Cannabinoids, flavonoids, and stilbenoids have been identified in the annual dioecious plant Cannabis sativa L. Of these, the cannabinoids are the best known group of this plant’s natural products. Polyketide synthases (PKSs) are responsible for the biosynthesis of diverse secondary metabolites, including flavonoids and stilbenoids. Biosynthetically, the cannabinoids are polyketide substituted with terpenoid moiety. Using an RT-PCR homology search, PKS cDNAs were isolated from cannabis plants. The deduced amino acid sequences showed 51%-73% identity to other CHS/STS type sequences of the PKS family. Further, phylogenetic analysis revealed that these PKS cDNAs grouped with other non-chalcone-producing PKSs. Homology modeling analysis of these cannabis PKSs predicts a 3D overall fold, similar to alfalfa CHS2, with small steric differences on the residues that shape the active site of the cannabis PKSs.
In plants, polyketide synthases (PKSs) play an important role in the biosynthesis of a myriad of secondary metabolites (Schröder, 1997; Flores-Sanchez and Verpoorte, 2009). PKSs are a group of homodimeric condensing enzymes that catalyze the initial key reactions in the biosynthesis of several compounds, such as flavonoids and stilbenoids. PKSs are classified into three types (Hopwood and Sherman, 1990; Staunton and Weissman, 2001; Fischbach and Walsh, 2006). Chalcone synthase (CHS, EC 18.104.22.168) and stilbene synthase (STS, EC 22.214.171.124) are the most studied enzymes from the group of type III PKSs (Schröder, 2000; Austin and Noel, 2003). Plant PKSs have 44%-95% amino acid identity and are encoded by similarly structured genes. For example, CHSs from Petunia hybrida, Petroselinum hortense, Zea mays, Antirrhinum majus, and Hodeum vulgare, and STS from Arachis hypogaea have 70%-75% identity on the protein level and the CHS and STS genes contain an intron at the same conserved position (Schröder et al., 1988; Schröder and Schröder, 1990). Families of PKS genes have been reported in many plants, such as Daucus carota L. (Hirner and Seitz, 2000), Gerbera hydrida (Helariutta et al., 1996), Glycine max (Shimizu et al., 1999), Humulus lupulus (Novak et al., 2006), Hypericum androsaemum (Liu et al., 2003), Ipomoea purpurea (Durbin et al., 2000), Lycopersicon esculentum (O’Neill et al., 1990), Medicago sativa L. (Junghans et al., 1993), Petunia hybrida (Koes et al., 1989), Phaseolus vulgaris (Ryder et al., 1987), Pinus sylvestris L. (Preisig-Muller et al., 1999), Pisum sativum(Harker et al., 1990), Psilotum nudum (Yamazaki et al., 2001), Rheum palmatum (Abe et al., 2005), Rubus idaeus (Kumar and Ellis, 2003), Ruta graveolens (Springob et al., 2000), Saccharum spp. (Contessotto et al., 2001), Sorghum bicolor (Lo et al., 2002), Vitis vinifera(Wiese et al.., 1994; Goto-Yamamoto et al., 2002), and Zingiber officinale (Radhakrishnan and Soniya, 2009). Their expression is controlled differently and it has been suggested that PKSs have evolved by gene duplication and, subsequently, divergence by mutations, providing an adaptative differentiation to plants (Tropf et al., 1994; Durbin et al., 2000; Lukacin et al., 2001). As PKSs are in vital branch points for the biosynthesis of secondary metabolites, the presence of families of PKSs in one single species emphasizes the importance of their characterization to understand their functional divergence and their contribution to function(s) in different cell types of the plant.
Cannabis sativa L. is an annual dioecious plant from Central Asia. Several compounds have been identified in this plant. Cannabinoids are the best known group of natural products and more than 70 different cannabinoids have been found so far (ElSohly and Slade, 2005; Radwan et al., 2008). Several therapeutic effects of cannabinoids have been described (Williamson and Evans, 2000) and the discovery of an endocannabinoid system in mammals marked a renewed interest in these compounds (Mackie, 2008; Moreira and Lutz, 2008; Heifets and Castillo, 2009). However, other groups of secondary metabolites have also been described, such as flavonoids and stilbenoids (Flores-Sanchez and Verpoorte, 2008b). It is known that the PKSs CHS and STS catalyze the first committed step of the flavonoid and stilbenoid biosynthesis, respectively. Cannabinoid biosynthesis can be initiated by a PKS (Shoyama et al., 1975). Previously, a PKS cDNA was generated from C. sativa leaves. It encodes an enzyme with CHS, phlorisovalerophenone synthase (VPS), and isobutyrophenone synthase (BUS) activities, but lacking olivetolic acid synthase activity (Raharjo et al., 2004). The co-existence of cannabinoids, flavonoids, and stilbenoids in C. sativa could be correlated to different enzymes of the PKS family. Analyses of crude protein extracts from cannabis plant tissues have revealed the presence of PKS enzymatic activities. Multiple PKS activities were detected during the development and growth of glandular trichomes on bracts and the content analyses of cannabinoids and flavonoids revealed differences in their distribution in these glandular tissues (Flores-Sanchez and Verpoorte, 2008a). This report deals with the generation and molecular analyses of PKS cDNAs obtained from messenger RNA from glandular tissues of cannabis plants in order to obtain a PKS gene library for future studies. Homology modeling, motif, and phylogenetic analyses were used for an in silicio expression analysis.
Material and Methods
Seeds of Cannabis sativa, Skunk and Fourway drug-type varieties (The Sensi Seed Bank, Amsterdam, the Netherlands), and Kompolti fiber-type variety (Dr. D. Watson, HortaPharm, Amsterdam, the Netherlands) were germinated and 9-day-old seedlings were planted into 11 LC pots with soil (substrate 45 L, Holland Potgrond, Van der Knaap Group, Kwintsheul, the Netherlands) and maintained under a light intensity of 1930 lux, at 26 °C and a 60.02 ± 7.43% relative humidity (RH). After three weeks the small plants were transplanted into 10 L pots for continued growth until flowering. To initiate the flowering, two-month-old plants were transferred to a photoperiod chamber (12 h light, 27 °C, and 37.0 ± 11.6% RH). 3-month-old male plants were used to pollinate female plants. 5-day-old seedlings, young leaves from 13-week-old plants, female flowers in different stages of development, fruits from 18-day-old plants and male flowers from 4-month-old plants were harvested. Roots from 4-month-old female plants were harvested and washed with cold water to remove residual soil. In addition, cones of H. lupulus at different stages of development were collected in September 2004 from the Pharmacognosy gardens (Leiden University). All vegetal material was weighed and stored at -80 °C.
