cbd analysis hplc

December 15, 2021 By admin Off

Costs is another parameter for comparison these methods. Set up, maintenance and running costs are often important factors in selecting analytical techniques specially in industry settings. The running cost of LC, GC and HPLC are negligible but for equipment, LC is more expensive than GC and both are much more expensive than HPLC. Coupling of mass spectrometry with LC or GC can further increase the costs.

Matrix-Assisted Laser Desorption Ionization Mass Spectrometry ( MALDI-MS) is a new method which has been used in some studies for comparison with usual methods such as LCMS and GCMS in identification of cannabinoids metabolites (Beasley et al. 2016). Recently this method has attracted attention because compared to usual mentioned methods, the sample preparation is simpler, a narrower time frame of drug can be detected, and the sample amount is reduced. Beasley et al. (2016) have used MALDI-MS to detect the cannabinoids in a single hair sample. In this study, MALDI instrument was consist of MDS Sciex hybrid quadrupole time-of-flight mass spectrometer with an orthogonal MALDI ion source and a neodymium- doped yttrium aluminum garnet laser.

Around 144 cannabinoids have been identified in cannabis plant, among them tetrahydrocannabinol (THC) and cannabidiol (CBD) are the most prominent ones. Because of the legal restrictions on cannabis in many countries, it is difficult to obtain standards to use in research; nonetheless, it is important to develop a cannabinoid quantification technique with pharmaceutical applications for quality control of future therapeutic cannabinoids.

Although MS offers many benefits, the use of qTOF mass spectrometers is ideal when trying to differentiate between two compounds with different compositions but the same nominal mass (Citti et al. 2018). qTOF mass spectrometers can provide accurate mass identification with a threshold less than 5 ppm for precursor and product ions; this allows for differentiation between isomers of cannabinoids (Aizpurua-Olaizola et al. 2014; Citti et al. 2018) such as Δ8-tetrahydrocannabinol and Δ9-tetrahydrocannabinol which have the same m / z because these cannot be differentiated by MS (Citti et al. 2018). Such isomers may have different therapeutic properties and may need to be separated for manufacture, so it is important to adopt an analytical technique that can differentiate between them.

Leghissa et al. (2018b) used Multiple Reaction Monitoring (MRM) analysis of cannabis from a surrogate hops matrix by GC-MS with triple quadrupole mass spectrometry for the first time. They used silylated cannabinoids to avoid decarboxylation process due to high temperature in GC injection port. They found that this method is applicable to cannabinoids analysis from plant materials and cannabis products. The main achievement of their study is that, in this method, because the risk of interferences from the essential oils and waxes is reduced the extraction need less sample preparation in the laboratories compared to other techniques like SPE.

TLC (thin layer chromatography)

From Table 3, it is clearly obvious that C18 is the most popular column as it is mentioned earlier. The main difference between HPLC and GC is the operating temperature. That is why HPLC is used when preserving the acidic form of cannabinoids are matter. The only disadvantage of HPLC is, it is not able to analyse the volatile compounds like terpenes.

There are multiple benefits to using HPLC-MS/MS over other analytical methods presented in this review. For example, HPLC can differentiate between acidic and neutral cannabinoids, unlike GC (Romano and Hazekamp 2013). MS offers several benefits over other detection methods as well. For example, MS can differentiate between different cannabinoids based on the m/z value of their molecular ion. It offers more specificity compared to UV detectors and can analyze extracts from complex matrices, such as cannabis (Citti et al. 2018; Leghissa et al. 2018a).

There are other aspects which must be considered when selecting a quantification method such as method performance for different types of cannabinoids and also common interfering substances and impurities. Figure 2 shows suggested methods for cannabis compounds and their impurities. There is no evidence to support which analytical method works best for a specific cannabinoid. However, generally, LC is the preferred method for cannabinoids and GC for terpenes. GC does not have the capability to quantify the acidic form of cannabinoids unless through derivatization while terpenes cannot be detected by LC because they are volatile compounds. Another important factor to consider is analytical sample preparation which is the most time consuming and the most common cause for generating errors during analytical process. Sample preparation method, storage and handling are some of the parameters which can affect the results. For cannabis, the final sample should represent the original lot. So, not only the extraction method, but also the cultivation and processing steps play an important role in the analytical results.

Hazekamp (2013) used the TLC method both for polar and non-polar systems. They used reversed-phase silica gel plates and normal phase silica gel plates for non-polar and polar systems respectively. For more accurate results they used Fast-Blue B salt (4-benzoylamino-2, 5-diethoxy benzene diazonium chloride hemi salt) which is a suitable coloring agent for visualization of cannabinoids at TLC plates. Fast-Blue B can determine acetylcholinesterase, α- and β-glucosidase activity by changing to different colours which come from reacting of FBB with various compounds, however the colors depend on the concentration of constituents. As a result, they found that TLC is useful in rapid screening of many samples for the existence of cannabinoids, however, its performance is lower compared to other separation methods and the reproducibility of TLC depends on some parameters such as relative humidity (Romano and Hazekamp 2013).

