Biosynthesis of cannflavins A and B from Cannabis sativa L

Article abstract of DOI:10.1016/j.phytochem.2019.05.009

In addition to the psychoactive constituents that are typically associated with Cannabis sativa L., there exist numerous other specialized metabolites in this plant that are believed to contribute to its medicinal versatility. This study focused on two such compounds, known as cannflavin A and cannflavin B. These prenylated flavonoids specifically accumulate in C. sativa and are known to exhibit potent anti-inflammatory activity in various animal cell models. However, almost nothing is known about their biosynthesis. Using a combination of phylogenomic and biochemical approaches, an aromatic prenyltransferase from C. sativa (CsPT3) was identified that catalyzes the regiospecific addition of either geranyl diphosphate (GPP) or dimethylallyl diphosphate (DMAPP) to the methylated flavone, chrysoeriol, to produce cannflavins A and B, respectively. Further evidence is presented for an O-methyltransferase (CsOMT21) encoded within the C. sativa genome that specifically converts the widespread plant flavone known as luteolin to chrysoeriol, both of which accumulate in C. sativa. These results therefore imply the following reaction sequence for cannflavins A and B biosynthesis: luteolin ? chrysoeriol ? cannflavin A and cannflavin B. Taken together, the identification of these two unique enzymes represent a branch point from the general flavonoid pathway in C. sativa and offer a tractable route towards metabolic engineering strategies that are designed to produce these two medicinally relevant Cannabis compounds.

Full text of DOI:10.1016/j.phytochem.2019.05.009

Phytochemistry164(2019)162–171
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Phytochemistry
journal homepage: www.elsevier.com/locate/phytochem
Biosynthesis of cannflavins A and B from Cannabis sativa L
Kevin A Reaa, José A. Casarettoa, M. Sameer Al-Abdul-Wahidb, Arjun Sukumarana,
Jennifer Geddes-McAlistera, Steven J. Rothsteina, Tariq A. Akhtara,∗
a Department of Molecular and Cellular Biology, University of Guelph, Guelph, Ontario, N1G 2W1, Canada
b NMR Center, University of Guelph, Guelph, Ontario, N1G 2W1, Canada
A R T I C L E I N F O
A B S T R A C T
Keywords:
Cannabis sativa L. (Cannabaceae)
Cannflavin A
In addition to the psychoactive constituents that are typically associated with Cannabis sativa L., there exist
numerous other specialized metabolites in this plant that are believed to contribute to its medicinal versatility.
This study focused on two such compounds, known as cannflavin A and cannflavin B. These prenylated flavo-
noids specifically accumulate in C. sativa and are known to exhibit potent anti-inflammatory activity in various
animal cell models. However, almost nothing is known about their biosynthesis. Using a combination of phy-
logenomic and biochemical approaches, an aromatic prenyltransferase from C. sativa (CsPT3) was identified that
catalyzes the regiospecific addition of either geranyl diphosphate (GPP) or dimethylallyl diphosphate (DMAPP)
to the methylated flavone, chrysoeriol, to produce cannflavins A and B, respectively. Further evidence is pre-
sented for an O-methyltransferase (CsOMT21) encoded within the C. sativa genome that specifically converts the
widespread plant flavone known as luteolin to chrysoeriol, both of which accumulate in C. sativa. These results
therefore imply the following reaction sequence for cannflavins A and B biosynthesis: luteolin ► chrysoeriol ►
cannflavin A and cannflavin B. Taken together, the identification of these two unique enzymes represent a
branch point from the general flavonoid pathway in C. sativa and offer a tractable route towards metabolic
engineering strategies that are designed to produce these two medicinally relevant Cannabis compounds.
Cannflavin B
Prenyltransferase
O-methyltransferase
Flavonoid
Anti-inflammatory
1. Introduction
Pickens (1981) demonstrated that THC and CBD-free extracts from C.
sativa could reduce the cataleptic effects of THC in mice and that this
Since antiquity, Cannabis sativa L. (Cannabaceae) has been culti-
vated for a variety of industrial, medicinal and recreational uses. In
recent years, a worldwide socio-political movement aimed towards the
legalization of C. sativa has spurred a resurgence of interest in this
versatile plant and have afforded researchers the opportunity to explore
its metabolic diversity (McPartland and Russo, 2001; van Bakel et al.,
2011; Sawler et al., 2015; Booth et al., 2017; Russo and Marcu, 2017;
Sexton et al., 2018). Apart from the psychoactive Δ9-tetra-
hydrocannabinol (THC) and the pharmacologically related cannabi-
noids (ex. cannabidiol, CBD) that typically accumulate in a variety of C.
sativa cultivars, there exist a plethora of specialized metabolites in this
plant species that are believed to contribute to its medicinal properties
(Elsohly and Slade, 2005; Radwan et al., 2008a,b; Flores-Sanchez and
Verpoorte, 2008; McPartland and Russo, 2014; Booth et al., 2017). One
such class of compounds are the prenylated flavonoids, known as
cannflavins A and B (Barrett et al., 1985).
effect could be reversed by prostaglandin E2 (PGE2) administration.
Barrett and colleagues purportedly identified the causal agent in these
extracts as cannflavins A and B and verified that these prenylated fla-
vonoids could inhibit the production of PGE2 in human rheumatoid
synovial cells and provide anti-inflammatory benefits that were ap-
proximately thirty times more effective than aspirin (Barrett et al.,
1985, 1986). It was later demonstrated that the underlying basis for
their potent anti-inflammatory properties was that cannflavins A and B
act to inhibit the in vivo production of two pro-inflammatory mediators,
prostaglandin E2 and the leukotrienes (Werz et al., 2014). Surprisingly,
however, since these striking reports of two non-psychoactive con-
stituents from C. sativa that have medicinal potential, little attention
has been focused on how these unique flavonoids are actually synthe-
sized within C. sativa (Andre et al., 2016; Pollastro et al., 2018).
Cannflavins A and B belong to the class of plant flavonoids known as
flavones, which occur in several plant lineages (Winkel-Shirley, 2001;
Ross and Kasum, 2002; Andersen and Markham, 2005; Jiang et al.,
2016). Flavones perform a myriad of in planta functions that range from
The interest surrounding cannflavins A and B within the Cannabis
community stems from three seminal studies: First, Fairbairn and
∗ Corresponding author.
