrometry (MS). The sample introduction and nebulization systems for ICP systems are similar or identical to the nebulizers described previously for other emission spectrometers. The primary difference between ICP and other emission spectrometers is the excitation source and the detector. In ICP, the excitation source is plasma—a homogenous mixture of ionized gases (usually containing argon) composed of electrons, ions, and neutral species. ICP systems contain an ICP torch where argon moves an aerosolized sample through an applied Rf power supply. The plasma is ignited by a spark from a Tesla coil (see 

The early generations of ICP systems used a side-on or radial view of the plasma. The radial view takes a plasma cross-section from the side (

). Later, alternative configurations developed end-on or axial observation of the plasma along the entire length of the plasma (

Modern systems can have either type of viewing or be a dual configuration (combination of both axial and radial viewing). In dual view systems, the torch is oriented most often in a horizontal configuration and optics are used to create the axial and radial views. This horizontal configuration allows the optics to stay cleaner as the heat and exhaust of the torch vents away from the path of the optical components.

Radial plasma tends to have less sensitivity than the axial configuration but is preferred with samples with complex or difficult matrices. Samples with large amounts of organic compounds or dissolved solids perform better with a radial view opposed to the axial view. One of the reasons is that an axial view boosts sensitivity of all the elements in the plasma including the background and can be susceptible to interference.

An interference is any effect that can alter the measurement of the target elements. In ICP spectroscopy there are three important types of interference: physical, chemical, and spectral.

 are the differences between the samples being tested and any standards, calibration curves or blanks that can alter the sample introduction and nebulization. Most of the difference stems from the makeup of the matrices of the sample and standards. Changes in density, viscosity, and dissolved organic matter from either the matrix or, as a result of any digestion or preservation techniques, can create these physical interferences. Some of these difficulties can be addressed by diluting samples, using internal standards, standard addition, and matrix matching.

 are differences in the way in which a sample, standard, or element reacts in the plasma. These interferences can result in ionization effects, formation of molecules, or changes to the plasma. Some elements, such as alkali metals, often are seen to have ionization interactions resulting in interferences. Chemical interferences can be addressed by adding ionization buffers.

 are changes to the optical or detector spectral response from background shifts, interference from other elements in the sample, or spectral overlaps. In background shifts, the baseline or background of the sample is different between the sample injection and injections of standards, calibrants, or blanks. These shifts can occur because of the presence of an element in one of the injections that is not present or in the same concentration as the other injections.

 is the difference in the baseline, either higher or lower than the target sample’s baseline in the immediate area of a targeted spectra (see 

). A sloping background shift is also often due to the presence or concentration of another element that causes the baseline or background to slope either upward or downward in the area of a target spectrum (see 

). Complex background shifts are interferences from multiple sources or elements creating increased noise in the baseline interfering with the detection of the target spectrum.

Another type of spectral interference are partial or full spectral overlaps (see 

). Direct overlaps occur when the wavelengths between two elements are too close to fully resolve. Despite the fact that no two elements have exactly the same spectral emission, there are many situations where the width of the spectral lines or resolution of the optical system can create overlap. When these overlaps occur, a correction called an 

 (IEC) may be applied. The correction applies a ratio based on the concentration of the interfering element. To properly use the IEC, the concentration or contribution of the interfering element must be measured at another wavelength and a correction factor applied. Each suspected interfering element must be measured and used in the correction. Complexity of the sample can cause the use of an IEC to be cumbersome and may be limited by the optics of the system.

Spectral interferences can be mitigated by understanding the composition of your samples and running single element standards for would be targets and interferences. Many analytical methods include standard wavelength suggestions that can be adjusted for specific types of matrices. For unknown samples, it may be necessary to run some preliminary semi-quantitative runs to determine the overall composition before fine tuning analysis for the targets of primary interest. Once the probable composition is known, wavelengths can be selected to minimize interferences. Many references can be found for the most common wavelengths for ICP amenable elements including an online tool kit from myperiodictable.us (see 