Different detection techniques can be used in conjunction with High Performance Liquid Chromatography (HPLC) to analyze cannabinoids. Common detection methods include mass spectrometry (MS) and ultraviolet (UV) absorbance (190 to 400 nm) (Aizpurua-Olaizola et al. 2014; Leghissa et al. 2018a). UV detection is much less expensive and more straightforward than MS detection (Leghissa et al. 2018a). Acidic cannabinoids show absorption peaks at around 270 nm and 310 nm while neutral cannabinoids show absorption peaks at about 220 nm (Citti et al. 2016; Hazekamp et al. 2005). Citti, Ciccarella (Aminah Jatoi et al. 2002) developed a rapid HPLC technique with UV detection (HPLC-UV) to qualify and quantify major cannabinoids (CBDA, CBD, CBN, THC, and THCA) in cannabis extracts. However, absorption profiles from UV detection do not contain enough information to be used in isolation to accurately identify cannabinoids (Leghissa et al. 2018a). Much more information can be obtained by diode array detection (DAD), which covers the visible and UV spectrum. DAD can help to improve specificity because acidic and neutral cannabinoids have different absorption spectrums (Aminah Jatoi et al. 2002; Leghissa et al. 2018a). Thus, Peschel, Politi (Andreae et al. 2015) used HPLC-DAD to differentiate between Cannabis sativa chemotypes, extracts of different polarity, and to profile extracts.

Hazekamp et al. (2005) measured cannabinoids with FTIR. They added KBr to the ethanolic solution of cannabinoids followed by vacuum ethanol evaporation because KBr does not show any absorption spectrum in IR region. Additionally, KBr has a 100% transmission window in the range of wave number at the FTIR spectroscopy. The IR spectra were measured in the range of 500 to 4000 cm − 1 . Compared to UV spectra, IR spectra presented more absorbance peaks (Hazekamp et al. 2005). Mutje et al. (2007) showed the existence of carbonyl and ester groups by the FTIR peak at 1775 and 1725 cm − 1 in composite samples of cannabis extract.

To find relevant articles for this narrative review paper, a combination of keywords such as medicinal cannabis, analytical, quantification and cannabinoids were searched for in PubMed, EMBASE, MEDLINE, Google Scholar and Cochrane Library (Wiley) databases.

In the Table 2, nitrogen and helium are the carrier gases. In many studies, it is proved that nitrogen has the best efficacy, but it is time consuming. On the other hand, by using helium, the process is rapid and efficient, but the price is not affordable. The Initial and end temperatures, the type of columns and thedrawback are almost similar to GC-MS.

Based on the information presented in this review, the ideal cannabinoid quantification method is HPLC- MS/MS for the cannabinoids.

Nonetheless, all light absorbance detectors lack the specificity of MS (Citti et al. 2018; Leghissa et al. 2018a), which is particularly useful in analyzing extracts from complex matrices such as cannabis . However, some cannabinoids, such as CBG and CBD are difficult to separate using UV detection especially in concentrations greater than 10% in the extract (Citti et al. 2018; De Backer et al. 2009). In the case of CBG and CBD, MS is preferred because it can differentiate between different cannabinoids based on the m/z value of their molecular ion (Citti et al. 2018). M/z value is not always unique, however; in an ongoing study, Citi, Braghiroli (Beal et al. 1995) found five cannabinoids with the same m/z of 315.2294; this value matches that of THC and CBD in Bediol® oil and ethanol extracts. Because some of these cannabinoids coelute, analysis of these compounds is difficult.


In another study, Gas Chromatography with Vacuum Ultraviolet spectroscopy (GC-VUV) was used which is gas chromatography with vacuum ultraviolet spectroscopy. The detection of cannabinoids and the cannabinoid metabolites was fast and simple, so that it can be used in rapid detection of them even without having a baseline for cannabinoids for comparison. This method has just one disadvantage which is high limit of detection (LODs). Due to this drawback, detecting analytes in biological matrices cannot be accomplished without pretreatments (Leghissa et al. 2018c).

To find relevant papers for this narrative review paper many data bases have been reviewed for 8 months. A combination of keywords such as medicinal cannabis, analytical, quantification and cannabinoids were searched. Papers from 1967 to 2019 from PubMed, EMBASE, MEDLINE, Google Scholar and Cochrane Library (Wiley) databases have been searched in English. In the next step, papers have been scanned to discard irrelevant papers. Those papers which were relevant went through for more investigations in details. In total, the number of papers which have been read were about 75 including around 15 irrelevant papers.

Cannabis sativa L. is an annual herbaceous flowering plant indigenous to eastern Asia (De Backer et al. 2009). The phenotypes of cannabis are highly variable and the plant is accepted to have two subspecies: C. sativa subsp. sativa and C. sativa subsp. indica (Hillig and Mahlberg 2004; Knight et al. 2010). A third subspecies, C. sativa subsp. ruderalis, has been identified; however, it is not broadly recognized (Fischedick et al. 2010a; Hillig and Mahlberg 2004). Cannabis has been used for its therapeutic properties for thousands of years and it was introduced in western medicine in the nineteenth century until its prohibition in the US from mid-1930s (Aizpurua-Olaizola et al. 2014).

GC is normally coupled with mass spectrometry (MS) or flame ionization detection (FID) to detect and quantify cannabinoids (Citti et al. 2018; Hazekamp et al. 2009) (Tables 1 and 2). MS employs standardized electron ionization to fragment analytes, permitting the use of compound libraries to identify the parent analyte. FID provides more accurate cannabinoid quantification because it makes use of relatively cheap authentic standards while mass spectrometry usually requires equivalent deuterated standards, which are expensive and not available for all cannabinoids (Citti et al. 2018; Hazekamp et al. 2009).

