Analytical Techniques for Phytocannabinoid Profiling of Cannabis and Cannabis-Based Products—A Comprehensive Review

Abstract

Cannabis is gaining increasing attention due to the high pharmacological potential and updated legislation authorizing multiple uses. The development of time- and cost-efficient analytical methods is of crucial importance for phytocannabinoid profiling. This review aims to capture the versatility of analytical methods for phytocannabinoid profiling of cannabis and cannabis-based products in the past four decades (1980–2021). The thorough overview of more than 220 scientific papers reporting different analytical techniques for phytocannabinoid profiling points out their respective advantages and drawbacks in terms of their complexity, duration, selectivity, sensitivity and robustness for their specific application, along with the most widely used sample preparation strategies. In particular, chromatographic and spectroscopic methods, are presented and discussed. Acquired knowledge of phytocannabinoid profile became extremely relevant and further enhanced chemotaxonomic classification, cultivation set-ups examination, association of medical and adverse health effects with potency and/or interplay of certain phytocannabinoids and other active constituents, quality control (QC), and stability studies, as well as development and harmonization of global quality standards. Further improvement in phytocannabinoid profiling should be focused on untargeted analysis using orthogonal analytical methods, which, joined with cheminformatics approaches for compound identification and MSLs, would lead to the identification of a multitude of new phytocannabinoids.

Full text


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Citation: Stefkov, G.; Cvetkovikj
Karanfilova, I.; Stoilkovska
Gjorgievska, V.; Trajkovska, A.;
Geskovski, N.; Karapandzova, M.;
Kulevanova, S. Analytical Techniques
for Phytocannabinoid Profiling of
Cannabis and Cannabis-Based
Products—A Comprehensive Review.
Molecules 2022,27, 975. https://
doi.org/10.3390/molecules27030975
Academic Editors: Neda
MIMICA-DUKI ´
C and Biljana Božin
Received: 24 November 2021
Accepted: 9 January 2022
Published: 1 February 2022
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4.0/).
molecules
Review
Analytical Techniques for Phytocannabinoid Profiling
of Cannabis and Cannabis-Based Products—A
Comprehensive Review
Gjoshe Stefkov 1, Ivana Cvetkovikj Karanfilova 1, * , Veronika Stoilkovska Gjorgievska 1, Ana Trajkovska 1,
Nikola Geskovski 2, Marija Karapandzova 1and Svetlana Kulevanova 1
1Institute of Pharmacognosy, Faculty of Pharmacy, Ss. Cyril and Methodius University, Bul. Majka Tereza 47,
1000 Skopje, North Macedonia; gost@ff.ukim.edu.mk (G.S.); vsgjorgievska@ff.ukim.edu.mk (V.S.G.);
ana.trajkovska@ff.ukim.edu.mk (A.T.); Marija_Karapandzova@ff.ukim.edu.mk (M.K.);
svku@ff.ukim.edu.mk (S.K.)
2
Institute of Pharmaceutical Technology, Faculty of Pharmacy, Ss. Cyril and Methodius University, Bul. Majka
Tereza 47, 1000 Skopje, North Macedonia; ngeskovski@ff.ukim.edu.mk
*Correspondence: ivanacvetkovikj@gmail.com; Tel.: +389-2-3126-032 (ext. 103); Fax: +389-2-3123-054
Abstract:
Cannabis is gaining increasing attention due to the high pharmacological potential and
updated legislation authorizing multiple uses. The development of time- and cost-efficient analytical
methods is of crucial importance for phytocannabinoid profiling. This review aims to capture the
versatility of analytical methods for phytocannabinoid profiling of cannabis and cannabis-based
products in the past four decades (1980–2021). The thorough overview of more than 220 scientific
papers reporting different analytical techniques for phytocannabinoid profiling points out their
respective advantages and drawbacks in terms of their complexity, duration, selectivity, sensitivity
and robustness for their specific application, along with the most widely used sample preparation
strategies. In particular, chromatographic and spectroscopic methods, are presented and discussed.
Acquired knowledge of phytocannabinoid profile became extremely relevant and further enhanced
chemotaxonomic classification, cultivation set-ups examination, association of medical and adverse
health effects with potency and/or interplay of certain phytocannabinoids and other active con-
stituents, quality control (QC), and stability studies, as well as development and harmonization of
global quality standards. Further improvement in phytocannabinoid profiling should be focused
on untargeted analysis using orthogonal analytical methods, which, joined with cheminformatics
approaches for compound identification and MSLs, would lead to the identification of a multitude of
new phytocannabinoids.
Keywords: Cannabis sativa; sample preparation; analysis; quality control
1. Introduction
Cannabis sativa L. (C. sativa L.),from the family Cannabaceae, is the most widely
cultivated, trafficked, consumed and investigated, yet most notorious and controversial,
plant in the world [
1
,
2
]. It is one of the oldest known crops to humanity, with first records
of use dating to 3000 B.C. [
3
], and one of the most commonly used plants for industrial
and medical purposes, with a global legal market expected to reach 147 billion USD by the
end of 2027 but also the world’s most widespread drug of abuse [
1
], comprising around
200 million global users.
1.1. Botany of C. sativa
C. sativa is an annual dioecious plant with histaminate male and pistillate female
flowers on separate plants. It grows up to 5 m height, with serrated leaves with a distinct
vein pattern that extends to their tips [
4
]. The inflorescences of the female plants produce
Molecules 2022,27, 975. https://doi.org/10.3390/molecules27030975 https://www.mdpi.com/journal/molecules

Molecules 2022,27, 975 2 of 42
several individual bunches of flowers, a large cluster on the upper torso and various small
clusters in each branch, covered by trichome glands containing resin rich in phytocannabi-
noids and terpenoids. Phytocannabinoids are mainly accumulated in the glands of both
capitate stalked and capitate-sessile trichomes, but mostly in the latter [
4
]. C. sativa was
first classified in 1753 by the Swedish botanist Carolus Linnaeus (Carl Von Linné). More
than 2 centuries later, despite its wide use, C. sativa is considered a plant with inconclusive
taxonomic organization and evolutionary history that are the subject of constant scientific
debates [
3
,
5
,
6
]. The United Nations Office on Drugs and Crime (UNODC) [
5
] considers that
the plant has only one recognized species, C. sativa L. [
5
–
7
], although other reported taxa for
this genus, such as C. sativa subsp. sativa,C. sativa subsp. indica,
C. sativa subsp. ruderalis
,
C. sativa subsp. spontanea, and C. sativa subsp. kafiristanca are currently recognized as
subspecies [
3
,
5
–
7
]. Today, due to the difficulty in distinguishing cannabis species either
morphologically or chemically, and given the continuous changes occurring in subspecies
according to the cultivation environment, the designation C. sativa is considered suitable
for all plants from the genus [3,5].
1.2. Phytocannabinoids
A wide variety of chemical constituents, i.e., more than 750 compounds, have been
identified in C. sativa, belonging to the various classes of natural products such as mono-
and sesquiterpenes, flavonoids, steroidsand nitrogen-containing compounds [
8
]. Among
them, more than 100 are classified as phytocannabinoids [
9
], the family of plant-derived
C
21
or C
22
terpenophenolic compounds, including analogues and metabolites. They are
synthesized in secreting cells of glandular trichomes in a biosynthetic pathway from ger-
anyl pyrophosphate (GPP) as the parent precursor of both phytocannabinoids and terpenes.
By coupling with olivetolic acid or divarinic acid, C
5
or C
3
cannabinoid acids are produced,
respectively [
10
]. Most phytocannabinoids naturally occur as acidic precursors in unfertil-
ized female flowers prior to senescence [
2
], of which delta-9-tetrahydrocannabinolic acid
(
∆9
-THCA), cannabidiolic acid (CBDA), cannabigerolic acid (CBGA) and cannabichromenic
acid (CBCA) are most abundant, with cannabidivarinic acid (CBDVA) and tetrahydro-
cannabivarinic acid (THCVA) as less abundant [
11
]. Lower phytocannabinoids content is
found in leavesand stems, while absent from roots and seeds.
Cannabinoid acids are converted to their neutral counterparts by decarboxylation in-
duced by heat or age. Cannabidiol (CBD), the first cannabinoid was isolated from
C. sativa
in
1963 [
12
], -delta-9–tetrahydrocannabinol (
∆9
-THC), the second cannabinoid from
C. sativa
in 1964 [
13
], delta-8-tetrahydrocannabinol (
∆8
-THC), the third, cannabigerol (CBG), iso-
lated in 1964, followed by cannabichromene (CBC), isolated in 1966 [
14
], cannabidivarin
(CBDV) [
15
] and tetrahydrocannabivarin (THCV) [
16
], which are formed from CBDA,
∆9
-THCA, CBGA, CBCA, CBDVA and THCVA, respectively [
17
]. Oxidative degradation
of
∆9
-THC results in the formation of cannabinol (CBN), while isomerization leads to
the formation of the more stable, but less active isomer of
∆9
-THC,
∆8
-THC.
∆9
-THCA
can degrade to cannabinolic acid (CBNA) and further to CBN. Molecular and structural
formula, molecular mass and major fragments as well as UV-VIS spectrum and mid-IR
spectra of major phytocannabinoids are presented in Table 1and Figure 1.

Molecules 2022,27, 975 3 of 42
Table 1. Formula, MS and UV data of major phytocannabinoids.
Compound [18] Molecular Formula and Mr [18]
[M-H]−
[MF1-H]−
[MF2-H]−
[MF3-H]−
[19]
Structure [19]
UV-VIS Spectra [18]
Acidic HPLC Systems/
Basic HPLC Systems
1H NMR in Deuterated
Chloroform [4,20]
∆9-THC C21H30 O2
314.472
C21H29 O2, 313.2173
C16H21 O2, 245.1547
C12H15 O2, 191.1078
C11H15 O22, 179.1067
R1-C5H11, R2-H, R3-H
3.20 (1H, dm, 10.9 Hz) 6.31 (1H, q,
1.6 Hz) 1.68 (3H, s) 2.16 (2H, m) 1.90
(1H, m), 1.40 (m) 1.69 (m) 1.41 (3H, s)
1.09 (3H, s) 6.14 (1H, d, 1.6Hz) 6.27 (1H,
d, 1.6 Hz) 2.42 (2H, t, 7.3 Hz, 1.6 Hz),
1.55 (2H, q, 7.8 Hz) 1.29 (m) 1.29 (m) d
0.87 (3H, t, 7.0 Hz) 4.87 (1H, s)
∆8-THC C21H30 O2
314.472
C21H29 O2, 313.2173
C16H21 O2, 245.1547
C12H15 O2, 191.1078
C11H15 O2, 179.1067
-
THV C19H26 O2
286.418
C19H25 O2, 285.1860
C14H17 O2, 217.1234
C10H11 O2, 163.0765
C9H11O2, 151.0765
R1-C3H17, R2-H, R3-H
-
-
CBD C21H30 O2
314.472
C21H29 O2, 313.2173
C16H21 O2, 245.1547
C12H15 O2, 191.1078
C11H15 O2, 179.1067
R1-C5H11, R2-H, R3-H
3.90 (1H, dm, 11.8Hz) 5.57 (1H, s) 2.21
(1H, m), 2.09 (1H, m) 1.84 (m) 2.40 (m)
1.79 (3H, s) 4.64 (trans, 1H, m), 4.54 (cis,
1H, m) 1.66 (3H, s) 6.26 (1H, brs) 6.16
(1H, brs) 2.43 (2H, t, 7.5Hz) 1.55 (2H, q,
7.6Hz) 1.29 (m) 1.29 (m) 0.88 (3H, t,
6.8Hz) 5.99 (1H, s) 5.02 (1H, s)

Molecules 2022,27, 975 4 of 42
Table 1. Cont.
Compound [18] Molecular Formula and Mr [18]
[M-H]−
[MF1-H]−
[MF2-H]−
[MF3-H]−
[19]
Structure [19]
UV-VIS Spectra [18]
Acidic HPLC Systems/
Basic HPLC Systems
1H NMR in Deuterated
Chloroform [4,20]
CBN C21H26 O2
310.440
C21H25 O2, 309.1860
C19H19 O2, 279.1391
C12H11 O2, 171.0815
R1-C5H11, R2-H, R3-H, R4-H
8.16 (1H, s) 2.38 (3H, s) 7.07 (1H, d,
7.9Hz) 7.14 (1H, d, 7.9Hz) 1.60 (6H, s)
1.60 (6H, s) 6.29 (1H, d, 1.1Hz) 6.44 (1H,
d, 1.1Hz) 2.50 (2H, t, 7.5Hz) 1.63 (m)
1.32 (m) g 1.32 (m) g 0.89 (3H, t,
6.8Hz)5.13 (1H, s)
CBG C21H32 O2
316.488
C21H31 O2, 315.2329
C16H21 O2, 245.1547
C12H15 O2, 191.1078
C11H15 O2, 179.1067
R1-C5H11, R2-H, R3-H, R4-H
6.26 (2H, s) d 6. (2H, s) d 3.41 (2H, d,
7.0 Hz) 5.29 (1H, m) 1.82 (3H, s) 2.09
(4H, m) 2.09 (4H, m) 5.07 (1H, m) 1.60
(3H, s) 1.69 (3H, s) 2.45 (2H, t, 7.5 Hz)
1.56 (2H, q, 7.8 Hz) 1.33 (4H, m) 1.33
(4H, m) 0.90 (3H, t, 6.9 Hz) 5.36 (2H, s)
CBC C21H30 O2
314.172
C12H29 O2, 313.2173
C16H19 O2, 243.1391
C12H15 O2, 191.1078
C11H15 O2, 179.1067
R1-C5H11, R2-H
N/A
CBL C21H30 O2
314.472
C21H29 O2, 313.2173
C16H19 O2, 243.1391
C12H15 O2, 191.1078
C11H15 O2, 179.1067
N/A N/A

Molecules 2022,27, 975 5 of 42
Table 1. Cont.
Compound [18] Molecular Formula and Mr [18]
[M-H]−
[MF1-H]−
[MF2-H]−
[MF3-H]−
[19]
Structure [19]
UV-VIS Spectra [18]
Acidic HPLC Systems/
Basic HPLC Systems
1H NMR in Deuterated
Chloroform [4,20]
∆9-THCA C22H30 O24
358.482
C22H29 O4, 357.2071
C21H30 O2, 245.1547
C12H15 O2, 191.1078
C11H15 O22, 179.1067
R1-C5H11, R2-COOH, R3-H
3.23 (1H, dm, 7.0 Hz), 6.39(1H, brs), 1.68
(3H, s), 2.17 (2H, m) 1.92 (1H, m) 1.35
(m) 1.67 (m) 1.44 (3H, s) 1.11 (3H, s) 6.26
(1H, s) 2.94 (1H, m) 2.78 (1H, m) 1.57
(2H, 1.35 (m) 1.35 (m) 0.90 (3H, t,
6.9 Hz) 12.19 (1H, s)
∆9-THCA-C4 C21H28 O4
344.455
C21H27 O4, 343.1915
C15H19 O2, 231.1391
C11H13 O2, 177.0921
C10H13 O2, 165.0921
R1-C4H9, R2-COOH, R3-H
N/A
THVA C20H26 O4
330.428
C20H25 O4, 329.1758
C14H17 O2, 217.1234
C10H11 O2, 163.0765
C9H11O2, 151.0765
R1-C3H7, R2-COOH, R3-H
N/A

Molecules 2022,27, 975 6 of 42
Table 1. Cont.
Compound [18] Molecular Formula and Mr [18]
[M-H]−
[MF1-H]−
[MF2-H]−
[MF3-H]−
[19]
Structure [19]
UV-VIS Spectra [18]
Acidic HPLC Systems/
Basic HPLC Systems
1H NMR in Deuterated
Chloroform [4,20]
CBDA C22H30 O4
358.482
C22H29 O4, 357.2071
C16H21 02, 245.1547
C12H15 02, 191.1078
C11H15 02, 179.1067
R1-C5H11, R2-COOH, R3-H
N/A
CBNA C22H26 O4
354.450
C22H25 O4, 353.175
C19H19 02, 279.1391
C12H11 02, 171.0815
C21H25 02, 309.1860
R
1
-C
5
H
11
, R
2
-COOH, R
3
-H, R
4
-H
N/A
CBGA C22H32 O4
360.498
C22H31 O4, 359.2228
C16H21 O2, 245.1547
C12H15 O2, 191.1078
C11H15 O2, 179.1067
R
1
-C
5
H
11
, R
2
-COOH, R
3
-H, R
4
-H
N/A

Molecules 2022,27, 975 7 of 42
Table 1. Cont.
Compound [18] Molecular Formula and Mr [18]
[M-H]−
[MF1-H]−
[MF2-H]−
[MF3-H]−
[19]
Structure [19]
UV-VIS Spectra [18]
Acidic HPLC Systems/
Basic HPLC Systems
1H NMR in Deuterated
Chloroform [4,20]
CBCA C22H30 O4
358.482
C22H29 O4, 357.2071
C16H19 O2, 243.1391
C12H15 O2, 191.1078
C11H15 O2, 179.1067
R1-C5H11, R2-COOH,
N/A
CBLA C22H30 O4
358.482
-
-
-
-
N/A

Molecules 2022,27, 975 8 of 42
Figure 1.
Mid
−
IR spectra from the main cannabinoids; THCA, THC, CBDA, CBD were adapted
from [21,22]; CBN, CBGA and CBG were adapted from [18].
1.3. Use of C. sativa
For more than 12,000 years, Cannabis spp. is used as a source of textile fiber and food
worldwide [
23
]. The earliest data for medical use of Cannabis by the Assyrians goes back
to 3000 B.C. Follow-up records date from around 2700 B.C. in China, where C. sativa was
used as a medicine for menstrual fatigue, rheumatism, malaria, constipation and other
conditions. It was later used by other ancient civilizations-the Egyptians (ca. 1700 B.C.),
the Indians (ca. 1600 B.C.), the Persians (ca. 750 B.C.), the Greeks and the Romans (ca.
450 B.C.
). Historically, Cannabis was also used for additional indications such as glaucoma,
anal fissures, diarrhoea, as obstetric aid and as anxiety relief [
23
]. The plant was introduced
to the modern western medicine in the early 19th century, and was mainly indicated for
treating pain, glaucoma, nausea, depression and neuralgia [24].
Today, medical use of C. sativa includes multiple indications supported by reliable
clinical evidence. Such are the treatment of chronic pain, mutiple sclerosis, resistant epilepsy
and chemotherapy-associated nausea and vomiting, apetite and weight loss associated with
HIV/AIDS, Tourette syndrome, anxiety disorders, sleep disorders, post-traumatic stress
disorder and schizophrenia [
1
,
25
,
26
]. Multiple additional health benefits of C. sativa extracts
are reported in
in vitro
and
in vivo
trials, such as lowering blood cholesterol, triglycerides
and blood pressure, antioxidant and antimicrobial activity [
27
]. C. sativa and its extracts
are also used in the treatment of dermatitis and degenerative imunological diseases, both
as supplements and as traditional medications [
25
]. Cannabis oils, oral solutions, oil-like
concentrates and tinctures used orally and sublingually, lotions, balms, creams, bath salts,
salves, gels, patches and other topical products as well as rectal and vaginal products
(suppositories, tablets) are the most common pharmaceutical dosage forms employed [
26
].
Moreover, hemp is additionally used in food and beverage production, as hempseeds
are shown to have great nutritional value: high content of protein, containingnine essential
amino acids, dietary fibers and an ideal ratio of
ω
-6:
ω
-3 fatty acids (3:1). The same applies
to hempseed flour and oils, which have high content of proteins, insoluble fibers and
polyunsaturated fatty acids. The “cannabis edibles” are the latest type of cannabis-based
products that recently became popular. Chewing gums, lollipops, caramel hard candy,
berry gummies, lozenges, candy bars, jam, tea, soda, coffee, water, honey, etc., containing
∆9-THC- and/or CBD-dominant extracts and concentrates are widely marketed [28].