Isolation of glandular hairs and lupulin glands
Six grams of frozen female flowers containing 17-, 23-, 35-, and 47-day-old glandular trichomes from cannabis plants were removed by shaking frozen material through a tea leaf sieve and collecting it in a mortar containing liquid N2, where it was immediately used for RNA extraction. The effectiveness of this method is comparable to the method reported by Yerger et al. (1992), which consists of shaking the tissues with powdered dry ice and sieving. For lupulin glands, frozen cones of hop were ground in liquid nitrogen using a frozen mortar and pestle only to long enough to separate the bracteoles and then were shaken using the same system as for cannabis glandular hairs.
Total RNA and mRNA isolation
For total RNA isolation from flowers, leaves, roots, seedlings, fruits, glandular hairs, glandular lupulins, and hop cones, frozen tissues (0.1-0.5 g) were ground to a fine powder in a liquid-nitrogen-cooled mortar, suspended and vortexed in 0.5 mL of extraction buffer (0.35 M glycine, 0.048 M NaOH, 0.34 M NaCl, 0.04 M EDTA, and 4% SDS) and 0.5 mL of water-saturated phenol. The suspension was centrifuged at 1,400 rpm for 2 min to separate the phenol and water phases. The RNA was precipitated in 1/3 volume 8 M LiCl at 4 °C overnight. The RNA was collected by centrifugation at 14,000 rpm for 10 min and resuspended in 0.1 mL of H2O. The suspension was heated at 60 °C for 20 min and centrifuged. A total of 5 μL of 3 M Na-acetate (pH 4.88) was added to the supernatant to initiate the precipitation with 0.25 mL of 100% EtOH at -20 °C for 30 min and centrifuged at 14,000 rpm for 7 min. The pellet was washed with 250 μL of 70% EtOH, centrifuged for 2 min at 14,000 rpm, dried at 60 °C for 15 min, dissolved in 50 μL of H2O, and incubated at 50 °C for 10 min.
Alternatively, Micro-Fast Track 2.0 kit and Trizol Reagent (Invitrogen, Carlsband, CA, USA) were also used for mRNA and total RNA isolation following the manufacturer’s instructions. Isolated RNA was stored at -80 °C.
Degenerated primers, HubF (5′-GAGTGGGGYCA RCCCAART-3′), HubR (5′-CCACCIGGRTGWGYAAT CCA-3′), CHSF (5′-GGYTGYIIIGSYRGTGGMA-3′), CHSR (5′-CCIGGYCCRAASCCRAA-3′), STSF (5′-GGITGCIIIGCIGGIGGMAC-3′), and STSR (5′-CCIGGI CCRAAICCRAA-3′) (Biolegio BV, Malden, the Netherlands) were designed based on CHS, STS, and stilbene carboxylate synthase (STCS) sequences from H. lupulus, A. hypogaea, Rheum tataricum, Pinus strobus, V. vinifera and H. macrophylla. For primers HubF, HubR, CHSF, and CHSR, the conserved regions were from CHS and VPS (GenBank accession nos. AJ304877, AB061021, AB061022, AJ430353, and AB047593), while for STSF and STSR they were from STS and STCS (GenBank accession nos. AB027606, AF508150, Z46915, AY059639, AF456446). RT-PCR was performed with total RNA or mRNA as a template using different combinations of primers. Reverse transcription was performed at 50 °C for 1 h followed by deactivation of the ThermoScript Reverse Transcriptase (Invitrogen) at 85 °C for 5 min. The PCR conditions were: 5 cycles of denaturation for 45 s at 94 °C, 1 min of annealing at 40 °C, 1 min of DNA synthesis at 72 °C, followed by 5 cycles with annealing at 41 °C and 5 cycles with annealing at 43 °C, and ending with 35 cycles with annealing at 45 °C. A Perkin Elmer DNA Thermal Cycler 480 and a Taq PCR Core kit (QIAGEN, Hilden, Germany) were used. A final extension step for 10 min at 72 °C was included. The PCR products were separated on 1.5% agarose gel, visualized under UV light, and recovered using Zymoclean Gel DNA Recovery kit (Zymo Research, Orange, CA, USA) or QIAquick PCR Purification kit (QIAGEN) according to the manufacturer’s instructions.
For generation of 5′ and 3′ end cDNAs, we used total RNA, gene specific primers, and a SMART RACE kit (ClonTech, Palo Alto, CA, USA). The cycling parameters were: 94 °C for 1 min followed by 35 cycles at 94 °C for 35 s, annealing temperature for 35 s and 72 °C for 3 min. A final elongation step for 10 min at 72 °C was included. Gene-specific, amplification, and sequencing primers and annealing temperatures are shown in Table S1 (Supplementary Material). The PCR products were separated on 1.5% agarose gel and visualized under UV light. For generation of complete sequences, total RNA and amplification primers were used. Nested amplifications were made with gene-specific primers to select PKS sequences for sequencing. PKS full-length cDNAs were re-sequenced with sequencing primer in order to confirm that the ORFs of the sequences were correct. The corresponding amplification products were ligated into pGEM-T vector and cloned into JM109 cells according to the manufacturer’s instructions (Promega, Madison WI, USA). Plasmids containing the inserted fragment were sequenced (BaseClear, Leiden, the Netherlands).
The PKS 3D models were generated by the web server Geno3D (Combet et al., 2002), using as template the X-ray crystal structures of M. sativa CHS2 (Protein Bank accession nos. 1BI5.pdb, 1CHW.pdb, and 1CMl.pdb). The models were based on the sequence homology of residues Arg5-Ile383 of the PKSs PKSG1, PKSG2, PKSG4, PKSG5, and PKSF3. The VPS model was based on the sequence homology of the residues Val4-Val390. The corresponding Ramachandran plots confirm that the majority of residues grouped in the energetically allowed regions. All models were displayed and analyzed by the program DeepView – Swiss-PdbViewer (Guex and Peitsch, 1997).