The medicinal compounds from cannabis are mostly concentrated in the female flowers of this dioecious species (Fischedick et al. 2010a). The so-called resin is the source of a wide variety of terpenoids and cannabinoids (Fischedick et al. 2010a). The therapeutic properties of cannabis are attributed to cannabinoids (Hazekamp et al. 2014). Cannabinoids are found in the resin produced by the trichomes which are widely distributed on both the male and female plants however are most highly concentrated on the female flowers of the cannabis plant (Citti et al. 2018; De Backer et al. 2009). Cannabinoids are terpenophenolic compounds unique to cannabis ( Hillig 2004 ) . To date, 144 cannabinoids have been identified (Hazekamp et al. 2014). The two cannabinoids most well known for their therapeutic properties are tetrahydrocannabinol (THC) and cannabidiol (CBD) (Aizpurua-Olaizola et al. 2016; Hillig 2004). THC and CBD are the neutral homologs of tetrahydrocannabinolic acid (THCA) and cannabidiol acid (CBDA) respectively (Aizpurua-Olaizola et al. 2016). A conventional classification model of cannabinoids is due to their chemical contents dividing them to eleven subclasses including cannabigerol (CBG), tetrahydrocannabinol (THC), cannabidiol (CBD), cannabichromene (CBC), cannabinol (CBN), (−)-Δ8- trans -tetrahydrocannabinol (Δ8-THC), cannabicyclol (CBL), cannabinodiol (CBND), cannabielsoin (CBE), cannabitriol (CBT) and miscellaneous (Berman et al. 2018) (Fig. 1).

The most common cannabinoid quantification techniques include gas chromatography (GC) and high-performance liquid chromatography (HPLC). GC is often used in conjunction with mass spectrometry (MS) or flame ionization detection (FID). The major advantage of GC is terpenes quantification however, for evaluating acidic cannabinoids it needs to be derivatised. The main advantage of HPLC is the ability to quantify both acidic and neutral forms of cannabinoids without derivatisation which is often with MS or ultraviolet (UV) detectors.

HPLC-electrospray ionization-quadrupole time of flight (HPLC-ESI-qTOF) is very effective in identifying complex and common compounds and can identify the main component of the sample in addition to enhancing the signal to noise ratio in the peaks (Aminah Jatoi et al. 2002). Citti, Ciccarella (Aminah Jatoi et al. 2002) analyzed cannabinoid concentrations in olive oil, ethanol, and supercritical CO 2 and found that UV-DAD and qTOF detectors produced similar results, thus suggesting that these two detection systems are equally useful in cannabinoid analysis. Pellati and Brighenti (Brighenti et al. 2017; Pellati et al. 2018) used HPLC-ESI-MS both in positive and negative ion mode for the analysis of cannabinoids. By developing HPLC methods, they improved resolution, peak shape, and separation performance together with the improvement of the ionization in HPLC-ESI-MS (Brighenti et al. 2017; Pellati et al. 2018).


Ultra-performance liquid chromatography allows researchers to use a thinner column compared to HPLC, and it can be used for particles less than 2 μm which leads to better separation with higher speed than conventional HPLC. Additionally, Aizpurua-Olaizola, Omar (Borgelt et al. 2013) identified seven unknown minor cannabinoids using UPLC-quadrupole time of flight mass spectrometry (UPLC-qTOF). Jung, Meyer (Brighenti et al. 2017) also implemented qTOF in their study to isolate and identify THCA and 12 of its metabolites in rat urine using LC-MS, LC-MS/MS and LC-qTOF MS. The use of qTOF results in increased accuracy of the detected ions, and when analysing extracts acquired from complex matrices using MS/MS allows for increased cannabinoid specificity (Leghissa et al. 2018a).

Another alternative to GC and HPLC is NMR (Citti et al. 2018; Hazekamp et al. 2014). NMR is accurate and reproducible and unlike GS and HPLC, NMR is not sensitive to impurities, such a chlorophyll or lipids present in the sample (Hazekamp et al. 2014). Hazekamp, Choi (Casiraghi et al. 2018) developed a method for cannabinoid quantification using 1 H-NMR that does not require chromatographic purification and has a 5-min final analysis time. In that study, they analysed singlets in the δ 4.0–7.0 range in the 1 H-NMR spectrum and found that their technique was appropriate for the quantification of CBDA, THCA, CBG, CBGA, and possibly other cannabinoids as well. One of the major advantages of this technique is that reference standards are not required, meaning that this method can quantify cannabinoids that lack pre-existing reference standards and therefore cannot be analysed by other techniques. Although the results from NMR are promising, one major disadvantage to NMR is that high resolution instruments are very expensive (Citti et al. 2018).

Because consumers have limited means to analyse the chemical composition of the cannabis products, consumers may be inadvertently purchasing products with undesired properties given that different cannabinoids produce different effects (Fischedick et al. 2010b). As a result, it is important to implement methods of quality control so that consumers can be certain that what they are consuming will have the desired effects (Dussy et al. 2005; Fischedick et al. 2010a; Fischedick et al. 2010b). As cannabis use becomes progressively accepted, it becomes increasingly important to quantify the cannabinoid profile and content of cannabis preparations to ensure the uniformity and quality of the preparations (Omar et al. 2014).

Given that cannabis preparations from the same cannabis strain can vary by as much as 25% in cannabinoids composition, there is a clear need to develop an effective and efficient cannabinoid quantification technique so that clinicians can be certain about the chemical properties of the products they are administering (Hazekamp and Fischedick 2012). This literature review has explored a variety of cannabinoid quantification techniques.