Molecules 2022,27, 975 9 of 42
Today, cannabis is the most commonly used illicit drug worldwide, despite the strict
international control for more than eight decades [9].
1.4. Legal Aspect of C. sativa
C. sativa and/or cannabis-based products have been legalized for medical use in
41 countries
(23 in Europe) between 2012–2021. As C. sativa and cannabis-based products
are classified based on the
∆9
-THC content with psychotropic properties, and most of
them contain drug-type C. sativa extracts, various legal limitations in many countries
still exist. In total, 50 countries in Europe, Asia, North and South America use the plant
for industrial purposes [
29
]. Cultivation and supply of 69 C. sativa varieties [
30
] with
∆9
-THC content not exceeding 0.2% is legal in EU [
31
], with some exclusions (Czech
Republic and Austria, <0.3%, Switzerland 1.0%).The industrial use of C. sativa is focused on
production of >2500 products used in agriculture, textile production, recycling, automotive
industry, furniture production, paper industry, production of construction materials, energy
production, personal care products and medical supplements [29].
Cultivation and use of marijuana, the crude drug derived from C. sativa for recreational
purposes, is not legalized in Europe, but it is decriminalized in 32 countries worldwide
(
16 in Europe
), with various limitations regarding the amount of dry marijuana, number of
cultivated plants and punishing public consumption [32].
Several Pharmacopoeias, including the German pharmacopoeia (DAB), Swiss pharma-
copoeia (Ph.Helv.), European Pharmacopoeia (Ph.Eur.), and the American Herbal Pharma-
copoeia (AHP) comprise monographs defining Cannabis flowers (“Cannabis inflorescence”)
as herbal substance that consists of whole or crushed, flowering, dried shoot tips of the
female plants of C. sativa L. (Cannabaceae) that contain 90.0–110.0% of the amounts of
phytocannabinoids specified in the label, such as
∆9
-THC and CBD, as well as cannabinoid
carboxylic acids such as
∆9
-THCA and CBDA, calculated as
∆9
-THC or CBD, based on the
dried drug [33–36].
Due to legal issues, chemotype classification of C. sativa is nowadays much more
significant. Depending on the content of
∆9
-THC and CBD, authorities have classi-
fied generally three chemotypes of C. sativa L.:
∆9
-THC-predominant type, i.e., drug-
type (CBD/
∆9
-THC = 0.00–0.005), CBD-predominant type, i.e., fiber type (“hemp” type)
(
CBD/∆9-THC = 15.0–25.0
) and intermediate chemotype (CBD/
∆9
-THC = 0.5–3.0) [
37
].
AHP proposes more comprehensive chemotype classification. Six chemotypes are defined:
(1) type I-Drug (0.5–15%
∆9
-THC; 0.01–0.16%CBD and 50:1
∆9
-THC/CBD ratio); type
II-Intermediate (0.5–5%
∆9
-THC; 0.9–7.3% CBD and 0.25/~2
∆9
-THC/CBD ratio); type
III-Fiber (0.05–0.70%
∆9
-THC; 1.0–13.6% CBD and <1:5
∆9
-THC/CBD ratio); type IV-CBG
(<0.05%
∆9
-THC; <0.5% CBD); type V-non-cannabinoid (
∆9
-THC = 0; CBD = 0) [
33
]. In
addition, DAB defines a discontinued cannabis extract-Cannabis extractum normatum as
an extract from whole or shredded, flowering, dried shoot tips of the female plants of
C. sativa L.
that contains
∆9
-THC at least 1% and at most 25% (m/m) for the extract and 90.0
to 110.0% of the nominal salary specified in the label, and CBD maximum 10.0% (m/m)
for the extract and 90.0 to 110.0% of the nominal content stated in the label [
38
]. High
within-chemotype variability is recorded, due to changes in growing and storage condi-
tions, such as environmental factors of cultivation (climates and elevation of cultivated
area), the development stage of the plant at harvest time and genetic characteristics of
seed-stocks [2,17].
1.5. Incentive for Investigating Phytocannabinoids in C. sativa and Cannabis-Based Products
Under the pressure of its criminal association, the chemical constitution, pharma-
cological effects, genetic structure, evolutionary and domestication history of remained
poorly understood until the last decade of the 20th century. Authorized investigations
related to C. sativa were either forensic studies to aid law enforcement or medical and
social research specifically intended to document and reduce harmful effects [
6
]. Since the
last decade of the 20th century, a great urge for more thorough investigation of C. sativa

Molecules 2022,27, 975 10 of 42
appeared, mainly as a result of the resurrection of production of C. sativa for non-narcotic
and medical purposes and the growing tolerance of the extremely widespread recreational
use. This increased attention C. sativa and cannabis-based products gained due to their high
pharmacological potential, updated legislation authorizing many different uses, and, thus,
the emerging need to control their quality. This imposes a great challenge for academics,
particularly in the field of natural products, from which a contribution to improve and
standardize the extraction and characterization of the bioactive compounds from C. sativa
species is expected.
Scientific and technological development in regards to C. sativa began, highlighting the
need of sensitive, specific and robust analytical methods for identification and quantifica-
tion of the active constituents of C. sativa. Chemical profiling of the plant became extremely
relevant, since the acquired knowledge further enhanced: (1) chemotaxonomic classifica-
tion; (2) cultivation set-ups examination, and thus adjustment of cultivation conditions
and breeding methodologies in order to produce C. sativa varieties with fit-for-purpose
physicochemical properties; (3) investigation of potency of seized samples, thus discover-
ing sources of interconnected illegal production and trafficking; (4) association of medical
and adverse health effects with potency and/or interplay of certain phytocannabinoids
and other active C. sativa constituents and (5) QC of medical cannabis and final medical
cannabis-based products and potency examination [1,2].
The multitude of cannabis-based products including cannabis extracts, oils, resins,
pharmaceutical dosage forms, cannabis-infused edibles and beverages are obliged to
comply with the national regulatories’ quality control regulations, especially in terms
of phytocannabinoids content. Accurate qualitative and quantitative analyses of the phy-
tocannabinoids content and chemical profile in cannabis plants are extremely relevant, in
order to associate medicinal and possible adverse health effects with the potency of certain
phytocannabinoids and other compounds, such as terpenoids [
2
]. One of the most relevant
problems in analytical determinations for QC, especially when there are legal problems
related with quantitation, such as for cannabis, relates to the proficiency of laboratories.
Both qualitative and quantitative determinations require carrying out of standardized
assays that meet the analytical criteria approved by the relevant control authorities.
This review aims to capture the versatility of analytical methods for natural phyto-
cannabinoids profiling in cannabis and cannabis-based products in the past four decades
(1980–2021). As such, this thorough overview is first of its kind. Other most recent reviews
cover either a shorter time period, i.e., 2002–2016 [
1
], 2010–2016 [
2
], 2009–2019 [
39
], fo-
cus on both plant materials and biological matrices [
1
,
39
], describe multiple [
2
] or single
(GC) [
39
] instrumental analytical platforms, omit cannabis-based products [
4
] or, apart
from phytocannabinoids, also include profiling of other bioactive C. sativa constituents [
1
].
2. Analytical Methods for Phytocannabinoid Profiling
The overview of the analytical methods for phytocannabinoid profiling used in phyto-
cannabinoid profiling of cannabis and cannabis-based products is schematically presented
in Figure 2.
2.1. Sample Preparation Techniques
Rapid and simple extraction methods are essential for time- and material-efficient high-
throughput phytocannabinoid profiling. Optimization of the three key parameters during
extraction is crucial for the overall analytical method [
40
], which are: (1) the granulometry of
the solid sample; (2) the system temperature and (3) the affinity of the extraction liquid towards
compounds of interest [
41
]. C. sativa and cannabis-based products are very complex and
inhomogeneous matrices, as their different parts may have different cannabinoid profiles
due to the variety of phytocannabinoids, terpenes and other volatile compounds and
high sugar and fat content. Thus, extraction of phytocannabinoids from plant material
and cannabis-based products in an efficient and consistent manner, with acquisition of
accurate and reliable potency data can be a challenging task. In addition, there are no

Molecules 2022,27, 975 11 of 42
standardized preparation procedures for the hemp-based infusions (hemp leaf, hemp-
based tea mixtures), and the cannabinoid content could be significantly affected by the
infusion preparation procedure [
42
]. Finally, there is large and unpredictable variability of
the average composition as a result of genetic and environmental differences, making the
efforts for standardization of sample preparation techniques an ultimatum [43].
Initial steps of sample preparation include mechanical preparation aiming to increase
the contact surface between the solvent and the active ingredients. It is of special importance
for cannabis plant material, since, despite that most of the active resin is claimed to occur in
the superficial glandular trichomes, significant amounts are found in non-glandular tissues.
Consequently, immersion of unbroken fresh plant material would give unsatisfactory ex-
traction [
44
]. Cannabis or its resins are reduced to small pieces by a grater [
5
] or spatula [
45
],
grinded or pulverized, while cannabis oils are directly proceeded to instrumental analysis.
Figure 2.
Analytical methods used in phytocannabinoid profiling of cannabis and cannabis-
based products.
Manual pulverization and homogenization of the dried plant material can be per-
formed using mortar and pestle [
46
–
50
], metal spoon [
51
] or glass rod [
52
], by cutting the
plant material [
53
] or crushing and riddling (0.5 mm) [
54
] or by manual grinder [
50
,
55
].
According to the UNODC [
5
], dried herbal cannabis material and cannabis resins should be
pulverized by a cutter (at high revolution speed, i.e., 100 rps) and sieved (mesh size 1 mm).
Plant samples can also be homogenized in a crucible [
56
], in laboratory blender, usually
to 60–80 mesh (177–250
µ
m) particle size, but sometimes a larger (
≤
335
µ
m) or smaller
(100–150
µ
m) particle size is required [
57
–
59
]. Unlike classical, mechanical hand grinders
and electrical grinders, some sample preparation methods employ superfine grinding of
cannabis plant material [
60
–
62
]. However, manual grinding with a handheld herb grinder
resulted in higher yield of total phytocannabinoids (17.5
±
0.5%) than with electric blender
(12.0
±
0.3%). The minimization of analyte loss using manual grinder is attributed to the
adhesion of cannabis resin to the blades and plastic housing surface of a plastic blender
during the high-speed pulverization [
50
]. Mechanical grinding-activation in an intensity
planetary vibrational mill [
62
,
63
], ball mill [
55
], knife mill [
64
] or freeze mill [
65
] are also
applied. Instead of drying, fresh cannabis plant material can be frozen with liquid nitrogen

Molecules 2022,27, 975 12 of 42
and crushed [
66
,
67
] or frozen, lyophilised at
−
50
◦
C and grounded by hand [
68
] or in a
mill [
69
]. Comparison of coarse homogenization by sieving through a 1-mm mesh and fine
homogenization with a ball mill revealed better extraction efficiency for CBDA and THCA
for the finely powdered plant material, and no difference for the neutral phytocannabinoids
(CBD, ∆9-THC) [55].
In the next step, thermal processing occurs, which aims to remove moisture, usually
to 8–13% residual humidity, as recommended by UNODC [
5
], achieved by drying at room
temperature for several days or at 70
◦
C until the leaves become brittle or, according to
EC, within 48 h using any method below 70
◦
C [
70
]. Lower drying temperatures should be
avoided, as they result in mould growth [
71
]. Despite this, drying is frequently performed
at variety of temperatures and durations, such as at 135
◦
C, 2 h, 120
◦
C [
72
], 103
◦
C,
4h[73]
,
65
◦
C, 16 h [
74
] or 48 h [
75
,
76
], 60
◦
C, 12 h [
77
], 40–50
◦
C [
54
], 40
◦
C,72 h [
78
], 48 h [
17
,
79
] or
24 h [
74
,
80
,
81
], 38
◦
C, 4–8 h [
82
], 35
◦
C, 24 h [
83
], 30–40
◦
C, 1–2 days [
46
], 30
◦
C [
57
] on forced
ventilation oven, by natural ventilation at 32
◦
C for 60 h [
84
], at 30
◦
C overnight [
52
] or for
4 h [
80
] or air-dried at room temperature (20–22
◦
C) for 24 h [
81
], for 3 days [
44
,
85
], 6 days
after harvesting [86], for 4 weeks [87] or until a residual humidity ≤12% is achieved [47].
Dried and mechanically processed samples are extracted using maceration, LLE,
PLE, HS-SPME, SFE or FUSE. Other extraction techniques, including ultrasonic assisted
extraction (UAE), microwave assisted extraction (MAE), dynamic maceration (DM) and
accelerated solvent extraction (ASE), are faster and use less extraction fluids than the
“classic” maceration. The number of the consecutive extractions did not have significant
effects on total phytocannabinoid yield. Yet, phytocannabinoids yield after sonication was
found to be slightly lower than the yield obtained by one-day DM [
50
]. Phytocannabinoid
extraction was omitted in only one study, which, consequently, reported low sensitivity
(Table S1); thus difficulties occurred during quantification of trace phytocannabinoidsin
cannabis plant tissues [
88
]. The summary of the properties of the most frequently used
sample preparation techniques are given in Table 2.
The conventional sample preparation methods for cannabis plant material are macera-
tion and LLE using versatile organic solvents with great affinity towards phytocannabinoids.
Although universal and simple, they are time-consuming and not environment-friendly, as
they require large quantity of organic solvents.
UNODC recommends maceration prior to GC-flame ionization detector (FID) analysis;
0.2 g dried and homogenized herbal cannabis, 0.1 g cannabis resin or 0.05 g cannabis oil is
extracted with internal standard (IS) solution of tribenzylamine in 96% EtOH (
0.5 mg/mL
)
for 15 min in an ultrasonic bath [
5
]. DAB’s Cannabis flos monograph proposes extraction
with EtOH (96%, v/v), while the AHP’s Cannabis inflorescence monograph proposes
extraction with MeOH/CHCl
3
(9:1, v/v) [
33
]. EC recommends maceration of 0.1 g semi-fine
powdered herbal cannabis with IS solution of 35 mg of squalane/100 mL hexane [70].
The most commonly used solvent and solvent mixtures for extraction of phytocannabi-
noids from cannabis and cannabis-based products are given in Table 3.
Table 2.
Sample preparation techniques for phytocannabinoid profiling of cannabis and cannabis-
based products.
Sample Preparation Technique Advantages Disadvantages
LLE
- variety of solvents and solvent
mixtures with appropriate
extraction efficiencies;
- appropriate for all matrices
- low price
- high solvent consumption
PLE
- possibility to perform
decarboxylation in situ
- greatreproducibility
- low price
- miscelanous scientific finidings
regarding the ability of PLE to
extract thermolabile compounds

Molecules 2022,27, 975 13 of 42
Table 2. Cont.
Sample Preparation Technique Advantages Disadvantages
HS-SPME
-
programmable automated operation;
- improved chromatographic
peak shape;
- reduction of matrix interferences
- specific to GC-based methods only;
- applicable mostly for simple
matrices (herbal material)
SFE
- “green” extraction method;
- ensures stability of thermolabile
and light-sensitive
phytocannabinoids;
- high extraction yields;
- ability to separate
phytocannabinoids from terpenes
- rarely used
- high price
FUSE, UAE - low solvent and energy
consumption
- applicable mostly for simple
matrices (herbal material)
SPE
- most suitable for food matrices
and extracts
- “green” extraction technique
- laborious and time-consuming
- high price
MHD
-
simultaneous extraction of terpenes
and phytocannabinoids
- simultaneous decarboxylation
-
more commonly used for extraction
of essential oils
- high price
CPE
- analyte extraction and
preconcentration in a single,
solvent-free step
- avoidance of analyte loss during
solvent evaporation
- low price
- low extraction efficiency for
phytocannabinoids
- time consuming
CPC - allows for large-scale extraction of
phytocannabinoids with high
efficiency
- high solvent consumption and
waste generation
- high price
Table 3.
Most commonly used solvents and solvent mixtures in maceration and LLE of phytocannabinoids.
Solvent/Solvent Mixture References
MeCN [89,90]
MeCN + 1% acetic acid [65]
MeCN saturated with n-hexane [91]
MeOH [38,42,50,54,56,68,78,79,85,89,92–101]
absolute ethanol (99.7%, v/v) [10,49,51,53,66,90,102–108]
EtOH(96%, v/v) [40,54,55,66,90,92,100,109–113]
isopropanol [63,108]
cyclohexane [82,114]
EtAc [69,89,115–117]
CHCl3[44,52,58,77,118–120]
n-hexane [40,47,54,66,75,76,78,86,87,101,121–125]
light petroleum [46]
petroleum ether [126–128]
toluene [129]
benzene [130]

Molecules 2022,27, 975 14 of 42
Table 3. Cont.
Solvent/Solvent Mixture References
CCl4(later evaporated and extracts reconstituted in chloroform) [131]
MeCN/MeOH (8:2, v/v) [132]
hexane/isopropanol (9:1, v/v) [57,94,106,133]
hexane/EtAc (9:1, v/v), (7:3, v/v), (6:4, v/v) [54,57,66,94,104]
hexane/CHCl3(1:1, v/v) [134,135]
MeOH/CHCl3(4:1, v/v) [48,136]
MeOH/CHCl3(9:1, v/v), (99:1, v/v) [57,67,86,106,137]
MeOH/hexane (9:1, v/v) [138]
petroleum ether/MeOH (9:1, v/v) [45]
EtOH/H2O (1:1, v/v) [133]
KOH in MeOH and hexane/EtAc (9:1, v/v) [139]
IS (tribenzylamine) in 96% EtOH [57]
IS (tribenzylamine) in MeCN [140]
IS (nonadecane) in EtOH [138]
IS (diphenylhydramine) in EtOH [74]
IS (4-androstene-3,17-dione) in EtOH [9,137]
IS (docosane) in petroleum ether [128,141]
IS (nonadecane) in MeOH/CHCl3(9:1, v/v) [67]
IS (squalane) in hexane [76,89,142]
IS (chrysene-d12) in hexane [71]
IS (ketamine hydrochloride) in MeCN [124]
IS (4-androstene-3,17-dione) in MeOH/CHCl3(9:1, v/v) [9,75,86,87,143–145]
Absolute EtOH is the most preferred organic solvent for maceration and LLE due to its
great affinity for phytocannabinoid structure [
54
,
115
,
146
] that leads to high extraction effi-
ciencies. EtOH is, however, known to co-extract significant amount of pigments and ballast
from cannabis plant material, much more than CHCl
3
, enhancing matrix interferences [
44
].
To avoid this, n-hexane is used [
87
]. For highly aqueous cannabis-based products, such
as coffee beverages, 100% MeCN is preferred LLE solvent [
90
]. CHCl
3
is preferred for
extraction of dried glandular plant material (leaves), cannabis resin and reefers (mixtures of
tobacco with powdered resin or herbal cannabis), yielding 99% extraction efficiency, much
higher than light petroleum [
44
]. Hexane was found to be the worst performer in terms of
recovery of all phytocannabinoids, while EtOH was shown to possess the most appropriate
polarity for cannabinoid compounds [40].
Prior to TLC/HPTLC analysis, maceration and/or LLE is the extraction technique of
choice. Phytocannabinoids are extracted from cannabis plant material using MeOH, accord-
ing to Ph.Eur., Ph.Helv. and DAB [
34
–
36
] and published studies [
147
], dichloromethane,
according to AHP [
33
], solvent of choice (petroleum ether, MeOH, n-hexane, toluene,
CHCl
3
and solvent mixtures, e.g., MeOH/CHCl
3
(9:1, v/v)), according to UNODC [
5
]. Prior
to maceration/LLE, lactose-containing cannabis products (cream cheese, butter, coffee
beverages with milk) require lactase pre-treatment/lactose hydrolysis in order to avoid
matrix interferences [90].
Maceration and LLE of semisolid fatty/oily matrices of cannabis-based products,
such as butter, margarine, chocolate bars, nonpolar topical ointments and balms requires
warming of the samples on a hot plate after addition of the extraction solvent in order to
melt the matrix. Prior to maceration/LLE, lactose-containing cannabis products (cream
cheese, butter, coffee beverages with milk) require lactase pre-treatment/lactose hydrolysis
in order to avoid matrix interferences [
90
]. For high sugar and carbohydrate matrices
including hard candies, honey and fruit preserves, a matrix trapping effect occurs, resulting

Molecules 2022,27, 975 15 of 42
in the formation of glassy, impervious precipitates which are hard to extract. To avoid
this, the aqueous portion of the extractant is added first, followed by warming the matrix
in order to dissolve the sugars and carbohydrates, and addition of ACN (to 83–91% final
proportion) [
90
]. Maceration and LLE of cannabis-based products is further aggravated by
the presence of glycerine and propylene glycol, especially in oral supplements and vape
products. They interfere cannabinoid profiling significantly. High levels of co-extracted
glycerine or propylene glycol may swamp the silylating derivatization agents, disabling
complete derivatization of CBD to CBD-2TMS, along with undesired side conversion of
∆8
-THC and
∆9
-THC. The problem can be avoided with CAN extraction, as sugars and
glycerine have much lower solubility in ACN than in EtOH [90].
While most of the methods include vortexing, ultrasonication in bath and centrifu-
gation immediately after solvent addition, older methods include immersion, e.g., for
1h[
9
,
75
,
121
,
143
,
144
], overnight [
121
], 10-days soaking [
123
], several hours’ maceration [
87
]
or heating near boiling for few hours in solvent, followed by separation of the liquid ex-
tract by filtration [
108
]. Only one study evaluated the shaking time during extraction
with MeOH/CHCl
3
(9:1, v/v) at room temperature in a range from 10 min to overnight
shaking and showed that 20 min of shaking gained sufficient extraction efficiency for
∆9
-THC, CBD and CBN [
101
]. Finally, exhaustive extractions in Soxhlet apparatus are
rarely performed [
95
,
117
]. In terms of temperature conditions, extraction is performed with
highest efficiency at room temperature, with rare exemptions (e.g., at 4
◦
C) [
77
], in order
to avoid conversion of phytocannabinoids from plant material [
1
]. If additional heating is
required (e.g., when no preliminary decarboxylation was performed), hot extraction (e.g.,
at 70–78 ◦C) is performed by sonication [92] or in Soxhlet apparatus [95,117].
After extraction of cannabis plant material, additional filtrationis performed, us-
ing sintered glass disc [
44
], sterile cotton plugs [
9
,
75
,
143
,
144
], PFTE syringe filters, with
0.22 µm [68,111]
or 0.45
µ
m pore size [
56
,
94
,
95
,
108
], millipore filter (0.45
µ
m) [
50
,
84
], nylon
filters (0.45
µ
m) [
133
], membrane filter [
49
], PVD filter (0.22
µ
m) [
113
] or, alternatively,
drying on MgSO
4
and filtration [
67
,
138
]. The insoluble plant material can also be re-
moved by vacuum filtration. The filtered extract is usually dried under nitrogen and
dissolved in IS (androst-4-ene-3,17-dione) in EtOH [
58
,
77
], or in anhydrous pyridine [
119
].
Alternatively, IS can be added directly to the supernatant after filtration [
44
] or centrifuga-
tion [
10
,
103
,
106
,
111
]. As filtration cannot be performed efficiently for cannabis resin extracts
due to filter clogging by felt of trichomes and other vegetable debris [
44
], centrifugation is
only performed.
Solvent exchange may also be included as final step, by diluting extracts with solvents
and solvent mixtures more similar to the mobile phases employed, in case of (HP)LC
analysis. For example, cannabis-infused chocolate is first soaked in IS solutions and
isopropyl alcohol, extracted with MeCN + 1% acetic acid and finally diluted with MeCN
prior to LC-MS/MS or MeCN/H2O (75:25, v/v) prior to HPLC-UV analysis [65].
Solvent exchange is also performed prior to GC analysis, especially when derivatiza-
tion is performed [
7
,
80
,
87
,
104
,
121
,
123
,
133
]. Extracts are reduced to dryness usually under
gentle N
2
steam, causing least damage to total extracted amounts of phytocannabinoids
and terpenoids than drying in rotary evaporator or in a speedvac, the latter reducing the
concentrations of
∆9
-THC and CBG for two-thirds [
66
]. Reduced extracts can be dissolved
in solvents (e.g., pyridine and benzene [
116
], CHCl
3
[
5
], MeCN [
116
], dry EtAc [
55
]), in a
mixture of derivatization agent and solvent (e.g., toluene and BSTFA [
94
], pyridine and
BSTFA + 1% TMCS [
118
], pyridine, isooctane and MSTFA [
99
], pyridine and BSTFA [
90
],
pyridine and MSTFA + 1% TMCS [
84
]) or directly in the derivatization agent [
80
,
99
,
100
,
133
].
As silylation agents are harmful for GC injection port and column, additional evaporation
to dryness is frequently performed, followed by dissolution in solvent, e.g., n-hexane [
84
]
or MeCN [
95
]. If no derivatization is performed, EtAc [
120
,
126
,
127
] and EtOH [
66
,
123
] are
the most commonly used solvents for reconstitution prior to GC analysis.
Before instrumental analysis, decarboxylation may be also introduced, initiating ther-
mal degradation of phytocannabinoid acids to neutral counterparts for the purpose of