E-mail address: takhtar@uoguelph.ca (T.A. Akhtar).
https://doi.org/10.1016/j.phytochem.2019.05.009
Received 24 July 2018; Received in revised form 19 April 2019; Accepted 10 May 2019
0031-9422/©2019TheAuthors.PublishedbyElsevierLtd.ThisisanopenaccessarticleundertheCCBYlicense
(http://creativecommons.org/licenses/BY/4.0/).

K.A. Rea, et al.
Phytochemistry164(2019)162–171
regulators of auxin transport to mediators in plant-pathogen interac-
tions (Johnson et al., 2007; Zhang et al., 2009; Falcone Ferreyra et al.,
2012). In addition, the dietary consumption of various plant flavones is
well established to offer neuroprotective, antioxidant, and anticancer
properties in several animal models (Pietta, 2000; Nabavi et al., 2015;
Madunić et al., 2018). While the flavone biosynthetic pathway has been
extensively studied in several plants, almost nothing is known about
this process in cannabis (Flores-Sanchez and Verpoorte, 2008; Andre
et al., 2016). It is expected that the core flavone pathway is similarly
embedded within C. sativa, given that this plant also accumulates three
widespread flavones (apigenin, luteolin and chrysoeriol) as well as their
glycosylated derivatives (Turner et al., 1980; Radwan et al., 2008a,b).
Cannflavins A and B appear to be Cannabis specific (Vanhoenacker
et al., 2002; Ross et al., 2005; Werz et al., 2014) and their unique
bioactivity appear to be linked to two key modifications of their parent
flavone backbone: First is their distinct prenylation pattern in which a
prenyl side-chain, in the form of a geranyl (C10) or a dimethylallyl (C5)
group, are affixed to the 6 position of the flavone A-ring, respectively
(Barrett et al., 1985; Choi et al., 2004). These prenyl moieties impart
lipophilicity to the parent flavone, which are believed to enhance up-
take and bioaccumulation into cells and promote their interaction with
membrane-bound enzymes and receptors that are involved in numerous
cell-signalling pathways (Milligan et al., 1999; Botta et al., 2005;
Wätjen et al., 2007; Werz et al., 2014; Vochyánová et al., 2017).
Second, both cannflavins A and B are modified at the 3′ position of the
flavone B-ring with a methoxy group, which also increases lipophilicity
and may therefore enhance their cellular retention and access to var-
ious cellular targets (Ibrahim, 2005; Walle, 2007; Berim and Gang,
2016). In which order these two unique modifications (prenylation and
methoxylation) of the parent flavone occur en route to cannflavin A and
cannflavin B biosynthesis, however, is not known.
defined by the specific branch of aromatic metabolism in which they
participate. The eight CsPTs occupy three of these six groups: CsPT2
and CsPT6 reside in a unique clade of prenyltransferases (Group 2)
which have been shown to participate in the tocopherol biosynthetic
pathway (Collakova and DellaPenna, 2001; Savidge et al., 2002; Tian
et al., 2007). CsPT5 appears to be orthologous to homogentisate sola-
nesyltransferases (Group V) that function in plastoquinone biosynthesis
(Venkatesh et al., 2006; Tian et al., 2007). The five remaining CsPTs
(CsPT1, 3, 4, 7, and 8) formed a third and distantly related group
(Group VI) that includes two prenyltransferases from Humulus lupulus
(hops), which are involved in the aromatic prenylation reactions re-
quired for terpenophenolic biosynthesis (Nagel et al., 2008; Tsurumaru
et al., 2012; Li et al., 2015). Surprisingly, this analysis revealed that
none of the CsPTs were closely related to any of the flavonoid or cou-
marin prenyltransferases (Groups I and IV, respectively) that have been
previously identified in various plant species (Sasaki et al., 2008, 2011;
Akashi et al., 2009; Shen et al., 2012; Wang et al., 2014; Munakata
et al., 2016; Yoneyama et al., 2016; Yang et al., 2018). Interestingly, in
silico analysis of each CsPT predicted that they are all targeted to
plastids (Table S2). We return to this point below.
2.2. Biochemical characterization of recombinant CsPTs identifies a
regiospecific chrysoeriol-6-prenyltransferase
In pursuit of identifying a flavone prenyltransferase(s) that is in-
volved in cannflavin A and B biosynthesis, we focused on the sixth
group of plant PTs that was identified from our phylogenetic analysis,
which included CsPT1, 3, 4, 7, and 8. These particular CsPTs drew our
attention for two reasons: First, they represent the only pre-
nyltransferases in our search of the C. sativa genome that are evolu-
tionarily distinct from those CsPTs that appear to function in tocopherol
and plastoquinone biosynthesis, which are central plant compounds
(Sattler et al., 2004; Kriese et al., 2004). Second, this group of CsPTs are
closely related to two enzymes from hops (a Cannabaceae family re-
lative), which are believed to prenylate naringenin chalcone, a wide-
spread intermediate in the general flavonoid pathway (Nagel et al.,
2008; Li et al., 2015).
We therefore introduced each of these CsPTs into a well-established
yeast expression system that is typically used to characterize this class
of enzymes (Sasaki et al., 2011; Shen et al., 2012; Li et al., 2014; Wang
et al., 2014). Principle component analysis of whole cell proteomes
from the strains expressing each of the five CsPTs revealed a clear
clustering that was separate from the strain harboring the vector alone
negative control (Figure S2). Moreover, a Pearson correlation analysis
followed by Hierarchical clustering by Euclidean distance confirmed
distinct clustering between the strains expressing the CsPTs compared
to the strain harboring the empty vector negative control, which to-
gether indirectly indicate that each CsPT is active and expressed in this
system. As a positive control for prenyltransferase enzyme activity, we
also introduced the open-reading frame of GuA6DT from G. uralensis
into this host yeast strain, which was previously shown to prenylate not
only apigenin, but a variety of flavones at position 6 of the A ring using
dimethylallyl diphosphate (DMAPP) as a substrate (Li et al., 2014). As
expected, in assays with microsomes expressing GuA6DT together with
apigenin and DMAPP as a prenyl donor, we observed a single reaction
product whose mass-to-charge ratio (m/z 339) was consistent with 6-
dimethylallyl apigenin (Fig. S2). As previously demonstrated, luteolin
and chrysoeriol were also converted by GuA6DT to their corresponding
6-dimethylallyl flavones using DMAPP as a substrate (Li et al., 2014),
and these monoprenylated flavones were subsequently purified by
HPLC to serve as prenylflavone standards in subsequent exploratory
assays with recombinant CsPTs (Fig. S3). Accordingly, the microsomal
fractions from each yeast strain expressing the open-reading frames of
CsPT1, 3, 4, 7, and 8 were recovered and tested for prenyltransferase
activity with three flavone substrates that are present in C. sativa: api-
genin, chrysoeriol, and luteolin (McPartland and Russo, 2001;
We therefore reasoned that the biosynthesis of cannflavins A and B
occurs via a specific branch point from the conserved C. sativa flavone
biosynthetic pathway in which a central flavone must be methoxylated
at the 3′ position of the flavone B-ring and prenylated at the 6 position
of the A-ring. Using a combination of phylogenomic and biochemical
approaches, we report the identification and characterization of two
enzymes that catalyze these penultimate and final steps of cannflavin A
and cannflavin B synthesis in C. sativa.