A variety of analytical techniques have been developed for quantification and qualification cannabinoids and other compounds in cannabis plant. Advances in analytical methods have also resulted in detection of various compounds from cannabis extracts in the last decade (eg terpenes). The purpose of this literature review is to explore cannabinoid quantification techniques and subsequently suggest an optimal method for pharmaceutical grade quantification.

Liquid chromatography (LC) often employs electrospray ionization (ESI) and atmospheric-pressure chemical ionization (APCI) as ionization sources (Grauwiler et al. 2007). These usually only generate a protonated molecule without diagnostic fragmentation; therefore MS/MS is required to obtain diagnostic information when using LC. Additionally, because cannabinoids have phenolic and carboxylic functional groups that are not ionized effectively using ESI or APCI, GC-MS may offer greater sensitivity that LC-MS (Leghissa et al. 2018a). GC × GC provides greater separation power and analysis speed compared to coupled-column techniques such as liquid chromatography-mass spectrometry (LC-MS) (Dallüge et al. 2003).

The most common cannabinoids and their conversion pathway by decarboxylation because of heat or aging. C BGA can convert to CBDA and THCA by CBDA synthase and THCA synthase, respectively. CBGA: cannabigerolic acid, CBG: cannabigerol, CBDA: cannabidiolic acid, CBD: cannabidiol, THCA: tetrahydrocannabinolic acid, THC: tetrahyrocannabinol, CBN: cannabinol ( Fathordoobady et al. 2019 )

GC-MS is often employed for cannabinoid quantification (Ciolino et al. 2018; Groger et al. 2008; Hazekamp et al. 2009; Jung et al. 2009; Omar et al. 2013; Omar et al. 2014). However, quantification of cannabinoids via GC requires a derivatization step to avoid the decarboxylation of acidic cannabinoids (Citti et al. 2018; De Backer et al. 2009; Grauwiler et al. 2007; Hazekamp et al. 2009; Leghissa et al. 2018a). Performing GC without derivatization requires the calculation of total cannabinoid content from a combination of acidic and neutral cannabinoid content which can be an uncertain process (Dussy et al. 2005). HPLC-DAD and HPLC-UV provide alternatives to GC analysis but these detection techniques lack specificity and sensitivity (Galal et al. 2009; Grauwiler et al. 2007). The literature suggests that HPLC-MS/MS using ESI and APCI methods provide enough specificity and sensitivity to quantify cannabinoid content in all cannabis extracts (Aizpurua-Olaizola et al. 2014; Citti et al. 2018; Grauwiler et al. 2007).

The optimized method is able to separate cannabidivarin, cannabidiolic acid, cannabigerolic acid, cannabigerol, cannabidiol, cannabinol, Δ9-tetrahydrocannabinol, and tetrahydrocannabinolic acid within 10 min. For all target analytes, the %-Bias at the lower and upper calibration range varied from − 1.3 to 10.3% and from − 3.9 to 8.6%, respectively. The most suitable agent for extracting cannabis plant samples was evaluated to be a mixture of acetonitrile and water in a ratio 1:1. The extraction efficiency was more than 95% for all analytes in recreational hemp samples. Stability studies on acidic cannabinoids showed a high likeliness of decarboxylation at 100 °C and aromatization after exposure to UV light, respectively. A modified loss on drying method revealed a moisture content between 4 and 10%. The developed method was successfully applied to measure the cannabinoid content in recreational and forensic hemp samples representing broad range of cannabinoid amounts and patterns.

The main scope of this study was the development and validation of a fast and convenient UV-detector-based RP-HPLC method for the fast quantification of cannabinoids in CBD samples and forensic cannabis samples. The present study examines further pre-analytical conditions and the analytical stability of cannabinoids under different stress conditions. Eight authentic CBD-hemp materials and 12 forensic cannabis samples offering a wide range of cannabinoid patterns were analyzed. Results of the overall THC-content of forensic samples were compared with gas chromatographic method (U.N.O.o. Drugs, Crime 2013), the formerly gold standard in cannabinoid analysis. Additionally, a modified loss on drying method was applied to determine the moisture content of all cannabis samples. Finally, the developed method was transferred easily to an ultra-high performance liquid chromatography (UHPLC) device using know metrics, thus further reducing analysis time from 10 to less than 5 min.

The present work proposes validated methods for the determination of cannabinoids in cannabis samples. The use of RP-HPLC-UV renders this method broadly applicable and allows the detection of acidic precursor in even less time compared to GC-based methods.


We report the successful development and validation of an accurate and broadly applicable reversed-phase high-performance liquid chromatography (RP-HPLC) method coupled to an ultra violet (UV) detector including an optimized extraction procedure for the separation and quantification of eight different cannabinoids.

All preliminary extraction experiments were performed using sample (A). Twenty milligrams of sample was extracted with 2.5 mL solvent in a cooled ultrasonic bath. Afterwards, samples were centrifuged at 10 °C for 15 min at 4000 rpm. Supernatant was filtered using a PFTE filter (0.45 μm, Machery Nagel) prior to analysis and tenfold diluted with solvent. Recovery effect (RE) was tested at QC low level using three independent spiked hop samples.