Molecules 2022,27, 975 16 of 42
accurate phytocannabinoid profiling and potency examination. Dried extracts are most
commonly heated at 150–210
◦
C for 10–30 min and reconstituted in the same solvent or
solvent mixtures [
7
,
52
,
60
,
101
,
128
], or, alternatively, at 50
◦
C for 180 min and then 145
◦
C
for 15 min [
69
]. A total 15 min of decarboxylation at a temperature range of 120–180
◦
C
showed that maximum yield is achieved at 140–160
◦
C, with no significant within-range
differences [
115
]. Decarboxylation temperatures higher than 160
◦
C should be avoided, as
∆9
-THC is oxidized to CBN [
115
] and isomerised to
∆8
-THC [
87
]. However, it is almost
impossible for decarboxylation to yield 100%, which initiates significant discrepancies
in potency data. Laboratories quantifying the total
∆9
-THC as the sum of the
∆9
-THC
already present in the plant and
∆9
-THCA get higher values than laboratories that perform
decarboxylation prior to instrumental analysis [115].
Once prepared, cannabis extract should be stored in light-protected conditions, at
room temperature, refrigerated at 4
◦
C [
17
,
48
,
84
,
126
], at
−
20
◦
C [
53
,
103
] or at
−
80
◦
C [
78
]
prior to instrumental analysis.
HS-SPME is a solvent-free sample preparation method used for analysis of phytocannabi-
noids inthe headspace oversolutions or solid samples [
5
]. Multiple factors affect the extraction
efficiency during HS-SPME, including SPME fiber coating, exposure temperature, extraction
time and desorption time. Evaluation of the effect of the fiber coating on extraction efficiencies
of
∆9
-THC, CBD and CBN from herbal cannabis samples showed that among polydimethyl-
siloxane (PDMS) 100
µ
m, PDMS/divinylbenzene (PDMS/DVB)
65 µm
, Carboxen
®
/PDMS
and divinylbenzene/Carboxen
®
/PDMS (DVB/Carboxen
®
/PDMS) 50/30
µ
m, PDMS 100
µ
m
performed optimally in general, although the PDMS/DVB fiber provided higher extraction
efficiency for CBD, due to its higher polarity and affinity to PDMS/DVB. Among the three
exposure temperatures (80
◦
C, 90
◦
C, 150
◦
C), 150
◦
C was optimal, simultaneously promoting
volatilization and decarboxylation [
59
,
148
]. Extraction time depends upon matrix viscosity
and lipophilicity that define the speed of diffusion of analytes from the liquid to the gas phase
and, as a result, HS-SPME rate and efficiency. HS-SPME is more appropriate for simpler matri-
ces (e.g., cannabis tea), as the extraction recoveries are proportional to the sample amount [
99
],
while complex liquid- and protein-containing matrices cause significant matrix retention and
lower recoveries, with higher LODs and lower method precision. For fatty/oily matrices,
such as versatile hemp foods, alkaline hydrolysis with NaOH and Na
2
CO
3
is performed prior
to HS-SPME in order to saponify the matrix lipids and reduce lipid matrix interferences [
99
].
Therefore, extraction time varies depending upon the sample matrix, from 10 min for herbal
cannabis [
59
,
148
] to 25 min for different hemp food products [
99
]. Finally, desorption time
depends upon analytes’ lipophilicities [
148
]. It is superior to LLE (n-hexane/EtAc (9:1, v/v)) in
terms of chromatographic peak shape and matrix interferences, despite the good agreement
of achieved LODs in food samples [99].
SFE uses supercritical fluids (SCFs) and liquefied gases as green solvents for extraction
and fractionation of complex samples. SFE offers low solvent consumption and ensures
stability of thermolabile and light-sensitive compounds. For the purpose of phytocannabi-
noid profiling, SFE is rarely used in sample preparation; its main purpose is to separate
the aromatic fraction for further analysis. It is usually performed using supercritical CO
2
(SC-CO
2
) as attractive SCF with solvent strength tuned by sensitive changes in temperature
and pressure above the critical point (31.1
◦
C, 73.7 bar), conditions that are experimentally
easy to reach [
4
], along with the low cost, short processing time and low environmental
impact [
149
]. However, SC-CO
2
is a low polarity solvent that poorly dissolves phyto-
cannabinoids; therefore, employment of co-solvent, usually H
2
O, alcohols and acids, to
improve the overall extraction rate of phytocannabinoids is required. EtOH (5–20% in
CO
2
) is most commonly used co-solvent, added in constant flow [
40
,
43
,
143
,
150
,
151
] or in
pulses [
143
]. Higher SC-CO
2
pressures offer lower extraction selectivity, but high initial
extraction rate, apparent solubility and total yield, which are also a function of tempera-
ture, exposure time and phytocannabinoid content of the plant material [
143
]. A 90–94%
extraction yields for
∆9
-THC, CBD and CBN are achieved at 100 bar, 35
◦
C and 1 mL/min
flow during 10 min [
43
,
150
]; at 340 bar, 55
◦
C and 200 g/min maximal yields up to 92%

Molecules 2022,27, 975 17 of 42
are reported [
143
], along with satisfactory yields at milder conditions, (37
◦
C, 250 bar). Ex-
traction efficiency was further improved by washing of the extract with fresh SC-CO
2
and
addition of a cold separator (separating chamber) immediately after the sample containing
chamber [149].
When compared to DM, UAE and MAE with same extraction solvent (EtOH) in same
w/v ratio to sample, no significant difference existed between SFE and UAE, with the lowest
extraction yields for CBD, CBDA and CBGA. DM and MAE showed higher yield for CBD
and CBGA, but DM was selected as the optimal sample preparation technique (EtOH,
room ◦C
, 45 min). In case of MAE, increased CBD yield was accompanied by decreased
CBDA yield, suggesting partial decarboxylation due to high extraction temperature [142].
FUSE is used for phytocannabinoid extraction from herbal cannabis, employing cy-
clohexane/isopropanol (1:1, v/v) in an ice-water media in order to avoid degradation and
solvent evaporation, followed by centrifugation and filtration through 0.45
µ
m nylon fil-
ter [
43
]. FUSE is slightly more efficient than SFE for extraction of
∆9
-THC, CBD and CBN, as
80% of the phytocannabinoids are extracted at the first extraction, while <40% are extracted
after the third extraction with pure SC-CO
2,
and more than 90% are extracted with the first
extraction in the presence of co-solvent. Therefore, adding the ability of SFE to separate
terpenes from phytocannabinoids, and therefore minimize matrix interferences, SFE was
selected as more optimal sample preparation technique [43].
PLE is one of the fastest and most efficient extraction techniques for plant metabolites.
High extraction yields are achieved under pressure, using the extractant at a temperature
above its normal boiling point, thus increasing its diffusion into the plant matrix. PLE on
herbal cannabis is performed using n-hexane as extraction solvent at 100
◦
C and 40 bar for
15 min. MeOH and n-hexane are found equally efficient for
∆9
-THC and CBN, but not for
∆9
-THCA, as it is less soluble in n-hexane [
95
]. PLE with hot water, i.e., pressurized hot
water extraction (PHWE) is used to yield CBD-rich extracts while supressing the THC and
CBN content. Here, decarboxylation is performed in situ, i.e., in the extraction cell, heated
in the oven prior to the dynamic extraction [80].
SPE with QuEchERS is used for purification of honey extracts, with high recover-
ies for CBDA, CBGA,
∆9
-THCA, CBG, CBD and
∆9
-THC and low intra- and inter-day
variability. The method was more efficient than UAE with H
2
O at 40
◦
C, yielding ho-
mogeneous solution with no phase separation or solid residues, followed by extraction
of phytocannabinoids from the aqueous phase through LLE with n-hexane or EtAc, ob-
serving higher phytocannabinoid yield in n-hexane extracts [
40
]. QuEChERS is also used
for purification of MeCN extracts of hemp seeds, hempseed oil, hemp proteins, raw and
skimmed milk, coffee and chocolate, using reaction mixture of MgSO
4
/NaCl/C
6
H
5
Na
3
O
7
×
2H
2
O/disodium hydrogen citrate sesquihydrate (4:1:1:0.5, w/w); with further supernatant
dilution with MeCN/H
2
O (1:1) or with H
2
O. d-SPE with MgSO
4
, different combinations
of C
18
, primary secondary amines and zirconia-coated silica sorbents were also evaluated.
When PSA is used, cannabinoids were trapped by interactions with amines, resulting in
low recoveries; other combinations achieved satisfactory extraction [144].
Cloud point extraction (CPE) involves employment of non-ionic surfactant, salt
(Na
2
SO
4
) and deionized water to extract
∆9
-THC from cannabis resin using heating
(
40–90 ◦C
) and centrifugation. Despite the low extraction efficiency (60%), CPE offers
many advantages, such as the possibility of extraction and pre-concentration of analytes in
a single, solvent-free step and avoidance of analyte loss during solvent evaporation [145].
Centrifugal partition chromatography (CPC) is a liquid-liquid partitioning technique
in which the stationary phase is immobilized by centrifugation force, while the mobile phase
is pumped through at high flow rates. Compounds are separated based on the differences in
partition coefficients. CPC allows large-scale extraction of phytocannabinoids, i.e.,
∆9
-THC,
CBD, CBN, CBG,
∆9
-THCA, CBGA and CBDA with high efficiency (>90%) from herbal
cannabis material using two-phase system n-hexane/MeOH/H
2
O with 25 mM formic acid
(5:3:2, v/v/v) for
∆9
-THCA, CBGA and CBDA and hexane/acetone/MeCN (5:2:3, v/v/v) [
87
].

Molecules 2022,27, 975 18 of 42
Microwave-assisted hydrodistillation (MAHD) can provide a volatile hydrodistillate
that is rich in monoterpenes, sesquiterpenes, and a small amount of phytocannabinoids.
The optimized MAHD procedure in a pilot-scale reactor yielded 0.35
±
0.02% w/w of
hydrodistillate, while conventional hydrodistillation gave only 0.12
±
0.01%, w/w (in
relation to dry inflorescence mass). During MAHD, phytocannabinoid decarboxylation
inside the residual matrix was around 70% (69.01
±
0.98% and 74.32
±
1.02% for THC and
CBD respectively) [
146
]. In other studies, MAHD resulting essential oils are dominant in
CBD content (2.11–20.06 mg/g); interestingly, the essential oils from dried plant material
also contain CBDV, CBL and cannabicitran (CBT) [152].
2.2. Instrumental Analysis
Influenced by the intense scientific and technological development in regard to
C. sativa
cultivation, analytical platforms for phytocannabinoid profiling in cannabis and
cannabis-based products have intensively evolved over the last four decades (Figure 3).
GC- and LC-based methods as most commonly used, have achieved comparable accuracy,
selectivity, linearity, sensitivity and precision in phytocannabinoid profiling and are both
used in routine and investigational analysis of cannabis and cannabis-based products.
Figure 3. Prevalence of analytical techniques used for phytocannabinoid analyses.
Despite the lack of standardization process for analysis of phytocannabinoids omit-
ting the comparison of reliability of measurement among analytical platforms, a recent
interlaboratory study concluded that GC-MS is the most accurate and robust analytical
method for phytocannabinoid profiling, performing much better than GC-FID and UHPLC-
MS/MS [55].
The advantages and disadvantages of the most frequently used analytical techniques
for analysis of cannabis and different products are given in Table 4.

Molecules 2022,27, 975 19 of 42
2.2.1. GC-Based Methods
GC coupled to versatile detectors and mass analyzers is one of the oldest, but still
the most preferred and researched analytical platforms for phytocannabinoid profiling in
both plant material and biological matrices due to its robustness, reproducibility, sensitivity
and speed [
1
,
42
,
58
,
153
]. As such, GC methods are officially employed by authorities for
phytocannabinoid profiling [
1
], including the predominant phytocannabinoids (
∆9
-THC,
CBD and CBN) and quantification of
∆9
-THC/CBD ratio. This analytical platform is also
used for terpene profiling, pesticide screening and residual solvents analysis, which affords
potential benefits to regulatory bodies and cannabis industry [39].
By combining short columns, fast oven temperature ramps, high carrier gas linear
velocities, narrow columns, hydrogen carrier gas and low film thickness, fast and robust GC
methods are generated, appropriate for phytocannabinoid profiling in both research and
monitoring purposes [
39
]. The access to the well-established MSLs, such as the National
Institute of Standards and Technology (NIST) Mass Spectral Library and the Wiley Registry
Mass Spectral Library eases compound identification through GC-MS analysis. Here,
phytocannabinoids identification is performed by comparison of acquired MS or MS/MS
spectrum to spectra present MSLs, and further confirmed by analysis of analytical standards.
Although the main employment of GC-based platforms is for profiling of terpenes, it has
been extensively used for phytocannabinoid profiling as well (Table S1).
•Derivatizationof Phytocannabinoids
First action, after injection of the sample in the injector port of the GC (regardless of
employed detector), is vaporization that is achieved at temperature ranges
250–290 ◦C
and
causes in situ decarboxylation of acidic phytocannabinoids (
∆9
-THCA, CBDA, CBGA)
to the corresponding neutral phytocannabinoids (
∆9
-THC, CBD, CBN) prior to chro-
matographic separation. Thus, acidic and neutral phytocannabinoids are not distinguish-
able, but rather the result is the sum of neutral cannabinoid present in the extract and
neutral cannabinoid generated during decarboxylation. The issue is of no concern for
studies aiming to quantify total THC (
∆9
-THC and
∆9
-THCA) levels, as it is the case
with most of the GC-based studies included in this review without prior derivatiza-
tion [
9
,
17
,
18
,
45
,
47
–
49
,
52
,
54
,
56
,
58
,
59
,
63
,
67
,
72
–
76
,
79
,
81
,
82
,
85
,
86
,
89
,
93
,
96
,
98
,
101
–
103
,
105
,
108
,
115
,
121
–
124
,
128
,
138
,
139
,
141
,
148
,
150
,
152
,
154
–
160
] (Table S1). However, derivatization is
of great importance for studies aiming more thorough phytocannabinoid profiling. In
order to prevent their degradation and achieve profiling of the native chemical constitu-
tion of the cannabis material, a reaction of derivatization has to be performed prior to
GC analysis. In such a way, derivatization improves the limited volatility and thermal
stability of phytocannabinoids, and thus their amenability to GC analysis, which further
improves peak shape, peak resolution (especially for CBC and CBD) and sensitivity [
161
].
This comes at the price of increased analyses cost, duration [
2
] and measurement uncer-
tainty, as derivatization yields are sometimes highly variable and seldomly obtain 100% for
phytocannabinoids [23,91,122], making quantification results speculative.
Silylation is the most common derivatization reaction performed. It involves substitu-
tion of a hydrogen atom that is bound to a hetero atom (such as -OH, -COOH, -NH
2
, =NH,
and -SH) by a silyl group, i.e., a trimethylsilyl (TMS) or tert-butyldimethylsilyl group (TB-
DMS). The resulting TMS/TBDMS derivatives have lower polarity and increased thermal
and catalytic stability and GC amenability. However, they can be thermally degraded in
injector port and/or column system [162].

Molecules 2022,27, 975 20 of 42
Table 4. Analytical techniques for phytocannabinoid profiling of cannabis and cannabis-based product.
Analytical Techniques Advantages Disadvantages Note
GC-FID
- More accurate cannabinoid quantification
than GCMS
- Terpenes and residual solvents
- High resolution
- Time-consuming derivatization for acidic
cannabinoids - Gold standard technique for forensic purposes
GC-MSD - -Compound libraries to identify the parent analyte
- Higher specificity
- Sensitive
- Use of equivalent deuterated standards (expensive
and not available for all cannabinoids) /
GC-QQQ/QTOF
- -Simultaneous analysis of cannabinoids, terpenes
and residues of pesticides
- Highest sensitivity
- Analysis of “Unknowns”
/ /
(HP)TLC
- Rapid screening of many samples to confirm the
existence of cannabinoids, provide better resolution
and generate reports for more convenient
documentation for peer review of casework in
crime labs
- Lower performance compared to other separation
techniques
- Reproducibility depends of humidity - Compulsory method for identification
HPLC-UV/DAD - Quantification of both acidic and neutral forms of
phytocannabinoids
- The complex composition of the cannabis material
leads to significant peak overlap of the
phytocannabinoids
- Only target analytes can be determined, not
full spectrum
- Limited use for analyses of biological samples the
complex composition of the cannabis material leads
to significant peak overlap of the phytocannabinoids
- Only target analytes can be determined, not
full spectrum
- Limited use for analyses of biological samples
/
HPLC-QQQ - Fingerprinting with excellent sensitivity and
selectivity of complex matrices - Set-up of QQQ instruments require careful tuning
and optimization (require time and effort) - Often are used for simultaneous pesticide and
mycotoxins/aflatoxins analysis
HPLC-Q-Exactive Orbitrap®- High selectivity of complex matrices
- Confirm analyte structure
- Analysis of “Un-knowns”

Molecules 2022,27, 975 21 of 42
Table 4. Cont.
Analytical Techniques Advantages Disadvantages Note
SFC - Green technique suitable for thermally
labile compounds - Availability of SFC equipment /
NMR - Not sensitive to ballast compounds (chlorophylls
and lipids) reference standards are not required - High cost of this analyser /
RAMAN - Rapid, versatile and non-invasive qualitative and
quantitative profiling growth staging of cannabis
plant and extracts
/ /
FTIR, NIR, MIR
- Chemically fingerprint substances
-
Analysis of heterogeneous substances like cannabis
samples and to determine the potency of
cannabis flower
-
Rapid on-site use for monitoring growth and curing
processes of cannabis
- Should be combined with chemometrics
- Less accurate for potency analyses /

Molecules 2022,27, 975 22 of 42
Versatile silylation agents are used, ordered by reactivity: hexamethyldisilazine
(HMDS) [
88
], N-methyl-(trimethylsilyl) trifluoroacetamide (MSTFA) [
153
], N, O-bistrifluoro-
acetamide (BSTFA) [
90
], alone or accompanied by a catalyst, usually 1% trimethylchlorosi-
lane (TMCS) [
58
,
99
,
100
,
123
,
124
,
133
] for TMS derivatization and N-tert-butyldimethylsilyl-
N-methyltrifluoroacetamide (MTBSTFA) alone or with 1% tert-butyldimethylchlorosilane
(t-BDMCS) as a catalyst for TBDMS derivatization. A study comparing the derivati-
zation efficiency of different alkylsilylation agents, HMDS + trifluoroacetic acid (TFA),
MSTFA, activated MSTFA, MSTFA + 1% TMCS, BSTFA, BSTFA + 1% TMCS, MTBSTFA and
MTBSTFA + 1% t-BDMSC
, using pyridine as additional catalyst for CBC, CBD, CBG, CBN,
∆9
-THC and
∆9
-THCA in standard solutions and plant matrix concluded that maximum
responses are obtained with HMDS + TFA. Responses were not influenced by catalysts,
as well as reaction solvent (pyridine, EtAc, MeCN) [
88
]. Other less frequently exploited
derivatization is methylation of the hydroxyl groups of the phytocannabinoids using
trimethylsulfonium hydroxide (TMSH) [
97
]. Formation of alkylboronate-TMS derivatives
using alkylboronic acid and BSTFA + TMCS showed that methyl- and n-butylboronates
yielded derivatives that were stable for several weeks at 4
◦
C had improved GC peaks
when compared to TMS derivatives, and MS characteristics, with preserved fragmentation
patterns of the underivatized compound [116].
•GC Columns
Various GC-based methods became popular and widely used during the 40-year span
of phytocannabinoids profiling (Table S1). E.C. recommends use of glass capillary column
25 m
long and 0.22 mm wide, impregnated with 5% non-polar phenyl-methyl-siloxane phase,
that would allow good separation of phytocannabinoids [
70
]. Older GC-based methods
employ glass columns packed with 2% OV-17 on Chromosorb WHP (mesh
100–120
) [
119
],
2% OV-17 on 100–200 mesh GasChrom Q) [
98
,
123
,
140
], 2% OV-17 [
58
] and glass column
packed with 3% SE-30 on 100/200 mesh Gas Chrom Q [
116
]. Later, they were replaced by
capillary columns with cross-linked and bonded stationary phases with various polarity.
Here, analytes are separated based on differences in polarity, molecular mass and boiling
point. As most frequently, phytocannabinoid profiling studies are focused on the most promi-
nent phytocannabinoids, that are
∆8
-THC,
∆9
-THC, CBC, CBD, CBDA, CBDV, CBG, CBGA,
CBN,
∆9
-THCA and THCV. They contain aromatic, alkyl and alcohol moieties; it is expected
that the proportion of phenyl groups in mixed dimethylpolysiloxane-silphenylene or mixed
dimethylpolysiloxane-dimethyl-dimephenyl stationary phases to have an impact on their chro-
matographic separation. Wide employed thin-filmed capillary columns with non-polar station-
ary phases are used, such as 5%-diphenyl-dimethylpolysiloxane columns, including HP-5 (for
FID) or HP-5MS
(for MS) [50,52,57,58,62,70,85,89,91,94,101,103,104,126,133,141,142,154,156]
,
DB-5MS [9,99,108,128,151,163,164]
, Rxi-5MS [
73
], Mega-5MS [
146
], BP-5 [
45
], RTX-5 [
94
,
161
],
MDN-5S [
91
], SE-52 [
128
], ZB-5 [
95
], Zebron ZB-5HT Inferno [
102
] and SLB-5MS [
152
]. 100%
dimethylpolysiloxane columns, such as HP-1 [
10
,
109
,
112
,
165
], SPB-1 [
85
], OV-1 [
74
] and DB-
1 [
17
,
23
,
75
,
77
,
84
,
144
,
148
] are preferred for more successful separation of CBC and CBD, apart
from all other phytocannabinoids, or are used only for separation of CBC and CBD [
10
,
109
,
112
].
Such columns are appropriate for analysis of CBDV, THCV, CBD, CBC,
∆8
-THC,
∆9
-THC,
CBN but not for CBG [
163
]. Simultaneous injection on two column with different polarities,
i.e., a medium-polar (HP-50+) and non-polar column (DB-1MS) is another option for better
separation of CBC and CBD [18].
Columns with intermediate polarity, such as Rxi-35Sil MS [
90
] and DB-35MS [
163
]
(35% (-phenyl)methylpolysiloxane) and Zebron ZB-624 (6%- cyanopropyl-phenyl, 94-
dimethylpolysiloxane) [
106
] are also used for cannabinoid separation. Rxi-35Sil MS may
offer slightly wider retention window than a 5% silphenylene phase (DG-5MS) and 5%
diphenyl phase (HP-5MS) [
90
]. However, columns with 35% (phenyl)methylpolysiloxane
stationary phase might produce multiple peaks for CBC, one of which coeluted with THCV.
Low to mid-polarity columns, such as DB-170 (14%-cyanopropyl-phenyl)-methylpolysiloxa-
ne) are not appropriate, as they produce low responses, distorted peak shape and tail-
ing [
163
]. Despite this, the nonpolar 100% dimethylpolysiloxane column DB-1HT was used

Molecules 2022,27, 975 23 of 42
for successful separation of
∆9
-THC, CBD, CBC, CBN and other minor cannabinoids, along
with terpenes, sesquiterpenes, sterols, diglycerides and triglycerides [166].
•GC Detectors
A large proportion of the profiling studies employ FID [10,17,49,52,55,56,58,70,73–76,
79
,
82
,
85
,
93
–
96
,
98
,
101
–
103
,
106
,
108
,
115
,
118
,
119
,
138
,
141
,
157
–
159
] or dual FID [
9
,
75
,
121
,
144
].
FID offers more accurate quantitative response with respect to MS, but at the price of
lower sensitivity and specificity. The need of analogue analytical standards only makes
FID low-cost and simpler analyzer when compared to MS, which needs the use of the
corresponding deuterated analytical standards [
1
]. Moreover, GC-FID methods are more
robust than GC-MS methods both in full scan and SIM mode [
107
]. However, due to its
inability to discriminate CBD and CBC, FID is often replaced by MS detectors for more
thorough cannabinoid profiling.
GC-MS is the most researched analytical platform for cannabinoid profiling to
date [1,42,153,167]
. Electron impact ionization (EI) is the most commonly used ioniza-
tion technique in cannabinoid profiling, while others, i.e., chemical ionization (CI) in
positive/negative mode and atmospheric pressure ionization (API) are preferred in forensic
analysis of phytocannabinoids in biological samples. Mass selective detector
(MSD) [45,47,49,55,59,74,81,86,89,90,94,96,121–124,128,139,146,148,154,160,166]
, single qu-
adrupole (Q) [
57
,
66
,
76
,
163
] and triple quadrupole (QQQ) are the most frequently used
configurations. Improved sensitivity, specificity and reproducibility, with reduced noise
level and interferences are achieved by monitoring specific ions (selected ion monitor-
ing, SIM mode, MSDs and Qs) or fragmentation reactions (multiple reaction monitoring
mode, MRM mode, QQQ), then using the analyzers in scan mode. Ion trap (IT) is sel-
domly used [
23
,
70
,
151
], despite its advantageous ability to acquire structural information
by higher fragmentation (MS
3
, MS
4. . .
MS
n
). Quadrupole-time-of-flight (Q-TOF) mass
analyzer is rarely used for quantification purposes, but rather for untargeted cannabinoid
profiling of C. sativa extracts [
78
]. Rarely, FID and Q are simultaneously used by installing
a “Y” splitting unit at the column outlet [
152
]. Finally, a novel variation of GC-EI-MS,
cold EI, based on interfacing GC and MS with supersonic molecular beams (SMB) in a
fly-through ion source was recently successfully employed for an inaccurate, sensitive,
reproducible and comprehensive full (including phytocannabinoid) profiling of herbal
cannabis extracts [166].
A relatively novel GC-based analytical platform employs analyser based on vacuum
UV (VUV) operating in the UV/VUV spectral range (120–240 nm). UV/VUV absorption
events are very sensitive for differentiating isomers (positional isomers and diastereomers).
This, together with the ability to deconvolute overlapping spectra [
4
], which significantly
shortens analysis time, makes GC-UV/VUV potentially favourized analytical platform
over GC-MS. Almost all phytocannabinoids exhibit maximum absorbance in the region
170–200 nm, with no overlap among different phytocannabinoids and significant spectral
differentiability. This is especially important for CBC and CBD. However, this method
reports high LODs and, thus, cannot be employed for phytocannabinoid profiling in
biological matrices and cannabis-based products, but is sufficiently sensitive for plant
matrices [161].
Two-dimensional (2D) gas chromatography (GC
×
GC) is reported to offer better
chromatographic resolution of phytocannabinoids [
11
,
56
,
150
,
160
]. Despite this, it is rarely
used for phytocannabinoid profiling; chemical fingerprinting and classification are more
commonly performed GC x GC is usually coupled to MS [
56
,
160
] or to FID/MS [
150
]. Com-
bination of columns of different polarity is used, as, for example, 100% dimethylpolysilox-
ane and polyethylene glycol in sol-gel matrix [
56
], or a non-polar column (e.g., DB-5 or
HP-5MS) for first-dimension separation and medium-polarity column (e.g., DB-17) in the
second dimension [150,160].

