Cannabinoid synthases and osmoprotective metabolites accumulate in the exudates of Cannabis sativa L. glandular trichomes

Accepted Manuscript Title: Cannabinoid synthases and osmoprotective metabolites accumulate in the exudates of Cannabis sativa L. glandular trichomes Authors: Paweł Rodziewicz, Stefan Loroch, Łukasz Marczak, Albert Sickmann, Oliver Kayser PII: DOI: Reference:

S0168-9452(18)31486-9 https://doi.org/10.1016/j.plantsci.2019.04.008 PSL 10118

To appear in:

Plant Science

Received date: Revised date: Accepted date:

11 December 2018 9 April 2019 10 April 2019

Please cite this article as: Rodziewicz P, Loroch S, Marczak Ł, Sickmann A, Kayser O, Cannabinoid synthases and osmoprotective metabolites accumulate in the exudates of Cannabis sativa L. glandular trichomes, Plant Science (2019), https://doi.org/10.1016/j.plantsci.2019.04.008 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Cannabinoid synthases and osmoprotective metabolites accumulate in the exudates of Cannabis sativa L. glandular trichomes Paweł Rodziewicz1, Stefan Loroch2, Łukasz Marczak3, Albert Sickmann2,4,5, Oliver Kayser1* 1

Department of Technical Biochemistry, Technical University Dortmund, Emil-Figge-Str. 66

44227 Dortmund, Germany Leibniz-Institut für Analytische Wissenschaften - ISAS – e.V., Bunsen-Kirchhoff-Str. 11,

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2

44139 Dortmund, Germany

European Centre for Bioinformatics and Genomics, Institute of Bioorganic Chemistry PAS,

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3

Piotrowo 2, 60-965 Poznan, Poland 4

Medizinische Fakultät, Ruhr-Universität Bochum, 44801 Bochum, Germany.

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Department of Chemistry, College of Physical Sciences, University of Aberdeen, Aberdeen,

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AB24 3FX, United Kingdom *

Address:

Oliver Kayser

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TU Dortmund University

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corresponding author

Technical Biochemistry

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Emil-Figge-Str. 66 44227 Dortmund Germany

(+49) 231 755 7487

e-mail:

[email protected]

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Highlights

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Tel:

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 

Cannabinoid synthases accumulate in cannabis glandular trichome exudates during the flowering period. The extracted cannabis exudates retain catalytic activity ex vivo under trichomemimicking conditions.

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Cannabis glandular trichome exudates contain hydrophobic and amphiphilic compounds, but also hydrophilic metabolites with osmoprotective properties.

Abstract

Cannabinoids are terpenophenolic compounds produced by Cannabis sativa L., which accumulate in storage cavities of glandular trichomes as a part of the exudates. We investigated if tetrahydrocannabinolic acid synthase and cannabidiolic acid synthase, which are involved in the last step of cannabinoid biosynthesis, are also secreted into Cannabis

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trichome exudates. The exudates were collected by microsuction from storage cavities of

Cannabis glandular trichomes and were subjected for proteomic and metabolomic analyses.

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The catalytic activity of the exudates was documented by cannabigerolic acid

biotransformation studies under hydrophobic conditions. Electrophoretic separations revealed protein bands at ~65 kDa, which were further identified as tetrahydrocannabinolic acid synthase and cannabidiolic acid synthase. The accumulation of the enzymes in trichome

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exudates increased substantially during the flowering period in the drug-type Cannabis plants.

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The content of cannabinoids increased significantly after incubating hexane-diluted trichome exudates with cannabigerolic acid. In this study, we showed that Cannabis glandular

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trichomes secrete and accumulate cannabinoid synthases in storage cavities, and the enzymes

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able to convert cannabigerolic acid under hydrophobic trichome-mimicking conditions. Metabolite profiling of the exudates revealed compounds with hydrophilic, osmoprotective

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and amphiphilic properties, which may play a role in providing a necessary aqueous microenvironment, which enables enzyme solubility and biocatalysis under hydrophobic conditions of glandular trichomes.

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Keywords

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cannabinoids; Cannabis sativa; cannabidiolic acid synthase; glandular trichomes;

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osmolytes; tetrahydrocannabinolic acid synthase.

1. Introduction

The complex biochemistry of glandular trichomes results in the biosynthesis of various classes of secondary metabolites, including terpenoids, alkaloids, phenylpropanoids, cannabinoids, flavonoids, fatty acids, acyl sugars, and other [1]. The compounds produced in glandular trichomes act as a chemical weapon against pathogens, but also play a vital role in

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attracting pollinators [2,3]. From a humankind perspective, these substances are used as fragrances, food additives, natural pesticides and fungicides, but foremost they constitute an

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important source of pharmaceuticals [1,4]. The excellent example of the latter is antimalarial

drug artemisinin from Artemisia annua L. [5]. Also, cannabinoids from Cannabis sativa L. have become promising therapeutic agents for treating cancer, inflammation, appetite disorders, and neurological diseases [6]. However, most of the known medicinal effects are

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associated with decarboxylated forms of two major cannabinoids found in the plant – the

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psychoactive tetrahydrocannabinol (THC) and its non-psychoactive isomer cannabidiol (CBD) [7].

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The biosynthesis of cannabinoids is largely separated from primary cellular

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metabolism and takes place in glandular trichomes [8–10]. The capitate-stalked glandular trichomes, which are abundantly present on Cannabis female inflorescences, comprise of one

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basal cell, several stalk cells, and secretory cells surrounded by a large sub-cuticular storage cavity [4,11]. This type of glandular trichomes also contains the highest concentrations of cannabinoids [12–14]. The precursors for cannabinoid biosynthesis originate from two distinct biosynthetic pathways: the polyketide pathway, in which olivetolic acid (OA) is

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produced [15], and the plastidal 2-C-methyl-D-erythritol 4-phosphate (MEP) pathway, which leads to the biosynthesis of geranyl diphosphate (GPP) [16]. In further steps OA is prenylated

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with GPP by aromatic membrane-bound prenyltransferase, leading to the formation of cannabigerolic acid (CBGA), which is a central precursor for cannabinoid biosynthesis [17].

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Tetrahydrocannabinolic acid synthase (THCAS) and cannabidiolic acid synthase (CBDAS) are the ultimate enzymes in the cannabinoid pathway, and catalyze the unique oxidative cyclization of CBGA, which results in the formation of tetrahydrocannabinolic acid (THCA) and cannabidiolic acid (CBDA), respectively (Fig. S1) [18,19]. Both enzymes belong to flavin-dependent berberine bridge enzyme protein family and represent two isoforms of the same synthase, arising from two codominant alleles at one locus [20]. Both enzymes have a

theoretical molecular mass of ~ 62 kDa, and share 84% identity in their amino acid sequences, including the N-terminal 28 amino acid secretion signal peptide [19,21–23]. Cannabinoids are toxic to plant cells and cause mitochondrial permeability transitions and DNA degradation, which ultimately induce apoptosis [24,25]. To avoid cellular damage cannabinoids are accumulated in storage cavities of glandular trichomes, which were also suggested as the precise site of the last step of their biosynthesis [9,14]. Although the secretory pathway for THCAS and CBDAS was confirmed in heterologous systems, and

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THCAS was immunolocalized in the storage cavities of glandular trichomes in transgenic

tobacco, the enzymes have never been identified in the exudates of Cannabis glandular

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trichomes [22,23,26]. In this study we report the identification of THCAS and CBDAS in

Cannabis glandular trichome exudates and provide the evidence for CBGA bioconversion under hydrophobic trichome-mimicking environment. We also report the metabolic profile of the trichome exudates, and possible components, which may play a role in providing

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hydrophilic conditions for enzyme solubility and biocatalysis under glandular trichome

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conditions.