Reversed-phase chromatography was done using a LaChrom Elite System (Hitachi, Ltd., Tokio, Japan) HPLC system consisting of a LaChrom Elite L-2200 autosampler, a LaChrom Elite L-2130 pump, a LaChrom Elite L-2350 column oven, and a LaChrom Elite L-2420 UV-VIS detector. For peak integration, Agilent EZChrom Elite was used. The final liquid chromatography analysis was performed on a Phenomenex Kinetex XB-C18 column (150 × 4.6 mm, 2.6 μm) applying gradient elution, using pure-water (with 0.1% FA) and acetonitrile (with 0.1% FA) as the organic phase. The injection volume was 15 μL, and the dwell volume of the HPLC system was 1.8 mL. The column-oven temperature was set to 50 °C, and the flow rate was 0.8 mL/min. Monitoring of all cannabinoids was done at λ = 220 nm.

Since centuries, Cannabis sativa L. ( C. sativa ) is used for industrial purposes but it is better known as illegal drug possessing psychotropic properties. However, C. sativa is also a highly decorated medicinal plant for the use as anticancer agent, for neuroprotection and as bone marrow stimulants (Velasco et al. 2016; Machado Rocha et al. 2008). With the legalization of cannabis for therapeutic use, the demand for pure and characterized samples has grown significantly (Corroon and Phillips 2018). Therefore, currently new pharmacopeial monograph s are in development for quality control of C. sativa -based medicinal products (Pavlovic et al. 2018). Besides the medical use, there is an enormous interest from consumers/patients in the utilization of low Δ9-tetrahydrocannabinol (THC) hemp for recreational use. In recent years, a kind of gold-rush fever is observed in Europe and all over the world and many new suppliers entered the market (Pellechia 2018). Since there is a complicated and different legislation for C. sativa products all over Europe, caution for quality control has to be taken. Although there is no upper limit for the cannabidiol (CBD) or cannabidiolic acid (CBDA) content in most European countries, maximum limits of THCor Δ9-tetrahydrocannabinolic acid (THCA) contents vary between 0.1 and 1% within Europe.

Loss on drying experiment of hop was performed in an oven (VD20 Binder, Huber) 105 °C for 2 h (Pharmacopoea europaea (Ph. Eur.) 2.2.32) (Ph. Eur., Loss on Drying (2.2.32) 2018). Cannabis samples were placed in weighing flasks and were dried to constant mass at 60 °C.

Stability of cannabinoids was tested in an oven (VD20 Binder, Huber) at 100 °C and under UV light (Honle, Sol 2, 350–700 nm). For the heat stability experiment, the sample was placed in weighing flasks. For the UV stability, one weighing flask was covered with aluminum foil and the other was exposed to UV light. For both stability experiments, 20 mg of sample was taken from each of the flasks at indicated time points and was analyzed. At indicated time points, 20 mg of sample was taken from each of the vials and analyzed. All stability experiments were performed in duplicate.

HPLC conditions.

Analytical standards were obtained from Lipomed (Reinach, Switzerlanf). Formic acid (FA), methanol (MeOH), ethanol (EtOH), and acetonitrile (ACN) were obtained from Merck (Darmstadt, Germany) and were of LCMS grade. Pure-water was generated from an in-house water purification system from Labtec (Villmergen, Switzerland). For all experiments, Gilson DIAMOND tips were used. Hop strobiles ( Humulus lupulus L . ) were obtained from local pharmacies. CBD-hemp tobacco samples were purchased from several licensed producers within Switzerland. The Forensic Institute Zurich (Zurich, Switzerland) provided 12 forensic cannabis samples.

Extraction efficiency (EE) was determined in triplicate extracting CBD and THC rich samples three times.

Reversed-phase chromatography was done using a HITACHI ChromasterUltra UHPLC system consisting of a 6270 autosampler, a 6310 column oven, a 6170 binary pump, and a 6430 Diode Array Detector. For peak integration, Agilent EZChrom Elite was used. The final liquid chromatography analysis was performed on a Phenomenex Kinetex XB-C18 column (150 × 2.1 mm, 1.7 μm) applying gradient elution, pure-water (with 0.1% formic acid), and acetonitrile (with 0.1% formic acid) as the organic phase. The injection volume was 5 μL, and the dwell volume of the UHPLC system was 0.7 mL. The column-oven temperature was set to 50 °C, and the flow rate was 0.8 mL/min. Monitoring of all cannabinoids was done at λ = 220 nm.

Biosynthetic pathway of selected cannabinoids.