2. Results and discussion
2.1. Phylogenetic analysis of C. sativa prenyltransferases
To synthesize cannflavins A and B, a prenyl moiety must be added to
position 6 of a flavone that typically accumulates in C. sativa. Therefore,
we first searched for gene sequences that were putatively annotated as
flavonoid or related aromatic prenyltransferases in the Transcriptome
Shotgun Assembly (TSA) database for C. sativa, which is accessible
through NCBI. A previously described flavone prenyltransferase from
Glycyrrhiza uralensis (GuA6DT; GenBank AIT11912.1) was used as a
query in these searches (Li et al., 2014). GuA6DT prenylates apigenin
which is a widespread plant flavone that also accumulates in C. sativa
(McPartland and Russo, 2001). This search uncovered eight full-length
cDNA sequences from C. sativa that exhibited 22–53% identity at the
amino acid level to GuA6DT and were putatively annotated as C. sativa
prenyltransferases (CsPT1-8; Fig. S1). One of the prenyltransferases that
were identified in this search (CsPT1) matched a previously reported
enzyme from C. sativa that is known to be involved in the prenylation of
olivetolic acid to cannabigerolic acid in the cannabinoid biosynthesis
pathway (Page and Boubakir, 2014). We next performed a phylogenetic
analysis that included CsPT1 and these seven other prenyltransferases
from C. sativa along with all known plant prenyltransferases that have
been previously shown to accommodate aromatic substrates (Fig. 1;
Fig. S1). This analysis demonstrated that plant aromatic pre-
nyltransferases fall into six distinct groups, which are conveniently
163

K.A. Rea, et al.
Phytochemistry164(2019)162–171
Fig. 1. Phylogenetic analysis of prenyltransferases from C. sativa. The eight putative CsPTs identified in the C. sativa genome were aligned with the amino acid
sequences of various plant aromatic prenyltransferases that have been previously identified using Clustal W and a Neighbor-joining phylogenetic tree (1000 re-
plicates) to illustrate their evolutionary relationships was then constructed using the MEGA 6.0 software package. Bootstrap values (> 60%) are indicated at the
nodes of each branch and the branch lengths are proportional to the number of amino acid substitutions per sequence. The sequences and species abbreviations for all
of these prenyltransferases are listed in Supplemental Fig. S1 along with their corresponding GenBank accession numbers. Note that the eight CsPTs (bolded)
distinctly fall into three groups, which are involved in either tocopherol (II), plastoquinone (V), or terpenophenolic (VI) biosynthesis.
a prenyl donor, and kinetic analysis demonstrated that chrysoeriol was
the preferred flavone substrate (Table 1). Assays with microsomes
containing CsPT8 revealed that this enzyme also prenylated apigenin
using DMAPP as a substrate, but not with any of the other flavones (Fig.
S4). IPP was not accommodated as a prenyl donor in assays with mi-
crosomes containing either CsPT3 or CsPT8. The prenylated product
from assays with CsPT8 did not match that of 6-dimethylallyl apigenin,
based on HPLC retention time (Fig. S4). On the other hand, the enzy-
matic product that was observed in assays with CsPT3, chrysoeriol and
DMAPP exhibited the same HPLC retention time and fragmentation
pattern, as determined by tandem mass spectrometry, to that of 6-di-
methylallyl chrysoeriol, also known as cannflavin B (Fig. 2A). We
therefore chose to focus solely on CsPT3. Interestingly, CsPT3 also ac-
commodated GPP as a prenyl donor in assays with apigenin and chry-
soeriol, and kinetic analysis again revealed that chrysoeriol was the
preferred substrate (Table 1). That CsPT3 accommodated both DMAPP
and GPP fits well with its predicted subcellular location, as the enzyme
would have access to these two end products of the plastidial methy-
lerythritol phosphate pathway. The reaction product that was observed
with assays including CsPT3, chrysoeriol and GPP exhibited less po-
larity than 6-dimethylallyl chrysoeriol/cannflavin B (as indicated by
HPLC-UV analysis) and exhibited a mass-to-charge ratio (m/z 437) that
was consistent with the addition of a geranyl moiety onto the chry-
soeriol backbone, and was therefore assumed to be cannflavin A
(Fig. 2B).
Table 1
Kinetic parameters of CsPT3 with various C. sativa flavone substrates.
Substrate
DMAPP
GPP
Km (μM)
Vmax (pmol
Km (μM)
Vmax (pmol
min−1 mg−1
)
min−1 mg−1
)
Apigenin
Chrysoeriol 37.9
Luteolin N.D.
141.7
15.6 1.2
0.0566
0.0585
49.4
35.6
N.D.
11.4 1.3
10.7 1.3
0.0993
0.122
5.59
1.3
N.D.
N.D.
Measurements were made at 37 °C in 100 mM Tris-HCl, pH 9.0 and 10 mM
MgCl2. Reactions were initiated by adding either DMAPP or GPP as the prenyl
donor substrate to each reaction. Data are the means of three independent
determinations
SE. Kinetic constants were not determined (N.D.) for assays
with luteolin as a flavone substrate due to low conversion rates.
