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Focus on diterpene biosynthesis
Diterpene biosynthesis usually occurs in chloroplasts, where most GGPP precursors are produced (Tholl, 2015). The various diterpene biosynthesis pathways are believed to have evolved from gibberellin biosynthesis (Peters, 2010). A majority of diterpenes, including sclareol, are labdane-type diterpenes. Formation of diterpenes with a hydrocarbon skeleton of the labdane type usually involves the cyclization of GGPP through the consecutive action of a class II diterpene synthase (Figure 10) and a class I diterpene synthase (Peters, 2010). The obtained bicyclic skeleton can be further modified, for example by the addition of hydroxyl groups mediated by diterpene synthases or cytochrome P450 monoxygenases (Zerbe and Bohlmann, 2015; Bathe and Tissier, 2019). Another cyclization reaction leads to tricyclic abietane diterpenes (Peters, 2010) (Figure 10).
Factors affecting terpene content
Specialized metabolite content varies at all scales, between species and populations, but also between individuals and between the different parts of the same plant (Moore et al., 2014). In a given species, populations displaying distinct metabolite profiles are referred to as chemotypes (Moore et al., 2014). Physiological variations, environmental conditions and genetic polymorphisms, along with the interactions of these three factors, are known to determine specialized metabolite content in plants (Figueiredo et al., 2008; Moore et al., 2014).
In a single plant, terpenoid production depends on the organ considered, on the developmental stage of this organ, and also on circadian and seasonal cycles (Moore et al., 2014). Important qualitative and quantitative variations are observed between the different organs of the plant. For example, the flowers of Lavandula pinnata produce more monoterpenes and less sesquiterpenes than leaves and stems. These differences are related to the important role played by volatile monoterpenoids in guiding pollinators to the flowers (Figueiredo et al., 2008). In Achillea millefolium, the amount of volatile terpenoids produced by flowers differs according to their developmental stage. The amount of sesquiterpene chamazulene present in the essential oil decreases with flower maturation, while the proportion of the monoterpenes 1,8-cineole and camphor increases (Figueiredo et al., 2008). Diurnal variations of light level impact MEP and MVA pathways in the opposite way: the MEP pathway is stimulated by exposure to light, whereas the MVA pathway is upregulated in the dark (Tholl, 2015). Seasonal variations in terpenoid content can also be observed in some species. In sea fennel (Crithmum maritimum), proportions of sabinene and γ-terpinene in the essential oil vary in opposite ways: sabinene is more abundant during the flowering season (between July and October), whereas γ-terpinene predominates during the rest of the year and peaks in April (Barroso et al., 1992) (Figure 11).
Molecular mechanisms of terpene biosynthesis regulation
Phytohormone signaling pathways and master transcriptional regulators of plant growth and development control terpenoid biosynthesis in response to environmental and developmental signals. In Arabidopsis thaliana, the upregulation of the MEP pathway and carotenoid biosynthesis in light conditions was shown to be performed through the action of phytochrome interacting factors (Toledo-Ortiz et al., 2010) and the biosynthesis of floral sesquiterpenes is induced by master regulators of flower maturation in response to jasmonates, auxin and gibberellins (Tholl, 2015). Jasmonates stimulate terpenoid biosynthesis in many other plants (Goossens et al., 2016); for example, they induce artemisinin biosynthesis in Artemisia annua (Chen et al., 2017b). Genes involved in the same terpene biosynthesis pathway are often organized as gene clusters in plant genomes, an organization which facilitates their transcriptional coregulation (Töpfer et al., 2017). Positive transcriptional regulators of terpene biosynthesis genes have been identified in several plant species and generally belong to WRKY, AP2/ERF, bHLH or basic leucine zipper (bZIP) transcription factor families (Lu et al., 2016). For example, AaWRKY1, AaERF1, AaERF2, AaORA1 and AabZIP1 transcription factors positively regulate the biosynthesis of the sesquiterpene artemisinin in Artemisia annua (Yu et al., 2011; Lu et al., 2013; Han et al., 2014; Zhang et al., 2015a), TcWRKY1 stimulates the production of the diterpene paclitaxel in Taxus chinensis (Li et al., 2013) and the bZIP transcription factor OsTGAP1 positively regulates the biosynthesis of diterpene phytoalexins in rice (Okada et al., 2009).