Cannabinoids belong to terpenophenolic compounds and are the main constituents of the cannabis plant. Terpenoids and phenols were also identified in the cannabis plant but are of lower pharmacological importance (Pavlovic et al. 2018). Cannabigerolic acid (CBGA) is the starting point in the biosynthetic pathway of all cannabinoids, which are synthesized in vivo in a carboxylated form (Fig. 1). In the plant, CBDA and THCA are synthesized by enzymatic catalyzed reactions. However, ex vivo stress conditions like heat and UV light decompose these precursors to their decarboxylated form: CBGA ➔ cannabigerol (CBG), CBDA➔ CBD and THCA ➔ THC, respectively (Citti et al. 2018a; Sirikantaramas and Taura 2017). Under UV light, Δ9-THC is further aromatized to cannabinol (CBN). THC and CBD are two main biomarkers in commercial available hemp samples. THC is mostly responsible for psychotropic activities whereas CBD is more anxiolytic and sleep inducing. In comparison to THC, CBD is not considered a controlled substance. Numerous reports have been published for the qualitative and quantitative analysis of cannabinoids in cannabis and its preparations. This study will therefore focus on those substances for possible therapeutic use such as CBD, Δ9-THC, CBG, CBN, cannabidivarin (CBDV), cannabichromene, and tetrahydrocannabivarin (Amada et al. 2013; Thomas et al. 2007). Several comprehensive reviews of the chemical analysis of cannabis plants, corresponding preparations, and forensic specimens were presented in the past (Citti et al. 2018b; Wang et al. 2017; Patel et al. 2017; ElSohly and Salem 2000). The most widespread techniques applied for separation were gas chromatography (GC) with and without derivatization, high-performance liquid chromatography (HPLC), and to a lesser extend supercritical fluid chromatography (Wang et al. 2016; U.N.O.o. Drugs, Crime 2013). The GC method is still officially employed by the authorities for the determination of cannabinoids. But it is obvious that acidic forms are not accessible without prior derivatization, and further conversion of THCA to THC is not quantitative at all (Dussy et al. 2005). Some researchers postulate that an accurate cannabinoid profile should be evaluated by determining the acid and neutral forms separately (Pavlovic et al. 2018; Citti et al. 2018b; Calvi et al. 2018; Ambach et al. 2014). On the other hand, LC-based procedures render the derivatization step superfluous and enable the detection of the heat-labile acid precursor in less time. However, determining chromatographic conditions is more challenging. Additionally, the pre-analytical phase has to be taken into account for method development and validation. Extraction, storage conditions, and stability determination play a pivotal role in the analysis of C. sativa L.-derived products (Dussy et al. 2005; Brighenti et al. 2017; Mudge et al. 2017).

Authentic samples were extracted and quantified applying the developed and validated method. All samples (20 mg) were analyzed in duplicate. The final amount of analyte [%] was calculated using the dilution factor given by the procedure and the weighed amount of plant sample. The determined concentration of the authentic forensic samples was compared to those obtained by established GC-FID-based method as described previously (U.N.O.o. Drugs, Crime 2013). (Details can be found in Additional file 1).

Currently, an increasing demand of cannabis-derived products for recreational and medical use is observed. Therefore, the reliable and fast quantification of cannabinoids in hemp samples is essential for the control of product from Cannabis sativa , L. strains. In general, gas chromatography (GC) is the method of choice for the quantification of cannabinoids whereas this method is time consuming and the detection of acidic precursor is not feasible without derivatization.

Commercially available 1 mg/mL methanolic solutions of all analytes were used as stock solutions for calibration and QC spiking solutions. Four different concentrations of the analytes in the range of 1 –100 μg/mL were chosen. Working solutions were prepared by serial dilution from each stock solution in methanol. QC low and QC high samples were analyzed in duplicate on each of 6 days. Accuracy was given in terms of bias as the percent deviation of the mean calculated concentration compared to the theoretical value. Intra-day and inter-day imprecision was calculated as relative standard deviation (RSD) according to Peters et al. (2009). Phenprocoumon was used as internal standard (IS) at a final concentration of 200 μg/mL.

Reversed phase chromatography RP-HPLC was chosen for the separation of eight cannabinoids. The focus was set on C18 columns, since these were the most commonly used in the literature. Several C18 columns with different eluent compositions, flow rates, and column temperatures were tested (Additional file 1: Table S1). A baseline separation of all analytes was finally achieved using the Kinetex XB-C18 HPLC column (2.6 μm, 150 × 4.6 mm,) with H 2 O/0.1% FA and ACN/0.1% FA as solvent. Flow rate and temperature was set to 0.8 mL/min and 50 °C, respectively (Additional file 1: Table S2). Selected cannabinoids were separated within 10 min under HPLC conditions (Fig. 2). After cleaning and reequilibration, complete run time of this method was 20 min. The resolution of all analytes was at least R s > 1.7 and therefore in the acceptable range for quantification. The asymmetry factor of all peaks is between 1.2 and 1.5 (Table 1). Although for peaks 3 and 4, resolution and asymmetry factor were not in the optimal range, validation data in terms of bias and imprecision for CBGA(3) and CBG(4) were acceptable. The same column was used by Mudge et al. (2017)and De Backer et al. (2009) (Citti et al. 2018b) to separate the same number of cannabinoids, but with separation times of 14 min and 20 min, respectively. As internal standard (IS), phenprocoumon was used. Under selected chromatographic conditions, a clear separation between the IS and all cannabinoids was achieved. Finally, the developed HPLC method was transferred to an UHPLC system coupled to a diode array detector (DAD). The chemistry of the column (Kinetex C18, 1.7 μm, 150 × 2.1 mm) was similar to the HPLC column, and the same mobile phases were used. The injection volume was reduced to 5 μL. Target analytes were separated in less than 5 min (Additional file 1: Figure S5). The resolution of all peaks was above 1.5 and the asymmetry (10%) between 0.9 and 1.1.

Chromatographic method development was performed on a Shimadzu Nexera (Kyoto, Japan) using an Evoke C18, 15 cm x 4.6 mm column packed with 3 µm fully porous particles from Regis Technologies, Inc (Morton Grove, IL, USA). Reversed-phase conditions were screened using different organic modifiers (methanol and acetonitrile) in both isocratic and gradient modes of operation. Acid additives (formic acid and trifluoroacetic acid) were also investigated and found important in achieving good peak shape for the carboxylated species (e.g. CBCA, CBDA, etc.). The conditions that resulted in the most baseline resolved peaks and served as the foundation for further method development are listed in Table 1.