Brenneisen, 2007; Radwan et al., 2008a,b). As potential prenyl donors,
DMAPP, IPP and GPP were also included as co-substrates in each of
these enzyme assays. The reaction products from these assays were
extracted with ethyl acetate, analyzed by reverse-phase HPLC, and
compared to the authentic prenylated flavones that were isolated from
in vitro assays with GuA6DT (see above). No detectable prenylated
flavone products were observed in assays with microsomes obtained
from yeast cells harbouring the empty vector or with CsPT1, 4, or 7
under the above conditions. However, this analysis revealed that mi-
crosomes containing CsPT3 readily converted apigenin and chrysoeriol,
but not luteolin, to their corresponding prenylflavones using DMAPP as
The structures of the two enzymatic products that were obtained
164

K.A. Rea, et al.
Phytochemistry164(2019)162–171
Fig. 3. Key HMBC correlations for cannflavin A (above) and cannflavin B
(below). Bold arrows indicate correlations that were observed from both the 5-
OH hydroxyl proton and the 1ꞌꞌ proton on the geranyl or prenyl group to the
same carbon resonance.
Table 2
NMR assignments for cannflavin A and cannflavin B.
Position
Cannflavin A
Cannflavin B
∂
H
∂
C
∂
H
∂
C
2
3
4
5
6
7
8
9
10
1'
2'
3'
4'
–
6.68
–
–
164.4
104.2
182.8
159.8
112.0
162.1
93.9
156.3
104.9
123.4
110.0
148.5
151.0
115.8
121.0
21.8
–
6.69
–
–
–
–
6.62
–
–
164.7
104.5
183.2
160.2
112.3
162.3
94.1
156.6
105.3
123.8
110.5
148.8
151.3
116.3
121.3
22.0
123.2
131.7
17.9
25.9
–
–
6.62
–
–
–
7.60
–
–
7.00
7.57
3.36
5.29
–
1.96
2.05
5.07
–
1.59
1.54
1.79
3.98
13.30
Fig. 2. Evidence for cannflavin
A and B biosynthesis by CsPT3. (A)
Representative HPLC chromatograms are shown for authentic 6-dimethylallyl
chrysoeriol/cannflavin B (top) and for the separation of the extracted enzymatic
reaction products from assays with recombinant CsPT3, chrysoeriol, and
DMAPP (bottom). Note that in the enzyme assays, chrysoeriol is converted to a
product that shares the same HPLC retention time and CID-Q-TOF mass spectra
(inset) with 6-dimethylallyl chrysoeriol/cannflavin B. (B) A representative
HPLC chromatogram illustrating the enzymatic reaction products that were
extracted from assays conducted with CsPT3, chrysoeriol, and GPP. Note the
conversion of chrysoeriol to a product with greater hydrophobicity than 6-di-
methylallyl chrysoeriol/cannflavinB and which exhibits a Q-TOF mass spectra
(inset) consistent with geranylated chrysoeriol ([M+H]+ 437).
–
7.61
–
–
5'
6'
7.00
7.59
3.36
5.28
–
1.78
1.65
–
–
–
–
–
1''
2''
3''
4''
5''
6''
7''
8''
9''
10''
O-CH3
5-OH
122.9
135.0
40.1
27.6
124.8
131.3
25.3
17.3
16.0
–
–
–
–
56.6
–
from assays with CsPT3, chrysoeriol, and either DMAPP or GPP as
prenyl donors, were further analyzed by 1H and 13C NMR. The 1H NMR
spectra of cannflavin A exhibited three peaks of area 3H in the region
between 1.5 and 2.0 ppm and 2 peaks of area 1H in the region between
5.0 and 5.5 ppm, consistent with three methyl groups and two vinylic
protons, respectively, suggesting the presence of a geranyl group. The
1H NMR spectra of cannflavin B exhibited only two peaks of area 3H in
the region between 1.5 and 2.0 ppm and a single peak of area 1H in the
region between 5.0 and 5.5 ppm, consistent with the two methyl groups
and one vinylic proton of a single prenyl group. COSY, TOCSY, and
HMBC spectra also demonstrated peak patterns that were consistent
with a geranyl group for cannflavin A and a prenyl group for cannflavin
B (Crombie and Crombie, 1982; Barrett et al., 1986; Choi et al., 2004).
Initial NMR assignments for both cannflavin A and B were performed
using the COSY, TOCSY, and HSQC spectra to assign all proton re-
sonances as well as those of proton-bearing carbons and an HMBC ex-
periment was then used to complete the carbon assignments (Fig. 3). In
both cannflavin A and cannflavin B, HMBC correlations were observed
56.2
–
3.99
13.30
from both the 5-OH hydroxyl proton and the 1ꞌꞌ proton on the geranyl
or prenyl group to the same carbon resonance (shown as bold arrows in
Fig. 3). The 5-OH hydroxy proton is too far away from the carbon at
position 8 for an HMBC correlation to be observed and we therefore
reasoned that the geranylation/prenylation must be at the 6 position.
The complete 1H and 13C-NMR assignments are shown in Table 2.
Taken together, these results suggest that CsPT3 appears to be the sole
member of the CsPT family that prenylates chrysoeriol, using either
GPP or DMAPP, in the final steps of cannflavin A and cannflavin B
biosynthesis, respectively.
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Phytochemistry164(2019)162–171
2.3. Phylogenetic analysis of C. sativa O-methyltransferases involved in the
methylation of luteolin to chryseoriol
into E. coli cells as N-terminal fusion proteins with a His6 tag and
subsequently assayed desalted protein extracts from cells expressing
each protein for O-methyltransferase enzyme activity. We chose to test
three flavones (apigenin, luteolin, and chrysoeriol) and two flavonols
(quercetin and kaempferol) that typically accumulate in C. sativa as
potential substrates for each CsOMT, along with the universal methyl
donor 14C-labeled S-adenosyl methionine as a co-substrate (Fig. 5A).