In plants, MVA and MEP pathways are compartmentalized. Therefore, they can be differentially regulated, and controlled regulatory crosstalks between terpenoid biosynthesis pathways have been reported (Tholl, 2015). Notably, terpenoid biosynthesis has been shown to be regulated at the post-translational level by protein prenylation (Liao et al., 2016). For instance, the production of the MVA-derived sesquiterpene capsidiol in tobacco relies on MEP-dependent protein geranylgeranylation (Huchelmann et al., 2014) and in Catharanthus roseus, MVA-derived farnesylated proteins regulate the expression of genes involved in MEP-dependent monoterpenoid biosynthesis (Courdavault et al., 2005).
Capitate and peltate glandular trichomes of the Lamiaceae
In Angiosperms, terpenoid production is often localized in specialized secretory structures. For example, in Hevea brasiliensis, polyisoprene components of natural rubber are produced in specialized ducts known as laticifers (Lange, 2015). Lamiaceae are generally characterized by the presence of epidermal secretory structures called glandular trichomes, which produce various compounds mainly involved in pollinator attraction or defense against herbivores (Werker, 1993). Terpenoids secreted by glandular trichomes are major components of the essential oil which can be extracted by hydrodistillation. Glandular trichomes of the Lamiaceae are multicellular and composed of 3 parts: a base, a stalk and a gland (or glandular head) (Figure 12). The gland is responsible for the secretion of specialized metabolites, the stalk is the structure bearing the gland, and the base connects the stalk to surrounding epidermal cells (Lange, 2015). Both vegetative and reproductive organs of the Lamiaceae harbor glandular trichomes already at early development stages (Werker, 1993).
Essential oil production in aerial parts
So far, terpene production by clary sage and its regulation have mainly been investigated through the study of essential oil yield and composition. Although clary sage oil chemical composition shows important variations between studies, linalool and its acetylated derivative linalyl acetate are generally reported as being its main constituents (Lattoo et al., 2006; Schmiderer et al., 2008; Wagner et al., 2012; Sharopov and Setzer, 2012; Nasermoadeli and Rowshan, 2013). The composition of clary sage essential oils produced in different countries of the Mediterranean basin have been compared and chemotypes have been identified according to the proportion of each compound in the essential oil (Sharopov and Setzer, 2012). Variations observed between clary sage chemotypes can probably be attributed to natural genetic diversity and to the influence of growth conditions. The impact of environmental conditions is illustrated by a study demonstrating that clary sage essential oil production is affected by shade level (Kumar et al., 2013). Shade level increases when the distance between plants decreases; in the study, it was artificially fixed using green shading nets. Essential oil yield was lower in shade conditions compared to open-sun conditions, and oil composition varied upon shade level: linalool content was highest in open-sun conditions and decreased with shade, whereas the opposite was observed for the sesquiterpene germacrene D (Kumar et al., 2013).
A clary sage glabrous mutant produces less sclareol and linalyl acetate
A clary sage “glabrous” line was identified in a collection of clary sage mutant lines obtained by EMS-generated random mutagenesis (Drevensek et al., unpublished data) (Figure 23a). According to careful numbering of different trichome types (Drevensek et al., unpublished data), the “glabrous” aspect of calyces from these plants is essentially due to a drastic reduction of the number of large capitate glandular trichomes, while the number of peltate and small capitate glandular trichomes remains unchanged (Figure 23b). Metabolite quantification by GC-MS indicates that sclareol and linalyl acetate content is strongly reduced in glabrous calyces compared to wild-type calyces (Figure 23c-d). This lower metabolite content is likely due to the nearly absence of large capitate glandular trichomes, an idea reinforcing the hypothesis that capitate glandular trichomes are primarily responsible for calyx sclareol production. However, distinct EMS-induced point mutations could be the cause of the decrease in metabolite content and the reduced density of capitate glandular trichomes, respectively. The analysis of the co-segregation of these two phenotypes and the cloning of their causal mutations should shed light on this question and lead to insights in the understanding of glandular trichome development and sclareol production in clary sage.