Examining Energy, Wavelengths, and Optics

At the base of all ICP-optical emission spectrometry (OES) detection is the ability of the system to measure the change in energy resulting in specific wavelengths detected by an optical detector. If you remember previous discussions of the electromagnetic spectrum, the larger the wavelength the lower the energy state. In elements, the more transitions of energy state the more energy is released and detectable as lower wavelengths. So, a transition of an electron through two or more orbitals has more energy and a shorter wavelength than a transition of an electron through a single orbital (see 

The ability of an ICP-OES system to detect and isolate elements and their wavelengths depends on the system’s optics and detector. A typical ICP-OES system must be able to produce wavelengths over a wide range of energies, typically from approximately 150–850 nm, and have resolution typically better than 0.03 nm over several wavelengths. The emitted energy from a single element must be first differentiated and then isolated to programmed wavelengths. This task is accomplished by a series of slits, mirrors, diffusion gratings, filters, and prisms. There are two basic types of optical configurations: monochromator and polychromator. A monochromator is usually a sealed system pumped with a gas such as argon where there is one entrance and one exit to the optical system allowing for the isolation of one element and wavelength at a given point in time to pass through to a detector. The angle of the grating can be altered quickly, allowing for scanning of more than one wavelength during sampling but it is limited by the sample and sampling rate. There are two common mount configurations for a monochromator: Czerny-Turner and Ebert. The Czerny-Turner possesses two concave mirrors while the Ebert mount has one large concave mirror that spans between the entrance and exit slits. Monochromators with their ability to examine one wavelength after another are known as sequential configurations.

A polychromator is a system that has an entrance slit and multiple exit slits leading to more than one detector and allowing for the detection of more elements and more than one wavelength in a set time period. This allows for more information to be collected across multiple elements, but also will reduce resolution of any one element or wavelength since the signal is split up into dozens of wavelengths limited by the number of detectors. This multi-element capacity makes polychromators known as a simultaneous configuration. The polychromators are less flexible than monochromators and are set to fixed wavelengths where the monochromators can be change quickly and frequently (see 

Both types of systems use a diffraction grating to split the signal into selected wavelengths. The basic principle is that incident light hits the highly polished and etched mirror finish of the diffraction grating and splits into component wavelengths. The surface of the diffraction grating mirror is finely etched with lines closely spaced together resulting in several hundred to thousands of lines per millimeter. As the light strikes the surface of the grating, it is diffracted at specific angles depending upon the wavelength of light and the density of lines.

Another type of grating common in ICP-OES analysis is an echelle grating. This grating has a course set of lines compared to the other diffraction grating and separates light of a variety of wavelengths and produces overlapping spectra. A second device (either another grating or prism) further cross disperses the light into a two-dimensional array called an echellogram.This type of grating diffracts lights and focuses it on the slits. This type of grating is often used in polychromator systems.

As with all the other analytical systems we have examined the last component of the analytical system is the detector. In ICP systems, since the resulting energy are wavelengths of light the detector will use some combination of a photomultiplier, photo diode array, or solid-state device (similarly to the same components discussed in the previous columns regarding chromatography and spectrophotometry). In the past, analytes were measured by photomultipliers. Modern systems rely on technology called solid-state charge transfer devices (CTDs). These devices have almost completely replaced photomultipliers. There are several types of CTDs used to process signals including charge injection (CID) and charge coupled (CCD) devices. The CID and CCD technology is based on the light sensitivity properties of silica and have the benefit of being two-dimensional detectors compared to the older photodiode arrays and photomultipliers. The silicon atom structure forms crystalline lattice with each silicon atom bonding to the adjacent atoms to form a three-dimensional structure. When photons break one of the bonds it results in a release of an electron and leaves a hole in the lattice called the electron-hole pair (see 

). The released electron moves in the opposite direction while the hole moves with the energy direction causing a charge.

A two-dimensional silicon wafer composed of elements called pixels stores and detects photonic charge. CIDs allow activation of random and individual pixels with an electrode at each pixel site. CIDs have a very light sensitive surface divided into thousands of pixels with rows and columns of electrodes that allow for detection of wavelengths from about 160–900 nm. The integration of CID pixels means that the charge is not destroyed. CIDs have higher noise and require extreme cool as with liquid nitrogen to reduce noise.