Molecules 2022,27, 975 24 of 42
2.2.2. LC-Based Methods
LC-based methods are recently becoming methods of choice for qualitative and quan-
titative phytocannabinoid profiling. The simplified sample preparation and the low tem-
peratures, high pressure and high flow rates used during TLC, HPTLC, HPLC and UHPLC,
and the recently emerging supercritical fluid chromatography (SFC) analysis allow sample
preservation without decarboxylation and decomposition, reliable separation of neutral
and acidic phytocannabinoid species and, thus, direct identification and quantification of
both neutral and acidic forms of phytocannabinoids in the extracted samples [
18
,
165
]. The
simplified sample preparation methods, along with avoidance of analytes loss favourized
LC over GC in cannabinoid profiling [66].
•TLC and HPTLC Methods
TLC is an attractive method for analyses of herbal drug constituents [
168
], and es-
pecially suitable method for the purpose of preliminary semi-quantitative screening of
cannabinoid content in routine tests [
164
]. It is method of choice for identification of
cannabis flowers in all Cannabis flos monographs (DAB, AHP, Ph.Helv., Ph.Eur.) [
33
–
36
] and
DAB’s Cannabis extractum normatum monograph [
38
]. Using TLC, cannabinoid identifica-
tion is performed by comparing retardation factors (R
F
s) of analytes with R
F
s of standards
on a TLC plate developed with appropriate mobile phase, whereas visual evaluation
is obtained by dipping or spraying the TLC plate into/with the appropriate visualiza-
tion reagent under UV light, or under daylight. In DAB [
36
] and Ph.Eur. [
35
] analytical
monographs of cannabis flower, a MeOH extract of 0.1 g pulverized drug is identified by
comparison of RFs of analytes to reference solutions of CBD and ∆9-THCA (5 mg each) in
MeOH. Solutions are applied on TLC C
18
silica gel F
254
plate (2 to 10
µ
m) and developed
with H
2
O/glacial acetic acid/MeOH (15:15:70, v/v/v). After air drying, the TLC plate is
sprayed with vanillin reagent, dried at 100–105
◦
C for 15 min and examined on daylight.
Identical procedure applies to Cannabis extractum normatum according to DAB [
38
]. Ph.Helv.
includes filtration of MeOH extract of cannabis flower through membrane filter (0.45
µ
m)
as an additional step prior to application to the C
18
silica gel F
254
plate and reference
solution of CBDA in MeCN and
∆9
-THCA in 2-isopropanol, following the identical pro-
cedure for development and detection [
34
]. AHP’s cannabis flower monograph employs
TLC C
18
F
254
plate with MeOH/H
2
O with 1% glacial acetic acid (75:25, v/v) as a mobile
phase for identification of CBC,
∆9
-THC, CBN, CBG, CBD THCV,
∆9
-THCA and CBDA in
dichloromethane extract of 0.1 g pulverized drug. Visualization is performed using Fast
Blue reagent and vanillin/H
2
SO
4
under UV (254 nm) [
33
]. UNODC suggests maceration in
ultrasound bath with 10 mL of solvent (MeOH, petroleum ether, n-hexane, toluene, CHCl
3
or solvent combinations–MeOH:CHCl
3
(9:1, v/v) for 15 min at room
◦
C, using three sys-
tems for elution of HPTLC silica gel plates (A: petroleum ether 60/90/diethyl ether (80:20,
v/v); B: cyclohexane/di-isopropyl ether/diethylamine (52:40:8, v/v) and C (for cannabinoid
acids): n-hexane:dioxane/MeOH (70:20:10, v/v)). Fast Blue reagent BB or RR in MeOH or
MeOH:H2O is used as spaying reagent using visualization method 1 or method 2 [5].
The accuracy, repeatability and the acceptable LODs and LOQs in the linear dynamic
range of this methodology makes TLC methods attractive for fingerprinting cannabis [
4
];
however, such parameters are fairly low compared to the more sophisticated LC analytical
platforms. The “classic” TLC became further less utilized due to its inconvenience to
document for peer review, poor resolution due to systematic errors rising from hand-
spotting, temperature/humidity control and imprecise RFmeasurement [167,169].
Recent advances in TLC are in the development of HPTLC methods. Such reliable
methods could offer advantages over both HPLC and GC techniques for cannabis profiling,
including its ability to analyse multiple samples simultaneously and the consequently
lower running costs and analysis runtime. Further, automation of sample application
in HPTLC methods eliminate systematic errors, provide better resolution and generate
reports for more convenient documentation for peer review of casework [
170
]. In that spirit,
normal-phase HPTLC with an automated spotter is shown to achieve better separation

Molecules 2022,27, 975 25 of 42
than TLC for the main neutral phytocannabinoids. The method is comparable within a
small degree of error (±0.5%) to a validated HPLC method [110].
Older methods use plates spread with a layer of a slurry of alumina/CaSO
4
/H
2
O
(
22.0 g
:3.0 g:50 mL), activated at 110
◦
C for 30 min and stored under anhydrous CaCl
2
[
131
],
silica gel G plates [
129
], precoated silica gel G plates [
171
] and silica gel G layers im-
pregnated with dimethylformamide [
120
]. Reverse phase (RP)-TLC is performed using
RP-18 HPTLC plates [
129
] and RP-C
18
bonded silica gel F plates [
18
]. The more recent
TLC/HPTLC methods most commonly use HPTLC silica gel 60 F
254
plates for successful
separation of 11 phytocannabinoids (
∆9
-THC, CBD, CBN, CBC, THCV,
∆8
-THC, CBDV,
CBG, CBGA, CBDA,
∆9
-THCA) [
127
] or of
∆9
-THC, CBD, CBN and CBG [
172
], or for
separation of
∆9
-THC, CBD and CBN only [
125
], silica gel 60 [
110
], silica gel 60F [
135
] or
silica gel plates [
5
,
126
]. For some methods, for instance, the type of TLC plate was not
clearly defined [130,147].
Most suitable mobile phases include xylene/hexane/diethylamine (25:10:1, v/v/v) [
127
],
CHCl
3
, with plate prewashing with MeOH [
110
], hexane/diethyl ether (80:20, v/v), which
allowed clear separation between
∆8
-THC,
∆9
-THC, CBD and CBN [
125
], cyclohexane/
toluene/diethylether (75:15:10, v/v/v) [
126
], benzene/n-hexane/diethylamine (25:10:1,
v/v/v) [
130
], benzene/chloroform (50:50, v/v) [
131
], diethylether/petroleum ether (1:4,
v/v) [
120
], benzene/n-hexane/diethtylamine (25:5:0.5, v/v/v) [
147
], n-hexane/CHCl
3
/
dioxane (89:8.75:2.25, v/v/v) [
135
], benzene, benzene/n-hexane (6:4, v/v), benzene/
n-hexane/diethylamine (70:25:5, v/v/v) [
129
], MeOH/dioxane/hexane (1:2:7, v/v/v), hex-
ane/EtAc (4:1, v/v), hexane/diethylether (4:1, v/v) [
171
], hexane/ethyl ether (8:2, v/v) [
172
],
petroleum ether/deithylether (8:2, v/v), cyclohexane/diisopropyl ether/deithylamine
(52:40:8, v/v/v) or n-hexane/dioxane/MeOH (7:2:1, v/v/v) [
5
]. For 2D TLC, first n-heptane/
dichloromethane/butan-2-one (83:5:12, v/v/v) was used for first and n-hexane/acetone
(86:14, v/v) for second development after 90◦rotation [129]. RP-TLC employs MeCN:H2O
(9:1, v/v) [129] or MeOH/5% acetic acid (19:1, v/v) [18] as mobile phase.
Evaluation of 10 mobile phases (hexane/diethylether (80:20, v/v), toluene, n-heptane/
diethyl ether/formic acid (75:25:0.3, v/v/v), CHCl
3
, hexane/acetone (87:13, v/v), benzene,
xylene/hexane/diethylamine (25:10:1, v/v/v), 4–8% diethyamine in toluene, MeOH/H
2
O
with 0.1% glacial acetic acid (75:25, v/v) and hexane/acetone (75:25, v/v) on Silica gel 60 F
254
plate showed that xylene/hexane/diethylamine (25:10:1, v/v/v) allows most precise bands
and best separation of
∆9
-THC, CBD and CBN, but without migration of
∆9
-THCA, CBDA
and CBGA. The cannabinoid acids were successfully separated using n-heptane/diethyl
ether/formic acid (75:25:0.3, v/v/v) on C
18
F
254
plate and EtOH/H
2
O with 0.1% glacial
acetic acid (75:25, v/v) on RP-C
18
F
254
plate [
127
]. Another study showed that, when using
alkanes as eluents (isooctane, heptane, hexane and pentane/diethylether (90:10, v/v), the
capability to separate
∆9
-THC, CBD and CBN decreased as the length of the carbon-bearing
chain increases [125].
Visualization of (HP)TLC plates is usually performed using 0.1% aqueous solution
of Fast Blue B salt reagent [
110
,
125
–
127
,
130
,
147
], alone, under white light (254 nm and
366 nm) [
127
], under UV (254 nm) [
131
], as solution in 0.1M NaOH [
5
] or under UV
(
206 nm
) [
110
] or as 0.5% aqueous solution, followed by 0.1M NaOH [
23
,
125
,
173
]. Prior to
Fast Blue B, diethylamine can be applied (50 mg
·
L
−1
H
2
O + 20 mL MeOH) [
5
]. RP-TLC
plates are visualized using Fast Blue B in 0.1M NaOH or in 50 g
·
L
−1
H
2
O/acetone (9:1,
v/v) [
129
]. Fast Blue RR was better for visualization of
∆9
-THC, CBD, CBG and CBN than
Fast Blue B salt [
172
]. As qualitative evaluation for the presence of cannabinoids during
(HP)TLC analysis is based on color determination, it is often subject of analysts’ erroneous
determination. Recent studies made the pioneering efforts in developing a method for
standardizing and naming colors using the Sci-Chromus
®
software, that significantly
reduced the subjectivity of the color names in identifying
∆9
-THC, CBD, CBN and CBG in
cannabis extracts [172].
Apart from TLC and HPTLC, other planar chromatography methods are seldomly
used, such as optimum performance laminar chromatography (OPLC) and automated

Molecules 2022,27, 975 26 of 42
multiple development (AMD) for phytocannabinoid profiling, despite their greater repro-
ducibility due to complete automation. Moreover, OPLC offers extension as semiprepar-
ative technique for sample purification, while AMD offers best resolution. The only
reported employment of OPLC in AMD in cannabinoid profiling is in hexane extracts
of cannabis resins (dried and reconstituted in toluene) and in hexane extracts of cannabis
resin. OPLC was performed for determination of
∆9
-THC, CBD and CBN on HTSorb
BSLA 011 and HT Sorb BSLA 003 columns using isooctane/diethylether (90:10, v/v) as
eluent. Semi-preparative OPLC was performed for isolation of CBD from cannabis resin
using hexane/diethylether (80:20, v/v). Using AMD, separation was performed on HPTLC
with the elution gradient 1C acetone (100, v/v), diisopropylether (100, v/v), hexane (100,
v/v), hexane (100, v/v) and hexane (100, v/v) during 20 steps. For both OPLC and AMD,
visualisation is performed with Fast Blue B salt reagent [125].
•HPLC Methods
The HPLC technique is gaining popularity as the main choice for fingerprinting
study for the quality control of herbal drugs [
174
], thus enabling chemical characterization
of herbal medicines [
175
]. HPLC methods offer larger linear ranges and more consistent
calibration curves for all phytocannabinoids in regards to GC-based methods [
163
]. In terms
of reliability, reproducibility and sensitivity, it was shown that high-resolution GC/FID
and HPLC-UV methods for quantification of
∆9
-THC, CBD and CBN are comparable [
98
].
•HPLC Mobile Phases
Most of the mobile phases used in HPLC/DAD analysis of phytocannabinoids consisted
of buffered aqueous solutions of ammonium acetate [
176
], ammonium
formate [83,142]
, formic
acid [11,40,50,60–63,78,173,177–182]
, acetic acid [
19
,
183
,
184
], o-phosphoric
acid [165,185]
or 5%
MeCN/80% MeCN with 0.1% o-phosphoric acid [
69
]. Acidic conditions are preferred for
cannabinoid acids (∆9-THCA, CBDA, CBGA).
Pharmacopoeial methods (DAB, Ph.Helv., Ph.Eur.) for assay of Cannabis flos and
Cannabis extractum normatum focus on the five main phytocannabinoids: CBDA, CBD, CBN,
∆9
-THC and
∆9
-THCA using aqueous solution of 85% o-phosphoric acid and MeCN as
mobile phases [34–36,38]. AHP’s Cannabis flos monography recommends identical mobile
phases for quantification of the major phytocannabinoids (
∆9
-THCA,
∆9
-THC, CBD, CBDA,
CBG, CBGA and CBN) (Table S2) [33].
Binary mobile phase system consisted of H
2
O/MeOH (10:90, v/v) [
186
], H
2
O/MeOH
(17/83, v/v) [
187
] and H
2
O + 0.1% formic acid/MeOH + 0.1%; formic acid is most com-
monly used for phytocannabinoid profiling [
11
,
63
,
65
,
147
,
185
,
188
]. Other binary systems
that provide good peak shape and improved resolution are also used, such as H
2
O +
0.1% formic acid/MeCN + 0.1% formic acid [
40
,
55
,
64
,
80
,
109
,
134
,
144
,
182
,
183
,
185
,
188
,
189
],
H
2
O + 0.1% TFA/MeOH + 0.1% TFA [
68
], MeCN/H
2
O (75:25, + 0.05% formic acid,
v/v)/isopropanol:MeCN (80:20 + 0.05% formic acid, v/v) [
183
,
184
], 5% MeCN + 0.1% formic
acid/MeCN + 5% H
2
O + 0.1% formic acid [
78
], H
2
O + 0.1% formic acid/MeCN [
27
,
63
,
185
],
5% MeCN + 0.1% formic acid/80% MeCN + 0.1% formic acid [
69
] and MeCN/H
2
O + 0.85%
phosphoric acid [42].
Other buffering solutions are less frequently used, such as 0.1% acetic acid in a tertiary
system, e.g., H
2
O + 0.1% acetic acid/MeCN + 0.1% acetic acid/MeOH [
19
], MeCN/0.5%
acetic acid (66:34, v/v) [
190
], ammonium formate in a binary system MeOH/H
2
O + 50 mM
ammonium formate (pH 5.19) [
83
], 5 mM ammonium formate + 0.1% HCOOH/MeCN +
0.1% HCOOH [65,100] or 5 mM ammonium formate/MeCN + 0.1% HCOOH [92].
Decrease in buffer concentration from 50 mM to 25 mM eliminated baseline drifting,
thus avoiding decrease in UV absorption. As ammonium formate causes co-elution of
∆9-THCA
with CBG, it is preferably replaced by ammonium acetate (25 mM, pH 4.75),
which offers more reproducible, reliable and rugged chromatographic separation, espe-
cially between CBG and
∆9
-THCA with improved peak shape and, thus, sensitivity. Better
reproducibility is achieved using H
2
O/MeCN (15:85, v/v) + 50 mM phosphoric acid than
with 0.1% formic acid as buffering solution [
185
]. Fast separation of 10 phytocannabi-

Molecules 2022,27, 975 27 of 42
noids in less than 8.5 min using binary system H
2
O + 0.085% phosphoric acid/MeCN +
0.085% phosphoric acid as mobile phase was achieved, that, together with employment
of RP-C
18
column prevented co-elution of CBD and THCV and both isomers,
∆9
-THC
and
∆8-THC [165]
. Another, less frequently used buffer is triethylammonium phosphate
(TMAP) in MilliQ, with MeCN in isocratic elution programme [191].
•HPLC Columns
The multitude of LC-based methods for phytocannabinoid profiling use similar
columns; it is the variation of the instrumental conditions that produces superior quan-
tification approaches. Only one methods use direct injection [
184
], bypassing the column.
Most of the published methods employ columns with normal phase C
18
stationary phase
(Ascentis Express C
18
[
40
,
178
], Luna C
18
[
69
,
177
], Kinetex C
18
[
11
,
19
,
71
], Luna Omega
Polar C
18
[
58
,
114
,
118
,
139
], Luna Omega PS C
18
[
27
], XTerra MS C
18
[
83
], Acquity UPLC
BEH C
18
[
181
], Acquity BEH Shield RP18 [
144
], MacMod ACE5 C
18
-AR [
190
], ACE 3 C
18
-
PFP [
187
], ACE Excel 3 C
18
[
111
], Poroshell 120 SB-C
18
[
182
], Poroshell 120 EC-C
18
[
188
,
189
]
or 120 SB-C
18
[
34
–
36
,
38
], Shim-pack XR-ODSII RP C
18
[
149
], Nucleodur
®
C
18
Gravity [
185
],
Zorbax Eclipse Plus C
18
[
92
,
183
], Zorbax Eclipse XDBC
18
[
186
], Zorbax SB-C
18
[
133
], At-
lantis T3 C
18
[
42
], with [
11
,
19
,
34
–
36
,
38
,
83
,
188
,
189
] or without [
43
,
184
,
185
,
188
–
190
] guard
column or C
18
guard cartridge [
177
] that allow reliable separation and quantification of a
wide range of phytocannabinoids (focused on, but not limited to, CBDV, CBDA, CBGA,
CBG, CBD, THCV, CBN,
∆9
-THC,
∆8
-THC, CBC,
∆9
-THCA). Raptor ARC-18 is the C
18
column with the widest applicability in phytocannabinoid profiling, offering the most ap-
propriate separation of 17 phytocannabinoids (CBG, CBD, CBN,
∆9
-THC,
∆8
-THC, THCA,
THCV, THCVA, CBC, CBCA, CBGA, CBDA, CBL, CBLA, CBDV, CBDVA, CBLA) at a
runtime suitable for commercial environment. It improves peak resolution issue of some of
the aforementioned columns, such as Kinetex C
18
, Luna C
18
, Luna Polar C
18
and Raptor
C
18
[
100
] and has been shown to, together with Raptor ARC-18 EXP guard column, be
suitable for analysis of pesticides, mycotoxins and cannabinoids [65].
The latest research points out the importance of core-shell technology columns (e.g.,
Poroshell) for separation of 96 phytocannabinoids by ESI-LC/MS [
19
]. C
18
columns with
advanced bonding of the trifunctional C
18
phase and end-capping process, such as Acquity
UPLC
®
HSS T3, are also used [
183
,
184
]. RP-C
18
columns, most commonly Synergi Hydro
RP C
18
[
63
,
65
,
66
], RP-C
18
Hydro [
61
,
62
], Mediterranea RP-C
18
[
78
], LiChrospher 60, RP-
Select B [
191
], RP-C
18
[
165
], with [
60
–
63
,
191
] or without [
78
,
165
] C
18
guard column are also
used. Other columns, such as Ascentis Express RP-amide are also used [64].
A better chromatographic performance (in terms of both resolution and sensitivity), a
shorter analysis time (10 min vs. 12 min) and a considerable saving of solvent consumed
while working at a flow rate of 0.5 mL/min instead of 1.5 mL/min, was observed while
working on a fuse-core stable bond (SB) RP-C
18
column rather than fully porous RP-
C
18
column [
182
]. Another study evaluated three different columns, RP-C
18
, fused-core
stable bond (SB) RP-C
18
and fused-core end-capped (EC) RP-C
18
with numerous mobile
phases and gradient conditions in an attempt to shorten the run time and to increase the
separation of 8 phytocannabinoids (CBDA, CBGA, CBD, CBG, CBN,
∆9
-THCA,
∆9
-THC
and
∆8
-THCA). The SB RP-C
18
core shell column provided the best performance due to
significant improvement in separation and symmetry of chromatographic peaks with a
baseline separation between CBD and CBG within 20 min shorter run time [176].
•HPLC Detectors
For the purpose of phytocannabinoid profiling, (U)HPLC analytical platforms are
coupled to UV, DAD, PAD or MS. Phytocannabinoids have low molar absorptivity, which
results in relatively low sensitivity of LC methods employing UV and DAD. This re-
stricts employment of DAD detection to low wavelengths where there is often strong
background absorbance from the eluent components, especially during gradient elution
experiments [
176
]. Additionally, UV/DAD methods often have low specificity for some
phytocannabinoids, e.g., CBDA and CBGA, due to similar UV/DAD spectra [178].