Results and Discussion
Amplification of cannabis PKS cDNAs
RNA isolated from glandular hairs of cannabis flowers and plant tissues was used as a template for reverse transcription-polymerase chain reaction (RT-PCR) amplification of segments of PKS mRNAs. RNA from hop tissues was used as a positive control. The degenerated primers corresponded to conserved regions surrounding Gln 119, the catalytic domain around Cys 164, a region surrounding His 303, and the C-terminal region of the selected protein sequences from CHS, STS, and STCS. Amplification with different primer combinations yielded products of expected size (Figure S1, Supplementary Material). Nucleotide sequence analysis showed open reading frames (ORFs) encoding for PKS proteins. Amplifications derived from mRNA of leaves, seedlings, glandular hairs, and female and male flowers showed 76%-78% homology with CHS3 from H. lupulus and 66%-69% homology with the known cannabis CHS-type PKS (Raharjo et al., 2004). This CHS-type PKS was also identified in female and male flowers and in glandular hairs. Partial sequences of VPS and CHS2 from the hop cone’s secretory glands (also called lupulin glands) were obtained. It is known that VPS and CHS1 are expressed in lupulin glands (Okada and Ito, 2001; Matousek et al., 2002a,b) and the presence of a gene family of VPS, as well as one of CHS, has been suggested. Supplementary Figure S2 shows the strategy to obtain the full-length cDNAs of the likely PKS gene(s) differing from the earlier CHS-type PKS gene.
Nucleotide and protein sequence analyses
Several full-length PKS cDNAs containing ORFs of 1158 bp were obtained (Table 1). The main difference among these PKS cDNAs was the size of untranslated regions (UTRs). Several studies suggest that the untranslated regions (UTRs) are important for the control of gene expression in plants at the post-transcriptional level. Stability (Feldbrugge et al., 2001; Schwartz et al., 2006; Vaucheret, 2006), transport (Li and Hunt, 1997; Siomi, 2000), and translation (Klaff et al., 1996; Guo et al., 2000; Geslain and Ribas de Pouplana, 2004) of RNA depend on the 5′-UTR, the 3′-UTR, or both. We believe that variation in the size of UTRs from these PKS cDNAs could be the result of alternative transcription initiation and polyadenylation sites from post-transcriptional processing of PKS pre-mRNAs (Dean et al., 1986; Joshi, 1987; Rothnie, 1996). The nucleotide sequence data were deposited at GenBank database with the GenBank accession numbers EU551163 (PKSG1), EU551164 (PKSG2), EU551162 (PKSF3), EU551165 (PKSG4), and EU551166 (PKSG5).
The ORFs encode proteins of 385 amino acids with a calculated MW of approximately 42 kDa and a pI ranging from 5.98 to 6.09 (Table 2). The predicted amino acid sequences have more than 97% homology (Figure 1). In the plant H. lupulus, three CHS 1 mRNA sequences were isolated that shared more than 99% and 98% identity on nucleotide and amino acid levels, respectively. They are clearly homologous to the original CHS 1 sequence (AJ304877), forming a CHS 1 oligofamily (Matousek et al., 2006). The presence of this oligofamily could promote differences in concentration of prenylflavonoids in different varieties of H. lupulus (De Keukeleire et al., 2003; Matousek et al., 2005, 2010).
According to the percentage of identity at amino acid level (Table S2), our five PKSs showed to have more homology with the CHSs 3, 4, and VPS from H. lupulus than other PKSs (70%-73%). Conserved amino acid residues present in type III PKSs are also preserved in the amino acid sequences from our PKSs (Figure 1). The catalytic triad (Cys157, His297, and Ans330), the “gatekeeper” phenylalanines (Phe208 and Phe259), and Met130, which ties one catalytic site to the other one on the homodimeric complex, as well as Gly250, which determines the elongation cavity volume of the active site, are strictly preserved when compared to CHS2 from alfalfa (Ferrer et al., 1999; Jez et al., 2000a; Jez et al., 2001). The GFGPG loop, which is important for the cyclization reactions in CHS/STS-type PKSs (Suh et al., 2000), is also preserved on our PKSs. In the starter substrate-binding pocket, the amino acid residues Ser126 and Ser332 are also preserved as on alfalfa CHS2, but Glu185, Thr187 and Thr190 are replaced by an Asp, a Met and a Leu, respectively. Only in PKSG2 is the residue Thr187 preserved as on alfalfa CHS2. In the PKS 2-pyrone synthase (2PS), the amino acid residue Thr190 is replaced by a Leu. All these amino acid residues are important for the selectivity of the starter substrate. In alfalfa CHS2, the catalytic efficiency of the p-coumaroyl-CoA-binding pocket was affected by replacement of these residues (Jez et al., 2000b). The replacement of Thr197 by Leu slightly reduced its catalytic efficiency for the substrate p-coumaroyl-CoA. However, it was increased for the substrate acetyl-CoA. It was found that the change of three amino acid residues (Thr197Leu, Gly256Leu, and Ser338Ile) converts a CHS activity to 2PS activity. In our PKSs, the substrate-binding pocket could be slightly different from that of the alfalfa CHS2 by changes from polar to nonpolar amino acid residues (Thr187Met and Thr190Leu) and from length and bulkiness of the side chain residues (Glu185Asp185). Although the residues that shape the geometry of the active site (Pro131, Gly156, Gly160, Asp210, Gly256, Pro298, Gly299, Gly300, Gly329, Gly368, Pro369, and Gly370) are preserved on alfalfa CHS2 (Ferrer et al., 1999), Leu209 is replaced by the amino acid Ile.