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2.1. Chemicals

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2. Material and Methods

All chemicals used for the experiments were of analytical grade. Solvents used for chromatography and mass spectrometry analysis were of LC-MS and GC-MS grade or higher. CBGA was purchased from Taros Chemicals (Dortmund, Germany). Purity by 99% is

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documented.

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2.2. Plant material and cultivation

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Female plants of Cannabis sativa L. strain Euphoria (Royal Queen Seeds, UK) and

variety Finola (Finola Oy, Finland) were used for the experiments. The drug-type Cannabis (Euphoria) is a chemotype II plant with 1:1 THC:CBD ratio, and medicinal value. Finola is an oilseed hemp, and belongs to chemotype III, with CBD as a dominant cannabinoid, and with only minor amounts of THC [27,28]. To ensure uniform genetic background, plants were generated from cuttings obtained from mother plants. Cannabis plants were cultivated in plant growth chamber (CLF Plant Climatics GmbH, Germany) under controlled conditions with

18/6 h of light/dark at 25/21 °C (day/night) temperature and 70% humidity. Plants were grown in a semi-hydroponic system with the Flora Series nutrient solution (General Hydroponics, USA), applied accordingly to manufacturer’s instruction. After 8 weeks of vegetative growth, flowering phase was induced by changing light regime to 12/12 h, which lasted for 8 weeks. All plant handling and experimental procedures were carried out under the license No. 4584916 issued by Federal Institute for Drugs and Medical Devices (BfArM,

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Germany).

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2.3. Microsuction of glandular trichome storage cavities and sample preparations

Cannabis leaves containing glandular trichomes were used for experiments. The leaves were attached to a microscope slide with a tape. The exudates were collected individually from capitate-stalked glandular trichome storage cavities (gtsc) under the stereomicroscope

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(model SMZ800, Nikon, Japan) using thin glass capillary (inner diameter 100 µm) attached to

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the cell micromanipulator TransferMan 2 (Eppendorf, Germany). For proteomic and metabolomic studies the collected material was transferred into glass vials containing 1.5 ml

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of ice-cold acetone. To optimize the transfer of the exudates, the material was displaced into

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the solvent each time after collecting the content from 10 gtsc, until the desired amount of the exudates was accumulated in the vial. To identify and monitor the accumulation of the

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secreted proteinaceous component, several samplings of the exudates were made. The exudates from 800 gtsc (both strains) and 1000 gtsc (Euphoria) were collected at 7th week of the flowering period. The time point was selected based on the previous results [29]. The samples were centrifuged (16,000 g, 4 °C, 10 min), and the supernatant was removed. The

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remaining pellet was dissolved in 5 µl of sample buffer and submitted for SDS-PAGE. The detected protein bands on the gel prepared from 800 gtsc were further analyzed with LC-MS.

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The sample containing 1000 gtsc was submitted to western blot analysis using primary polyclonal antibody specific for THCAS (Davids Biotechnologie GmbH, Germany). The of

the

peptide

selected

as

the

antibody

epitope

was

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sequence

SKHIPNNVANPKLVYTQHDQL. The secondary antibody was an alkaline phosphateconjugated goat anti-rabbit IgG (Sigma, Germany). To monitor the accumulation profile of THCAS and CBDAS, protein samples were prepared from 500 gtsc collected in triplicates at early-mid (5th week) and late (8th week) flowering stages from Euphoria strain. The samples were digested in-solution and analyzed with LC-MS. In each case, the amounts of sampled gtsc were sufficient to visualize and detect the secreted enzymes. The metabolite profile of the

exudates was determined from the extracts collected from 1000 gtsc, which ensured sufficient concentration of metabolites and their identification by GC-MS. The biotransformation study was based on the samples containing lower amounts of gtsc (200) to minimalize the content of naturally occurring cannabinoids and other metabolites, and to ensure detectable differences in cannabinoid concentration after enzymatic conversion of the substrate.

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2.4. Trichome isolation and analysis

Apart from glandular trichome exudates, whole trichomes were also analyzed.

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Trichomes were extracted as described by Yerger et al. [31] with certain modifications. Briefly, 5 grams of Cannabis (Euphoria) fresh flowers at 7th week of the flowering stage were

put in liquid nitrogen and transferred into 50 ml falcon tube and approximately 5 cm 3 of fine powdered dry ice was added. The tube was capped with a 140 µm nylon mesh (Merck,

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Germany) and the content was shaken by hand for 1 min into a small beaker placed on dry

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ice. The acquired trichomes were transferred into 2 ml tube and stored in -80 °C. Proteins were extracted from 100 mg of trichomes by phenolic method described by Hurkman and

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Tanaka [32] and submitted for SDS PAGE and mass spectrometry identification or western

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blot analysis with the antibody specific for THCAS.

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2.5. CBGA biotransformation assay in hydrophobic conditions

The exudates extracted from 200 gtsc in triplicates were transferred immediately into 100 µl vials containing 20 µl of ice-cold hexane. This solvent was used to provide

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hydrophobic, trichome-mimicking environment, and it is known to have only a minor effect on protein denaturation [33]. To avoid water condensation, the probes were placed in 1.5 ml

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tubes and were kept on ice. The vials were kept closed, and each time before opening and transferring the portion of the sampled exudates they were dried with kimwipes. Each of the

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acquired samples was divided into two, each containing 9 µl of collected material. Subsequently, 1 µl of hexane (control) or 1µl of 100 µM CBGA in hexane (assay) was added; 10 µl of 10 µM CBGA in hexane was used as a negative control. The samples were put in a plastic rack, covered with aluminum foil and placed in incubator. This approach ensured even distribution of the temperature and prevented hexane condensation under the vial caps. The samples were incubated in 37 °C for 30 min followed by centrifugation (16,000 g, 1 min) and

vacuum concentration. The samples were stored in -80 °C until GC-MS analysis. Prior analysis the samples were dissolved in 30 µl of hexane.