After extracting and quantifying the amount of radiolabel within the
enzymatic products that were obtained from these initial assays, it was
determined that only CsOMT6 and CsOMT21 exhibited appreciable
OMT activity with the flavonoid substrates that were provided. Re-
combinant CsOMT6 exhibited strict substrate specificity towards
quercetin (Fig. S6), while CsOMT21 methylated luteolin primarily
(Fig. 5B), yet also accommodated quercetin as a substrate, albeit with
less efficiency (57% activity compared to luteolin). Notably, neither
apigenin, nor chrysoeriol, nor kaempferol, which lack a free 3′-hydroxyl
group on their B-ring, were used as substrates by either enzyme. This
observation therefore lends further support to the view (Lam et al.,
2007) that CsOMT6, CsOMT21 and the other type 1 OMTs that are
present in group three (defined by our phylogenetic analysis) en-
compass an evolutionarily conserved group of enzymes with re-
giospecificity for 3′-hydroxyl groups on a variety of flavonoid com-
pounds. It also implies that CsOMT21 is a 3ꞌ-O-methyltransferase and
likely catalyzes the penultimate step in cannflavin A and B biosynthesis
by converting luteolin to chrysoeriol. To test this possibility, re-
combinant CsOMT21 was purified via Ni2+ affinity chromatography
(Fig. 5B) and assayed with luteolin and unlabelled S-adenosyl methio-
nine as co-substrates. Indeed, the identity of the reaction product was
confirmed to be chrysoeriol, based on its identical HPLC retention time
and mass spectral fragmentation pattern with the authentic standard
(Fig. 5C). Recombinant CsOMT21 exhibited Michaelian kinetics with
kinetic constants that were similar to those that have been previously
reported for this class of enzymes (Fig. 5D). While our analysis cannot
exclude the possibility that the other twenty-one CsOMT family
member(s) may convert luteolin to chrysoeriol, these results do provide
evidence for a branch point from the general flavonoid pathway that is
present in C. sativa, whereby luteolin can be first methylated to chry-
soeriol en route towards cannflavin A and cannflavin B synthesis.
The observation that CsPT3 preferentially prenylates chrysoeriol, in
vitro, and that prenylated luteolin is apparently absent in extracts from
C. sativa implies, a priori, that methylation of luteolin to chrysoeriol
must occur first in the cannflavin A and/or B pathway. We reasoned
that the alleged enzyme that methylates luteolin at the 3′-hydroxyl
position of the flavone B-ring to yield chrysoeriol would likely fall into
the class of S-adenosyl-L-methionine (AdoMet)-dependent O-methyl-
transferases (OMTs), which are widely distributed throughout the plant
kingdom (Ibrahim et al., 1998; Ibrahim, 2005; Kim et al., 2010). We
focused our initial searches of the TSA database for C. sativa on type 1
OMTs, which specifically methylate hydroxyl moieties of phenylpro-
panoid-based compounds (Noel et al., 2003). Using a previously char-
acterized flavonoid-O-methyltransferase from Oryza sativa (OsOMT9)
that methylates the 3′-hydroxyl group on a variety of flavonoids as a
query (Kim et al., 2006), this search uncovered 40 unique nucleotide
sequences corresponding to partial and/or full-length transcripts that
were loosely annotated as ‘caffeic acid-O-methyltransferases’. We next
compared these transcript sequences via BLASTn searches against the
Cannabis whole genome contig database to confirm their corresponding
full-length open reading frames. This analysis revealed twenty-four
unique protein sequences (Fig. S5) which were subsequently annotated
as C. sativa O-methyltransferases (CsOMT1-24).
A phylogenetic analysis of the CsOMT family was then performed to
establish their evolutionary relatedness to various plant OMTs that have
been previously identified to act on aromatic substrates. This analysis
revealed that type 1 CsOMTs are distributed into four general groups
(Fig. 4). It should be noted however, as Schrӧder et al. (2002) and Lam
et al. (2007) previously pointed out, that assigning substrate preference
based on sequence similarity alone for this class of plant enzymes is
precarious. For example, the first group of type 1 plant OMTs depicted
in our phylogenetic analysis includes enzymes that utilize a broad array
of aromatic substrates - from simple phenolic compounds, such as
chavicol, guaiacol and orcinol (Gang et al., 2002; Scalliet et al., 2006;
Akhtar et al., 2013), to more complex heterocyclic aromatics, such as
homoeriodictoyl, myricetin, and resveratrol (Schrӧder et al., 2004;
Schmidin et al., 2008; Schmidt et al., 2011). We found nine CsOMT
family members present within this group. The second group of type 1
OMTs appear specific to the Cannabaceae family and include seven
CsOMTs along with two OMTs from Humulus lupulus that are involved
in the synthesis xanthohumol (Nagel et al., 2008). The third and fourth
groups represent two closely related sister clades of type 1 plant OMTs
and contain the remaining members of the CsOMT family. Strikingly, all
plant OMTs that are known to methylate the 3′-hydroxyl position of
various flavonoids are confined to group three and include re-
presentatives from Arabidopsis, peppermint, rice, wheat, and American
golden saxifrage (Gauthier et al., 1996; Muzac et al., 2000; Willitis
et al., 2004; Kim et al., 2006; Zhou et al., 2006). We found three
CsOMTs (CsOMT6, 12 and 21) that fell into this group.
3. Concluding remarks
Guided by previously published metabolomic data from C. sativa, a
targeted phylogenomics approach combined with in-vitro biochemical
assays was employed in this study to explore the biosynthetic pathway
towards cannflavin A and B (Fig. 6). We provide evidence that a unique
branch point from the general plant flavonoid pathway has evolved in
C. sativa in which the widespread plant flavone, luteolin, is converted
into cannflavin A and B via regiospecific methylation and prenylation
reactions. The identification of these two enzymatic steps opens new
opportunities for the metabolic engineering of cannflavin A and cann-
flavin B biosynthesis and underscores the value of phylogenomics
driven gene discovery, an approach that is largely underutilized in the
Cannabis research space.
2.4. Identification and biochemical characterization of a luteolin O-
methyltransferase
4. Experimental
While recognizing that phylogenetic-driven predictions of plant
OMT function has its caveats, we were nevertheless intrigued by the
presence of CsOMT6, 12 and 21 amongst a group of evolutionary
conserved OMTs that exhibit regioselective methylation activity for the
3′-position on a variety of flavonoids. We therefore chose to survey the
enzymatic activities of these three CsOMTs to elucidate if their encoded
proteins could methylate luteolin at the 3′-hydroxyl position of the
flavone B-ring to yield chrysoeriol. In accordance with our hypothesis,
this reaction would represent the penultimate step in cannflavin A and
B biosynthesis.