Sclareol is mainly produced by glandular trichomes
Glandular trichomes of the Lamiaceae are known to be specialized in the production of essential oil terpenes, mainly mono- and sesquiterpenes (Werker, 1993; Lange and Turner, 2013). In Nicotiana species, glandular trichomes produce diterpenes, for example sclareol in Nicotiana glutinosa (Guo and Wagner, 1995). These elements led the scientific community to assume that sclareol accumulated in clary sage calyces is produced by glandular trichomes. We provide here several lines of evidence reinforcing this hypothesis. First, spots of [linalyl acetate-K]+ and [(sclareol)2-K]+ ions were detected on calyx samples by mass spectrometry imaging and coincide with capitate glandular trichomes (Figure 19 and 20). [Sclareol-K]+ ions were also detected on the rest of the epidermis (Figure 19 and 20). However, [(sclareol)2-K]+ ions probably form only where sclareol is most abundant and may thus indicate the main sites of sclareol production. Secondly, a clary sage line with fewer capitate glandular trichomes produces less linalyl acetate and sclareol (Figure 23). Due to the presence of thousands of EMS-induced point mutations in this line, the two phenotypes may be caused by distinct mutations, but we can suppose that they have the same causal mutation. Since peltate and small capitate trichomes are unchanged compared to the wild-type (Figure 23b), this mutation may be located in a gene encoding a transcription factor specifically inducing large capitate trichome development. Taken together, these results support the idea that glandular trichomes are the main site of sclareol production. This theory is further underpinned by gene expression analyses performed on isolated glandular trichomes, showing that sclareol-synthase and LPP-synthase genes are more expressed in isolated glandular heads compared to the rest of the calyx (Drevensek et al., unpublished data) (Figure 24). Sclareol present on the major part of calyx epidermis, detected in the form of [sclareol-K]+ ions, could initially be produced in glandular heads and spread over pavement cells along with the rest of glandular head content. This process of accumulation was shown to occur in the case of the diterpene duvatrienediol in Nicotiana tabacum (Chang and Grunwald, 1980; Keene and Wagner, 1985). Tobacco capitate glandular trichomes are known to regularly exude their content (Tissier, 2012) and our scanning electron microscopy images highlight the presence of glandular head content dripping down clary sage trichome stalks (Figure 21).
Table of contents :