Molecules 2022,27, 975 28 of 42
Phytocannabinoids have different UV behaviour on the basis of their chemical struc-
ture. Cannabinoid acids (CBDA and CBGA) are characterized by three absorption maxima
(
λmax
), one stronger at 220–223 nm, the second at 266–270 nm and the third one around
305 nm, while neutral phytocannabinoids (CBD and CBG) show a first
λmax
at
210–215 nm
and an additional one at 270 nm. Generally, the ranges 190–600 nm and 200–650 nm are
most commonly used for UV acquisition, while two wavelengths are selected–210 nm for
neutral phytocannabinoids and 220 nm for cananbinolic acids [
43
,
72
,
185
]. For DAD, a
narrower range is selected (i.e., 200–400 nm [
83
], 190–500 nm [
182
]). Single wavelength can
be selected for evaluation of multiple phytocannabinoids in hemp seed oils [
179
], in plant
material (230 nm) [68], 214 nm [188,189] or 220 nm in plant material and resins [69,165] or
cannabis extracts [
27
], in commercial veterinary supplements (225 nm) [
192
] or for quan-
tification of
∆9
-THC,
∆9
-THCA, CBN and CBD (210 nm) [
191
] or
∆9
-THC and
∆9
-THCA
in plant material (211 nm and 220 nm) [
185
]. For wide range methods, such as cannabis-
based medical extracts [
182
], cannabis-infused cholcolate [
65
] and 17 phytocannabinoids
in cannabis inflorescences and oils [
100
], 228 nm has been shown to be the most suitable.
Multiple detection wavelengths, e.g., 220 nm, 240 nm, 270 nm and 307 nm are also used for
phytocannabinoids profiling [111,190]. Evaluation of these detection wavelengths for five
phytocannabinoids (CBD, CBDA, CBN,
∆9
-THC and
∆9
-THCA) in versatile cannabis-based
products was performed. While none of the phytocannabinoids showed
λmax
at 240 nm,
this wavelength tended to equalize the response (slope) across the five phytocannabinoids,
except for CBN, which has the highest response. The highest response for all five com-
ponents was found with 220 nm and was useful for low level quantification (Table S2).
270 nm
and 307 nm provided high selectivity for CBDA and
∆9
-THCA, which was used to
minimize or eliminate detection of matrix interferences, as needed. Detection wavelengths
in the range of 270–280 nm are inappropriate for CBD, CBN, and
∆9
-THC due to retention
time (Rt) interferences [90].
Although many studies employ UV/DAD for separation of the major phytocannabi-
noids (CBDA, CBD, CBN,
∆9
-THC and
∆9
-THCA), most do not account for interference
from minor phytocannabinoids (e.g, CBNA). Such interference is of special importance dur-
ing profiling of concentrates where minor phytocannabinoids can be enriched to detectable
levels. Additionally, some terpenes absorb UV light at the same wavelength as phyto-
cannabinoids. All these issues decrease sensitivity, specificity and selectivity of UV/DAD
methods, which are easily overcome by employment of MS detection. Detection of neutral
phytocannabinoids based on their ability to absorb fluorescence under the acidic conditions
used in RP-LC is only recently reported [
111
] as a fast, low-cost and selective alternative,
but without the ability to detect phytocannabinoid acids and a somewhat narrower linear
range than DAD due to saturation at high concentrations.
Most commonly used MS analyzers include simple Q spectrometers [
181
], QQQ
analyzers [
11
,
58
,
68
,
97
,
114
,
147
,
193
,
194
], IT analyzers [
28
,
43
,
72
,
185
], QTRAP [
67
,
159
,
191
,
195
]
or high resolution-accurate mass MS (HRAM-MS) analysers, such as Q-TOF [
83
,
88
,
139
,
189
]
and Q-ExactiveTM Orbitrap [19,63,65,66].
Phytocannabinoid profiling is most commonly performed using ESI, rarely using
variations such as dual source [
78
] or heated ESI source [
19
,
63
]. Acquisitions are per-
formed in positive (+) [
88
,
147
,
189
,
191
,
193
,
194
], negative (
−
) [
134
] or both (+) and (
−
)
modes [27,40,55,62–64,92,144,178,181]
. In fact, all major phytocannabinoids are detected
in both ionization modes, except for THCV [
78
]. However, it is noted that acidic phyto-
cannabinoids (CBDA, THCA, CBCA, CBGA, CBNA, CBCVA) give better signals in the
(
−
) mode, while neutral phytocannabinoids are better ionized in (+) mode (CBDV, CBG,
CBD, CBC, CBDA, CBDVA, CBGA,
∆9
-THCA,
∆9
-THC and
∆8
-THC) [
43
,
58
,
83
]. ESI (
−
)
provides identification and quantification of additional minor phytocannabinoids, such as
CBGA methyl ester (CBGMA) [
40
] and improves identification accuracy for two neutral
phytocannabinoids (CBL and CBN) [78].
Atmospheric pressure chemical ionization (APCI) is also used in (+) [
11
] and (
−
)
mode [
69
] for phytocannabinoid profiling in plant material. Formation of (+) charged

Molecules 2022,27, 975 29 of 42
sodium adducts [M+Na]
+
instead of the precursor ion (M+ H)
+
is observed in (+) ionization
mode. This feature increases sensitivity and allows accurate identification of 7 phyto-
cannabinoids (cannabicoumaric acid, CBCA, CBGA, CBGAM, 10-ethoxy-9-hydroxy-
∆6α
-
THC, 4-acetoxycannabichromeand
∆9
-THCA-C4). APCI (
−
) is shown to be suitable for
CBD, CBG and CBGA [69].
MS information from HRAM-MS acquisitions, that is, the accurate mass and mass
fragmentation patterns, is used for untargeted phytocannabinoid
profiling [19,63,65,66,83]
.
Compound identification is performed using one or more MSLs, such as mzCloud (High-
Chem LLC, Bratislava, Slovakia) [
196
], in-house MSL, such as the recent MSL of LC-MS/MS
spectra of 94 phytocannabinoids accompanied with metadata (names, R
t
s, accurate masses,
fragmentation patterns and fragments structures) [
19
], compound DBs, such as and Chem-
Spider [
197
] and Human Metabolome DB (HMDB) [
198
] or cheminformatics software, such
as Compound Discoverer (Thermo Fischer Scientific, Waltham, MA, USA).
HPLC-UV high-resolution MS (HRMS) is employed for simultaneous quantification of
the two main impurities in “pure” commercial CBG samples (cannabigerovarin (CBGV) and
cannabigerobutol (CBGB)) with subsequent confirmation by comparison with synthesized
compounds [193].
•Matrix Effect
The sample, i.e., matrix type greatly affects selection of extraction technique, extraction
solvent(s), HPLC column, mobile phase and detection method, which further enhances
method sensitivity, selectivity and specificity. All three validation parameters are affected
by the presence of matrix constituents that co-extracts with phytocannabinoids, causing
signal alteration (suppression or enhancement). In that spirit, matrix effect is frequently
reported during LC-MS-based phytocannabinoid profiling. Cannabis plant material is
complex matrix with high fat, pigment and polar compounds content, being flavonoids
and terpenes most prominent. Cannabis-based products are much more versatile in terms
of fat-, sugar- and polar-interferences content, thus being more prominent to expressing
significant matrix suppression during instrumental analysis.
Significant polar matrix interferences are reported to occur when EtOH is used for
extraction of phytocannabinoids from honey, which also co-extracts several interfering
matrix components, such as flavonoids [
194
]. Matrix effect is also examined during phyto-
cannabinoid profiling of commercial products including oils, creams, and plant material.
No significant matrix effect is observed in oil for
∆9
-THC, CBD, CBDA and
∆9
-THCA
(110.4
±
116.0%, 105.4
±
112.2%, 96.3
±
117.8% and 92.7
±
107.8%, respectively); acceptable
matrix effects for
∆9
-THC and CBD in plant material (102.0
±
112.8% and 91.2
±
129.4%,
respectively) and in creams (79.4
±
93.1% and 83.8
±
100.2%, respectively). Matrix effect of
CBDA was especially pronounced in plant material, with signal enhancement particularly
at low concentrations [173].
2.2.3. SFC Methods
Despite being efficient, SFC is an analytical technique that has still not been fully
exploited for the analysis, separation and quantification of cannabis plants and cannabis-
based products, compared to GC and LC. The limited number of studies available report
SFC as a fast (8–10 min runtime), cost-effective method with high specificity and separation
power for phytocannabinoid profiling [
195
,
199
,
200
]. Prior to SFC analysis, derivatization
and/or decarboxylation of phytocannabinoids is not required, thus reducing the risk of
sample contamination (unlike GC). Additionally, SFC allows separation of the neutral from
the acidic phytocannabinoids, simultaneously due to the properties of the supercritical
fluids, offers shorter analysis time, better resolution and definitive identification in a
single chromatogram of cannabis products when compared to both GC-MS and HPLC
methods [
199
]. UHPSFC is shown to offer greater selectivity than UHPLC, but at the price
of lower sensitivity, as a result of considerable variation of the refractive index of CO
2
,
resulting in greater baseline noise [188].

Molecules 2022,27, 975 30 of 42
Recently, the development of ultra-high performance SFC (UHPSFC) improved re-
solving power and efficiency, such that SFC has regained its popularity as an alternative
phytocannabinoid profiling of cannabis plants and cannabis-based products. SFC combined
with a 2
µ
m particle size column offers rapid separation and when coupled to UV or MS
detection, offers highly efficient analysis of the main phytocannabinoids, using inexpensive
and environmentally friendly SC-CO
2
as solvent. On the other hand, UHPSFC is considered
as highly orthogonal technique which provides different elution order and relative retention
of the investigated components compared to UHPLC; therefore, in combination with MS, it
could increase the discrimination power of phytocannabinoids in complex matrices.
For the purpose of phytocannabinoid profiling, SFC using cyanopropyl silica packed
column was employed [
199
], while the reported UHPLSFC methods used BEH 2-EP (2-
ethylpyridine) column [
195
,
200
]. Evaluation of columns with different stationary phases,
Torus 1-AA (1-aminoanthracene), Viridis BEH-2EP (ethyl-pyridine) and Torus Diol (OH)
revealed that the latter column achieved the highest number of (although not completely
separated) peaks; thus, it is most appropriate for routine phytocannabinoid profiling [
188
].
SC-CO
2
is used with MeOH as co-solvent, with a constant (2%) [
188
] or gradually in-
creasing concentration from 2% to 7% [
199
], often with the addition of 0.1% formic acid
to improve peak shape of cannabinoid acids [
188
]. UHPSFC methods employ SC-CO
2
with isopropanol/MeCN (80:20, v/v) with 1% H2O and linear gradient [195,200]. SFC and
UHPSFC are usually coupled either to PDA detectors [
188
] or to mass analyzers, such as
APCI-QQQ [
199
] and ESI-Q mass analyser [
195
,
200
]. The reported linearity, sensitivity
and specificity confirm the potential of SFC and UHPSFC to become the main profiling
analytical platforms instead of GC- and LC-based methods in near future.
2.2.4. Vibrational Spectroscopy Methods
In the past decade, vibrational spectroscopy techniques (IR, NIR, MIR, FTIR and
Raman) emerged as process analytical tool in the pharmaceutical industry for monitoring
various quality attributes, and as such were recognized by United States Food and Drug
Administration (FDA) [
201
]. Their ability for high-throughput screening of large volume
of sample for a short period of time can exert sampling-based errors and provide rapid,
versatile and non-invasive approach in qualitative and quantitative profiling and growth
staging of cannabis plant and extracts (Table S3).
The vibrational IR and Raman spectroscopy are considered complementary techniques;
even though both are relying on different physical processes, their main observations are
focused on light-induced molecular excitation [
202
]. Vibrational spectroscopy is based
on sample’s absorption of light at a defined wavelength range, which occurs as a conse-
quence of the vibrational features of the sample that result in the formation of overtones
and combination bands that form the spectrum. The relationship between the spectrum
and the physicochemical properties of the sample are mathematically modelled using
various multivariate regression methods (principal component analysis (PCA) and incre-
ment PCA (iPCA) [
203
], alone [
204
] or together with partial least-square (PLS) regression
analysis [205,206]
, based on which the compound concentration in the sample is predicted.
NIR spectroscopy, in conjunction with multivariate data analysis, is a widely accepted
approach for the abovementioned analysis. Quantitative data generated by NIR strongly
agree with UHPLC-UV data, confirming the potential of employment of NIR spectroscopy
in routine monitoring of cannabis plant material and cannabis resins [
189
]. Dispersive NIR
and FT-NIR methods were developed for quantification of eight different phytocannabi-
noids (CBDV,
∆9
-THCV, CBD, CBC,
∆8
-THC,
∆9
-THC, CBG and CBN) in ground leaves
and inflorescences from C. sativa [
205
] and for discriminating illegal and legal cannabis
varieties [
207
]. A similar NIR method demonstrated sensitivity and specificity for CBD
quantification in different liquid pharmaceutical products, thus showcasing its potential
as a fast method for monitoring of CBD in the production process [
206
]. Moreover, an
NIR method for the growth staging of Cannabis plants was reported as being sensitive to
concentrations of phytocannabinoids and volatile substances in the samples, which are also

Molecules 2022,27, 975 31 of 42
correlated to the plant age, thus justifying the feasibility of the method for growth staging of
cannabis [203]. Dispersive NIR using a scanning monochromator [205], FT-NIR spectrom-
eter with integrated Michelson interferometer and a highly sensitive PbS detector [
205
],
FT-NIR spectrometer based on measurement by diffuse reflectance [
206
], NIR spectrometer
with In-Ga-As detector [
203
], and a handheld NIR with In-Ga-As array detector [
189
] were
so far employed in phytocannabinoid profiling in cannabis ground leaves and inflores-
cences [
205
], in cannabis seeds [
203
] and in liquid pharmaceutical products (medium-chain
triglyceride and propylene glycol-based formulations) [
206
]. Two handheld NIR devices
(NIR-S-GI and MicroNIR) are used for in-field determination of
∆9
-THC content in cannabis
inflorescences and cannabis resins. For this purpose, spectrophotometers with a larger
sample analysis window are more appropriate for highly heterogenous samples, such as
whole cannabis inflorescences [189].
The literature data regarding phytocanabinnoids’ structural and molecular analysis,
to date, is very scarce, lacking band assignation and in-depth structural analysis of the
molecules [
23
,
208
,
209
] despite the documented ability to provide chemical fingerprinting
and qualitative profiling of phytocannabinoids, especially of FTIR in quantification of
biological compounds in complex matrices [
210
]. However, the interest in qualitative
profiling of cannabis (identification/classification) and quantitative profiling of the main
phytocannabinoids is slowly gaining momentum in the last few years (Supporting infor-
mation, Table S3). A recent study for ATR mid-IR quantification of
∆9
-THC and CBD in
cannabis flowers and extracts described a stepwise approach in developing multivariate
quantification models accompanied by detailed band assignment of the mentioned phyto-
cannabinoids, both for pure compounds and analysed samples (complex matrices) [
21
]. In
a further study by the same group, the potential of ATR mid-IR as a process analytical tool
(PAT) for continuous monitoring of ∆9-THCA decarboxylation was showcased [22].
The new portable vibrational spectroscopy apparatus versions are very applicable,
especially for continuous monitoring of the main phytocannabinoids in all growth stages of
cannabis and cannabis-based products manufacturing. Thus, a handheld Raman spectrom-
eter in conjunction with orthogonal PLS-DA was utilized to construct a classification model
for discriminating
∆9
-THCA rich, CBD rich Cannabis plants and hemp (low
∆9
-THCA,
CBDA and CBD) [
211
,
212
]. Reference standards from the main phytocannabinoids were
used to perform a detailed band assignation of the spectra that were further correlated
with the loading plot of the multivariate models. In both cases, favourable accuracy de-
scriptors for the classification models were reported, thus showcasing the applicability
of the portable Raman device for accurate and fast Cannabis plant classification. Raman
spectroscopy is a highly sensitive analytical technique, that, due to the variety of monochro-
matic light sources, and the emergence of surface-enhanced Raman scattering (SERS),
stimulated resonance and coherent anti-Stokes Raman scattering (CARS) offers greatly
enhanced capability and resolution, especially in its imaging mode. CARS imaging at
different Raman vibrations, known as hyperspectral CARS imaging, is a spectroscopic
imaging technique with high-resolution capabilities for a chemical distinction that employs
sophisticated data processing methods [
208
]. This analytical method was used to analyze
the secondary metabolites (
∆9
-THCA and CBDA) in glandular cannabis trichomes with
distinct spatial resolution, without the need to extract the resin [
213
]. To get additional
morphological data, the authors superimposed the image over a single photon fluorescence
and SEM image of the trichomes. The similarity of the chemical fingerprints of the distinct
regions with the secondary metabolites was determined with hierarchical clustering analy-
sis (HCA). The proposed methodology enables an easy discrimination between trichomes
with high-∆9-THCA and high-CBDA content.
2.2.5. Other Analytical Techniques
•CE
CE is analytical platform used in cannabinoid profiling in only one study [
68
]. MeCN-
based background electrolyte (with 6.5 mM NaOH) in the presence of
β
-cyclodextrin

Molecules 2022,27, 975 32 of 42
(
β
CD), improving orthogonal separation media by transiently interacting with compounds
based on their geometry and polarity, was used to separate 14 phytocannabinoids (CBG,
CBGA, CBD, CBDA, CBN,
∆9
-THC, CBC, CBCA,
∆9
-THCA, THCV, CBDV and CBGVA) by
constant transition between the background electrolyte and
β
CD. CE performed better than
HPLC-DAD in terms of selectivity and runtime, but with significantly lower sensitivity.
CE’s variety, capillary electromatography (CEC), coupling the benefits of CE and
HPLC methods, has been used in phytocannabinoid profiling coupled with UV PDA
detector in only one study [
214
]. Baseline separation of seven phytocannabinoids (CBG,
CBD, CBN,
∆9
-THC,
∆8
-THC, CBC,
∆9
-THCA) was achieved by using C
18
column and
MeCN/25 mM phosphate buffer (75:25, v/v) as mobile phase for analysis of MeOH/CHCl
3
(9:1, v/v)cannabis plant material extracts, while improved sensitivity is achieved using UV
cell with extended path length and injection size.
•NMR Spectroscopy
NMR-based methods are superior for the purpose of 3D-structure elucidation, espe-
cially
ζ
-resolving,
1
H-
1
H COSY and
1
H-
13
C heteronuclear multiple quantum coherence
(HMQC) and heteronuclear multiple bond correlation (HMBC) spectroscopy. The complete
1
H- and
13
C-NMR assignments of the major Cannabis constituents,
∆9
-THC,
∆9
-THCA,
∆8
-THC, CBG, CBN, CBD, CBDA, cannflavin A and cannflavin B have been determined
on the basis of one- and two-dimensional NMR spectra, including
1
H- and
13
C-NMR,
1
H-
1
H-COSY, HMQC and HMBC [
20
]. However, they are seldomly used for quantification
purposes, due to the laborious separation and isolation steps required, where significant
loss of mass can occur and where there is low sensitivity [
4
,
215
]. Additional issues are the
high instrumental costs and necessity of highly specialized personnel. Despite that, NMR
is considered as a highly accurate, reproducible and fast technique [1].
For the purpose of phytocannabinoid profiling, NMR spectroscopy is used as (semi)qu-
antitative method alone [
20
,
209
] or as an orthogonal technique to LC [
216
–
218
] or GC [
219
]
for the purpose of qualitative peak assignment of major phytocannabinoids [
20
], chem-
ical and morphological examination [
220
], chemotaxonomic classification [
20
,
219
,
220
],
metabolomics-based chemovar distinction [
51
] or quantitative analysis of cannabis plant
material without the need of pre-purification step [
221
], chromatographic separation or use
of certified reference standards [
219
]. Cryogenic NMR spectroscopy combines improved
sensitivity and noise reduction with a cryogenic cooling system for the receiver coil and
preamplifiers. Its improved spectral quality is employed in compound identification from
mass limited samples and as orthogonal analytical technique to HPLC in phytocannabi-
noid profiling in laser-micro dissected samples of capitate-stalked and capitate-sessile
trichomes [220].
3. Conclusions and Future Directions
Scientific and technological advancements in cultivation, manufacturing, recreational,
industrial and medical use of cannabis, as well as updated legislation, led to the develop-
ment of multitude of analytical methods for phytocannabinoid profiling. Matrix nature
greatly affects selection of extraction technique, sample preparation and analytical method
due to the fact that significant matrix interferences can occur and aggravate the overall
analysis of target phytocannabinoids. Sample preparation for phytocannabinoid profiling
in the past four decades is mainly based on versatile types of accelerated maceration, such
as SLE, LLE, PLE, SPE, USE, FUSE and MAHD. Recently, new trends have enlightened
environmental-friendly techniques, such as easily-automatable HS-SPME and SFE, which
adds speed, repeatability and reproducibility to the analyses. From the multitude of analyt-
ical platforms, TLC and HPTLC, HPLC-DAD, GC and LC coupled with mass spectrometry
(MS or MS/MS), are most commonly used; however, recently emerging techniques are
NMR and vibrational spectroscopy methods, such as IR, NIR, FTIR and FT-NIR. TLC,
together with HPTLC, which is a suitable method for screening of samples and is included
in the pharmacopoeias in the identification methods. Cannabinoid profiling for research,
industrial and QC purposes is based mostly on two analytical platforms: GC and LC. GC

Molecules 2022,27, 975 33 of 42
coupled to versatile detectors and mass analyzers is one of the oldest, the most preferred
and researched analytical platforms for phytocannabinoid profiling due to its robustness,
reproducibility, sensitivity and speed.
As a result of the advancement of computational tools, mass spectral libraries (MSLs),
public compound repositories and compound databases (DBs), as well as various advanced
detection techniques, GC has become the analytical platform in forensic, pharmacokinetic
and phytochemical analysis of natural phytocannabinoids. As such, GC methods are also
officially employed by authorities for terpene profiling, pesticide screening and residual
solvents analysis, which affords potential benefits to regulatory bodies and cannabis indus-
try. High pressure and high flow rates used during TLC, HPTLC, HPLC and UHPLC, and
the recently emerging SFC technique allow sample preservation without decarboxylation
and decomposition, reliable separation of neutral and acidic cannabinoid species and, thus,
direct identification and quantification of both neutral and acidic forms of phytocannabi-
noids in the extracted samples. In the past decades, LC has become an analytical platform
of choice (HPLC-DAD and LC-MS) in first line for potency studies but also for untargeted
analysis of cannabis and cannabis-based products. Despite the fact that vibrational spec-
troscopy methods, such as NIR, FTIR, FT-NIR and Raman spectroscopy are reserved for
structural elucidation, in the last few years there is an evident trend of their utilization for
rapid quantitative phytocannabinoid profiling. Although these vibrational spectroscopy
techniques can provide rapid, versatile and non-invasive approach in qualitative and quan-
titative profiling and growth staging of cannabis plant and extracts, they demonstrate far
higher LOD and LOQ than the described chromatography-‘wet’ methods, they are fast,
inexpensive, non-destructive and require minimum (e.g., drying, grinding) or no sample
preparation. Regardless of high instrumental costs and necessity of highly specialized
personnel, NMR is considered as a highly accurate, reproducible and fast technique that
offers quantitative analysis of cannabis without the need of pre-purification step, chromato-
graphic separation or use of certified reference standards. Even though there are currently
various well-established methods available for chemical analyses of phytocannabinoids,
there is still a need for adaptations and enhancement of these methods in the light of new
scientific evidence regarding the plant and its plant metabolites, especially taking into
account the pharmacological activity and its medical use, association of medical and ad-
verse health effects with potency and/or interplay of certain phytocannabinoids and other
active constituents, quality control and stability studies of cannabis and cannabis-based
products. Further advancements in phytocannabinoid profiling should move towards un-
targeted analysis of cannabis plant material and cannabis-based products using orthogonal
analytical methods. By employment of cheminformatics approaches for small molecule
identification and MSLs, a multitude of new phytocannabinoids and other compounds is
expected to be identified in the near future, thus allowing access to complete and accurate
phytocannabinoid and terpene profiles.
Supplementary Materials:
The following are available online. Table S1: GC-based analytical methods
for cannabinoid profiling [
5
,
9
,
10
,
17
,
18
,
43
–
49
,
51
–
59
,
66
,
67
,
70
–
75
,
77
–
79
,
81
,
82
,
84
–
91
,
93
–
99
,
101
–
108
,
114
–
116
,
118
,
119
,
121
–
124
,
128
,
132
,
136
–
141
,
148
,
150
,
152
–
158
,
160
,
161
,
163
,
166
,
216
,
222
–
224
];
Table S2: LC-based
ana-
lytical methods for cannabinoid profiling [
11
,
19
,
27
,
40
,
42
,
50
,
55
,
60
–
65
,
68
,
69
,
78
,
80
,
83
,
92
,
100
,
107
,
109
,
111
,
113
,
133
,
134
,
142
,
144
,
149
,
165
,
173
,
176
–
180
,
182
,
183
,
185
–
188
,
190
–
192
,
194
,
200
,
215
,
217
];
Table S3: Vibrational
spectroscopy-based analytical methods in conjunction with multivariate data analysis for phytocannabi-
noid profiling and/or classification of cannabis plant material [21,189,203–207,211,212,218].
Author Contributions:
Conceptualization, G.S. and S.K.; methodology, I.C.K.; investigation, V.S.G.
and A.T.; data curation, I.C.K., V.S.G. and A.T.; writing—original draft preparation, I.C.K. and N.G.;
writing—review and editing G.S., M.K. and S.K.; supervision, G.S. and S.K; project administration,
G.S.; funding acquisition, S.K. and G.S. All authors have read and agreed to the published version of
the manuscript.
Funding: This research received no external funding.
Conflicts of Interest: The authors declare no conflict of interest.