CCDs have a more systematic charge output measurement from the pixels. The charge is transferred sequentially from each pixel to a buffer then to a recorder. Pixels are processed in rows or columns that are read by the CCD. As the pixels are read, the charge is destroyed. These devices are very fast, but require cooling with a cooling system such as a Peltier unit.

Some devices combine features of both the CID and CCD. Systems such as a segmented array charge coupled device (SCD) uses an echelle array and is designed with thousands of pixels arranged in small subarrays (about 200) positioned on the silicon wafer producing an echellogram. These subarrays mirror the most important ICP spectral lines of about 70 common elements. The subarrays each have their own electrode and can be read in any order. These systems have higher sensitivity than the other types of CCD systems.

Inductively coupled plasma spectroscopy is a widely used instrumentation in inorganic analysis. The technique has existed for many years with great improvements on range of analytes and limits of detection that benefit the analyst. There are many choices in the individual components of the system that can be made to further refine and target analysis—from the sample induction to the torch, optics, and detector—the choice of configuration is important to improving your methodology. By understanding the physical configurations and their use, the analyst has another tool to improve their workflow.

Atkins, P. Energy and Elements, Part I: Understanding Atomic Spectroscopy, 

Boss, C. B.; Fredeen, Instrumentation and Techniques in Inductively Coupled Plasma Optical Emission Spectrometry, 

Neubauer, K., How to Choose the Correct Wavelength in ICP-OES, 

 is a Senior Applications Scientist with Spex, an Antylia Scientific company and has been a member of many cannabis advisory committees and working groups for cannabis including NACRW, AOAC and ASTM.

Atkins, P., Energy and Elements, Part II: A Closer Look at ICP

Navigating the Lab

Here we continue the discussion of energy and elements used in atomic spectroscopy by looking at one of the most commonly used analytical techniques in the elemental analytical laboratory—inductively coupled plasma-atomic emission spectroscopy (ICP-AES).

Energy and Elements, Part II: A Closer Look at ICP

In 2018, the Hemp Farming Act which proposed that hemp be removed from Schedule I and be reclassified as any other agricultural commodity was adopted into the 2018 Farm Bill, making hemp federally legal (1). Even though hemp is now legal in the US, its production is highly regulated and hemp testing laboratories must be aware of the relevant laws and regulations they must adhere to. This article delineates the nuts and bolts of the United States Department of Agriculture’s (USDA’s) hemp testing rules and what laboratories should know regarding hemp testing.

 that contains less than 0.3% of the psychoactive cannabinoid tetrahydrocannabinol (THC) on a dry weight basis. 

 containing more than 0.3% THC is classified as cannabis.

Here is where the line is drawn between hemp, which is legal, and cannabis, which remains a schedule I substance and is therefore illegal. The Controlled Substances Act (CSA) prohibits the “manufacture, distribution, dispensation, and possession of Schedule I substances except for federal government-approved research studies” (2). This delineates the need and relevance of hemp testing laboratories; to ensure that the 0.3% threshold for THC is not exceeded.

The Agricultural Improvement Act of 2018 (2018 Farm Bill) removed hemp from the Drug Enforcement Administration (DEA) list of Schedule I controlled substances by adopting the Hemp Act (3). Consequently, the US Domestic Hemp Production Program was created to provide regulatory oversight for the hemp industry. The program mandated the USDA to create a regulatory framework and issue guidance to regulate the production of hemp. Following this, the USDA issued the Interim Final Rule (IFR) on hemp, which was published on October 31, 2019 (4). The US Food and Drug Administration (FDA) also allowed a 60-day comment period for the IFR. The 2020 growing season was used to test drive the IFR and together with the submitted comments from the public, the Final Rule (FR) was published in 2021 (5).

The Final Rule (FR) on hemp was published in January 2021 and took effect on March 22, 2021. It covers several key factors on hemp testing, including licensing requirements for hemp testing laboratories, sampling and testing procedures, handling of violations, reporting of test results, and the disposal of noncompliant crops. For hemp testing laboratories, the most critical consideration from the FR is the accurate testing of hemp samples and the calculation of THC concentration on a dry weight basis (5).



Grand View Research placed the value of the global hemp market at $4.13 billion in 2021. The value is expected to hit $16.75 billion by 2030, growing at a compound annual growth rate (CAGR) of 16.8% (6). This signals the growing demand for hemp testing services in the next couple of years to meet the regulatory requirements defined by the FR.