2.6. SDS-polyacrylamide gel electrophoresis and sample preparation of gel bands for LC-MS

SDS-polyacrylamide gel electrophoresis was performed using Mini-Protean II system

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(BioRad, USA) and "in-house" prepared 12% acrylamide gels (7 cm x 10 cm x 0.1 cm). The electrophoretic separations were performed at 80V for 20 min., and subsequently at 100 V for

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50 min. at room temperature. The gels were stained overnight with colloidal solution of Coomassie Brilliant Blue and the excess dye from the gel matrix was removed with deionized water [30]. Protein bands were excised from SDS-gels and digested with trypsin (Trypsin Gold, Promega, USA) following the protocol described by Shevchenko and coworkers [34]

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with certain modifications. Briefly, each excised gel piece was washed two times with 50 µl

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50 mM ammonium bicarbonate (ABC) followed by 50 µl 25 mM ABC in 50% acetonitrile (ACN). Subsequently, the samples were incubated in 10 mM dithiothreitol (DTT) at 56°C for

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30 min and next in 30 mM iodoacetamide (IAA) in the dark. The gel pieces were washed

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again two times and were dried in a vacuum centrifuge. For digestion, 5 µl of trypsin solution (12.5 ng/µl in 50 mM ABC, gold/MS grade, Promega, USA) was added and the samples were

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incubated overnight at 37°C. The peptides were eluted twice with 15 µl of 0.1% of trifluoroacetic acid (TFA) following 15 µl of 0.1% TFA/60% ACN and finally dried under vacuum.

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2.7. Sample preparation of trichome exudates for LC-MS and GC-MS analysis

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Samples obtained from week 5th and 8th were digested in-solution. Briefly, samples

were centrifuged (20,000g, 4 °C, 15 min) and the precipitate was re-solubilized in 2 µl of 2 M

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guanidinium hydrochloride and 10 mM DTT. Cysteines were reduced by incubation at 56°C for 30 min and alkylated by adding 1 µl 60 mM IAA and incubation for 30 min in the dark. Twelve µl 50 mM ABC/1mM CaCl2 and 1 µl of 10 ng/µl trypsin (sequencing grade, Promega, USA) were added. The samples were incubated for 14 h at 37°C. One µl of 1 % TFA was added to stop the reaction Prior GC-MS analysis the samples containing exudates collected from 1000 gtsc were vacuum concentrated. The first sample was dissolved in 60 µl of hexane and subjected

directly to GC-MS analysis. The second sample was dissolved in 20 µl of 20 mg/ml methoxyamine hydrochloride solution in pyridine, and the trimethylsilyl derivatization of polar metabolites was performed for 1.5 h followed by 30-min reaction with 40 µl of Nmethyl-N-(trimethylsilyl)trifluoroacetamide (MSTFA). Both reactions were performed at 37 °C.

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2.8. LC-MS

The digested samples were subjected to nanoLC-MS/MS analysis using LTQ Orbitrap

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Velos Pro or LTQ Orbitrap Elite mass spectrometer, online coupled to a U3000 nanoHPLC

(Thermo Scientific, USA). Samples were loaded onto a C18 trap-column (5 µm, 100 µm × 2 cm) at a flow rate of 12 µl/min in 0.1 % TFA. After 10 min, the trap column was switched inline with the C18 main column (3 µm, 75 µm × 50 cm) and peptides were separated at 270

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nl/min and 60 °C using a 45 min linear ACN gradient in the presence of 0.1% formic acid.

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The column effluent was introduced to the MS using a nano-ESI source equipped with silica coated PicoTip (360 µm outer and 10 µm inner diameters at the tip; New Objective,

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Switzerland) operated at 1.5 kV and at 275 °C transfer tube temperature. Data dependent

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acquisition was performed using a topN collision induced dissociation method. Survey scans were acquired in the Orbitrap (300-2000 m/z) with a resolution of 60,000 at m/z 400 using a

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target value of 1E6 (maximum fill time 100 ms). The most intense ions were isolated and fragmented in the linear ion trap (target value 1E4, maximum fill time 100 ms) using 35% normalized collision energy. The lock mass at m/z = 371.101236 was used for internal calibration and precursor ions were excluded for at least 5 sec from re-fragmentation using the

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dynamic exclusion option. Raw files of gel bands were searched using Mascot 2.4.1, raw files of trichome extracts were searched using Sequest and Mascot 2.6.1 implemented in Proteome

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Discoverer v1.4 against the UniProt "Viridiplantae" database (March 2015, 36,399 target sequences). Error tolerances were set to 0.5 Da and 10 ppm for MS and MS/MS and oxidation

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of Met was set as variable and carbamidomethylation of Cys as static modification. Results were filtered for ≤ 1 % false discovery rate on the level of peptide spectrum matches using the target decoy PSM validator (gel bands) or Percolator v2.04 (trichome extracts). For quantification, peak areas of the first three isotope peaks were determined from extracted ion chromatograms (± 20 ppm) using XCalibur v2.2.44 (Thermo Scientific, USA). THCAS was quantified via the identified unique peptide [HIPNNVANPK+2H]2+, CBDAS was quantified via the summed area of the two identified unique peptides [NEQSIPPLPR+2H]2+ and

[TLVDPNNFFR+2H]2+ since other unique peptides were too low in intensity for robust quantification.

2.10. GC-MS

The samples prepared for metabolite profiling and the samples resulting from CBGA biotransformation studies were subjected to GC-MS analysis using TRACE 1310 gas

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chromatograph connected to TSQ8000 triplequad MS (both Thermo Scientific, USA). A DB5 bonded-phase fused-silica capillary column (30 m length, 0.25 mm inner diameter, 0.25 µm

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film thickness) (J&W Scientific Co., USA) was used for separation. The GC oven temperature program was as follows: 2 min at 70 °C, raised by 10 °C/min to 300 °C and held for 10 min at 300 °C. The total time of GC analysis was 36 min. Helium was used as the

carrier gas at a flow rate of 1 ml/min. One microliter of each sample was injected in splitless

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mode. The initial injector temperature was 40 °C for 0.1 min and after that time raised by 600

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°C/min to 350 °C. The septum purge flow rate was 3 ml/min and the purge was turned on after 60 s. The transfer line and ion source temperatures were set to 250 °C. In-source

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fragmentation was performed with 70 eV energy. Mass spectra were recorded in the mass

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range 35–650 m/z. Data acquisition, peak detection, mass spectrum deconvolution, and

3. Results

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library search were done in XCalibur v.2.2 SP1 48 (Thermo Scientific, USA).

3.1. Identification of THCAS and CBDAS in the storage cavities of capitate-stalked

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glandular trichomes

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The microsuction technique enabled us to precisely collect the exudates from

glandular trichome storage cavities without disrupting the secretory cells (Fig. 1). Similar

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technique was already applied to collect the glandular trichome exudates from wild potato for proteomic studies [35]. The electrophoretic separation of the samples prepared from glandular trichome exudates combined with mass spectrometry analysis enabled identification of THCAS and CBDAS in Euphoria and Finola plants, respectively (Fig. 2, Tab. 1). In the exudates collected from Euphoria strain, THCAS was also detected with the antibody (Fig. 3A). In turn, from the SDS-gel prepared from the whole-trichome protein extract, THCAS was identified with mass spectrometry at two masses: ~65 kDa (less abundant) and ~74 kDa

(more abundant) (Fig. 3C, Tab. 1). In the parallel sample subjected to western blot analysis, the signal which corresponded to THCAS was detected only at ~65 kDa, and not at ~74 kDa (Fig. 3B). The putative secretory signal peptide at the N-terminal end of ~74 kDa THCAS could hinder the antibody capacity to bind its target sequence, localized directly after the signal peptide. The other signals detected with the antibody in the whole trichome extract were identified as olivetolic acid cyclase (protein bands 5-7) (Fig. 3B,C, Tab. 1), which share slight sequence similarity with THCAS near the peptide selected as epitope, which could be

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the reason for unspecific binding.