The CHS-based homology modeling predicted that our cannabis PKSs would have the same three-dimensional overall fold as alfalfa CHS2 (Figure S3). A schematic representation of the residues that shape the geometry of the active site of cannabis PKSs is shown in Figure 2. The models suggest small differences in the local reorientation of the residues that shape the active site of the cannabis PKSs. The substrate and product specificity of the enzyme reaction can be affected by the steric modulation of the active-site architecture (Ferrer et al., 1999; Jez et al., 2000a,b, 2001; Suh et al., 2000). It can be inferred from these models that cannabis PKSs could have differences in substrate specificity or in catalytic efficiency (kcat/KM). In PKF3 and PKSG5 the Phe208 and Phe259, which are situated at the active site entrance, seem to be closer together than those in the other PKSs. In PKSG1, PKSG2, and PKSG4, the active site entrance looks narrower than that at CHS2. Jez et al. (2002) reported that the replacement F265V increased the preference for aliphatic CoA starters in CHS2, but the replacement F215S yielded a CHS mutant that accepts N-methylanthraniloyl-CoA as a substrate. In acridone synthase (ACS) from Ruta graveolens, the exchange of Val265Phe reduced the catalytic activity and shifted the starter substrate preference (N-methylanthraniloyl-CoA) to p-coumaroyl-CoA, whereas a triple replacement from the residues Ser132Thr, Ala133Ser, and Val265Phe transformed the ACS to a functional CHS (Lukacin et al., 2001, 2005). On the other hand, it has been suggested that Phe215 may help orient substrates at the active site during elongation of the polyketide intermediate and that the position of the CoA’s terminal thiol may affect the conformations of Phe215 and Phe265 (Jez et al., 2000a). A PKS isolated from Polygonum cuspidatum showed a pH-dependent activity and its gatekeepers, Phe215 and Phe265, are replaced by Leu and Cys, respectively. It showed a preference for aromatic CoA esters and could not accept isobutyryl-CoA, isovaleryl-CoA, or acetyl-CoA as substrates (Ma et al., 2009). The residues Leu214 and Phe215 are replaced by Ile214 and Leu215 in benzalacetone synthase (BAS). These residues are involved in the formation of benzalacetone in R. palmatum (Abe et al., 2003). On the other hand, Thr197, Gly256, and Ser338, which are replaced by Ala, Leu, and Thr, respectively, in aloesone synthase (ALS) have different roles in the formation of the heptakedite aloesone in R. palmatum. Gly256 determines starter substrate selectivity, Thr197 controls polyketide chain length, and Ser338 guides the linear polyketide intermediate into the pocket and leads the formation of aloesone (Abe et al., 2006). In cannabis PKSs, the residues Ser126, Leu190, Gly250, and Ser332 have changes in their orientation (Figure 2). As was mentioned above, these minor changes could have drastic effects on the enzymatic activity of each cannabis PKS.
Motif analyses predicted PKSG1, 2, 4, 5 and PKSF3 to be non-secretory proteins with a putative cytoplasmic location. In addition, potential residues for post-translational modifications, such as phosphorylation and glycosylation, were also predicted. Biochemical analyses are required to prove that these PKSs have a cytoplasmic localization and can be modified by glycosylation and/or phosphorylation.
A PKS family in cannabis plants
We characterized five PKS cDNAs, four from glandular hairs (PKSG1, PKSG2, PKSG4, and PKSG5) and one from seedlings (PKSF3). The last one was also identified in male and female flowers by RT-PCR and sequencing, while the expression of PKSG2 was also detected in leaves. Although a low expression of the known cannabis CHS-type PKS was reported in female flowers, glandular hairs, leaves, and roots (Raharjo et al., 2004), we detected by RT-PCR that it is also expressed in male flowers. Southern blot analyses of C. sativa genomic DNA showed that at least four homologous PKS genes are present (Raharjo et al., 2004). A phylogenetic analysis (Figure 3) of our cannabis PKSs revealed that they group together with other non-chalcone and non-stilbene forming enzymes and appear to be most closely related to the CHSs 2, 3, 4, and VPS from H. lupulus, while the known cannabis CHS-type PKS groups with chalcone-forming enzymes and is most closely related with H. lupulus CHS1, of which expression is highly specific in the lupulin glands during the cone maturation, but also it can be detected on all the plant (Matousek et al., 2002a). CHS2has been detected from hop leaf and lupulin fractions and CHS4 and VPS have been detected from the glandular tissue of hop cones, although no expression for CHS3 has been found on any hop tissue (Okada and Ito, 2001; Novak et al., 2003; Okada et al., 2004). In regard to enzyme activities, CHS1 shows CHS activity with p-coumaroyl-CoA. VPS, CHS2, and CHS4 use isovaleryl-CoA and isobutyryl-CoA, but CHS2 and CHS4 reactions form byproducts. No reaction products have been detected for CHS3 enzyme activity (Okada et al., 2004; Novak et al., 2006). Probably, the right substrates have not been identified for this enzyme yet.
A comparison of the 3D model of our PKSs, VPS, and alfalfa CHS2 predicted variations in the orientation of the active site residues which suggests a different substrate specificity regarding VPS (Figure 4).
The isolation and identification of PKSs with different enzymatic activity in one plant species has been reported, as well as the occurrence of PKS gene families in a species (Rolfs and Kindl, 1984; Zheng et al., 2001; Samappito et al., 2002; Matousek et al., 2006). A number of points suggest the participation of several PKSs in the secondary metabolism of this plant. These are: the CHS- and STS-type, and olivetol-forming PKS activities from protein crude extracts from C. sativa (Flores-Sanchez and Verpoorte, 2008a), the expression and partial characterization of a PKS cDNA from leaves with CHS-type activities (Raharjo et al., 2004), the characterization of four PKS cDNAs generated from mRNA of a glandular hair mixture and one from mRNA of seedlings, which is also expressed in female and male flowers (this study), and the small gene family of PKS detected in genomic DNA (Raharjo et al., 2004). Recently, the crystallization of a cannabis PKS, called hexanoyl triacetic acid lactone (HTAL) or olivetol synthase (OLS), condensing malonyl-CoA and hexanoyl-CoA to form hexanoyl triacetic acid lactone or olivetol, was reported (Taguchi et al., 2008; Marks et al., 2009; Taura et al., 2009). It has been proposed that pyrones or polyketide free acid intermediates undergo spontaneous cyclization to yield alkylresorcinolic acids or stilbenecarboxylic acids (Akiyama et al., 1999); or that post-PKS modifying enzymes are required to form them (Austin and Noel, 2003; Eckermann et al., 2003; Flores-Sanchez and Verpoorte, 2009). The homology of this protein with our PKSs was more than 97%. Although the differences in the amino acid residues from our PKSs and HTAL/OLS are small (Figure 1), probably because of the varieties of cannabis plant used, a complete biochemical characterization of the proteins encoded by PKSG1, PKSG2, PKSF3, PKSG4, and PKSG5 is required to study and understand their function and diversity, as well as to learn more about signals or factors that could control their transcription and translation.