Not many laboratories are offering hemp testing services at the moment because the industry is still new, and the regulatory requirements are stringent. For example, hemp testing can only be offered by laboratories that are registered with the DEA. Thus, there’s a large opportunity of venturing into hemp testing and owning the space before it gets crowded.

Hemp testing laboratories as well as those aspiring to venture into the niche need to be conversant with the relevant rules as defined in the Final Rule. They also need to leverage the best technology to get ahead of the regulatory curve and become profitable in the soonest time possible. Here are the nine rules that apply to hemp testing.

Nine Cardinal Rules for Hemp Testing Laboratories

Hemp testing laboratories are of course free to test several other compounds as well. This means that they need to adhere to all the rules and regulations that apply to laboratories. However, there are some rules and regulations that are specific to hemp testing and must be adhered to as well. This begins with how hemp samples should be received, tested, and disposed of as well as how the test results should be disseminated. Some of the rules for hemp testing laboratories are below.

All laboratories that test for hemp must be registered with the DEA. Registration information can be accessed from the DEA’s Diversion Control Division website. The FR states that “Registration is necessary because laboratories could potentially handle cannabis that tests above 0.3% THC on a dry weight basis, which is, by definition, marijuana and a Schedule I controlled substance.” Fortunately, hemp testing laboratories had until the final day of 2022 to comply with this requirement.

The laboratories will also need to open a Hemp eManagement Platform (HeMP) account that allows them to manage and submit hemp information and report to the USDA, replacing the need to submit via email, mail, or fax. The USDA has published a directory for laboratories that have been certified by the DEA to test hemp that customers and other stakeholders can easily access.

2. Measurement of Uncertainty Must Be Calculated and Reported

The measurement of uncertainty (MU), is a measurement that expresses an acceptable degree of variance in a test result. It characterizes the limit of dispersion in the values (7). Hemp testing laboratories are required to calculate and report the MU when testing for THC in a sample. Possible sources of uncertainty need to be identified and their level of contribution quantified. This will be used to determine the overall measurement of uncertainty. The method for calculating the MU needs to include all possible contributors of uncertainty including the possible decarboxylation of THC (8).

The FR is very specific as to the parts of hemp that can be used as samples for THC testing. This is because THC, as well as other phytochemicals, are present in high amounts in the flowers, but exist in minimal amounts in the stems. Fortunately, the FR gave a greater allowance for the parts of hemp that can be tested, as compared to the IFR. Laboratories can now test samples from the buds and leaves, for as long as they are taken from at least 5–8-in. away from the main stem. Samples can also be picked from the central cola or terminal bud.

Once an appropriate sample has been obtained, it should be dried within 24 h. Note that THC is usually reported on a dry weight basis. It is important to minimize the variance that would arise if some laboratories report the THC level using a wet sample while others report using a dry sample. If the sample cannot be dried within 24 h, then it should be frozen at a temperature of -20 °C or lower.

The laboratory should grind the sample to a fine consistency prior to testing it. This is important for homogeneity. The sample should then be divided into two parts. One part should be used for testing while the other should be retained in case a retest is needed. The sample preparation procedure should be well captured in the laboratory’s internal standard operating procedures (SOPs) (9).

5. Use of Post-Decarboxylation Testing Method

The FR requires that hemp testing laboratories use a post-decarboxylation method when testing for THC in a hemp sample. Decarboxylation is the process through which acidic cannabinoids lose their acidity and become activated after they are exposed to heat. Raw hemp contains THC (as well as other cannabinoids) in its acidic forms. When exposed to heat, tetrahydrocannabinolic acid (THCA) is converted to the psychoactive cannabinoid THC. A testing method that does not consider the decarboxylation process will fail to capture the true concentration of THC in a given sample.

Gas and liquid chromatography are the approved post-decarboxylation methods for testing THC in hemp. Laboratories are free to explore other testing methods that factor in the decarboxylation process. However, laboratories will need to seek approval in writing prior to using any other method to test for THC (9).