As shown in Figure 2, the addition of ammonium formate to mobile phase A results in reduced retention of the carboxylated cannabinoids while the decarboxylated species remain unaffected, thus baseline-resolving CBGA/CBG and THCVA/CBN. With 5 mM ammonium formate, the retention time of CBNA is shifted to 7.63 minutes and coelutes with exo-THC, an impurity formed in the synthesis of Δ9-THC (Figure 2b). By increasing the concentration to 10 mM ammonium formate, the retention of CBNA is further shifted, causing it to elute earlier than the THC isomers, but THCA-A is shifted into coeluting with CBC (Figure 2c). An intermediate concentration of 7.5 mM ammonium formate was found to provide baseline resolution of all 17 cannabinoids in the test mixture (Figure 2d). Table 2 shows how the retention times of the acidic cannabinoids change when the concentration of ammonium formate buffer in mobile phase A is varied.

Figure 3 plots the effect of varying the percentage and composition of mobile phase B (MPB) on the isocratic resolution of 1:2 Δ9-THC:Δ8-THC using the same Evoke C18, 15 cm x 4.6 mm column. Consider the analysis when performed with H2O/MPB = 10/90. The resolution of Δ9-THC and Δ8-THC is 1.06 when MPB = 100% acetonitrile. When MPB = 100% methanol, the resolution is 2.84. Maximum resolution (Rs = 3.12) is observed when MPB = 15:85 acetonitrile:methanol. That relatively minor improvement in resolution afforded by the blended MPB might suggest pure methanol to be the preferred organic modifier for this analysis, especially given the convenience of using a single solvent over pre-mixing a blend of acetonitrile:methanol or investing in alternative pumping instrumentation (e.g. quaternary pumps). With complex samples, though, care must be taken to observe how a desired change in selectivity can affect other analytes in the separation.

In some assays, analysts are concerned with improving the resolution of certain critical pairs. This may be especially true in cases where one component is far more abundant than the other. In the gradient separations shown in Figure 2, the resolutions between Δ9-THC and Δ8-THC are approximately 1.50. These isomers are neutral, and their retentions are largely unaffected by changes in mobile phase pH or ionic strength. Often, it is possible to improve resolution by running an isocratic analysis and by reducing eluent strength. In the case of Δ9-THC and Δ8-THC, the greatest effect is observed by changing the composition of mobile phase B with various ratios of acetonitrile and methanol.

The interplay of buffer concentration and pH was further investigated with respect to the retention time of one of the carboxylated species, CBNA. The conditions are outlined in Table 3. In the first three cases, mobile phase A was prepared with 0.1% formic acid and ammonium formate concentration of 0 mM, 5 mM, and 10 mM. The unadjusted pH values were measured as 2.7, 3.1, and 3.5, respectively. As described above, retention of CBNA decreased with increased buffer concentration (8.31 min, 7.76 min, and 7.34 min). In the fourth case, 10 mM ammonium formate was used in mobile phase A and no formic acid was used in either mobile phase A or B. CBNA is ionised under these conditions, and its retention was reduced to 1.64 minutes. In the fifth case, 10 mM ammonium formate was used and pH was adjusted with formic acid to a value of 3.1 in order to match the pH of mobile phase A when prepared with 0.1% formic acid and 5 mM ammonium formate, and the retention time of CBNA was 7.36 minutes. In the final case, 10 mM ammonium formate was adjusted to a pH value of 2.8, and the retention time was 7.52 minutes. Thus, it can be seen that the retention of the carboxylated, ionisable cannabinoids is a complex function of eluotropic strength, pH (and the corresponding protonation state of the analyte), and buffer concentration/ionic strength.

Figure 2a shows the baseline-subtracted chromatogram for the separation of the 17 cannabinoid test mixture using the conditions listed in Table 1. Baseline resolution is achieved for each of the component peaks with the exceptions of CBGA and CBG (Rs = 1.40), THCVA and CBN (Rs = 1.42), and the coelution of Δ8-THC and CBNA at 8.20 minutes. In an effort to improve the resolution of these pairs, the effect of adding ammonium formate, the ammonium salt of formic acid, to mobile phase A in concentrations ranging between 5 and 10 mM was investigated. The addition of ammonium formate to formic acid mobile phases increases the ionic strength as well as slightly raises the pH [6,7]. With 0.1% formic acid and ammonium formate concentrations of 0 mM, 5 mM, and 10 mM, the pH values of mobile phase A were measured to be 2.7, 3.1, and 3.5, respectively.

The global cannabis industry is growing rapidly, with many countries and US states adding regulatory frameworks for medical and recreational cannabis programs [1,2]. Quality control is an essential component in protecting the health and safety of the consumer in this emerging market, and there is increasing demand upon cannabis testing laboratories for analytical determination of multiple cannabinoids along with potential contaminants such as pesticides, mycotoxins, heavy metals, etc. Current regulations surrounding potency vary by jurisdiction, but usually require testing for the active forms of tetrahydrocannabinol (THC) and cannabidiol (CBD). In addition, many require testing for the acid forms, tetrahydrocannabinolic acid (THCA) and cannabidiolic acid (CBDA), along with other cannabinoids like cannabigerol (CBG), cannabigerolic acid (CBGA), tetrahydrocannabivarin (THCV), cannabichromene (CBC), cannabicyclol (CBL), and cannabinol (CBN). As regulations evolve, and as research interests in minor cannabinoids expand, it is important to have robust analytical methods in place that are capable of meeting those needs.