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3.2. Monitoring the level of cannabinoid synthases during flowering period

The multiple steps in gel-based protein digestion method generate material losses, which in turn may limit the rate of successful identification [36]. We applied in-solution

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protein digestion method and LC-MS analysis to monitor the level of cannabinoid synthases

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during mid and late flowering stages in glandular trichome exudates of Euphoria plants. Apart from THCAS, CBDAS was also identified from the samples collected at 5th and 8th week of

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the flowering stage (Fig. S2, S3, Tab. S1). We also detected minor contaminations such as

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glyceraldehyde-3-phosphate dehydrogenase (2 peptides) and histone H2B5 (1 peptide) (Fig. S4, Tab. S1). However, none of those proteins correlated with the abundance of THCAS or

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CBDAS and were only identified in some of the replicates, which could arise from minor cell rupture or contamination during extraction procedure. The accumulation of both cannabinoid synthases increased over the flowering period and the fold change of 8.7 and 4.6 for THCAS

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and CBDAS was noted, respectively (Fig. 4, Fig. S2, S3).

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3.3. Bioconversion of CBGA under hydrophobic conditions

The exudates extracted from the drug-type Cannabis (Euphoria) were diluted in

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hexane and submitted for bioconversion studies with CBGA as a substrate. Due to low amount of the collected material, isomeric and hydrophobic nature of the analyzed compounds, we chose GC-MS to evaluate the samples after bioassay. This analytical platform has already been proven as a valuable tool used for cannabinoid analysis, but high temperatures operating during GC analysis cause decarboxylation of the analyzed cannabinoids, and therefore only the ions of their neutral forms can be observed (Fig. S6)

[37,38]. After incubating trichome exudates with CBGA we noted 2.7 and 2.3-fold change in CBD and THC concentrations, respectively (Fig. 5A-D, Fig. S5).

3.4. Identification of metabolites in the exudates of Cannabis glandular trichomes

GC-MS analysis of the glandular trichome exudates from the drug-type Cannabis (Euphoria) enabled detection of more than 4000 compounds in both extracts and identification

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of 174 in total (Fig. S7, Tab. S2, S3). The vast majority of the annotated metabolites were hydrophobic, i.e. aliphatic hydrocarbons and terpenoids (Fig. 6, Tab. S2, S3). However,

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amphiphilic and hydrophilic compounds were also identified, for example, fatty acid amines,

fatty alcohols, carboxylic acids, amino acids, sugars and polyols (Fig. 6, Tab. S2, S3). The metabolites identified in hexane extract constituted 60% of the total ion current (TIC). Although the largest groups were terpenoids (43 compounds) and aliphatic hydrocarbons (35

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compounds), amphiphilic fatty acid amines (only 5 identified compounds) constituted 25.8%

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of TIC, and palmitamide and myristamide were the most abundant metabolites in that group (15.2% and 7.8% of TIC, respectively). Sesquiterpenes (29 compounds) constituted the

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largest class of secondary metabolites identified in trichome exudates with β-eudesmol, γ-

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eudesmol, α-farnesene, and (-)-aristolene as the most abundant components. Other terpenoid compounds identified in both extracts belonged to monoterpenoids, diterpenoids, and terpene

identified,

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alcohols. Cannabinoids constituted 12.9% of TIC in hexane extract and eight of them were including

CBG,

Δ9-THC,

Δ8-THC,

CBD,

Δ9-tetrahydrocannabivarin,

cannabichromene, cannabinol and cannabicyclol. In the sample subjected to derivatization the identified compounds constituted 30% of TIC and the most abundant metabolites were fatty

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acids: stearic acid (5% of TIC) and palmitic acid (4.2%), followed by cannabinoids: CBD (3.6%) and Δ9-THC (3.4%). Polyols (3% of TIC) were the most numerous class of

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metabolites with hydrophilic properties, including glycerol, which was the main component (2.6%), and also pinitol, galactinol, glyceric acid, threonic acid, mannitol, meso-erythritol,

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and myo-inositol. Other abundant hydrophiles included lactic acid (4.2% of TIC), glycine (1.9%), and sucrose (0.5%) (Tab. S3).

4. Discussion

4.1. THCAS and CBDAS accumulate in the exudates of glandular trichomes during flowering period

Cannabinoid synthases undergo structural changes during the secretory pathway, including cleaving out the secretion peptide and processing post-translational modifications, which may explain the differences in mass between the enzymes identified from various parts of Cannabis plant, i.e. exudates, whole glandular trichomes, and leaves [18,19,22,26]. Glycosylation is particularly found in secreted proteins and has a significant impact on their structure, solubility as well as activity [39]. Also, the conformation of the proteins with lower

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level of glycosylation is usually more hydrophobic, which, in turn, may lead to stronger intra-

protein electrostatic interactions in conditions with limited water activity [40]. Seven possible

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Asn N-glycosylation sites have been already confirmed for cannabinoid synthases, and the

enzymes were found to be highly glycosylated in heterologous systems [19,23,41]. However, after completing the secretory pathway both synthases had lower mass and were less glycosylated, but exhibited higher catalytic activity than their highly glycosylated

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counterparts, which suggest a regulatory role of that post-translation modification [23,42].

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The increase in the accumulation of cannabinoid synthases in the exudates of glandular trichomes observed from mid to late flowering period agrees with an increasing content of

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cannabinoids during the generative plant growth [7,43]. The ratio of THCAS to CBDAS did

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not change significantly during the flowering period. The relative abundance of the identified synthases in Euphoria strain reflected its chemotype, which is stable during the life cycle of

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Cannabis, and the switch from one to another is usually not observed [28,44]. The chemotype and the potency of the particular Cannabis strain, as well as the availability of the substrates used for cannabinoid production, may affect the expression of cannabinoid biosynthetic genes. For instance, the expression of the genes involved in cannabinoid pathway was

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reported to be consistent with the rate of cannabinoid and terpenoid production during early to mid-flowering periods in the flowers of high THCA-producing Cannabis strain (chemotype I)

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[45]. On the other hand, during mid to late flowering stages in moderately potent Cannabis strain (chemotype II), the abundance of transcripts of THCAS and CBDAS followed only

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slight changes, whereas the genes involved in the production of precursor compound, i.e. olivetolic acid, were down-regulated, but correlated with increasing content of cannabinoids [29].