The FR increased the negligence threshold for hemp testing from 0.5% to 1%. A hemp sample that tests above 0.3% is considered hot hemp. However, if the THC does not exceed 1%, it is not considered a negligent violation and hence is not penalized. However, the hot hemp still needs to be destroyed and disposed of. Previously, the FR allowed a negligence threshold of up to 0.5%. If a hemp sample tests above 1% THC, this is considered a negligent violation and subsequently the Licensed Producer (LP) is issued with a notice of violation. If an LP receives three consecutive notices within five years, the LP will be disallowed from taking part in the hemp program for the next five-year period.

Hemp testing laboratories are required to share the test results with the USDA, regardless of whether the THC limit is exceeded or not. The test results are shared through HeMP, so laboratories will need to create a HeMP account. The laboratories should also download Form 22 from the USDA’s Agricultural Marketing Services (AMS) website and use it to submit the THC results.

The test results must also be shared with the LP in a format that is most suitable for them. The results must be shared with the relevant state or tribe department of agriculture and should bear the stamp “official compliance.” Laboratories must store the test results for a minimum period of three years.

The laboratories can conduct a retest of the hemp sample at the request of the LP. The same procedure for testing, as stipulated in the SOP, should befollowed. The results should also be shared with the three parties: licensed producer, the USDA, and the relevant department of agriculture. The LP should bear the cost of the retest.

After testing, a laboratory is likely to come across some hot samples that will need to be destroyed and disposed of (10). Fortunately, the FR allows for the remediation of hemp, which means utilizing the hemp biomass. This prevents the LP from incurring huge losses. Different “dilution” methods can be used to remediate hemp. For example, the hemp can be shredded into biomass, and then a sample submitted for testing. The original testing method should be used for the retesting. The hemp biomass can be used for fiber or fuel if the THC level is compliant upon further testing.

As mentioned earlier, hemp testing is a potentially lucrative venture that has minimal competition at this point. That said, since the industry is being highly regulated there are several barriers to entry. Laboratories looking to venture into the industry should not be deterred by the hurdles, but should instead do all it takes to understand the legal landscape and implement measures to remain compliant with the relevant rules and regulations.

A LIMS Helps Hemp Testing Laboratories Meet Compliance Challenges Head On

A software as a service (SaaS) laboratory information management system (LIMS) for cannabidiol (CBD) and THC testing can help hemp testing laboratories get through the seemingly insurmountable compliance challenges (11). A LIMS helps laboratories track samples and manage test results while ensuring an optimum level of accuracy. A LIMS automates laboratory workflows, ensures efficiency, and minimizes errors. A LIMS supports the dissemination of test results to the relevant stakeholders while maintaining the record for the stipulated duration. A cloud-hosted LIMS enables remote access to laboratory data while providing enhanced data security (12). A cloud-based SaaS LIMS for CBD and THC testing reduces the operational costs of laboratories by eliminating the need to purchase IT infrastructure and hire dedicated IT personnel. Moreover, it enables laboratories to eliminate other upfront costs otherwise needed for deploying a traditional LIMS.

Hemp testing laboratories must meet the regulatory requirements set by the USDA that can be both specific and cumbersome. For example, all testing methods must take into account the decarboxylation of THCA to THC once the sample is exposed to heat. This restricts the testing methods that laboratories can use. Another challenge is balancing regulatory requirements with customer satisfaction. A hemp testing laboratory may develop a precise and accurate method to test THC that meets regulatory threshold for a well-defined measurement of uncertainty. However, this method may provide results that do not favor the customer, hence the laboratory can easily lose the customer on these grounds. At the end of the day, hemp testing laboratories must juggle between meeting regulatory requirements, satisfying customers, and at the same time ensuring the highest level of scientific accuracy. A cloud-based SaaS LIMS for CBD and THC testing enables hemp testing laboratories can meet compliance goals with ease while always assuring testing accuracy.

H.R.5485 - 115th Congress (2017-2018): Hemp Farming Act of 2018 

https://www.congress.gov/bill/115th-congress/house-bill/5485 

This article addresses the US Department of Agriculture’s hemp testing rules and what laboratories should know regarding hemp testing. The author provides a short tutorial for laboratories and includes a section how laboratory information management systems (LIMS) can help.

January/February 2023

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Delineating the Nuts and Bolts of the USDA’s Hemp Testing Rules