4.1. Chemicals and reagents
Authentic flavonoid standards for apigenin and luteolin were pur-
chased from Indofine Chemical Company, kaempherol and quercetin
were purchased from Sigma-Aldrich, and chrysoeriol was from Toronto
Research Chemicals. The trans-prenyl diphosphates, isopentenyl di-
phosphate, dimethyllalyl diphosphate, and geranyl diphosphate, were
obtained from Echelon Biosciences. Radiolableled S-[Methyl-14C] ade-
nosyl-L-methionine (58.0 mCi mmol−1
Synthetic drop-out media lacking histidine for culturing yeast was
)
was from PerkinElmer.
We first introduced the open-reading frames of CsOMT6, 12 and 21
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Phytochemistry164(2019)162–171
Fig. 4. Phylogenetic analysis of type 1 O-methyl-
transferases from C. sativa.
A neighbour-joining
phylogenetic tree (1000 replicates) of type 1 O-me-
thyltransferases from diverse plants was constructed
using the MEGA 6.0 software package. Included in
this analysis are twenty-four unique type 1 O-me-
thyltransferases (CsOMTs) that were identified in the
C. sativa genome. Branch lengths indicate the number
of amino acid substitutions per sequence and boot-
strap values (> 60%) are indicated next to each
branch. The amino acid sequences and species ab-
breviations for all of these type
1 o-methyl-
transferases are listed in Supplemental Fig. S1 along
with their corresponding GenBank accession num-
bers. Note that the CsOMTs (bolded) are uniformly
distributed amongst four main groups of plant type 1
proteins.
obtained from US Biological. All primers were synthesized by Sigma-
Aldrich and are listed in Table S1. All other chemicals were obtained
from Sigma-Aldrich, BioBasic, or Fisher Scientific.
4.3. Cloning and recombinant protein expression of CsOMTs in E. coli
The C. sativa L. O-methyltransferase open reading frames were
synthesized by Genscript. These cDNAs for CsOMT6, 12 and 21 were
amplified by PCR using the KOD Hot Start DNA polymerase (Novagen)
and then ligated between the NdeI/AseI and HindIII sites of the pET28b
vector system (Novagen) which introduces an N-terminal 6 x His tag to
each coding sequence. These constructs were then introduced into E.
coli BL21-CodonPlus (DE3)-RIPL cells. Bacterial cells expressing re-
combinant CsOMT6, 12 and 21 were cultured in LB media at 37 °C to an
OD600 of 0.6. Isopropyl-β-D-thiogalactoside was then added to a final
concentration of 1 mM and the cells were incubated at 16 °C for an
additional 18 h. The bacterial cells were collected by centrifugation, re-
suspended in buffer A (20 mM Tris-HCl, pH 8.0, 500 mM KCl), and then
disrupted by sonication. Crude protein extracts were centrifuged at
12,000×g for 10 min at 4 °C to remove unbroken cells and debris and
then applied to a 1 mL HisTrap HP column (GE Healthcare) equilibrated
in buffer A. Proteins bound to the Ni2+ affinity matrix were washed
with five column volumes of buffer A containing 20 mM imidazole,
eluted with one column volume of buffer A containing 400 mM imi-
dazole, and then immediately desalted on PD-10 columns (GE
Healthcare) equilibrated with 50 mM Tris-HCl, pH 7.5, 5 mM MgCl2,
and 10% (v/v) glycerol. Protein concentration was determined by the
method of Bradford (1976) using BSA as a standard.
4.2. Phylogenetic analysis
The DNA sequences encoding for putative type
1 O-methyl-
transferases (CsOMTs) and aromatic prenyltransferases (CsPTs) were
first identified in the Cannabis sativa genome available at NCBI (https://
blast.ncbi.nlm.nih.gov/). Search was performed in the Transcriptome
Shotgun Assembly (TSA) database available for the strain Purple Kush
(Bioproject Accession PRJNA74271; van Bakel et al., 2011) via
tBLASTn (https://blast.ncbi.nlm.nih.gov/Blast.cgi) for transcripts that
exhibited homology to the Oryza sativa O-methyltransferase, OsOMT9,
and the Glycyrrhiza uralensis prenyltransferase, Gu6ADT. These se-
quences were then matched against the Cannabis whole genome contig
database (https://www.ncbi.nlm.nih.gov/genome/?term = cannabis
+sativa; van Bakel et al., 2011), via BLASTn searches, to retrieve the
full-length open reading frames for the C. sativa OMT and PT gene fa-
milies (Supplemental Fig. S1 and S4). The in-silico assembled amino
acid sequences from these gene sequences were then used to construct
phylogenetic relationships using the MEGA software package (version
6.0) by the neighbor-joining method with bootstrap analysis of 1000
replicates.
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K.A. Rea, et al.
Phytochemistry164(2019)162–171
Figure 5. Enzymatic characterization of CsOMT21.
(A) Three flavones (apigenin, luteolin and chry-
soeriol) and two flavonols (quercetin and kaemp-
ferol) that typically accumulate in C. sativa were
tested as potential substrates. (B) Relative enzyme
activity and substrate preference. Recombinant
CsOMT21 was purified by Ni2+-affinity chromato-
graphy (inset). Proteins were resolved by SDS-PAGE
and stained with Coomassie Blue. Lanes 1 and 2
contain crude E. coli extracts containing CsOMT21
and the purified protein, respectively; Molecular
weights (kDa) are indicated. The selected flavones
and flavonols (as numbered above) were provided to
recombinant CsOMT21 in enzyme assays along with
[
14C]-AdoMet as
means SD from three independent experiments
a methyl donor. Data are the
and are presented as relative activity compared to
that observed with the preferred substrate, luteolin.
(C) Evidence for the conversion of luteolin to chry-
soeriol by CsOMT21. Enzyme assays with CsOMT21
together with luteolin and unlabelled AdoMet as co-
substrates were extracted and analyzed by HPLC.
Note the identical retention time and CID-Q-TOF
mass spectral fragmentation pattern of the enzymatic
product from these assays (bottom panel) with that of
an authentic chrysoeriol standard (top panel). (D)
Kinetic analysis of CsOMT21. Recombinant CsOMT21
was assayed under standard assay conditions at the
indicated concentrations of luteolin. Kinetic para-
meters were determined by non-linear regression
analysis using the Michaelis-Menten kinetics model
of the SigmaPlot 12.3 software.