Molecules 2022,27, 975 34 of 42
Sample Availability: Samples of the compounds are available from the authors.
References
1.
Citti, C.; Braghiroli, D.; Vandelli, M.A.; Cannazza, G. Pharmaceutical and biomedical analysis of cannabinoids: A critical review.
J. Pharm. Biomed. Anal. 2018,147, 565–579. [CrossRef] [PubMed]
2.
Borille, B.T.; González, M.; Steffens, L.; Ortiz, R.S.; Limberger, R.P. Cannabis sativa: A systematic review of plant analysis. Drug
Anal. Res. 2017,1, 1–23. [CrossRef]
3.
Sawler, J.; Stout, J.M.; Gardner, K.M.; Hudson, D.; Vidmar, J.; Butler, L.; Page, J.E.; Myles, S. The Genetic Structure of Marijuana
and Hemp. PLoS ONE 2015,10, e0133292. [CrossRef] [PubMed]
4.
Ramirez, C.L.; Fanovich, M.A.; Churio, M.S. Cannabinoids: Extraction Methods, Analysis, and Physicochemical Characterization.
In Studies in Natural Products Chemistry; Elsevier: Amsterdam, The Netherlands, 2019; pp. 143–173. [CrossRef]
5.
United Nations Office on Drugs and Crime. Recommended Methods for the Identification and Analysis of Cannabis and Cannabis
Products Manual for Use by National Drug Analysis Laboratories; United Nations: New York, NY, USA, 2009. Available online:
http://search.ebscohost.com/login.aspx?direct=true&scope=site&db=nlebk&db=nlabk&AN=450863 (accessed on 5 July 2020).
6. Small, E.; Cronquist, A. A Practical and Natural Taxonomy for Cannabis. Taxon 1976,25, 405–435. [CrossRef]
7.
Small, E. Evolution and Classification of Cannabis sativa (Marijuana, Hemp) in Relation to Human Utilization. Bot. Rev.
2015
,81,
189–294. [CrossRef]
8.
ElSohly, M.A.; Slade, D. Chemical constituents of marijuana: The complex mixture of natural cannabinoids. Life Sci.
2005
,78,
539–548. [CrossRef]
9.
Mehmedic, Z.; Chandra, S.; Slade, D.; Denham, H.; Foster, S.; Patel, A.S.; Ross, S.A.; Khan, I.A.; ElSohly, M.A. Potency Trends of
∆
9-THC and Other Cannabinoids in Confiscated Cannabis Preparations from 1993 to 2008*. J. Forensic Sci.
2010
,55, 1209–1217.
[CrossRef] [PubMed]
10.
de Meijer, E.P.M.; Bagatta, M.; Carboni, A.; Crucitiit, P.; Moliterni, C.V.M.; Ranalli, P.; Mandolino, G. The Inheritance of Chemical
Phenotype in Cannabis sativa L. Genetics 2003,163, 335–346. [CrossRef] [PubMed]
11.
Aizpurua-Olaizola, O.; Omar, J.; Navarro, P.; Olivares, M.; Etxebarria, N.; Usobiaga, A. Identification and quantification of
cannabinoids in Cannabis sativa L. plants by high performance liquid chromatography-mass spectrometry. Anal. Bioanal. Chem.
2014,406, 7549–7560. [CrossRef]
12. Mechoulam, R.; Shvo, Y. The structure of cannabidiol. Tetrahedron 1963,19, 2073–2078. [CrossRef]
13.
Gaoni, Y.; Mechoulam, R. Isolation, Structure, and Partial Synthesis of an Active Constituent of Hashish. J. Am. Chem. Soc.
1964
,
86, 1646–1647. [CrossRef]
14. Gaoni, Y.; Mechoulam, R. Cannabichromene, a new active principle in hashish. Chem. Commun. 1966, 20–21. [CrossRef]
15. Vollner, L.; Bieniek, D.; Korte, F. Cannabidivarin, ein neur Haschisch-Inhalsstoff. Tetrahedron Lett. 1969,10, 145–147. [CrossRef]
16.
Gill, E.W.; Paton, W.D.M.; Pertwee, R.G. Preliminary Experiments on the Chemistry and Pharmacology of Cannabis. Nature
1970
,
228, 134–136. [CrossRef]
17.
Tipparat, P.; Kunkaew, W.; Julsrigival, S.; Pinmanee, S.; Natakankitkul, S. Classification of Cannabis Plants Grown in Northern
Thailand using physico-chemical properties. J. Nat. Sci. Res. 2014,10, 46–54.
18.
Hazekamp, A.; Peltenburg, A.; Verpoorte, R.; Giroud, C. Chromatographic and Spectroscopic Data of Cannabinoids from Cannabis
sativa L. J. Liq. Chromatogr. Relat. Technol. 2005,28, 2361–2382. [CrossRef]
19.
Berman, P.; Futoran, K.; Lewitus, G.M.; Mukha, D.; Benami, M.; Shlomi, T.; Meiri, D. A new ESI-LC/MS approach for comprehen-
sive metabolic profiling of phytocannabinoids in Cannabis. Sci. Rep. 2018,8, 14280. [CrossRef]
20.
Choi, Y.H.; Hazekamp, A.; Peltenburg-Looman, A.M.G.; Frédérich, M.; Erkelens, C.; Lefeber, A.W.M.; Verpoorte, R. NMR
assignments of the major cannabinoids and cannabiflavonoids isolated from flowers of Cannabis sativa. Phytochem. Anal.
2004
,
15, 345–354. [CrossRef]
21.
Geskovski, N.; Stefkov, G.; Gigopulu, O.; Stefov, S.; Huck, C.W.; Makreski, P. Mid-infrared spectroscopy as process analytical
technology tool for estimation of THC and CBD content in Cannabis flowers and extracts. Spectrochim. Acta Part A Mol. Biomol.
Spectrosc. 2021,251, 119422. [CrossRef]
22.
Gigopulu, O.; Geskovski, N.; Stefkov, G.; Stoilkovska, V.; Spirevska, I.S.; Huck, C.W.; Makreski, P. Synergistic use of FTIR
spectroscopy and TG to elucidate the solid state THCA decarboxylation reaction kinetics in THCA standard and cannabis flower.
Anal. Chem. 2022,267, 18.
23.
Russo, E.B. History of Cannabis and Its Preparations in Saga, Science, and Sobriquet. Chem. Biodivers.
2007
,4, 1614–1648.
[CrossRef]
24. Pertwee, R.G. Handbook of Cannabis. In Oxford Scholarship; Oxford University Press: London, UK, 2014.
25.
Leonard, W.; Zhang, P.; Ying, D.; Fang, Z. Hempseed in food industry: Nutritional value, health benefits, and industrial
applications. Compr. Rev. Food Sci. Food Saf. 2020,19, 282–308. [CrossRef]
26.
Geshtakovska, G.; Stefkov, G. Routes of cannabis administration: A brief review. In Modern Pharmacist-Bridging Science with
Practice; Macedonian Pharmaceutical Bulletin; Macedonian Pharmaceutical Association: Ohrid, Macedonia, 2016; pp. 515–516.
27.
Muscarà, C.; Smeriglio, A.; Trombetta, D.; Mandalari, G.; la Camera, E.; Grassi, G.; Circosta, C. Phytochemical characterization
and biological properties of two standardized extracts from a non-psychotropic Cannabis sativa L. cannabidiol (CBD)-chemotype.
Phytother. Res. 2021,35, 5269–5281. [CrossRef]

Molecules 2022,27, 975 35 of 42
28.
Klein, R.F.X. Analysis of “Marijuana Edibles”—Food Products Containing Marijuana or Marijuana Extracts—An Overview,
Review, and Literature Survey. Microgram J. 2017,14, 24.
29. Johnson, R. Hemp as an Agricultural Commodity; Congressional Research Service: Washington, DC, USA, 2018.
30.
European Commission. Common Catalogue of Varieties of Agricultural Plant Species. In Official Journal of the European Union;
European Commission: Brussels, Belgium, 2012; p. 39.
31.
European Commission. Commission Regulation (EC) No. 327/2002. In Official Journal of the European Communities; European
Commission: Brussels, Belgium, 2002.
32.
Vereinte Nationen, Büro für Drogenkontrolle und Verbrechensbekämpfung. World Drug Report; Vereinte Nationen: New York,
NY, USA, 2018.
33.
Upton, R.; Craker, L.; ElSohly, M.; Romm, A.; Russo, E.; Sexton, M. American Herbal Pharmacopoeia, Cannabis Inflorescence: Cannabis
spp.; Standards of Identity, Analysis, and Quality Control; American Herbal Pharmacopoeia: Scotts Valley, CA, USA, 2013.
34.
Swissmedic Swiss Agency for Therapeutic Products Pharmacopoeia Division. Fleur de Cannabis. In Pharmacopoeia Helvetica, 11th
ed.; Supplement 11.3; Swissmedic Swiss Agency for Therapeutic Products Pharmacopoeia Division: Bern, Switzerland, 2019.
35.
Council of Europe. Cannabis flos Monograph N
◦
: 3028. In European Pharmacopoeia, 10th ed.; Council of Europe: Strasbourg,
France, 2020.
36.
Geschäftsstelle der Arzneibuch-Kommissionen. Bundesinstitut fur Arzneimittel und Miedizinprodukte, Monografie Cannabis-
blüten. In German Pharmacopoeia, 2020th ed.; Geschäftsstelle der Arzneibuch-Kommissionen: Bundesinstitut fur Arzneimittel und
Miedizinprodukte: Bonn, German, 2020.
37.
Fernandez, E.B.; Peterseil, V.; Hackl, G.; Menges, S.; de Meijer, E.; Staginnus, C. Distribution of Chemical Phenotypes (Chemotypes)
in European Agricultural Hemp (Cannabis sativa L.) Cultivars. J. Forensic Sci. 2020,65, 715–721. [CrossRef]
38.
Geschäftsstelle der Arzneibuch-Kommissionen. Bundesinstitut fur Arzneimittel und Miedizinprodukte, Monografie Cannabis
extractum normatum (Eingestellter Cannabis-Extrakt). In German Pharmacopoeia; Geschäftsstelle der Arzneibuch-Kommissionen,
Bundesinstitut fur Arzneimittel und Miedizinprodukte: Bern, Switzerland, 2020.
39.
Nahar, L.; Guo, M.; Sarker, S.D. Gas chromatographic analysis of naturally occurring cannabinoids: A review of literature
published during the past decade. Phytochem. Anal. 2020,31, 135–146. [CrossRef]
40.
Brighenti, V.; Pellati, F.; Steinbach, M.; Maran, D.; Benvenuti, S. Development of a new extraction technique and HPLC method for
the analysis of non-psychoactive cannabinoids in fibre-type Cannabis sativa L. (hemp). J. Pharm. Biomed. Anal.
2017
,143, 228–236.
[CrossRef]
41.
Naviglio, D.; Scarano, P.; Ciaravolo, M.; Gallo, M. Rapid Solid-Liquid Dynamic Extraction (RSLDE): A Powerful and Greener
Alternative to the Latest Solid-Liquid Extraction Techniques. Foods 2019,8, 245. [CrossRef]
42.
Knezevic, F.; Nikolai, A.; Marchart, R.; Sosa, S.; Tubaro, A.; Novak, J. Residues of herbal hemp leaf teas—How much of the
cannabinoids remain? Food Control. 2021,127, 108146. [CrossRef]
43.
Omar, J.; Olivares, M.; Alzaga, M.; Etxebarria, N. Optimisation and characterisation of marihuana extracts obtained by supercritical
fluid extraction and focused ultrasound extraction and retention time locking GC-MS: Gas Chromatography. J. Sep. Sci.
2013
,36,
1397–1404. [CrossRef] [PubMed]
44.
Fairbairn, J.W.; Liebmann, J.A. The extraction and estimation of the cannabinoids in Cannabis sativa L. and its products. J. Pharm.
Pharmacol. 1973,25, 150–155. [CrossRef] [PubMed]
45.
Lewis, R.; Ward, S.; Johnson, R.; Burns, D.T. Distribution of the principal cannabinoids within bars of compressed cannabis resin.
Anal. Chim. Acta. 2005,538, 399–405. [CrossRef]
46. UNODC. Bulletin on Narcotics; Issue 1—005, Office on Drugs and Crime: Vienna, Austria, 1971.
47.
Mechtler, K.; Bailer, J.; de Hueber, K. Variations of
∆
9-THC Content in Single Plants of Hemp Varieties. Ind. Crops Prod.
2004
,19,
19–24. [CrossRef]
48. Niesink, R.J.M.; Rigter, S.; Koeter, M.W.; Brunt, T.M. Potency trends of ∆9-tetrahydrocannabinol, cannabidiol and cannabinol in
cannabis in the Netherlands: 2005-15: Potency trends of Dutch cannabis. Addiction 2015,110, 1941–1950. [CrossRef]
49.
Tsumura, Y.; Aoki, R.; Tokieda, Y.; Akutsu, M.; Kawase, Y.; Kataoka, T.; Takagi, T.; Mizuno, T.; Fukada, M.; Fujii, H.; et al. A
survey of the potency of Japanese illicit cannabis in fiscal year 2010. Forensic Sci. Int. 2012,221, 77–83. [CrossRef]
50.
Jin, D.; Dai, K.; Xie, Z.; Chen, J. Secondary Metabolites Profiled in Cannabis Inflorescences, Leaves, Stem Barks, and Roots for
Medicinal Purposes. Sci. Rep. 2020,10, 3309. [CrossRef]
51.
Fischedick, J.T.; Hazekamp, A.; Erkelens, T.; Choi, Y.H.; Verpoorte, R. Metabolic fingerprinting of Cannabis sativa L., cannabinoids
and terpenoids for chemotaxonomic and drug standardization purposes. Phytochemistry 2010,71, 2058–2073. [CrossRef]
52.
Hillig, K.W.; Mahlberg, P.G. A chemotaxonomic analysis of cannabinoid variation in Cannabis (Cannabaceae). Am. J. Bot.
2004
,
91, 966–975. [CrossRef]
53.
Hazekamp, A.; Tejkalová, K.; Papadimitriou, S. Cannabis: From Cultivar to Chemovar II—A Metabolomics Approach to Cannabis
Classification. Cannabis Cannabinoid Res. 2016,1, 202–215. [CrossRef]
54.
Bruci, Z.; Papoutsis, I.; Athanaselis, S.; Nikolaou, P.; Pazari, E.; Spiliopoulou, C.; Vyshka, G. First systematic evaluation of the
potency of Cannabis sativa plants grown in Albania. Forensic Sci. Int. 2012,222, 40–46. [CrossRef]
55.
Béres, T.; ˇ
Cernochová, L.; Zeljkovi´c, S. ´
C.; Benická, S.; Gucký, T.; Berˇcák, M.; Tarkowski, P. Intralaboratory comparison of analytical
methods for quantification of major phytocannabinoids. Anal. Bioanal. Chem. 2019,411, 3069–3079. [CrossRef]

Molecules 2022,27, 975 36 of 42
56.
Gröger, T.; Schäffer, M.; Pütz, M.; Ahrens, B.; Drew, K.; Eschner, M.; Zimmermann, R. Application of two-dimensional gas chro-
matography combined with pixel-based chemometric processing for the chemical profiling of illicit drug samples.
J. Chromatogr. A
2008,1200, 8–16. [CrossRef]
57.
Janatová, A.; Fraˇnková, A.; Tlustoš, P.; Hamouz, K.; Božik, M.; Klouˇcek, P. Yield and cannabinoids contents in different cannabis
(Cannabis sativa L.) genotypes for medical use. Ind. Crops Prod. 2018,112, 363–367. [CrossRef]
58.
Field, B.I.; Arndt, R.R. Cannabinoid compounds in South African Cannabis sativa L. J. Pharm. Pharmacol.
1980
,32, 21–24. [CrossRef]
[PubMed]
59.
Ilias, Y.; Rudaz, S.; Mathieu, P.; Christen, P.; Veuthey, J.-L. Extraction and analysis of different Cannabis samples by headspace
solid-phase microextraction combined with gas chromatography-mass spectrometry. J. Sep. Sci. 2005,28, 2293–2300. [CrossRef]
[PubMed]
60.
Pavlovic, R.; Nenna, G.; Calvi, L.; Panseri, S.; Borgonovo, G.; Giupponi, L.; Cannazza, G.; Giorgi, A. Quality Traits of “Cannabidiol
Oils”: Cannabinoids Content, Terpene Fingerprint and Oxidation Stability of European Commercially Available Preparations.
Molecules 2018,23, 1230. [CrossRef]
61.
Giupponi, L.; Leoni, V.; Pavlovic, R.; Giorgi, A. Influence of Altitude on Phytochemical Composition of Hemp Inflorescence: A
Metabolomic Approach. Molecules 2020,25, 1381. [CrossRef]
62.
Pavlovic, R.; Panseri, S.; Giupponi, L.; Leoni, V.; Citti, C.; Cattaneo, C.; Cavaletto, M.; Giorgi, A. Phytochemical and Ecological
Analysis of Two Varieties of Hemp (Cannabis sativa L.) Grown in a Mountain Environment of Italian Alps. Front. Plant Sci.
2019
,
10, 1265. [CrossRef]
63.
Calvi, L.; Pentimalli, D.; Panseri, S.; Giupponi, L.; Gelmini, F.; Beretta, G.; Vitali, D.; Bruno, M.; Zilio, E.; Pavlovic, R.; et al.
Comprehensive quality evaluation of medical Cannabis sativa L. inflorescence and macerated oils based on HS-SPME coupled to
GC–MS and LC-HRMS (q-exactive orbitrap®) approach. J. Pharm. Biomed. Anal. 2018,150, 208–219. [CrossRef]
64.
Pisciottano, I.D.M.; Guadagnuolo, G.; Soprano, V.; Esposito, M.; Gallo, P. A survey of
∆
9-THC and relevant cannabinoids in
products from the Italian market: A study by LC–MS/MS of food, beverages and feed. Food Chem.
2021
,346, 128898. [CrossRef]
65.
Reyes-Garcés, N.; Myers, C. Analysis of the California list of pesticides, mycotoxins, and cannabinoids in chocolate using liquid
chromatography and low-pressure gas chromatography-based platforms. J. Sep. Sci. 2021,44, 2564–2576. [CrossRef]
66.
Namdar, D.; Mazuz, M.; Ion, A.; Koltai, H. Variation in the compositions of cannabinoid and terpenoids in Cannabis sativa
derived from inflorescence position along the stem and extraction methods. Ind. Crops Prod. 2018,113, 376–382. [CrossRef]
67.
Stambouli, H.; Bouri, A.E.; Bellimam, M.A.; Bouayoun, T.; Karni, N.E. Cultivation of Cannabis sativa L. in northern Morocco. Bull.
Narc. 2005,57, 79–118. [PubMed]
68.
Zaripov, E.A.; Lee, T.; Dou, Y.; Harris, C.S.; Egorov, A.; Berezovski, M.V. Single-Run Separation and Quantification of 14 Cannabi-
noids Using Capillary Electrophoresis. Separations 2021,8, 30. [CrossRef]
69.
Kalinová, J.P.; Vrchotová, N.; Tˇríska, J.; Hellerová, Š. Industrial hemp (Cannabis sativa L.) as a possible source of cannabidiol.
J. Cent. Eur. Agric. 2021,22, 110–118. [CrossRef]
70.
European Commission. Union method for the quantitative determination of the
∆
9-tetrahydrocannabinol content in hemp
varieties. Delegated regulation (EU) No 639/2014, Annex III as amended by Regulation (EU) 2017/1155. In Official Journal of the
European Communities; 2017; Volume 167, pp. 1–15. Available online: http://data.europa.eu/eli/reg_del/2017/1155/oj (accessed
on 8 February 2021).
71.
Hewavitharana, A.K.; Golding, G.; Tempany, G.; King, G.; Holling, N. Quantitative GC-MS Analysis of
∆
9-Tetrahydrocannabinol
in Fiber Hemp Varieties. J. Anal. Toxicol. 2005,29, 258–261. [CrossRef]
72.
Chandra, S.; Lata, H.; Mehmedic, Z.; Khan, I.; ElSohly, M. Assessment of Cannabinoids Content in Micropropagated Plants of
Cannabis sativa and Their Comparison with Conventionally Propagated Plants and Mother Plant during Developmental Stages of
Growth. Planta Med. 2010,76, 743–750. [CrossRef]
73.
Delta9-THC determination by the EU official method: Evaluation of Measurement Uncertainty and Compliance Assessment of
Hemp Samples|SpringerLink, (n.d.). Available online: https://link.springer.com/article/10.1007/s00216-021-03283-x (accessed
on 18 July 2021).
74.
Florian-Ramirez, N.M.; Parada-Alfonso, F.; Garzon-Mendez, W.F. Estudio del contenido de cannabinoids en muestras de
marihuana (Cannabis sativa L.) cultivadas en varies regiones de Colombia, Vitae. Rev. De La Faculdad De Quim. Farm.
2009
,16,
237–244.
75.
Pacifico, D.; Miselli, F.; Micheler, M.; Carboni, A.; Ranalli, P.; Mandolino, G. Genetics and Marker-assisted Selection of the
Chemotype in Cannabis sativa L. Mol. Breed. 2006,17, 257–268. [CrossRef]
76.
Pacifico, D.; Miselli, F.; Carboni, A.; Moschella, A.; Mandolino, G. Time course of cannabinoid accumulation and chemotype
development during the growth of Cannabis sativa L. Euphytica 2008,160, 231–240. [CrossRef]
77.
Turner, J.C.; Hemphill, J.K.; Mahlberg, P.G. Gland distribution and cannabinoid content in clones of Cannabis sativa L. Am. J. Bot.
1977,64, 687–693. [CrossRef]
78.
Delgado-Povedano, M.M.; Callado, C.S.; Priego-Capote, F.; Ferreiro-Vera, C. Untargeted characterization of extracts from Cannabis
sativa L. cultivars by gas and liquid chromatography coupled to mass spectrometry in high resolution mode. Talanta
2020
,208,
120384. [CrossRef]
79.
Tipparat, P.; Natakankitkul, S.; Chamnivikaipong, P.; Chutiwat, S. Characteristics of cannabinoids composition of Cannabis plants
grown in Northern Thailand and its forensic application. Forensic Sci. Int. 2012,215, 164–170. [CrossRef]