It is interesting that the cDNAs PKSG1, PKSG2, PKSG4, and PKSG5 were generated from a combination of trichomes at different development stages. Under our greenhouse conditions, the beginning of the development of the trichomes on the perigonial bracts of female flowers was observed from 15 to 18 days after transferring the plants to a photoperiod regime. A full development of the glandular trichomes with presence of resin was observed from 30-35 days after initiation of the photoperiod regime. In addition, for the gland trichome isolation, flowers from two varieties of drug type (Skunk and Fourway) were used. Mahlberg et al.(1984) reported a glandular secretory system formed by three different forms of glandular trichomes on the epidermis of the outer surface of bracts from female flowers in C. sativa. The identification of non-glandular trichomes was also reported. A higher content of cannabinoids was detected in capitate-stalked glands than in capitate-sessile glands and appeared to be related to the gland age and type of cannabis plant. Bulbous glands are the smallest and there is no direct evidence for the presence of cannabinoids in them yet.
Olivetolic acid, an alkylresorcinolic acid, is the first precursor in the biosynthesis of pentyl-cannabinoids, and the identification of methyl- (Vree et al., 1972), butyl- (Smith, 1997) and propyl-cannabinoids (Shoyama et al., 1977) in cannabis plants suggests the biosynthesis of several alkylresorcinolic acids with different lengths of side-chain moiety (Figure S4). It is known that the activated fatty acid units (fatty acid-CoAs) act as direct precursors that form the side-chain moiety of alkylresorcinols (Suzuki et al., 2003). Probably, more than one PKS-forming alkylresorcinolic acid or pyrone co-exist in cannabis plants. An analysis of cannabinoid content from our plant material showed the presence of THCA, a pentyl-cannabinoid, and THVA, a propyl-cannabinoid, in female flowers (Flores-Sanchez and Verpoorte, 2008a). As in H. lupulus, the presence of these PKSs could yield differences in the concentration of cannabinoids into becoming different varieties of C. sativa. Thus, the biochemical characterization of PKSG1, PKSG2, PKSG4, and PKSG5 will be carried out in the future in order to determine which PKS(s) is (are) involved in the pentyl-cannabinoid and/or propyl-cannabinoid pathways.
The following online material is available for this article:
Oligonucleotide primers and annealing temperatures used in this study.
Homology percentage of C. sativa PKS ORFs with CHSs, STSs, and STCS.
Positions of degenerate primers and of the amplified PCR products, and sizes of PCR products, relative to CHS3 from H. lupulus (GenBank accession no. AB061022). Closed arrow heads indicate the sense and position of the degenerate primers relative to the amino acid sequences of the PKSs CHS, STS, and STCS. Amino acid numbering relative to CHS3 from H. lupulus.
Outline of RT-PCR and RACE for generation of PKS full-length cDNAs. Closed head arrows indicate the sense of the primers. The 5′- and 3′-ends were amplified from mRNA. PF, sense degenerate primer; PR, antisense degenerate primer. For nested amplification, gene-specific primers and amplification primers were used as nested primers.
Structural comparison of alfalfa CHS2 crystal structure with the 3D models from the deduced amino acid sequences of cannabis PKS cDNAs. The active site residues are shown as blue backbones; in alfalfa CHS structure naringenin and malonyl-CoA are shown as red and dark red backbones.
Proposed substrates for cannabis alkylresorcinolic acid-forming PKSs.
This material is available as part of the online article from http://www.scielo.br/gmb.
I.J. Flores Sanchez received a partial grant from CONACYT (Mexico).
Associate Editor: Sandro José de Souza
- Abe I., Sano Y., Takahashi Y., Noguchi H. Site-directed mutagenesis of benzalacetone synthase. J Biol Chem. 2003;278:25218–25226. [PubMed]
- Abe I., Watanabe T., Noguchi H. Chalcone synthase superfamily of type III polyketide synthases from rhubarb (Rheum palmatum) Proc Japan Acad Ser B. 2005;81:434–440.
- Abe I., Watanabe T., Lou W., Noguchi H. Active site residues governing substrates selectivity and polyketide chain length in aloesone synthase. FEBS J. 2006;273:208–218. [PubMed]
- Akiyama T., Shibuya M., Liu H.M., Ebizuka Y. p-Coumaroyltriacetic acid synthase, a new homologue of chalcone synthase, from Hydrangea macrophylla var. thunbergii. Eur J Biochem. 1999;263:834–839. [PubMed]
- Austin M.B., Noel J.P. The chalcone synthase superfamily of type III polyketide synthases. Nat Prod Rep. 2003;20:79–110. [PubMed]
- Combet C., Jambon M., Deleage G., Geourjon C. Geno3D: Automatic comparative molecular modeling of protein. Bioinformatics. 2002;18:213–214. [PubMed]
- Contessotto M.G.G., Monteiro-Vitorello C.B., Mariani P.D.S.C., Coutinho L.L. A new member of the chalcone synthase (CHS) family in sugarcane. Genet Mol Biol. 2001;24:257–261.
- Dean C., Tamaki S., Dunsmuir P., Favreau M., Katayama C., Dooner H., Bedbrook J. mRNA transcripts of several plant genes are polyadenylated at multiple sites in vivo. Nucleic Acids Res. 1986;14:2229–2240. [PMC free article] [PubMed]
- De Keukeleire J., Ooms G., Heyerick A., Roldan-Ruiz I., van Bockstaele E., De Keukeleire D. Formation and accumulation of α-acids, β-acids, desmethylxanthohumol, and xanthohumol during flowering of hops (Humulus lupulusL. ) J Agric Food Chem. 2003;51:4436–4441. [PubMed]
- Durbin M.L., McCaig B., Clegg M.T. Molecular evolution of the chalcone synthase multigene family in the morning glory genome. Plant Mol Biol. 2000;42:79–92.[PubMed]
- Eckermann C., Schröder G., Eckermann S., Strack D., Schmidt J., Schneider B., Schröder J. Stilbenecarboxylate biosynthesis: A new function in the family of chalcone synthase-related proteins. Phytochemistry. 2003;62:271–286. [PubMed]
- ElSohly M.A., Slade D. Chemical constituents of marijuana: The complex mixture of natural cannabinoids. Life Sci. 2005;78:539–548. [PubMed]
- Feldbrugge M., Arizti P., Sullivan M.L., Zamore P.D., Belasco J.G., Green P.J. Comparative analyses of the plant mRNA-destabilizing element, DST, in mammalian and tobacco cells. Plant Mol Biol. 2001;49:215–223. [PubMed]
- Ferrer J.L., Jez J.M., Bowman M.E., Dixon R.A., Noel J.P. Structure of chalcone synthase and the molecular basis of plant polyketide biosynthesis. Nat Struct Biol. 1999;6:775–784. [PubMed]
- Fischbach M.A., Walsh C.T. Assembly-line enzymology for polyketide and nonribosomal peptide antibiotics: Logic, machinery, and mechanisms. Chem Rev. 2006;106:3468–3496. [PubMed]
- Flores-Sanchez I.J., Verpoorte R. PKS activities and biosynthesis of cannabinoids and flavonoids in Cannabis sativa L. plants. Plant Cell Physiol. 2008a;49:1767–1782.[PubMed]
- Flores-Sanchez I.J., Verpoorte R. Secondary metabolism in cannabis. Phytochem Rev. 2008b;7:615–639.