4.2. THCAS and CBDAS accumulated in glandular trichome exudates are catalytically active under hydrophobic conditions

In aqueous buffers cannabinoid synthases remain active only for a short time [19,23,41,42]. According to the proposed mechanism, the oxidative cyclization of CBGA relies on hydrophilic interactions between the substrate and the active center of the enzyme, but the reaction was found to be independent from molecular oxygen and co-produced hydrogen peroxide [18,26,46]. It was also suggested that in aqueous media THCAS may suffer from substrate/product inhibition since the hydrophobic molecules may slowly enter/leave the enzyme catalytic center from/into polar environment [47]. In the two-phase

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setup, which is closer to the conditions in glandular trichomes, the addition of hexane to

aqueous buffer positively influenced stability of THCAS and significantly prolonged its

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catalytic activity (from 15 min to 3 h) [47,48]. The limitation in mass transfer of the substrate

and the inhibition of the enzyme at the interface were suggested as the possible reasons for lower cannabinoid yield, which was, however, improved by exchanging the organic phase and adding higher amounts of CBGA [48]. The biosynthesis of metabolites in monophasic organic

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solvents was demonstrated for several enzymes, but mainly in the industrial context [49].

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However, it must be underlined that even in such conditions, the water shell around the enzyme has to be preserved since it has a critical impact on the ionization state of amino acid

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residues, and therefore on the protein function [40,49,50]. The remaining water layer is

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essential for biotransformation, and in organic solvents many enzymes showed activity only after adding low amount of water [51]. For example, polyphenol oxidase required 0.1% of

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water (v/v) to catalyse the oxidation of phenols to o-quinones in chloroform, but even after creating the hydration shell, the enzyme remained undissolved in organic phase [51,52]. The issue can be overcome by adding amphiphilic compounds to the reaction media and creating microemulsions or reverse micelles depending on the experimental setup [53]. These systems

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ensure high catalytic activity and since many enzymes act on the surface of various biological

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membranes they were also proposed as an in vitro model to study biocatalysis [54].

4.3. Osmoprotective metabolites secreted into glandular trichome exudates may play

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a role in providing appropriate environment for enzymatic reactions

Cannabinoid synthases exhibit similarities to other berberine bridge-like enzymes in terms of sequence and physicochemical properties [55]. Also, the structural characteristics of THCAS are rather typical for the enzymes soluble in aqueous buffers with polar amino acids exposed on the surface of the protein [46]. For example, both cannabinoid synthases share 47% sequence identity (65% similarity including substitutions) with nectarin V secreted into the

floral nectar of tobacco [56]. Floral nectars are hydrophilic solutions, which contain high concentrations of carbohydrates, but also amino acids and organic acids [57]. Apart from playing a role in attracting pollinators, these metabolites also constitute a working environment for the enzymes catalyzing ex vivo [58–60]. Osmoprotective compounds, such as sugars, polyols, and polar amino acids, are able to maintain the essential configuration of the enzymes by providing the hydration shell, especially in conditions with limited water activity [58,61,62]. For instance, in the mucilage of carnivorous sundew (Drosera capenis L.) myo-

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inositol plays a role as a cross-linker in forming hydrogen-bond network between polysaccharide strands, which constitute a hydrophilic matrix for the secreted digestive

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enzymes [63,64]. Osmolytes and amphiphiles were also found in the glandular secretions of coltsfoot (Tussilago farfara L.), but the extract was not tested for the presence of proteins

[65]. However, in case of other aromatic plants, such as thyme, sage, and rosemary, in addition to carbohydrates, the secreted proteinaceous component was also detected, but not

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yet identified [66–68]. In turn, in glandular trichomes of Solanum species, which also produce

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large amounts of osmolytes and amphiphilic compounds, polyphenol oxidase was found to be the main secreted enzymatic component [35,69–71]. Likewise, Cannabis exudates also

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constitute a mixture of proteins and metabolites with hydrophobic, amphiphilic, and

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hydrophilic properties. The histochemical analysis of the storage cavities of glandular trichomes of Cannabis, but also hops and Leonotis leonurus L., revealed ultrastructural

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system of microchannels and secretory vesicles [72–74]. Although, its role in trichome metabolism as well as the exact components were not precisely determined, it was found that the dense layer delimiting secretory vesicles comprise of pectin polysaccharides, and constitute an interface between hydrophilic and lipophilic regions, which further suggest

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emulsion properties of the exudates [73,74].

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5. Conclusions

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In this study we identified THCAS and CBDAS in the exudates of Cannabis glandular

trichomes, which accumulate during the flowering period, and remain catalytically active under hydrophobic conditions. Since these enzymes do not exhibit unusual properties for being solubilized under hydrophobic conditions, the compounds with hydrophilic and amphiphilic properties detected in the exudates may play a role in creating an aqueous microenvironment necessary for biocatalysis. More studies are needed to identify the exact role of the secreted compounds in trichome metabolism, the location of cannabinoid synthases

within the ultrastructural network of Cannabis exudates, the contribution of the secreted synthases to the overall production of cannabinoids, and also the nature of metabolite transport in glandular trichomes. More investigations should also concern structural changes cannabinoid synthases undergo during secretory pathway, and especially the significance of the post-translational modifications on the structure, activity and solubility of these enzymes. Further studies should also include analysis of trichome exudates from other plant species, which may result in identification of other secreted enzymes. Understanding the mechanisms

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of the biocatalysis in glandular trichomes will contribute to the development of these secretory structures as natural biosynthetic factories, and may also advance the field of

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biosynthesis of secondary metabolites in organic solvents.

Author Contributions

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PR and OK designed the research; PR, SL, and ŁM performed the experiments; PR,

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SL, ŁM collected, analyzed and interpreted the data. PR, SL, ŁM, AS, and OK wrote the

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Conflict of interest

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manuscript. All authors contributed to the final version of the manuscript.

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All authors declare no conflict of interest.

Acknowledgements

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We would like to thank Ingo Feldmann and Cornelia Schumbrutzki for their help with sample preparations for proteomic analyses, and Dr. Armin Quentmeier for providing the

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antibody. PR would like to thank Patricia Tkocz for the critical input during this work. LCMS analyses were performed at Leibniz-Institut für Analytische Wissenschaften - ISAS – e.V,

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Germany. GC-MS analyses were performed at European Centre for Bioinformatics and Genomics, Poland.

References

[1] A. Huchelmann, M. Boutry, C. Hachez, Plant glandular trichomes: natural cell factories of high biotechnological interest, Plant Physiol. 175 (2017) 6–22.