Molecules 2022,27, 975 37 of 42
80.
Nuapia, Y.; Maraba, K.; Tutu, H.; Chimuka, L.; Cukrowska, E. In Situ Decarboxylation-Pressurized Hot Water Extraction for
Selective Extraction of Cannabinoids from Cannabis sativa. Chemometric Approach. Molecules
2021
,26, 3343. [CrossRef]
[PubMed]
81.
Cadola, L.; Broséus, J.; Esseiva, P. Chemical profiling of different hashish seizures by gas chromatography–mass spectrometry
and statistical methodology: A case report. Forensic Sci. Int. 2013,232, e24–e27. [CrossRef] [PubMed]
82.
Zamengo, L.; Frison, G.; Bettin, C.; Sciarrone, R. Variability of cannabis potency in the Venice area (Italy): A survey over the
period 2010–2012: Variability of cannabis potency in the Venice area (Italy): A survey over the period 2010–2012. Drug Test. Anal.
2014,6, 46–51. [CrossRef] [PubMed]
83.
de Backer, B.; Debrus, B.; Lebrun, P.; Theunis, L.; Dubois, N.; Decock, L.; Verstraete, A.; Hubert, P.; Charlier, C. Innovative
development and validation of an HPLC/DAD method for the qualitative and quantitative determination of major cannabinoids
in cannabis plant material. J. Chromatogr. B 2009,877, 4115–4124. [CrossRef] [PubMed]
84.
Cardenia, V.; Toschi, T.G.; Scappini, S.; Rubino, R.C.; Rodriguez-Estrada, M.T. Development and validation of a Fast gas
chromatography/mass spectrometry method for the determination of cannabinoids in Cannabis sativa L. J. Food Drug Anal.
2018
,
26, 1283–1292. [CrossRef]
85. Kaa, E. Cannabis plants illicitly grown in Jutland (Denmark). Z. Rechtsmed. 1989,102, 367–375. [CrossRef]
86.
de CássiaMariotti, K.; Marcelo, M.C.A.; Ortiz, R.S.; Borille, B.T.; Reis, M.d.; Fett, M.S.; Ferrão, M.F.; Limberger, R.P. Seized cannabis
seeds cultivated in greenhouse: A chemical study by gas chromatography–mass spectrometry and chemometric analysis. Sci.
Justice 2016,56, 35–41. [CrossRef]
87.
Hazekamp, A.; Simons, R.; Peltenburg-Looman, A.; Sengers, M.; van Zweden, R.; Verpoorte, R. Preparative Isolation of
Cannabinoids from Cannabis sativa by Centrifugal Partition Chromatography. J. Liq. Chromatogr. Relat. Technol.
2004
,27,
2421–2439. [CrossRef]
88.
Fodor, B.; Boldizsár, I.; Molnár-Perl, I. Alkylsilyl speciation and direct sample preparation of plant cannabinoids prior to their
analysis by GC-MS. Anal. Chim. Acta 2018,1021, 51–59. [CrossRef] [PubMed]
89.
Jang, E.; Kim, H.; Jang, S.; Lee, J.; Baeck, S.; In, S.; Kim, E.; Kim, Y.; Han, E. Concentrations of THC, CBD, and CBN in commercial
hemp seeds and hempseed oil sold in Korea | Elsevier Enhanced Reader. Forensic Sci. Int. 2020,306, 110064. [CrossRef]
90.
Ciolino, L.A.; Ranieri, T.L.; Taylor, A.M. Commercial cannabis consumer products part 1: GC–MS qualitative analysis of cannabis
cannabinoids. Forensic Sci. Int. 2018,289, 429–437. [CrossRef]
91.
Yotoriyama, M.; Ishiharajima, E.; Kato, Y.; Nagato, A.; Sekita, S.; Watanabe, K.; Yamamoto, I. Identification and Determination of
Cannabinoids in both Commercially Available and Cannabis Oils Stored Long Term. J. Health Sci. 2005,51, 483–487. [CrossRef]
92.
Scheunemann, A.; Elsner, K.; Germerott, T.; Hess, C.; Zörntlein, S.; Röhrich, J. Extensive phytocannabinoid profiles of seized
cannabis and cannabis-based medicines—Identification of potential distinguishing markers. Forensic Sci. Int.
2021
,322, 110773.
[CrossRef] [PubMed]
93.
Pitts, J.E.; O’Neil, P.J.; Leggo, K.P. Variation in the THC Content of Illicitly Imported Cannabis* Products-1984–1989. J. Pharm.
Pharmacol. 1990,42, 817–820. [CrossRef] [PubMed]
94.
Casiraghi, A.; Roda, G.; Casagni, E.; Cristina, C.; Musazzi, U.; Franzè, S.; Rocco, P.; Giuliani, C.; Fico, G.; Minghetti, P.; et al.
Extraction Method and Analysis of Cannabinoids in Cannabis Olive Oil Preparations. Planta Med.
2018
,84, 242–249. [CrossRef]
95.
Wianowska, D.; Dawidowicz, A.L.; Kowalczyk, M. Transformations of Tetrahydrocannabinol, tetrahydrocannabinolic acid and
cannabinol during their extraction from Cannabis sativa L. J. Anal. Chem. 2015,70, 920–925. [CrossRef]
96.
Trofin, I.G.; Vlad, C.C.; Noja, V.V.; Dabija, G. Identification and characterization of special types of herbal Cannabis. UPB Sci. Bull.
2012,74, 13.
97.
Mölleken, H.; Husmann, H. Cannabinoids in seed extracts of Cannabis sativa cultivars. J. Int. Hemp Assoc.
1997
,4, 76–79.
Available online: http://www.druglibrary.org/olsen/hemp/IHA/jiha4201.html (accessed on 5 July 2020).
98.
Gambaro, V.; Dell’Acqua, L.; Farè, F.; Froldi, R.; Saligari, E.; Tassoni, G. Determination of primary active constituents in Cannabis
preparations by high-resolution gas chromatography/flame ionization detection and high-performance liquid chromatogra-
phy/UV detection. Anal. Chim. Acta 2002,468, 245–254. [CrossRef]
99.
Lachenmeier, D.W.; Kroener, L.; Musshoff, F.; Madea, B. Determination of cannabinoids in hemp food products by use of
headspace solid-phase microextraction and gas chromatography-mass spectrometry. Anal. Bioanal. Chem.
2004
,378, 183–189.
[CrossRef] [PubMed]
100.
Galettis, P.; Williams, M.; Gordon, R.; Martin, J.H. A Simple Isocratic HPLC Method for the Quantitation of 17 Cannabinoids.
Aust. J. Chem. 2021,74, 453. [CrossRef]
101.
de Oliveira, G.L.; Voloch, M.H.; Sztulman, G.B.; Neto, O.N.; Yonamine, M. Cannabinoid contents in cannabis products seized in
São Paulo, Brazil, 2006–2007. Forensic Toxicol. 2008,26, 31–35. [CrossRef]
102.
Potter, D.J.; Hammond, K.; Tuffnell, S.; Walker, C.; di Forti, M. Potency of
∆
9-tetrahydrocannabinol and other cannabinoids in
cannabis in England in 2016: Implications for public health and pharmacology. Drug Test. Anal.
2018
,10, 628–635. [CrossRef]
[PubMed]
103.
Hazekamp, A.; Fischedick, J.T. Cannabis—From cultivar to chemovar: Towards a better definition of Cannabis potency. Drug Test.
Anal. 2012,4, 660–667. [CrossRef] [PubMed]
104.
Potter, D.J.; Clark, P.; Brown, M.B. Potency of
∆
9–THC and Other Cannabinoids in Cannabis in England in 2005: Implications for
Psychoactivity and Pharmacology. J. Forensic Sci. 2008,53, 90–94. [CrossRef]

Molecules 2022,27, 975 38 of 42
105.
Knight, G.; Hansen, S.; Connor, M.; Poulsen, H.; McGovern, C.; Stacey, J. The results of an experimental indoor hydroponic
Cannabis growing study, using the ‘Screen of Green’ (ScrOG) method—Yield, tetrahydrocannabinol (THC) and DNA analysis.
Forensic Sci. Int. 2010,202, 36–44. [CrossRef]
106.
de Meijer, E.P.M.; Hammond, K.M.; Sutton, A. The inheritance of chemical phenotype in Cannabis sativa L. (IV): Cannabinoid-free
plants. Euphytica 2009,168, 95–112. [CrossRef]
107.
Florian-Ramirez, N.M.; Garzon-Mendez, W.F.; Parada-Alfonso, F. Gas Chromatography in Forensic Chemistry: Cannabinoids
Content in Marijuana Leaves (Cannabis sativa L.) from Colombia. In Gas Chromatography—Biochemicals Narcotics and Essential Oils;
Salih, B., Ed.; InTech: London, UK, 2012. [CrossRef]
108.
Poulsen, H.A.; Sutherland, G.J. The potency of cannabis in New Zealand from 1976 to 1996. Sci. Justice
2000
,40, 171–176.
[CrossRef]
109.
Tremlová, B.; Mikulášková, H.K.; Hajduchová, K.; Jancikova, S.; Kaczorová, D.; Zeljkovi´c, S. ´
C.; Dordevic, D. Influence of
Technological Maturity on the Secondary Metabolites of Hemp Concentrate (Cannabis sativa L.). Foods
2021
,10, 1418. [CrossRef]
110.
Fischedick, J.T.; Glas, R.; Hazekamp, A.; Verpoorte, R. A qualitative and quantitative HPTLC densitometry method for the
analysis of cannabinoids in Cannabis sativa L. Phytochem. Anal. 2009,20, 421–426. [CrossRef] [PubMed]
111.
Dubrow, G.A.; Pawar, R.S.; Srigley, C.; Sam, J.F.; Talavera, C.; Parker, C.H.; Noonan, G.O. A survey of cannabinoids and toxic
elements in hemp-derived products from the United States marketplace. J. Food Compos. Anal. 2021,97, 103800. [CrossRef]
112.
Glivar, T.; Eržen, J.; Kreft, S.; Zagožen, M.; ˇ
Cerenak, A.; ˇ
Ceh, B.; Benkovi´c, E.T. Cannabinoid content in industrial hemp (Cannabis
sativa L.) varieties grown in Slovenia. Ind. Crops Prod. 2020,145, 112082. [CrossRef]
113.
Saloner, A.; Bernstein, N. Nitrogen supply affects cannabinoid and terpenoid profile in medical cannabis (Cannabis sativa L.). Ind.
Crops Prod. 2021,167, 113516. [CrossRef]
114.
Zamengo, L.; Frison, G.; Bettin, C.; Sciarrone, R. Cannabis potency in the Venice area (Italy): Update 2013: Cannabis potency in
the Venice area (Italy): Update 2013. Drug Test. Anal. 2015,7, 255–258. [CrossRef] [PubMed]
115.
Dussy, F.E.; Hamberg, C.; Luginbühl, M.; Schwerzmann, T.; Briellmann, T.A. Isolation of
∆
9-THCA-A from hemp and analytical
aspects concerning the determination of ∆9-THC in cannabis products. Forensic Sci. Int. 2005,149, 3–10. [CrossRef] [PubMed]
116.
Harvey, D.J. Cyclic alkylboronates as derivatives for the characterization of cannabinolic acids by combined gas chromatography
and mass spectrometry. Biol. Mass Spectrom. 1977,4, 88–93. [CrossRef] [PubMed]
117.
Pandohee, J.; Holland, B.J.; Li, B.; Tsuzuki, T.; Stevenson, P.G.; Barnett, N.W.; Pearson, J.R.; Jones, O.A.H.; Conlan, X.A. Screening
of cannabinoids in industrial-grade hemp using two-dimensional liquid chromatography coupled with acidic potassium
permanganate chemiluminescence detection: Liquid Chromatography. J. Sep. Sci. 2015,38, 2024–2032. [CrossRef] [PubMed]
118.
Fetterman, P.S.; Doorenbos, N.J. A simple gas liquid chromatography procedure for determination of cannabinoldlc Acids In
Cannabis sativa L. Experimentia 1971,27, 988–990. [CrossRef] [PubMed]
119. Tucker, R.B.; Graham, B.F. Cannabinoid Content of Colombian Cannabis. Can. Soc. Forensic Sci. J. 1981,14, 41–45. [CrossRef]
120. Krishnamurty, H.G.; Kaushal, R. Analysis of Indian Marihuana. Indian J. Pharmacol. 1974,36, 152–154.
121.
Stefanidou, M.; Dona, A.; Athanaselis, S.; Papoutsis, I.; Koutselinis, A. The cannabinoid content of marihuana samples seized in
Greece and its forensic application. Forensic Sci. Int. 1998,95, 153–162. [CrossRef]
122.
Stefanidou, M.; Athanaselis, S.; Alevisopolous, G.; Papoutsis, I.; Koutselinis, A. Delta9-tetrahydrocannabinol content in cannabis
plants of Greek origin. Chem. Pharm. Bull. 2000,48, 743–745. [CrossRef]
123.
Tayyab, M.; Shahwar, D. GCMS analysis of Cannabis sativa L. from four different areas of Pakistan. Egypt. J. Forensic Sci.
2015
,5,
114–125. [CrossRef]
124.
Kladar, N.; ˇ
Coni´c, B.S.; Božin, B.; Torovi´c, L. European hemp-based food products—Health concerning cannabinoids exposure
assessment. Food Control. 2021,129, 108233. [CrossRef]
125.
Galand, N.; Ernouf, D.; Montigny, F.; Dollet, J.; Pothier, J. Separation and Identification of Cannabis Components by Different
Planar Chromatography Techniques (TLC, AMD, OPLC). J. Chromatogr. Sci. 2004,42, 130–134. [CrossRef]
126.
dos Santos, N.A.; Souza, L.M.; Domingos, E.; França, H.S.; Lacerda, V.; Beatriz, A.; Vaz, B.G.; Rodrigues, R.R.T.; Carvalho, V.V.;
Merlo, B.B.; et al. Evaluating the selectivity of colorimetric test (Fast Blue BB salt) for the cannabinoids identification in marijuana
street samples by UV–Vis, TLC, ESI(+)FT-ICR MS and ESI(+)MS/MS. Forensic Chem. 2016,1, 13–21. [CrossRef]
127.
Liu, Y.; Brettell, T.A.; Victoria, J.; Wood, M.R.; Staretz, M.E. High performance thin-layer chromatography (HPTLC) analysis of
cannabinoids in cannabis extracts. Forensic Chem. 2020,19, 1–11. [CrossRef]
128.
Caligiani, A.; Palla, G.; Bernardelli, B. GC-MS Analysis of Hashish Samples: A Case of Adulteration with Colophony. J. Forensic
Sci. 2006,51, 1096–1100. [CrossRef]
129.
Vollner, L.; Bieniek, D.; Korte, F. Review of analytical methods for identification and quantification of Cannabis products. Regul.
Toxicol. Pharmacol. 1986,6, 348–358. [CrossRef]
130.
Matsunaga, T.; Nagatomo, H.; Yamamoto, I.; Yoshimura, H. Identification and determination of cannabinoids in commercially
available cannabis seeds. Eisei Kagaku 1990,36, 545–547. [CrossRef]
131.
Tewari, S.N.; Harpalani, S.P.; Sharma, S.C. Separation and identification of the constituents of hashish (cannabis indica linn.) by
thin-layer chromatography and its application in forensic analysis. Mikrochim Acta 1974,62, 991–995. [CrossRef]
132.
Ibrahim, E.; Gul, W.; Gul, S.; Stamper, B.; Hadad, G.; Salam, R.A.; Ibrahim, A.; Ahmed, S.; Chandra, S.; Lata, H.; et al.
Determination of Acid and Neutral Cannabinoids in Extracts of Different Strains of Cannabis sativa Using GC-FID. Planta Med.
2018,84, 250–259. [CrossRef] [PubMed]

Molecules 2022,27, 975 39 of 42
133.
Schettino, L.; Prieto, M.; Benedé, J.L.; Chisvert, A.; Salvador, A. A Rapid and Sensitive Method for the Determination of
Cannabidiol in Cosmetic Products by Liquid Chromatography–Tandem Mass Spectrometry. Cosmetics 2021,8, 30. [CrossRef]
134.
Piccolella, S.; Formato, M.; Pecoraro, M.T.; Crescente, G.; Pacifico, S. Discrimination of CBD-, THC- and CBC-type acid cannabi-
noids through diagnostic ions by UHPLC-HR-MS/MS in negative ion mode. J. Pharm. Biomed. Anal.
2021
,201, 114125.
[CrossRef]
135. UNODC. Bulletin on Narcotics; Issue 2—008, United Nations, Office on Drugs and Crime: Vienna, Austria, 1994.
136.
Baker, P.B.; Taylor, B.J.; Gough, T.A. The tetrahydrocannabinol and tetrahydrocannabinolic acid content of cannabis products.
J. Pharm. Pharmacol. 1981,33, 369–372. [CrossRef] [PubMed]
137.
ElSohly, M.A.; Mehmedic, Z.; Foster, S.; Gon, C.; Chandra, S.; Church, J.C. Changes in Cannabis Potency Over the Last 2 Decades
(1995–2014): Analysis of Current Data in the United States. Biol. Psychiatry 2016,79, 613–619. [CrossRef] [PubMed]
138.
Stambouli, H.; el Bouri, A.; Bouayoun, T. Évolution de la teneur en
∆
9-THC dans les saisies de résines de cannabis au Maroc de
2005 à2014. Toxicol. Anal. Et Clin. 2016,28, 146–152. [CrossRef]
139.
Ross, S.A.; Mehmedic, Z.; Murphy, T.P.; Elsohly, M.A. GC-MS Analysis of the total delta9-THC content of bBoth drug- and
fiber-type cannabis seeds. J. Anal. Toxicol. 2000,24, 715–717. [CrossRef]
140.
Petrovi´c, M.; Debeljak, Ž.; Kezi´c, N.; Džidara, P. Relationship between cannabinoids content and composition of fatty acids in
hempseed oils. Food Chem. 2015,170, 218–225. [CrossRef]
141. Licata, M.; Verri, P.; Beduschi, G. δ9 THC content in illicit cannabis products. Ann. Ist Super Sanita 2005,41, 483–485.
142.
Swift, W.; Wong, A.; Li, K.M.; Arnold, J.C.; McGregor, I.S. Analysis of Cannabis Seizures in NSW, Australia: Cannabis Potency
and Cannabinoid Profile. PLoS ONE 2013,8, e70052. [CrossRef] [PubMed]
143.
Rovetto, L.J.; Aieta, N.V. Supercritical carbon dioxide extraction of cannabinoids from Cannabis sativa L. J. Supercrit. Fluids
2017
,
129, 16–27. [CrossRef]
144.
Christinat, N.; Savoy, M.-C.; Mottier, P. Development, validation and application of a LC-MS/MS method for quantification of
15 cannabinoids in food. Food Chem. 2020,318, 126469. [CrossRef]
145.
Ameur, S.; Haddou, B.; Derriche, Z.; Canselier, J.P.; Gourdon, C. Cloud point extraction of
∆
9-tetrahydrocannabinol from cannabis
resin. Anal. Bioanal. Chem. 2013,405, 3117–3123. [CrossRef] [PubMed]
146.
Gunjevi´c, V.; Grillo, G.; Carnaroglio, D.; Binello, A.; Barge, A.; Cravotto, G. Selective recovery of terpenes, polyphenols and
cannabinoids from Cannabis sativa L. inflorescences under microwaves. Ind. Crops Prod. 2021,162, 113247. [CrossRef]
147. Poddar, M.K.; Ghosh, J.J.; Datta, J. Micromethod for the estimation of Cannabis components. Sci. Cult. 1975,41, 492–495.
148.
Ilias, Y.; Rudaz, S.; Mathieu, P.; Veuthey, J.-L.; Christen, P. Analysis of Cannabis Material by Headspace Solid-Phase Microextraction
Combined with Gas Chromatography-Mass Spectrometry. Chim. Int. J. Chem. 2004,58, 219–221. [CrossRef]
149.
Qamar, S.; Manrique, Y.J.; Parekh, H.S.; Falconer, J.R. Development and Optimization of Supercritical Fluid Extraction Setup
Leading to Quantification of 11 Cannabinoids Derived from Medicinal Cannabis. Biology 2021,10, 481. [CrossRef]
150.
Omar, J.; Olivares, M.; Amigo, J.M.; Etxebarria, N. Resolution of co-eluting compounds of Cannabis Sativa in comprehensive
two-dimensional gas chromatography/mass spectrometry detection with Multivariate Curve Resolution-Alternating Least
Squares. Talanta 2014,121, 273–280. [CrossRef] [PubMed]
151.
Grijó, D.R.; Osorio, I.A.V.; Cardozo-Filho, L. Supercritical Extraction Strategies Using CO
2
and Ethanol to Obtain Cannabinoid
Compounds from Cannabis Hybrid Flowers. J. CO2 Util. 2019,30, 241–248. [CrossRef]
152.
Micalizzi, G.; Alibrando, F.; Vento, F.; Trovato, E.; Zoccali, M.; Guarnaccia, P.; Dugo, P.; Mondello, L. Development of a Novel
Microwave Distillation Technique for the Isolation of Cannabis sativa L. Essential Oil and Gas Chromatography Analyses for the
Comprehensive Characterization of Terpenes and Terpenoids, Including Their Enantio-Distribution. Molecules
2021
,26, 1588.
[CrossRef]
153.
Pellegrini, M.; Marchei, E.; Pacifici, R.; Pichini, S. A rapid and simple procedure for the determination of cannabinoids in hemp
food products by gas chromatography-mass spectrometry. J. Pharm. Biomed. Anal. 2005,36, 939–946. [CrossRef]
154.
Broséus, J.; Anglada, F.; Esseiva, P. The differentiation of fibre- and drug type Cannabis seedlings by gas chromatography/mass
spectrometry and chemometric tools. Forensic Sci. Int. 2010,200, 87–92. [CrossRef] [PubMed]
155.
Chandra, S.; Radwan, M.M.; Majumdar, C.G.; Church, J.C.; Freeman, T.P.; ElSohly, M.A. New trends in cannabis potency in USA
and Europe during the last decade (2008–2017). Eur. Arch. Psychiatry Clin. Neurosci. 2019,269, 5–15. [CrossRef] [PubMed]
156.
Qureshi, M.N.; Kanwal, F.; Siddique, M.; Akram, M. Estimation of Biologically Active Cannabinoids in Cannabis indica by Gas
Chromatography-mass Spectrometry (GC-MS). World Appl. Sci. J. 2012,19, 918–923. [CrossRef]
157.
Pijlman, F.; Rigter, S.; Hoek, J.; Goldschmidt, H.; Niesink, R. Strong increase in total delta-THC in cannabis preparations sold in
Dutch coffee shops. Addict. Biol. 2005,10, 171–180. [CrossRef]
158.
Vanhove, W.; van Damme, P.; Meert, N. Factors determining yield and quality of illicit indoor cannabis (Cannabis spp.) production.
Forensic Sci. Int. 2011,212, 158–163. [CrossRef]
159.
Thomsen, K.R.; Lindholst, C.; Thylstrup, B.; Kvamme, S.; Reitzel, L.A.; Worm-Leonhard, M.; Englund, A.; Freeman, T.P.; Hesse, M.
Changes in the composition of cannabis from 2000–2017 in Denmark: Analysis of confiscated samples of cannabis resin. Exp. Clin.
Psychopharmacol. 2019,27, 402–411. [CrossRef]
160.
dos Santos, N.; Tose, L.; da Silva, S.; Murgu, M.; Kuster, R.; Ortiz, R.; Camargo, F.; Vaz, B.; Lacerda, V., Jr.; Romão, W.
Analysis of Isomeric Cannabinoid Standards and Cannabis Products by UPLC-ESI-TWIM-MS: A Comparison with GC-MS and
GC ×GC-QMS. J. Braz. Chem. Soc. 2019,30, 60–70. [CrossRef]