- Flores-Sanchez I.J., Verpoorte R. Plant polyketide synthases: A fascinating group of enzymes. Plant Physiol Biochem. 2009;47:167–174. [PubMed]
- Geslain R., Ribas de Pouplana L. Regulation of RNA function by aminoacylation and edition? Trends Genet. 2004;20:604–610. [PubMed]
- Goto-Yamamoto N., Wan G.H., Masaki K., Kobayashi S. Structure and transcription of three chalcone synthase genes of grapevine (Vitis vinifera) Plant Sci. 2002;162:867–872.
- Guex N., Peitsch M.C. Swiss-Model and the Swiss-PdbViewer: An environment for comparative protein modeling. Electrophoresis. 1997;18:2714–2723. [PubMed]
- Guo L., Allen E., Miller W.A. Structure and function of a cap-independent translation element that functions in either the 3′ or the 5′ untranslated region. RNA. 2000;6:1808–1820. [PMC free article] [PubMed]
- Harker C.L., Ellis T.H.N., Coen E.S. Identification and genetic regulation of the chalcone synthase multigene family in pea. Plant Cell. 1990;2:185–194.[PMC free article] [PubMed]
- Heifets B.D., Castillo P.E. Endocannabinoid signaling and long-term synaptic plasticity. Annu Rev Physiol. 2009;71:283–306. [PMC free article] [PubMed]
- Helariutta Y., Kotilainen M., Elomaa P., Kalkkinen N., Bremer K., Teeri T., Albert V.A. Duplication and functional divergence in the chalcone synthase gene family of Asteraceae: Evolution with substrate change and catalytic simplification. Proc Natl Acad Sci USA. 1996;93:9033–9038. [PMC free article] [PubMed]
- Hirner A.A., Seitz H.U. Isoforms of chalcone synthase in Daucus carota L. and their differential expression in organs from the European wild carrot and in ultraviolet-A-irradiated cell cultures. Planta. 2000;210:993–998. [PubMed]
- Hopwood D.A., Sherman D.H. Molecular genetics of polyketides and its comparison to fatty acid biosynthesis. Annu Rev Genet. 1990;24:37–66. [PubMed]
- Jez J.M., Ferrer J.L., Bowman M.E., Dixon R.A., Noel J.P. Dissection of malonyl-coenzyme A decarboxylation from polyketide formation in the reaction mechanism of a plant polyketide synthase. Biochemistry. 2000a;39:890–902. [PubMed]
- Jez J.M., Austin M.B., Ferrer J.L., Bowman M.E., Schröder J., Noel J.P. Structural control of polyketide formation in plant-specific polyketide synthases. Chem Biol. 2000b;7:919–930. [PubMed]
- Jez J.M., Bowman M.E., Noel J.P. Structure-guided programming of polyketide chain-length determination in chalcone synthase. Biochemistry. 2001;40:14829–14838.[PubMed]
- Jez J.M., Bowman M.E., Noel J.P. Expanding the biosynthetic repertoire of plant type III polyketide synthases by altering starter molecule specificity. Proc Natl Acad Sci USA. 2002;99:5319–5324. [PMC free article] [PubMed]
- Joshi C.P. An inspection of the domain between putative TATA box and translation start site in 79 plant genes. Nucleic Acids Res. 1987;15:6643–6653. [PMC free article][PubMed]
- Junghans H., Dalkin K., Dixon R.A. Stress response in alfalfa (Medicago sativa L. ) 15: Characterization and expression patterns of members of a subset of the chalcone synthase multigene family. Plant Mol Biol. 1993;22:239–253. [PubMed]
- Klaff P., Riesner D., Steger G. RNA structure and the regulation of gene expression. Plant Mol Biol. 1996;32:89–196. [PubMed]
- Koes R.E., Spelt C.E., van den Elzen P.J.M., Mol J.N.M. Cloning and molecular characterization of the chalcone synthase multigene family of Petunia hybrida. Gene. 1989;81:245–257. [PubMed]
- Kumar A., Ellis B.E. A family of polyketide synthase genes expressed in ripening Rubus fruits. Phytochemistry. 2003;62:513–526. [PubMed]
- Li Q., Hunt A. The polyadenylation of RNA in plants. Plant Physiol. 1997;115:321–325. [PMC free article] [PubMed]
- Liu B., Falkenstein-Paul H., Schmidt W., Beerhues L. Benzophenone synthase and chalcone synthase from Hypericum androsaemum cell cultures: cDNA cloning, functional expression and site-directed mutagenesis of two polyketide synthases. Plant J. 2003;34:847–855. [PubMed]
- Lo C., Coolbaugh R.C., Nicholson R.L. Molecular characterization and in silicoexpression analysis of a chalcone synthase gene family in Sorghum bicolor. Physiol Mol Plant Pathol. 2002;61:179–188.