A

CC

EP

TE D

M

A

N

U

SC R

IP T

[2] A. Kessler, Defensive function of herbivore-induced plant volatile emissions in nature, Science. 291 (2001) 2141–2144. [3] I. Filella, C. Primante, J. Llusià, A.M. Martín González, R. Seco, G. Farré-Armengol, A. Rodrigo, J. Bosch, J. Peñuelas, Floral advertisement scent in a changing plant-pollinators market, Sci. Rep. 3 (2013) 3434. [4] J. Glas, B. Schimmel, J. Alba, R. Escobar-Bravo, R. Schuurink, M. Kant, Plant glandular trichomes as targets for breeding or engineering of resistance to herbivores, Int. J. Mol. Sci. 13 (2012) 17077–17103. [5] P.J. Weathers, P.R. Arsenault, P.S. Covello, A. McMickle, K.H. Teoh, D.W. Reed, Artemisinin production in Artemisia annua: studies in planta and results of a novel delivery method for treating malaria and other neglected diseases, Phytochem. Rev. 10 (2011) 173–183. [6] R.G. Pertwee, Cannabinoid pharmacology: the first 66 years, Br. J. Pharmacol. 147 (2006) 163–171. [7] O. Aizpurua-Olaizola, U. Soydaner, E. Öztürk, D. Schibano, Y. Simsir, P. Navarro, N. Etxebarria, A. Usobiaga, Evolution of the cannabinoid and terpene content during the growth of Cannabis sativa plants from different chemotypes, J. Nat. Prod. 79 (2016) 324–331. [8] P. Dayanandan, P.B. Kaufman, Trichomes of Cannabis sativa L. (Cannabaceae), Am. J. Bot. 63 (1976) 578–591. [9] E.-S. Kim, P.G. Mahlberg, Immunochemical localization of tetrahydrocannabinol (THC) in cryofixed glandular trichomes of Cannabis (Cannabaceae), Am. J. Bot. 84 (1997) 336–342. [10] J.C. Turner, J.K. Hemphill, P.G. Mahlberg, Quantitative determination of cannabinoids in individual glandular trichomes of Cannabis sativa L. (Cannabaceae), Am. J. Bot. 65 (1978) 1103–1106. [11] G.J. Wagner, Secreting glandular trichomes: more than just hairs, Plant Physiol. 96 (1991) 675–679. [12] C.T. Hammond, P.G. Mahlberg, Morphology of glandular hairs of Cannabis sativa from scanning electron microscopy, Am. J. Bot. 60 (1973) 524–528. [13] N. Happyana, S. Agnolet, R. Muntendam, A. Van Dam, B. Schneider, O. Kayser, Analysis of cannabinoids in laser-microdissected trichomes of medicinal Cannabis sativa using LCMS and cryogenic NMR, Phytochemistry. 87 (2013) 51–59. [14] P.G. Mahlberg, E.S. Kim, Accumulation of cannabinoids in glandular trichomes of Cannabis (Cannabaceae), J. Ind. Hemp. 9 (2004) 15–36. [15] S.J. Gagne, J.M. Stout, E. Liu, Z. Boubakir, S.M. Clark, J.E. Page, Identification of olivetolic acid cyclase from Cannabis sativa reveals a unique catalytic route to plant polyketides, Proc. Natl. Acad. Sci. 109 (2012) 12811–12816. [16] M. Phillips, P. Leon, A. Boronat, M. Rodriguez-Concepción, The plastidial MEP pathway: unified nomenclature and resources, Trends Plant Sci. 13 (2008) 619–623. [17] M. Fellermeier, M.H. Zenk, Prenylation of olivetolate by a hemp transferase yields cannabigerolic acid, the precursor of tetrahydrocannabinol, FEBS Lett. 427 (1998) 283– 285. [18] F. Taura, S. Morimoto, Y. Shoyama, R. Mechoulam, First direct evidence for the mechanism of Δ1-tetrahydrocannabinolic acid biosynthesis, J. Am. Chem. Soc. 117 (1995) 9766–9767. [19] F. Taura, S. Sirikantaramas, Y. Shoyama, K. Yoshikai, Y. Shoyama, S. Morimoto, Cannabidiolic-acid synthase, the chemotype-determining enzyme in the fiber-type Cannabis sativa, FEBS Lett. 581 (2007) 2929–2934.

A

CC

EP

TE D

M

A

N

U

SC R

IP T

[20] E.P.M. de Meijer, M. Bagatta, A. Carboni, P. Crucitti, V.M.C. Moliterni, P. Ranalli, G. Mandolino, The inheritance of chemical phenotype in Cannabis sativa L, Genetics. 163 (2003) 335–346. [21] C. Onofri, E.P.M. de Meijer, G. Mandolino, Sequence heterogeneity of cannabidiolicand tetrahydrocannabinolic acid-synthase in Cannabis sativa L. and its relationship with chemical phenotype, Phytochemistry. 116 (2015) 57–68. [22] S. Sirikantaramas, Tetrahydrocannabinolic acid synthase, the enzyme controlling marijuana psychoactivity, is secreted into the storage cavity of the glandular trichomes, Plant Cell Physiol. 46 (2005) 1578–1582. [23] S. Sirikantaramas, S. Morimoto, Y. Shoyama, Y. Ishikawa, Y. Wada, Y. Shoyama, F. Taura, The gene controlling marijuana psychoactivity: molecular cloning and heterologous expression of Δ1-tetrahydrocannabinolic acid synthase from Cannabis sativa L., J. Biol. Chem. 279 (2004) 39767–39774. [24] S. Morimoto, Y. Tanaka, K. Sasaki, H. Tanaka, T. Fukamizu, Y. Shoyama, Y. Shoyama, F. Taura, Identification and characterization of cannabinoids that induce cell death through mitochondrial permeability transition in Cannabis leaf cells, J. Biol. Chem. 282 (2007) 20739–20751. [25] Y. Shoyama, C. Sugawa, H. Tanaka, S. Morimoto, Cannabinoids act as necrosisinducing factors in Cannabis sativa, Plant Signal. Behav. 3 (2008) 1111–1112. [26] F. Taura, S. Morimoto, Y. Shoyama, Purification and characterization of cannabidiolicacid synthase from Cannabis sativa L.. Biochemical analysis of a novel enzyme that catalyzes the oxidocyclization of cannabigerolic acid to cannabidiolic acid, J. Biol. Chem. 271 (1996) 17411–17416. [27] D. Rotherham, S.A. Harbison, Differentiation of drug and non-drug Cannabis using a single nucleotide polymorphism (SNP) assay, Forensic Sci. Int. 207 (2011) 193–197. [28] G. Fournier, C. Richez-Dumanois, J. Duvezin, J.-P. Mathieu, M. Paris, Identification of a New Chemotype in Cannabis sativa : Cannabigerol - Dominant Plants, Biogenetic and Agronomic Prospects, Planta Med. 53 (1987) 277–280. [29] N. Happyana, O. Kayser, Monitoring metabolite profiles of Cannabis sativa L. trichomes during flowering period using 1H NMR-based metabolomics and real-time PCR, Planta Med. 82 (2016) 1217–1223. [30] V. Neuhoff, R. Stamm, H. Eibl, Clear background and highly sensitive protein staining with Coomassie Blue dyes in polyacrylamide gels: A systematic analysis, Electrophoresis. 6 (1985) 427–448. [31] E.H. Yerger, R.A. Grazzini, D. Hesk, D.L. Cox-Foster, R. Craig, R.O. Mumma, A rapid method for isolating glandular trichomes, Plant Physiol. 99 (1992) 1–7. [32] W.J. Hurkman, C.K. Tanaka, Solubilization of plant membrane proteins for analysis by two-dimensional gel electrophoresis, Plant Physiol. 81 (1986) 802–806. [33] C. Mattos, Proteins in organic solvents, Curr. Opin. Struct. Biol. 11 (2001) 761–764. [34] A. Shevchenko, M. Wilm, O. Vorm, M. Mann, Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels, Anal. Chem. 68 (1996) 850–858. [35] S.P. Kowalski, N.T. Eannetta, A.T. Hirzel, J.C. Steffens, Purification and characterization of polyphenol oxidase from glandular trichomes of Solanum berthaultii, Plant Physiol. 100 (1992) 677–684. [36] K.A. Resing, N.G. Ahn, Proteomics strategies for protein identification, FEBS Lett. 579 (2005) 885–889. [37] S.A. Ross, Z. Mehmedic, T.P. Murphy, M.A. Elsohly, GC-MS analysis of the total delta9-THC content of both drug- and fiber-type cannabis seeds, J. Anal. Toxicol. 24 (2000) 715–717.