Seventeen analytical reference cannabinoid standards (1 mg/mL) were acquired from Cerilliant (Round Rock, TX, USA) and combined to a final component concentration of approximately 59 µg/mL in 53:47 methanol:acetonitrile. The mixture was composed of Δ8-tetrahydrocannabinol (Δ8-THC), Δ9-tetrahydrocannabinol (Δ9-THC), cannabichromene (CBC), cannabichromenic acid (CBCA), cannabicyclol (CBL), cannabidiol (CBD), cannabidiolic acid (CBDA), cannabidivarin (CBDV), cannabidivarinic acid (CBDVA), cannabigerol (CBG), cannabigerolic acid (CBGA), cannabinol (CBN), cannabinolic acid (CBNA), exo-tetrahydrocannabinol (exo-THC), tetrahydrocannabinolic acid A (THCA-A), tetrahydrocannabivarin (THCV), and tetrahydrocannabivarinic acid (THCVA). The molecular structures of these cannabinoids are shown in Figure 1.

Acetonitrile and methanol are two of the most common organic modifiers used in reversed-phase HPLC, and many studies have detailed the differing and often complementary selectivities that they provide. Fundamental understandings of the solute-mobile phase, solute-stationary phase, and stationary phase-mobile phase molecular interactions can inform the strategies used in HPLC method development [8]. For instance, it has been noted that, depending on the modifier used and how it has partitioned or adsorbed into the stationary phase, differences in hydrophobicity, hydrogen-bonding, and dipole-type interactions can be observed [9,10]. When developing methods and selecting appropriate mobile phases, it can be useful to consult Snyder’s solvent selectivity triangle, which plots solvents according to their acidic, basic, and dipolar properties [11,12,13]. Solvents that feature one of those properties more prominently than the other two can be readily identified from the plot. For example, methanol has acidic properties, and acetonitrile has dipole properties. Since they are miscible, they can be mixed in any ratio to achieve intermediate or new solvent properties.

A brief example serves to illustrate that several parameters should be considered when developing a chromatographic method for the resolution of complex samples involving key critical pairs. Consider again the separation of 1:2 Δ9-THC:Δ8-THC in the presence of cannabicyclol (CBL). In Figure 3 it can be seen that the resolution of the THC isomers is superior with pure methanol than with pure acetonitrile as the organic modifier. As shown in Figure 4, though, if CBL is present, it coelutes with Δ8-THC in H2O/methanol = 10/90. CBL elutes well away from the critical pair if pure acetonitrile is used, but the THC isomers are insufficiently resolved (Rs = 1.06). A 50:50 blend of acetonitrile:methanol provides good resolution, with Rs > 2.5 for both pairs. So, while binary mobile phase systems are very common in reversed-phase HPLC separations, ternary mobile phases can provide access to unique selectivities based on the combination of acidic, basic, and dipolar properties of the mobile phases used.

Upon evaluating the molecules of interest in terms of their charges, polarities, and other functionalities, chromatographic method developers turn their focus to column and solvent selection, pH conditions, buffer selection and concentration, temperature, etc. Specific approaches can differ depending upon the primary goals of a separation. For example, if comprehensive characterisation of a complex sample is desired, approaches to maximising overall separation at the expense of analysis time may be acceptable. If, on the other hand, resolution of only a particular critical pair is required, speed and selectivity (for the crucial pair) may be the primary focus. With these concerns in mind, we set out to develop an HPLC method capable of fully resolving 17 cannabinoids in a minimal amount of time. Additionally, a second objective concerning the improved resolution of a specific critical pair of THC isomers (Δ8-THC and Δ9-THC) was explored.

It should be noted that since ammonium formate is added only to the aqueous component of the mobile phase, the total ionic strength changes throughout the gradient runtime. For example, when 7.5 mM ammonium formate in mobile phase A is used in the gradient listed in Table 1, the total concentration on the column changes from 1.875 mM to 0.75 mM over the course of the 15 minute run. Nevertheless, with approximately 5 minute re-equilibration, run-to-run results were found to be reproducible. With real world samples, such as plant extracts, matrix effects may prove to be a concern. Although not determined here, the gradient method may permit some flexibility to build in a weaker solvent hold prior to the 75-90% MPB gradient in order to clear matrix interferences. Analysts must also assess whether other endogenous cannabis compounds, such as terpenes and terpenoids, potentially interfere with identification of cannabinoids.

Historically, gas chromatography/mass spectrometry (GC/MS) has been used for the separation and quantification of cannabinoids and other compounds of interest in cannabis analysis. With GC, though, care must be taken to avoid decarboxylation of acidic species during the heated injection. High pressure liquid chromatography (HPLC) methods permit analysts to eschew many sample preparation and derivatisation steps and have become the preferred approaches to cannabis potency analysis [3,4,5]. In general, all approaches to HPLC method development look to balance several elements, among which are the ultimate goals of the analysis, resolution of target compounds and potential interferences, speed, and assay robustness.

To recap, we developed an HPLC method that fully resolves 17 cannabinoids by using screening runs that altered concentrations of organic and acid modifiers and provided the foundation for further development. The addition of ammonium formate to mobile phase A gave a means to shift the retentions of the carboxylated species relative to the neutral ones, and an optimised concentration allowed for the baseline resolution of all cannabinoids in the test mixture. In addition, the use of a ternary mobile phase system (water, methanol, acetonitrile) was shown to improve the resolution of THC isomers while permitting the flexibility to avoid potential interferences.