- Lukacin R., Schreiner S., Matern U. Transformation of acridone synthase to chalcone synthase. FEBS Lett. 2001;508:413–417. [PubMed]
- Lukacin R., Schreiner S., Silber K., Matern U. Starter substrate specificities of wild-type and mutant polyketide synthases from Rutaceae. Phytochemistry. 2005;66:277–284. [PubMed]
- Ma L.Q., Pang X.B., Shen H.Y., Pu G.B., Wang H.H., Lei C.Y., Wang H., Li G.F., Liu B.Y., Ye H.C. A novel type III polyketide synthase encoded by a three-intron gene from Polygonum cuspidatum. Planta. 2009;229:457–469. [PubMed]
- Mackie K. Signaling via CNS cannabinoid receptors. Mol Cell Endocrinol. 2008;286:S60–S65. [PMC free article] [PubMed]
- Mahlberg P.G., Hammond C.T., Turner J.C., Hemphill J.K. Structure, development and composition of glandular trichomes of Cannabis sativa L. In: Rodriguez E., Healey P.L., Mehta I., editors. Biology and Chemistry of Plant Trichomes. Plenum Press: New York; 1984. pp. 23–51.
- Marks M.D., Tian L., Wenger J.P., Omburo S.N., Soto-Fuentes W., He J., Gang D.R., Weiblen G.D., Dixon R.A. Identification of candidate genes affecting Δ9-tetrahydrocannabinol biosynthesis in Cannabis sativa. J Exp Bot. 2009;60:3715–3726.[PMC free article] [PubMed]
- Matousek J., Novak P., Briza J., Patzak J., Niedermeierova H. Cloning and characterization of chs-specific DNA and cDNA sequences from hop (Humulus lupulus L. ) Plant Sci. 2002a;162:1007–1018.
- Matousek J., Novak P., Patzak J., Briza J., Krofta K. Analysis of true chalcone synthase from Humulus lupulus L. and biotechnology aspects of medicinal hops. Rostl Vyroba. 2002b;48:7–14.
- Matousek J., Vrba L., Skopek J., Novak P., Patzak J., De Keukeleire J., Skopek J., Heyerick A., Roldan-Ruiz I., De Keukeleire D. Cloning and molecular analysis of the regulatory factor HIMyb1 in hop (Humulus lupulus L. ) and the potential of hop to produce bioactive prenylated flavonoids. J Agric Food Chem. 2005;53:4793–4798.[PubMed]
- Matousek J., Vrba L., Skopek J., Orctova L., Pesina K., Heyerick A., Baulcombe D., De Keukeleire D. Sequence analysis of a “true’ chalcone synthase (chs H1) oligofamily from hop (Humulus lupulus L. ) and PAP1 activation of chs H1 in heterologous systems. J Agric Food Chem. 2006;54:7606–7615. [PubMed]
- Matousek J., Kocabek T., Patzak J., Stehlik J., Fussy Z., Krofta K., Heyerick A., Roldan-Ruiz I., Maloukh L., De Keukeleire D. Cloning and molecular analysis of HIbZip1 and HIbZip2 transcription factors putatively involved in the regulation of the lupulin metabolome in hop (Humulus lupulus L. ) J Agric Food Chem. 2010;58:902–912. [PubMed]
- Moreira F.A., Lutz B. The endocannabinoid system: Emotion, learning and addiction. Addict Biol. 2008;13:196–212. [PubMed]
- Novak P., Matousek J., Briza J. Valerophenone synthase-like chalcone synthase homologues in Humulus lupulus. Biol Plant. 2003;46:375–381.
- Novak P., Krofta K., Matousek J. Chalcone synthase homologues from Humulus lupulus: Some enzymatic properties and expression. Biol Plant. 2006;50:48–54.
- Okada Y., Ito K. Cloning and analyses of valerophenone synthase gene expressed specifically in lupulin gland of Hop (H. lupulus L. ) Biosci Biotechnol Biochem. 2001;65:150–155. [PubMed]
- Okada Y., Sano Y., Kaneko T., Abe I., Noguchi H., Ito K. Enzymatic reactions by five chalcone synthase homologs from hop (Humulus lupulus L. ) Biosci Biotechnol Biochem. 2004;68:1142–1145. [PubMed]
- O’Neill S.D., Tong Y., Sporlein B., Forkmann G., Yoder J.I. Molecular genetic analyses of chalcone synthase in Lycopersicon esculentum and an anthocyanin-deficient mutant. Mol Gen Genet. 1990;224:279–288. [PubMed]
- Preisig-Muller R., Schwekendiek A., Brehm I., Reif H.J., Kindl H. Characterization of a pine multigene family containing elicitor-responsive stilbene synthase genes. Plant Mol Biol. 1999;39:221–229. [PubMed]
- Radhakrishnan E.K., Soniya E.V. Molecular analysis of type III polyketide synthase (PKS) gene family from Zingiber officinale Rosc. Afr J Plant Sci. 2009;3:44–48.
- Radwan M.M., Ross S.A., Slade D., Ahmed S.A., Zulfiqar F., ElSohly M.A. Isolation and characterization of new cannabis constituents from a high potency variety. Planta Med. 2008;74:267–272. [PMC free article] [PubMed]
- Raharjo T.J., Chang W.T., Verberne M.C., Peltenburg-Looman A.M.G., Linthorst H.J.M., Verpoorte R. Cloning and over-expression of a cDNA encoding a polyketide synthase from Cannabis sativa. Plant Physiol Biochem. 2004;42:291–297. [PubMed]
- Rolfs C.H., Kindl H. Stilbene synthase and chalcone synthase: Two different constitutive enzymes in cultured cells of Picea excelsa. Plant Physiol. 1984;75:489–492. [PMC free article] [PubMed]
- Rothnie H.M. Plant mRNA 3′-end formation. Plant Mol Biol. 1996;32:43–61.[PubMed]
- Ryder T.B., Hedrick S.A., Bell J.N., Liang X., Clouse S.D., Lamb C.J. Organization and differential activation of a gene family encoding the plant defense enzyme chalcone synthase in Phaseolus vulgaris. Mol Gen Genet. 1987;210:219–233.[PubMed]
- Samappito S., Page J., Schmidt J., De-Eknamkul W., Kutchan T.M. Molecular characterization of root-specific chalcone synthases from Cassia alata. Planta. 2002;216:64–71. [PubMed]
- Schröder G., Brown J.W.S., Schröder J. Molecular analyses of resveratrol synthase: cDNA, genomic clones and relationship with chalcone synthase. Eur J Biochem. 1988;172:161–169. [PubMed]
- Schröder J. A family of plant-specific polyketide synthases: Facts and predictions. Trends Plant Sci. 1997;2:373–378.