A

CC

EP

TE D

M

A

N

U

SC R

IP T

[38] M. Tayyab, D. Shahwar, GCMS analysis of Cannabis sativa L. from four different areas of Pakistan, Egypt. J. Forensic Sci. 5 (2015) 114–125. [39] P. Goettig, Effects of glycosylation on the enzymatic activity and mechanisms of proteases, Int. J. Mol. Sci. 17 (2016) 1969. [40] A.M. Klibanov, Improving enzymes by using them in organic solvents, Nature. 409 (2001) 241–246. [41] B. Zirpel, F. Stehle, O. Kayser, Production of Δ9-tetrahydrocannabinolic acid from cannabigerolic acid by whole cells of Pichia (Komagataella) pastoris expressing Δ9tetrahydrocannabinolic acid synthase from Cannabis sativa L., Biotechnol. Lett. 37 (2015) 1869–1875. [42] F. Taura, E. Dono, S. Sirikantaramas, K. Yoshimura, Y. Shoyama, S. Morimoto, Production of Δ1-tetrahydrocannabinolic acid by the biosynthetic enzyme secreted from transgenic Pichia pastoris, Biochem. Biophys. Res. Commun. 361 (2007) 675–680. [43] O. Aizpurua-Olaizola, J. Omar, P. Navarro, M. Olivares, N. Etxebarria, A. Usobiaga, Identification and quantification of cannabinoids in Cannabis sativa L. plants by high performance liquid chromatography-mass spectrometry, Anal. Bioanal. Chem. 406 (2014) 7549–7560. [44] D. Pacifico, F. Miselli, A. Carboni, A. Moschella, G. Mandolino, Time course of cannabinoid accumulation and chemotype development during the growth of Cannabis sativa L, Euphytica. 160 (2008) 231–240. [45] H. van Bakel, J.M. Stout, A.G. Cote, C.M. Tallon, A.G. Sharpe, T.R. Hughes, J.E. Page, The draft genome and transcriptome of Cannabis sativa, Genome Biol. 12 (2011) R102. [46] Y. Shoyama, T. Tamada, K. Kurihara, A. Takeuchi, F. Taura, S. Arai, M. Blaber, Y. Shoyama, S. Morimoto, R. Kuroki, Structure and function of ∆1-tetrahydrocannabinolic acid (THCA) synthase, the enzyme controlling the psychoactivity of Cannabis sativa, J. Mol. Biol. 423 (2012) 96–105. [47] K. Lange, A. Schmid, M.K. Julsing, Δ9-Tetrahydrocannabinolic acid synthase production in Pichia pastoris enables chemical synthesis of cannabinoids, J. Biotechnol. 211 (2015) 68–76. [48] K. Lange, A. Schmid, M.K. Julsing, Δ9-Tetrahydrocannabinolic acid synthase: The application of a plant secondary metabolite enzyme in biocatalytic chemical synthesis, J. Biotechnol. 233 (2016) 42–48. [49] M.J. Liszka, M.E. Clark, E. Schneider, D.S. Clark, Nature versus nurture: Developing enzymes that function under extreme conditions, Annu. Rev. Chem. Biomol. Eng. 3 (2012) 77–102. [50] M.N. Gupta, Enzyme function in organic solvents, Eur. J. Biochem. 203 (1992) 25–32. [51] J.S. Dordick, Enzymatic catalysis in monophasic organic solvents, Enzyme Microb. Technol. 11 (1989) 194–211. [52] R.Z. Kazandjian, A.M. Klibanov, Regioselective oxidation of phenols catalyzed by polyphenol oxidase in chloroform, J. Am. Chem. Soc. 107 (1985) 5448–5450. [53] N.L. Klyachko, A.V. Levashov, Bioorganic synthesis in reverse micelles and related systems, Curr. Opin. Colloid Interface Sci. 8 (2003) 179–186. [54] K. Martinek, A. Levashov, Y. Khmelnitsky, N. Klyachko, I. Berezin, Colloidal solution of water in organic solvents: a microheterogeneous medium for enzymatic reactions, Science. 218 (1982) 889–891. [55] B. Daniel, B. Konrad, M. Toplak, M. Lahham, J. Messenlehner, A. Winkler, P. Macheroux, The family of berberine bridge enzyme-like enzymes: A treasure-trove of oxidative reactions, Arch. Biochem. Biophys. 632 (2017) 88–103. [56] C.J. Carter, R.W. Thornburg, Tobacco nectarin V is a flavin-containing berberine bridge enzyme-like protein with glucose oxidase activity, Plant Physiol. 134 (2004) 460–469.