4.4. Cloning and recombinant protein expression of CsPTs in yeast
additional 18 h at 28 °C to induce protein expression.
The C. sativa L. and G. uralensis prenyltransferase open reading
frames were synthesized by Genscript and included CsPT1, 3, 4, 7, 8
and GuA6DT (Li et al., 2014). These cDNAs were applied by PCR and
ligated between the BamHI and XhoI sites of pESC-HIS (Agilent). The
sequence-verified constructs were introduced into the Saccharomyces
cerevisiae YPH499 yeast strain (ura3–52 lys2–801amber ade2–101ochre
trp1–Δ63 his3–Δ200 leu2–Δ1) using the method outlined by Gietz and
Schiestl (2007) and transformants were selected on synthetic drop-out
media lacking histidine, supplemented with 0.67% yeast nitrogen base
and 2% glucose. For recombinant protein expression, the yeast trans-
formants were cultured as above at 28 °C to an OD600 of 1.0. Yeast cells
were then pelleted by centrifugation (5,000 x rpm, 10 min), washed
twice with sterile water, and re-suspended in the same media as above
containing 2% galactose instead of glucose. Cells were incubated for an
4.5. O-methyltransferase enzyme assays
Assays for determining O-methyltransferase enzyme activity were
performed using ∼2 μg of purified recombinant protein incubated in a
final reaction volume of 100 μL containing 1 mM substrate and 6.9 μM
S-[Methyl-14C] adenosyl-L-methionine in 50 mM Tris-HCl, pH 7.5,
5 mM MgCl2, and 10% (v/v) glycerol for 30 min at 37 °C. The enzymatic
products were extracted with four volumes of ethyl acetate and quan-
tified using a scintillation counter (Model LS6500, Beckman). For re-
action product identification, assays were scaled up to a final volume of
500 μL containing ∼50 μg of recombinant protein, 2 mM substrate and
2 mM S-adenosyl-L-methionine in 50 mM Tris-HCl, pH 7.5, 5 mM
MgCl2, and 10% (v/v) glycerol for 60 min at 37 °C. Enzymatic products
were extracted as above, evaporated to dryness under N2 gas, and
Fig. 6. Proposed biosynthetic pathway for cannflavin A and
B in Cannabis sativa. Solid arrows indicate core steps of the
flavone biosynthetic pathway for which enzymes have been
identified in a variety of plants. Dashed arrows represent
proposed enzymatic reactions based on the data from this
study. Enzyme abbreviations: PAL, phenylalanine ammonia-
lyase; C4H, cinnamic acid 4-hydroxylase; 4CL, 4-coumaric
acid:CoA ligase; CHS, chalcone synthase; CHI, chalcone
isomerase; FNS, flavone synthase; F3ʹH, flavonoid 3ʹ hy-
droxylase; CsOMT21, C. sativa O-methyltransferase 21;
CsPT3, C. sativa prenyltransferase 3.
168

K.A. Rea, et al.
Phytochemistry164(2019)162–171
resuspended in 100 μL of methanol. Samples were applied to
a
0.4 mL min−1. The mass spectrometer electrospray capillary voltage
was maintained at 4.0 kV and the drying gas temperature at 250 °C with
a flow rate of 8 L/min. Nebulizer pressure was 30 psi and the frag-
mentor was set to 160 V. Nitrogen was used as both nebulizing, drying
gas, and collision-induced dissociation gas. The mass-to-charge ratio
was scanned across a range of 100–3000 m/z in 4 GHz extended dy-
namic range positive-ion MS mode. The instrument was externally ca-
librated with the ESI TuneMix (Agilent). The sample injection volume
was 10 μl. Chromatograms were analyzed within Agilent Qualitative
Analysis software B 08.0 finding compounds by the Molecular Feature
algorithm and generating possible compound formulas including ele-
ments C, H, O, and N. Fragmentation patterns of the various parent
(molecular) ions were obtained using collision energies of 5, 10 and
20 eV, with 20 eV being optimal.
Spherisorb ODS2 reverse-phase column (250 mm × 4.6 mm, 5 μm;
Supelco) and resolved by HPLC using a non-linear potassium phosphate
buffer (50 mM, pH 3.0) and acetonitrile gradient at a flow rate of
1 mL min−1. The amount of acetonitrile in the mobile phase under the
starting conditions was 10% and then increased to 30% during the first
10 min. After being maintained at 30% for an additional 10 min, the
acetonitrile concentration was then increased by 5% increments every
five min during the next 30 min such that the final concentration was
60% at the 50 min mark. The eluted products were detected by ab-
sorption at 340 nm and quantified relative to authentic standards. Mass
spectral analysis of the enzymatic products was performed as described
below.
4.6. Microsome extraction and prenyltransferase enzyme assays
4.9. NMR characterization of enzymatic reaction products
Yeast cells expressing the various prenyltransferases were isolated
as described above. The cell pellets were re-suspended in 100 mM Tris-
HCl, pH 9.0 and disrupted with one-half volume of acid-washed glass
beads (425–600 μM, Sigma-Aldrich) for a total of four min (30 s vortex;
30 s on ice). Following lysis, cell debris and glass beads were removed
by centrifugation (1,500×g, 20 min, 4 °C) and microsomes were pel-
leted from the supernatant by ultracentrifugation (160,000×g, 90 min,
4 °C). The resulting supernatant was removed and the pelleted micro-
somes were then re-suspended in 100 mM Tris-HCl, pH 9.0 and protein
concentration was determined by the method of Bradford (1976) using
BSA as a standard. Prenyltransferase enzyme assays were conducted
with ∼200 μg of microsomal protein in a final reaction volume of
200 μL containing 200 μM of prenyl acceptor substrate and 400 μM
DMAPP, GPP or IPP in 100 mM Tris-HCl, pH 9.0 and 10 mM MgCl2.
Assays were allowed to proceed for 60 min at 37 °C and then terminated
with the addition of 10 μL of 20% formic acid. Prenylated reaction
products were extracted with two volumes of ethyl acetate, evaporated
to dryness under N2 gas, and then re-suspended in 100 μL of methanol.