- Schröder J. The family of chalcone synthase-related proteins: Functional diversity and evolution. In: Romeo J.T., Ibrahim R.K., Varin L., De Luca V., editors. Evolution of Metabolic Pathways. v. 34. Amsterdam: Pergamon Press; 2000. pp. 55–89.
- Schröder J., Schröder G. Stilbene and chalcone synthases: Related enzymes with functions in plant-specific pathways. Z Naturforsch C. 1990;45:1–8. [PubMed]
- Schwartz A.M., Komarova T.V., Skulachev M.V., Zvereva A.S., Dorokhov Y.L., Atabekov J.G. Stability of plant mRNA depends on the length of the 3′-untranslated region. Biochemistry. 2006;71:1377–1384. [PubMed]
- Shimizu T., Akada S., Senda M., Ishikawa R., Harada T., Niizeki M., Dube S.K. Enhanced expression and differential inducibility of soybean chalcone synthase genes by supplemental UV-B in dark-grown seedlings. Plant Mol Biol. 1999;39:785–795.[PubMed]
- Shoyama Y., Yagi M., Nishioka I. Biosynthesis of cannabinoid acids. Phytochemistry. 1975;14:2189–2192.
- Shoyama Y., Hirano H., Makino H., Umekita N., Nishioka I. Cannabis X: The isolation and structures of four new propyl cannabinoids acids, tetrahydrocannabivarinic acid, cannabidivarinic acid, cannabichromevarinic acid and cannabigerovarinic acid, from Thai cannabis, ‘Meao variant’. Chem Pharm Bull. 1977;25:2306–2311.
- Siomi M.C. The molecular mechanisms of messenger RNA nuclear export. Cell Struct Funct. 2000;25:227–235. [PubMed]
- Smith R.M. Identification of butyl cannabinoids in marijuana. J Forensic Sci. 1997;42:610–618.
- Springob K., Lukacin R., Ernwein C., Groning I., Matern U. Specificities of functionally expressed chalcone and acridone synthases from Ruta graveolens. Eur J Biochem. 2000;267:6552–6559. [PubMed]
- Staunton J., Weissman K.J. Polyketide biosynthesis: A millennium review. Nat Prod Rep. 2001;18:380–416. [PubMed]
- Suh D.Y., Fukuma K., Kagami J., Yamazaki Y., Shibuya M., Ebizuka Y., Sankawa U. Identification of amino acid residues important in the cyclization reactions of chalcone and stilbene synthases. Biochem J. 2000;350:229–235. [PMC free article] [PubMed]
- Suzuki Y., Kurano M., Esumi Y., Yamaguchi I., Doi Y. Biosynthesis of 5-alkylresorcinol in rice: Incorporation of a putative fatty acid unit in the 5-alkylresorcinol carbon chain. Bioorg Chem. 2003;31:437–452. [PubMed]
- Taguchi C., Taura F., Tamada T., Shoyama Y., Shoyama Y., Tanaka H., Kuroki R., Morimoto S. Crystallization and preliminary X-ray diffraction studies of polyketide synthase-1 (PKS-1) from Cannabis sativa. Acta Cryst F. 2008;64:217–220.[PMC free article] [PubMed]
- Taura F., Tanaka S., Taguchi C., Fukamizu T., Tanaka H., Shoyama Y., Morimoto S. Characterization of olivetol synthase, a polyketide synthase putatively involved in cannabinoid biosynthetic pathway. FEBS Lett. 2009;583:2061–2066. [PubMed]
- Tropf S., Lanz T., Schröder J., Schröder G. Evidence that stilbene synthases have developed from chalcone synthases several times in the course of evolution. J Mol Evol. 1994;38:610–618. [PubMed]
- Vaucheret H. Post-transcriptional small RNA pathways in plants: Mechanisms and regulations. Genes Dev. 2006;20:759–771. [PubMed]
- Vree T.B., Breimer D.D., van Ginneken C.A.M., van Rossum J.M. Identification in hashish of tetrahydrocannabinol, cannabidiol and cannabinol analogues with a methyl side-chain. J Pharm Pharmacol. 1972;24:7–12. [PubMed]
- Wiese W., Vornam B., Krause E., Kindl H. Structural organization and differential expression of three stilbene synthase genes located on a 13 kb grapevine DNA fragment. Plant Mol Biol. 1994;26:667–677. [PubMed]
- Williamson E.M., Evans F.J. Cannabinoids in clinical practice. Drugs. 2000;60:1303–1314. [PubMed]
- Yamazaki Y., Suh D.Y., Sitthithaworn W., Ishiguro K., Kobayashi Y., Shibuya M., Ebizuka Y., Sankawa U. Diverse chalcone synthase superfamily enzymes from the most primitive vascular plant, Psilotum nudum. Planta. 2001;214:75–84. [PubMed]
- Yerger E.H., Grazzini R.A., Hesk D., Cox-Foster D.L., Craig R., Mumma R.O. A rapid method for isolation glandular trichomes. Plant Physiol. 1992;99:1–7.[PMC free article] [PubMed]
- Zheng D., Schröder G., Schröder J., Hrazdina G. Molecular and biochemical characterization of three aromatic polyketide synthase genes from Rubus idaeus. Plant Mol Biol. 2001;46:1–15. [PubMed]
- Geno3D server. [May, 2007 ]. Available from: http://genoed-pbil.ibcp.fr.
- DeepView-the Swiss-Pdbviewer. [May, 2007 ]. Available from: http://www.expasy.org/spdbv/
- Motif analysis softwares. [May, 2007 ]. Available from: http://www.cbs.dtu.dk/services/,http://urgi.versailles.inra.fr/predator/andhttp://myhits.isb-sib.ch/cgi-bin/motif_scan/
- Stilbenecarboxylate information website. [June, 2008 ]. Available from: http://www.biologie.uni-freiburg.de/data/bio2/schroeder/stilbenecarboxylates.html.
- CLUSTALW program, v. 1.83. European Bioinformatics Institute; Available from: http://www.ebi.ac.uk/Tools/clustalw/index.html.
- TreeView program, v. 1.6.6. Available from: http://taxonomy.zoology.gla.ac.uk/rod/treeview.html.