A

CC

EP

TE D

M

A

N

U

SC R

IP T

[57] K. Tiedge, G. Lohaus, Nectar sugars and amino acids in day- and night-flowering Nicotiana species are more strongly shaped by pollinators’ preferences than organic acids and inorganic ions, PLOS ONE. 12 (2017) e0176865. [58] Y.H. Choi, J. van Spronsen, Y. Dai, M. Verberne, F. Hollmann, I.W.C.E. Arends, G.-J. Witkamp, R. Verpoorte, Are natural deep eutectic solvents the missing link in understanding cellular metabolism and physiology?, Plant Physiol. 156 (2011) 1701– 1705. [59] M.S. Hillwig, C. Kanobe, R.W. Thornburg, G.C. MacIntosh, Identification of S-RNase and peroxidase in petunia nectar, J. Plant Physiol. 168 (2011) 734–738. [60] M. Nepi, L. Bini, L. Bianchi, M. Puglia, M. Abate, G. Cai, Xylan-degrading enzymes in male and female flower nectar of Cucurbita pepo, Ann. Bot. 108 (2011) 521–527. [61] M.B. Burg, J.D. Ferraris, Intracellular organic osmolytes: function and regulation, J. Biol. Chem. 283 (2008) 7309–7313. [62] L.K. Shrestha, T. Sato, D. Varade, K. Aramaki, Effect of polyol on the structure of nonionic surfactant reverse micelles in glycerol monoisostearate/decane systems, Langmuir ACS J. Surf. Colloids. 26 (2010) 3115–3120. [63] Y. Huang, Y. Wang, L. Sun, R. Agrawal, M. Zhang, Sundew adhesive: a naturally occurring hydrogel, J. R. Soc. Interface. 12 (2015) 20150226. [64] T. Kokubun, Occurrence of myo-inositol and alkyl-substituted polysaccharide in the prey-trapping mucilage of Drosera capensis, Naturwissenschaften. 104 (2017) 83–83. [65] L.E. Muravnik, O.V. Kostina, A.L. Shavarda, Glandular trichomes of Tussilago farfara (Senecioneae, Asteraceae), Planta. 244 (2016) 737–752. [66] G. Serrato-Valenti, Structural and histochemical investigation of the glandular trichomes of Salvia aurea L. leaves, and chemical analysis of the essential oil, Ann. Bot. 79 (1997) 329–336. [67] M. Marin, V. Koko, S. Duletić-Laušević, P.D. Marin, D. Rančić, Z. Dajic-Stevanovic, Glandular trichomes on the leaves of Rosmarinus officinalis: Morphology, stereology and histochemistry, South Afr. J. Bot. 72 (2006) 378–382. [68] M. Marin, S. Budimir, D. Janosevic, P.D. Marin, S. Duletic-Lausevic, M. LjaljevicGrbic, Morphology, distribution, and histochemistry of trichomes of Thymus lykae Degen & Jav. (Lamiaceae), Arch. Biol. Sci. 60 (2008) 667–672. [69] J.D. Ryan, P. Gregory, W.M. Tingey, Phenolic oxidase activities in glandular trichomes of Solanum berthaultii, Phytochemistry. 21 (1982) 1885–1887. [70] H. Yu, S.P. Kowalski, J.C. Steffens, Comparison of polyphenol oxidase expression in glandular trichomes of Solanum and Lycopersicon species, Plant Physiol. 100 (1992) 1885–1890. [71] S.P. Slocombe, I. Schauvinhold, R.P. McQuinn, K. Besser, N.A. Welsby, A. Harper, N. Aziz, Y. Li, T.R. Larson, J. Giovannoni, R.A. Dixon, P. Broun, Transcriptomic and reverse genetic analyses of branched-chain fatty acid and acyl sugar production in Solanum pennellii and Nicotiana benthamiana, Plant Physiol. 148 (2008) 1830–1846. [72] M.M. Oliveira, M. Salomk, S. Pais, Glandular trichomes of Humulus lupulus var. Brewer’s Gold: Ontogeny and histochemical characterization of the secretion, Nord. J. Bot. 8 (1988) 349–359. [73] L. Ascensão, N. Marques, M.S. Pais, Peltate glandular trichomes of Leonotis leonurus leaves: ultrastructure and histochemical characterization of secretions, Int. J. Plant Sci. 158 (1997) 249–258. [74] E.S. Kim, P.G. Mahlberg, Secretory vesicle formation in the secretory cavity of glandular trichomes of Cannabis sativa L. (Cannabaceae), Mol. Cells. 15 (2003) 387– 395.

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Figure legends Figure 1. Capitate-stalked glandular trichome at 7th week of the flowering stage from the drug-type Cannabis (Euphoria) strain before (left) and after microsuction of the exudates (right). The applied technique enabled precise sampling of the exudates from storage cavities

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without disrupting the secretory cells.

Figure 2. Coomassie-stained electrophoretic separations of proteins extracted from the exudates collected from 800 glandular trichome storage cavities from Euphoria and Finola Cannabis plants. Protein bands indicated with numbers were identified as THCAS (1); and

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CBDAS (2). The numbering on the gels corresponds to Tab. 1.

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Figure 3. Western blot analysis of proteins extracted from the exudates collected from 1000

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storage cavities from the drug-type Cannabis (Euphoria) (A). Western blot analysis of

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proteins extracted from the whole glandular trichomes from Euphoria strain (B) and corresponding Coomassie-stained SDS gel prepared in parallel (C). The arrows with numbers

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(4-7) indicate protein bands which corresponded to detected protein signals on western blots. Protein bands (3-7) were processed for identification: 3, 4 - THCAS, 5-7 - olivetolic acid cyclase. The numbering on the gel corresponds to Tab. 1. Western blot analysis was carried out using polyclonal antibody specific for THCAS, although some unspecific binding to

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olivetolic acid cyclase was also observed.

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Figure 4. Accumulation profile of CBDAS and THCAS in the exudates of glandular

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trichomes isolated from the drug-type Cannabis (Euphoria) at the 5th and 8th week of the flowering period; n = 3; values represent mean ± SD. The proteins were quantified on the basis of identified unique peptides. The extracted ion chromatograms (XIC) of the analyzed

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samples and peak integration used for protein quantification for THCAS and CBDAS are

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shown in the Fig. S2 and S3, respectively.

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Figure 5. Total ion chromatograms showing peaks identified as CBD (1) and THC (2) in the

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exudates isolated from the drug-type Cannabis (Euphoria) glandular trichome storage cavities after incubation in hexane without CBGA (A), and with CBGA (B); negative control (C). The

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normalized intensity of THC and CBD in control samples and after incubation with CBGA (D); n = 3; values represent mean ± SD. Intensity was normalized (NL) to base peak. Total ion chromatograms in full range, and observed fragment ions for THC and CBD are shown in

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the Fig. S5 and S6, respectively.

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Figure 6. Metabolite classes identified in hexane and derivatized extracts of the exudates

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isolated from the drug-type Cannabis (Euphoria), and their contribution to total ion current. The total number of identified metabolites in each class is given in brackets. The lists of the

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respectively.

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metabolites identified in hexane and derivatized extracts are shown in the Tab. S3 and S4,

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Tables Table 1. Proteins identified from the in-gel digested samples prepared from Cannabis plants; Euphoria (bands 1, 3-7) and Finola (band 2). Seq. cov. [%]

Unique peptides

Peptides

PSMb

MW [kDa]

calc. pI

Accession

1121

34

10

20

52

61,9

8,95

Q8GTB6

2957

51

19

21

125

62,2

8,72

A6P6V9

848

33

6

10

24

61,9

8,95

Q8GTB6

657

22

4

10

24

61,9

102

22

2

2

3

12

102

37

3

3

4

12

7 Olivetolic acid cyclase 208 43 a Numbering refers to Fig. 2 and 3. b PSM - total number of identified peptide spectra matched for the protein.

4

4

9

12

Organism

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Tetrahydrocannabinolic acid synthase Cannabidiolic acid synthase Tetrahydrocannabinolic acid synthase Tetrahydrocannabinolic acid synthase Olivetolic acid cyclase

6

Olivetolic acid cyclase

1 2 3

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4

Cannabis sativa

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Score

Description

8,95

Q8GTB6

6,16

I6WU39

6,16

I6WU39

6,16

I6WU39

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Band no.a