The samples were applied to a Spherisorb ODS2 reverse-phase column
(250 mm × 4.6 mm, 5 μm; Supelco) and eluted with a 20 min linear
gradient from 45% to 95% methanol in water, pH 2.7 containing 0.1%
formic acid (v/v). The mobile phase was maintained at 100% methanol
for an additional 10 min. Products were detected by absorption at
340 nm and quantified relative to authentic standards.
The enzymatic reaction products from assays with CsPT3 were re-
solved by HPLC as described above. Compounds suspected to be
cannflavin A and B eluted at 23.35 min and 20.03 min, respectively,
and were subsequently collected. Approximately 0.5 mg of each com-
pound was evaporated to dryness under N2 gas, resuspended in acetone-
d6, and analyzed using 1H and 13C NMR. NMR spectra were collected on
a Bruker AVANCE III 600 MHz spectrometer equipped with a 5 mm TCI
cryoprobe. The sample temperature was regulated at 298
1 K. Peak
assignments for cannflavin A and B were determined using standard 2D
pulse sequences (COSY: cosygpqf, TOCSY: dipsi2gpphzs, HSQC:
hsqcetgpsisp2.2, HMBC: hmbcgpl2ndqf). The HMBC was collected with
768 increments in the indirect dimension; all other experiments were
collected with 256 indirect increments. The TOCSY mixing time was set
to 80 msec, and the HMBC coupling constant was set to 10 Hz.
Cannflavin A: 1H NMR (Acetone-d6, 600 MHz): δH 6.68 (1H, s, H-3),
6.62 (1H, s, H-8), 7.60 (1H, s, H-2′), 7.00 (1H, d, J = 8.3 Hz, H-5′), 7.57
(1H, d, J = 8.3 Hz, H-6’), 3.36 (2H, d, J = 7.1 Hz, H-1”), 5.29 (1H, dt,
J = 1.1 Hz, 7.2 Hz, H-2”), 1.96 (2H, t, J = 7.1 Hz, H-4”), 2.05 (2H, m,
H-5”), 5.07 (1H, t, J = 7.1 Hz, H-6”), 1.59 (3H, s, H-8”), 1.54 (3H, s, H-
9”), 1.79 (3H, s, H-10”), 3.98 (3H, s, O-CH3), 13.3 (0.5H, bs, 5-OH); 13
C
NMR (Acetone-d6, 150 MHz): δC 164.4 (C-2), 104.2 (C-3), 182.8 (C-4),
159.8 (C-5), 112.0 (C-6), 162.1 (C-7), 93.9 (C-8), 156.3 (C-9), 104.9 (C-
10), 123.4 (C-1′), 110.0 (C-2′), 148.5 (C-3′), 151.0 (C-4′), 115.8 (C-5′),
121.0 (C-6′), 21.8 (C-1"), 122.9 (C-2"), 135 (C-3"), 40.1 (C-4"), 27.6 (C-
5"), 124.8 (C-6"), 131.3 (C-7"), 25.3 (C-8"), 17.3 (C-9"), 16.0 (C-10"),
56.2 (O-CH3).
4.7. Synthesis of 6-dimethylallyl flavone standards
Cannflavin B: 1H NMR (Acetone-d6, 600 MHz): δH 6.69 (1H, s, H-3),
6.62 (1H, s, H-8), 7.61 (1H, d, J = 2.1 Hz, H-2′), 7.00 (1H, d,
J = 8.3 Hz, H-5′), 7.59 (1H, dd, J = 8.3 Hz, 2.1 Hz, H-6′), 3.36 (2H, d,
J = 7.2 Hz, H-1''), 5.28 (1H, m, H-2''), 1.78 (3H, s, H-4''), 1.65 (3H, d,
J = 0.9 Hz, H-5''), 3.99 (3H, s, O-CH3), 13.3 (0.5H, bs, 5-OH); 13C NMR
(Acetone-d6, 150 MHz): δC 164.7 (C-2), 104.5 (C-3), 183.2 (C-4), 160.2
(C-5), 112.3 (C-6), 162.3 (C-7), 94.1 (C-8), 156.6 (C-9), 105.3 (C-10),
123.8 (C-1′), 110.5 (C-2′), 148.8 (C-3′), 151.3 (C-4′), 116.3 (C-5′), 121.3
(C-6′), 22.0 (C-1''), 123.2 (C-2''), 131.7 (C-3''), 17.9 (C-4''), 25.9 (C-5''),
56.6 (O-CH3).
A flavonoid prenyltransferase from G. uralensis (GuA6DT) that cat-
alyzes the regiospecific addition of DMAPP onto position 6 of the A-ring
on a variety of flavones (Li et al., 2014) was expressed in yeast mi-
crosomes, as described above. According to the in vitro pre-
nyltransferase assay conditions outlined above, yeast microsomes ex-
pressing GuA6DT were supplied with apigenin, chrysoeriol, or luteolin
as flavone substrates, along with DMAPP as a prenyl donor. The en-
zymatic reaction products from these assays were resolved by HPLC and
compounds corresponding to 6-dimethylallyl apigenin, 6-dimethylallyl
chrysoeriol, and 6-dimethylallyl luteolin were collected off-line at re-
tention times of 19.58, 19.93, and 18.49 min, respectively.
Declarations of interest
4.8. Mass spectrometry analysis of enzymatic reaction products
K.R., J.A.C, S.J.R., and T.A.A. have interests in patent applications
associated with this work (United States provisional patent application
62-702-528).
The prenylated flavones that were produced by GuA6DT or CsPT3,
in vitro, were purified by HPLC as described above. Samples were then
subjected to liquid chromatography mass spectrometry analysis per-
formed on an Agilent 1200 HPLC liquid chromatograph interfaced with
an Agilent UHD 6530 Q-TOF mass spectrometer. A C18 cartridge
column (Agilent Rapid Resolution 2.1 × 30 mm, 3.5 μm) at 30 °C was
used with the following solvents 1:1 water and acetonitrile both with
0.1% formic acid. The first 2 and last 5 min of the isocratic flow were
sent to waste and not the spectrometer. The flow rate was maintained at
Acknowledgments
We thank Drs. Armen Charchoglyan and Dyanne Brewer for their
expertise with mass spectrometry analysis and Dr. Gale Bozzo for
fruitful discussions regarding flavonoid biochemistry.
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Phytochemistry164(2019)162–171
Appendix A. Supplementary data
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