Molecules 2022,27, 975 40 of 42
161.
Leghissa, A.; Smuts, J.; Qiu, C.; Hildenbrand, Z.L.; Schug, K.A. Detection of cannabinoids and cannabinoid metabolites using gas
chromatography with vacuum ultraviolet spectroscopy. Sep. Sci. Plus. 2018,1, 37–42. [CrossRef]
162.
Giese, M.W.; Lewis, M.A.; Giese, L.; Smith, K.M. Method for the Analysis of Cannabinoids and Terpenes in Cannabis. J. AOAC
Int. 2015,98, 1503–1522. [CrossRef]
163.
Trigg, S. Development of Gas and Liquid Chromatographic Methods for the Separation and Quantification of 11 Cannabinoids.
Bachelor Thesis, Murdoch University, Perth, Australia, 2017.
164.
Sherma, J.; Rabel, F. Thin layer chromatography in the analysis of cannabis and its components and synthetic cannabinoids. J. Liq.
Chromatogr. Relat. Technol. 2019,42, 613–628. [CrossRef]
165.
Mandrioli, M.; Tura, M.; Scotti, S.; Gallina, T. Fast Detection of 10 Cannabinoids by RP-HPLC-UV Method in Cannabis sativa L.
Molecules 2019,24, 2113. [CrossRef]
166.
Amirav, A.; Neumark, B.; Eren, K.J.M.; Fialkov, A.B.; Tal, N. Cannabis and its cannabinoids analysis by gas chromatography–mass
spectrometry with Cold EI. J. Mass Spectrom. 2021,6, 56. [CrossRef]
167. Kowalska, T.; Sajewicz, M.; Sherma, J. Planar Chromatography—Mass Spectrometry; CRC Press: Boca Raton, FL, USA, 2020.
168. Sherma, J.; Fried, B. (Eds.) Handbook of Thin-Layer Chromatography, 3rd ed.; rev.expanded; Marcel Dekker: New York, NY, USA, 2003.
169.
de Carvalho, T.C.; Tosato, F.; Souza, L.M.; Santos, H.; Merlo, B.B.; Ortiz, R.S.; Rodrigues, R.R.T.; Filgueiras, P.R.; França, H.S.;
Augusti, R.; et al. Thin layer chromatography coupled to paper spray ionization mass spectrometry for cocaine and its adulterants
analysis. Forensic Sci. Int. 2016,262, 56–65. [CrossRef]
170.
National Research Council (U.S.) (Ed.) Strengthening Forensic Science in the United States: A Path Forward; National Academies
Press: Washington, DC, USA, 2009.
171. Hanus, L. Our Experience with Identification and Separation of Compounds from Marijuana (Cannabis Sativa L.) by Thin Layer
Chromatography. Acta Univ. Palacki. Olomuc. Fac. Med. 1987,116, 15–24. [PubMed]
172.
Mano-Sousa, B.J.; Maia, G.A.S.; Lima, P.L.; Campos, V.A.; Negri, G.; Chequer, F.M.D.; Duarte-Almeida, J.M. Color determination
method and evaluation of methods for the detection of cannabinoids by thin-layer chromatography (TLC). J. Forensic Sci.
2021
,66,
854–865. [CrossRef]
173.
Meng, Q.; Buchanan, B.; Zuccolo, J.; Poulin, M.-M.; Gabriele, J.; Baranowski, D.C. A reliable and validated LC-MS/MS method for
the simultaneous quantification of 4 cannabinoids in 40 consumer products. PLoS ONE
2018
,13, e0196396. [CrossRef] [PubMed]
174.
Fan, X.-H.; Cheng, Y.-Y.; Ye, Z.-L.; Lin, R.-C.; Qian, Z.-Z. Multiple chromatographic fingerprinting and its application to the
quality control of herbal medicines. Anal. Chim. Acta 2006,555, 217–224. [CrossRef]
175.
Cheng, Y.; Chen, M.; Tong, W. An Approach to Comparative Analysis of Chromatographic Fingerprints for Assuring the Quality
of Botanical Drugs. J. Chem. Inf. Comput. Sci. 2003,43, 1068–1076. [CrossRef] [PubMed]
176.
Patel, B.; Wene, D.; Fan, Z. Qualitative and quantitative measurement of cannabinoids in cannabis using modified HPLC/DAD
method. J. Pharm. Biomed. Anal. 2017,146, 15–23. [CrossRef]
177.
Gul, W.; Gul, S.W.; Radwan, M.M.; Wanas, A.S.; Mehmedic, Z.; Khan, I.I.; Sharaf, M.H.M.; ElSohly, M.A. Determination of
11 Cannabinoids in Biomass and Extracts of Different Varieties of Cannabis Using High-Performance Liquid Chromatography.
J. AOAC Int. 2015,98, 1523–1528. [CrossRef] [PubMed]
178.
Pellati, F.; Brighenti, V.; Sperlea, J.; Marchetti, L.; Bertelli, D.; Benvenuti, S. New Methods for the Comprehensive Analysis of
Bioactive Compounds in Cannabis sativa L. (hemp). Molecules 2018,23, 2639. [CrossRef] [PubMed]
179.
Citti, C.; Pacchetti, B.; Vandelli, M.A.; Forni, F.; Cannazza, G. Analysis of cannabinoids in commercial hemp seed oil and
decarboxylation kinetics studies of cannabidiolic acid (CBDA). J. Pharm. Biomed. Anal. 2018,149, 532–540. [CrossRef] [PubMed]
180.
Pacifici, R.; Marchei, E.; Salvatore, F.; Guandalini, L.; Busardò, F.P.; Pichini, S. Evaluation of cannabinoids concentration and
stability in standardized preparations of cannabis tea and cannabis oil by ultra-high performance liquid chromatography tandem
mass spectrometry. Clin. Chem. Lab. Med. 2017,55, 1555–1563. [CrossRef] [PubMed]
181.
Yang, Y.; Lewis, M.M.; Bello, A.M.; Wasilewski, E.; Clarke, H.A.; Kotra, L.P. Cannabis sativa (Hemp) Seeds,
∆
9-
Tetrahydrocannabinol, and Potential Overdose. Cannabis Cannabinoid Res. 2017,2, 274–281. [CrossRef]
182.
Citti, C.; Ciccarella, G.; Braghiroli, D.; Parenti, C.; Vandelli, M.A.; Cannazza, G. Medicinal cannabis: Principal cannabinoids
concentration and their stability evaluated by a high performance liquid chromatography coupled to diode array and quadrupole
time of flight mass spectrometry method. J. Pharm. Biomed. Anal. 2016,128, 201–209. [CrossRef] [PubMed]
183.
Palermiti, A.; Cafaro, A.; Barco, S.; Bucchioni, P.; Franceschini, P.; Cusato, J.; de Nicolò, A.; Manca, A.; de Vivo, E.D.;
Russo, E.; et al.
Analysis of Cannabinoids Concentration in Cannabis Oil Galenic Preparations: Harmonization between Three Laboratories in
Northern Italy. Pharmaceuticals 2021,14, 462. [CrossRef] [PubMed]
184.
Carcieri, C.; Tomasello, C.; Simiele, M.; de Nicolò, A.; Avataneo, V.; Canzoneri, L.; Cusato, J.; di Perri, G.; D’Avolio, A.
Cannabinoids concentration variability in cannabis olive oil galenic preparations. J. Pharm. Pharmacol.
2018
,70, 143–149.
[CrossRef] [PubMed]
185.
Burnier, C.; Esseiva, P.; Roussel, C. Quantification of THC in Cannabis plants by fast-HPLC-DAD: A promising method for
routine analyses. Talanta 2019,192, 135–141. [CrossRef]
186.
Peace, M.R.; Butler, K.E.; Wolf, C.E.; Poklis, J.L.; Poklis, A. Evaluation of Two Commercially Available Cannabidiol Formulations
for Use in Electronic Cigarettes. Front. Pharmacol. 2016,7, 279. [CrossRef]

Molecules 2022,27, 975 41 of 42
187.
Yangsud, J.; Santasanasuwan, S.; Ahkkarachinoreh, P.; Maha, A.; Madaka, F.; Suksaeree, J.; Songsak, T.; Vutthipong, A.;
Monton, C.
Stability of cannabidiol,
∆
9-tetrahydrocannabinol, and cannabinol under stress conditions. Adv. Tradit. Med.
2021
,21, 475–484.
[CrossRef]
188.
Deidda, R.; Schelling, C.; Roussel, J.-M.; Dispas, A.; Bleye, C.D.; Ziemons, É.; Hubert, P.; Veuthey, J.-L. The analysis of cannabinoids
in cannabis samples by supercritical fluid chromatography and ultra-high-performance liquid chromatography: A comparison
study. Anal. Sci. Adv. 2021,2, 2–14. [CrossRef]
189.
Deidda, R.; Coppey, F.; Damergi, D.; Schelling, C.; Coïc, L.; Veuthey, J.-L.; Sacré, P.-Y.; de Bleye, C.; Hubert, P.; Esseiva, P.; et al.
New perspective for the in-field analysis of cannabis samples using handheld near-infrared spectroscopy: A case study focusing
on the determination of ∆9-tetrahydrocannabinol. J. Pharm. Biomed. Anal. 2021,202, 114150. [CrossRef]
190.
Ciolino, L.A.; Ranieri, T.L.; Taylor, A.M. Commercial cannabis consumer products part 2: HPLC-DAD quantitative analysis of
cannabis cannabinoids. Forensic Sci. Int. 2018,289, 438–447. [CrossRef]
191.
Ambach, L.; Penitschka, F.; Broillet, A.; König, S.; Weinmann, W.; Bernhard, W. Simultaneous quantification of delta-9-THC,
THC-acid A, CBN and CBD in seized drugs using HPLC-DAD. Forensic Sci. Int. 2014,243, 107–111. [CrossRef]
192.
Wakshlag, J.J.; Cital, S.; Eaton, S.J.; Prussin, R.; Hudalla, C. Cannabinoid, Terpene, and Heavy Metal Analysis of 29 Over-the-
Counter Commercial Veterinary Hemp Supplements. VMRR 2020,11, 45–55. [CrossRef]
193.
Tolomeo, F.; Russo, F.; Vandelli, M.A.; Biagini, G.; Capriotti, A.L.; Laganà, A.; Carbone, L.; Gigli, G.; Cannazza, G.; Citti, C.
HPLC-UV-HRMS analysis of cannabigerovarin and cannabigerobutol, the two impurities of cannabigerol extracted from hemp.
J. Pharm. Biomed. Anal. 2021,203, 114215. [CrossRef]
194.
Brighenti, V.; Licata, M.; Pedrazzi, T.; Maran, D.; Bertelli, D.; Pellati, F.; Benvenuti, S. Development of a new method for the analysis
of cannabinoids in honey by means of high-performance liquid chromatography coupled with electrospray ionisation-tandem
mass spectrometry detection. J. Chromatogr. A 2019,1597, 179–186. [CrossRef] [PubMed]
195.
Wang, M.; Wang, Y.-H.; Avula, B.; Radwan, M.M.; Wanas, A.S.; van Antwerp, J.; Parcher, J.F.; ElSohly, M.A.; Khan, I.A.
Decarboxylation Study of Acidic Cannabinoids: A Novel Approach Using Ultra-High-Performance Supercritical Fluid Chro-
matography/Photodiode Array-Mass Spectrometry. Cannabis Cannabinoid Res. 2016,1, 262–271. [CrossRef]
196.
mzCloud—Advanced Mass Spectral Database (n.d.). Available online: https://www.mzcloud.org/ (accessed on
28 October 2020
).
197.
Pence, H.E.; Williams, A. ChemSpider: An Online Chemical Information Resource. J. Chem. Educ.
2010
,87, 1123–1124. [CrossRef]
198.
Wishart, D.S.; Feunang, Y.D.; Marcu, A.; Guo, A.C.; Liang, K.; Vázquez-Fresno, R.; Sajed, T.; Johnson, D.; Li, C.; Karu, N.; et al.
HMDB 4.0: The human metabolome database for 2018. Nucleic Acids Res. 2018,46, D608–D617. [CrossRef]
199.
Bäckström, B.; Cole, M.D.; Carrott, M.J.; Jones, D.C.; Davidson, G.; Coleman, K. A preliminary study of the analysis of Cannabis
by supercritical fluid chromatography with atmospheric pressure chemical ionisation mass spectroscopic detection. Sci. Justice
1997,37, 91–97. [CrossRef]
200.
Wang, M.; Wang, Y.-H.; Avula, B.; Radwan, M.M.; Wanas, A.S.; Mehmedic, Z.; van Antwerp, J.; ElSohly, M.A.; Khan, I.A.
Quantitative Determination of Cannabinoids in Cannabis and Cannabis Products Using Ultra-High-Performance Supercritical
Fluid Chromatography and Diode Array/Mass Spectrometric Detection. J. Forensic Sci.
2017
,62, 602–611. [CrossRef] [PubMed]
201.
U.S. Department of Health and Human Services, Food and Drug Administration. Guidance for Industry PAT—A Framework for
Innovative Pharmaceutical Development, Manufacturing, and Quality Assurance; Food and Drug Administration: Silver Spring, MD,
USA, 2004.
202. Griffiths, P.R.; de Haseth, J.A. Fourier Transform Infrared Spectrometry, 2nd ed.; Wiley: Hoboken, NJ, USA, 2007.
203.
Borille, B.T.; Marcelo, M.C.A.; Ortiz, R.S.; Mariotti, K.d.; Ferrão, M.F.; Limberger, R.P. Near infrared spectroscopy combined with
chemometrics for growth stage classification of cannabis cultivated in a greenhouse from seized seeds. Spectrochim. Acta Part A:
Mol. Biomol. Spectrosc. 2017,173, 318–323. [CrossRef]
204.
Mendez, J.; Francisco, S. Optimization of Cannabis Grows Using Introduction Fourier Transform Mid-Infrared Spectroscopy; Perkin
Elmer Inc.: Waltham, MA, USA, 2016.
205.
Callado, C.S.; Núñez-Sánchez, N.; Casano, S.; Ferreiro-Vera, C. The potential of near infrared spectroscopy to estimate the content
of cannabinoids in Cannabis sativa L.: A comparative study. Talanta 2018,190, 147–157. [CrossRef]
206.
Grekopoulos, J.E. Construction and Validation of Quantification Methods for Determining the Cannabidiol Content in Liquid
Pharma-Grade Formulations by Means of Near-Infrared Spectroscopy and Partial Least Squares Regression. Med. Cannabis
Cannabinoids 2019,2, 43–55. [CrossRef]
207.
Duchateau, C.; Kauffmann, J.; Canfyn, M.; Stévigny, C.; de Braekeleer, K.; Deconinck, E. Discrimination of legal and illegal
Cannabis spp. according to European legislation using near infrared spectroscopy and chemometrics. Drug Test. Anal.
2020
,12,
1309–1319. [CrossRef]
208. Lu, G.; Fei, B. Medical hyperspectral imaging: A review. J. Biomed. Opt. 2014,19, 010901. [CrossRef] [PubMed]
209.
Politi, M.; Peschel, W.; Wilson, N.; Zloh, M.; Prieto, J.M.; Heinrich, M. Direct NMR analysis of cannabis water extracts and
tinctures and semi-quantitative data on ∆9-THC and ∆9-THC-acid. Phytochemistry 2008,69, 562–570. [CrossRef] [PubMed]
210. Martin, F.L. Shining a new light into molecular workings. Nat. Methods 2011,8, 385–387. [CrossRef]
211.
Sanchez, L.; Baltensperger, D.; Kurouski, D. Raman-Based Differentiation of Hemp, Cannabidiol-Rich Hemp, and Cannabis. Anal.
Chem. 2020,92, 7733–7737. [CrossRef] [PubMed]
212.
Sanchez, L.; Filter, C.; Baltensperger, D.; Kurouski, D. Confirmatory non-invasive and non-destructive differentiation between
hemp and cannabis using a hand-held Raman spectrometer. RSC Adv. 2020,10, 3212–3216. [CrossRef]

Molecules 2022,27, 975 42 of 42
213.
Ebersbach, P.; Stehle, F.; Kayser, O.; Freier, E. Chemical fingerprinting of single glandular trichomes of Cannabis sativa by
Coherent anti-Stokes Raman scattering (CARS) microscopy. BMC Plant Biol. 2018,18, 275. [CrossRef]
214.
Lurie, I.S.; Meyers, R.P.; Conver, T.S. Capillary Electrochromatography of Cannabinoids. Anal. Chem.
1998
,70, 3255–3260.
[CrossRef]
215.
Peschel, W.; Politi, M. 1 H NMR and HPLC/DAD for Cannabis sativa L. chemotype distinction, extract profiling and specification.
Talanta 2015,140, 150–165. [CrossRef]
216.
Isahq, M.S.; Afridi, M.S.; Ali, J.; Hussain, M.M.; Ahmad, S.; Kanwal, F. Proximate composition, phytochemical screening, GC-MS
studies of biologically active cannabinoids and antimicrobial activities of Cannabis indica. Asian Pac. J. Trop. Dis.
2015
,5, 897–902.
[CrossRef]
217.
Uziel, A.; Gelfand, A.; Amsalem, K.; Berman, P.; Lewitus, G.M.; Meiri, D.; Lewitus, D.Y. Full-Spectrum Cannabis Extract
Microdepots Support Controlled Release of Multiple Phytocannabinoids for Extended Therapeutic Effect. ACS Appl. Mater.
Interfaces 2020,12, 23707–23716. [CrossRef]
218.
Risoluti, R.; Gullifa, G.; Battistini, A.; Materazzi, S. The Detection of Cannabinoids in Veterinary Feeds by MicroNIR/Chemometrics:
A New Analytical Platform. Analyst 2020,145, 1777–1782. [CrossRef] [PubMed]
219.
Hazekamp, A.; Choi, Y.H.; Verpoorte, R. Quantitative Analysis of Cannabinoids from Cannabis sativa Using 1H-NMR. Chem.
Pharm. Bull. 2004,52, 718–721. [CrossRef] [PubMed]
220.
Happyana, N.; Agnolet, S.; Muntendam, R.; van Dam, A.; Schneider, B.; Kayser, O. Analysis of cannabinoids in laser-
microdissected trichomes of medicinal Cannabis sativa using LCMS and cryogenic NMR. Phytochemistry
2013
,87, 51–59.
[CrossRef]
221.
Choi, Y.H.; Kim, H.K.; Hazekamp, A.; Erkelens, C.; Lefeber, A.W.M.; Verpoorte, R. Metabolomic Differentiation of Cannabis sativa
Cultivars Using 1H NMR Spectroscopy and Principal Component Analysis. J. Nat. Prod.
2004
,67, 953–957. [CrossRef] [PubMed]
222.
van der Pol, P.; Liebregts, N.; de Graaf, R.; Korf, D.J.; van den Brink, W.; van Laar, M. Validation of Self-Reported Cannabis Dose
and Potency: An Ecological Study: Self-Reported Cannabis Dose and Potency. Addiction 2013,108, 1801–1808. [CrossRef]
223.
van der Pol, P.; Liebregts, N.; Brunt, T.; van Amsterdam, J.; de Graaf, R.; Korf, D.J.; van den Brink, W.; van Laar, M. Cross-Sectional
and Prospective Relation of Cannabis Potency, Dosing and Smoking Behaviour with Cannabis Dependence: An Ecological Study:
Cannabis Potency, Titration and Dependence. Addiction 2014,109, 1101–1109. [CrossRef]
224.
Leghissa, A. Method Development for Qualification and Quantification of Cannabinoids and Terpenes in Extracts by Gas
Chromatography-Mass Spectrometry. 91. Available online: https://rc.library.uta.edu/uta-ir/handle/10106/26115?show=full
(accessed on 8 January 2022).