Table of contents
Résumé en français
1. Introduction
2. Résultats
2.1. Localisation de la biosynthèse du sclaréol
2.2. Identification de l’origine métabolique du sclaréol par marquage isotopique
2.3. La biosynthèse du sclaréol est-elle régulée par les jasmonates ?
2.4. Etude de la diversité naturelle de la sauge sclarée
3. Revue de la littérature
4. Perspectives
Introduction
1. Ambergris and sclareol
1.1. The search for substitutes to ambergris
1.2. Interest in increasing sclareol global production
2. Clary sage
2.1. Botanical description of clary sage
2.2. Sclareol biological function in clary sage
2.3. Clary sage commercial use
2.4. First advances in clary sage genetics and biotechnology
2.5. Clary sage breeding in France
3. Terpene biosynthesis and function in plants
3.1. Specialized metabolism and terpenoids
3.2. The diverse biological functions of terpenes
3.3. Industrial exploitation of terpenes
3.4. Terpene biosynthesis
3.5. Terpene biosynthesis regulation
4. Glandular trichomes
4.1. Capitate and peltate glandular trichomes of the Lamiaceae
4.2. Clary sage glandular trichomes
4.3. Genetic control of glandular trichome development
5. Terpene production by clary sage
5.1. Essential oil production in aerial parts
5.2. Sclareol biosynthesis and secretion
5.3. Terpene production in roots
6. Aim of the project
Chapter 1 Mass spectrometry imaging to localize sclareol biosynthesis
1. Introduction
2. Materials and methods
2.1. Plant material
2.2. Scanning electron microscopy
2.3. Metabolite localization by LDI-FT-ICR
2.4. Sclareol and linalyl acetate quantification by GC-MS
3. Results
3.1. Different accumulation patterns for sclareol and linalyl acetate
3.2. Distinct roles of different trichome types in sclareol production
3.3. A clary sage glabrous mutant produces less sclareol and linalyl acetate
4. Discussion
4.1. Sclareol is mainly produced by glandular trichomes
4.2. Some sclareol could also be produced by other epidermal cells
4.3. Engineering higher glandular trichome density to increase sclareol yield
Chapter 2 A retro-biosynthetic approach to decipher sclareol metabolic origin
1. Introduction
1.1. Metabolic crosstalk between MVA and MEP pathways
1.2. Towards clary sage metabolic engineering
2. Materials and methods
2.1. Plant material
2.2. Treatment with MVA or MEP pathway-specific inhibitors
2.3. Sclareol and linalyl acetate quantification by GC-MS
2.4. Isotope labeling and analysis by 13C-NMR or GC-MS
3. Results
3.1. Sclareol and linalyl acetate are highly abundant terpenes of clary sage calyx surface
3.2. Sclareol and linalyl acetate are MEP-derived terpenes
3.3. Comparison of results obtained for sclareol and linalyl acetate
3.4. The case of the acetate group of linalyl acetate
3.5. The mixed origin of the sesquiterpene β-caryophyllene
4. Discussion
4.1. Biosynthetic origin of mono-, sesqui- and diterpenes in clary sage and other plants
4.2. Metabolic engineering of terpene production in plants
4.3. Perspectives for clary sage metabolic engineering
Chapter 3 Is sclareol biosynthesis regulated by jasmonates?
1. Introduction
1.1. Regulation of plant specialized metabolism by jasmonates
1.2. How to use jasmonate signaling to enhance the production of compounds of interest
1.3. Regulation of clary sage specialized metabolism by jasmonates: state of the art
2. Materials and methods
2.1. Calyx development stages
2.2. MeJA treatment by feeding
2.3. MeJA treatment by spraying
2.4. Sclareol and linalyl acetate quantification by GC-MS
3. Results
3.1. Regulation of sclareol biosynthesis during calyx development
3.2. MeJA treatment on flowers by feeding
3.3. MeJA treatment on flowers by spraying
4. Discussion
4.1. MeJA apparently does not impact sclareol production, but further protocol optimization is needed
4.2. Other potential ways to manipulate sclareol biosynthesis regulation
Chapter 4 Clary sage natural diversity
1. Introduction
1.1. The use of natural genetic diversity for breeding
1.2. Impact of environmental conditions on terpene production and genetic diversity
2. Materials and methods
2.1. Plant material
2.2. Genetic diversity analysis
2.3. Field trial
2.4. Sclareol and linalyl acetate quantification by GC-MS
3. Results
3.1. Croatian populations show genetic polymorphism in MEP pathway genes
3.2. Choice of the populations to be analyzed in the field
3.3. Comparison of flowering date and metabolite content of the different populations
4. Discussion
4.1. Complex relationships between genetic diversity, chemical diversity and environmental conditions
4.2. Perspectives for the study of clary sage natural diversity
4.3. Vatican White plants flower earlier and produce less specialized metabolites
Conclusion
1. Context and objectives of the project
2. Perspectives for research on sclareol production in clary sage
3. Perspectives for clary sage genetic improvement
Annex
Review: Genetic control of glandular trichome development
1. Introduction
1.1. Why study the genetic control of glandular trichome development?
1.2. Which species could serve as model(s)?
2. Description of glandular trichome morphology and development
3. Genes controlling glandular trichome initiation
3.1. Transcription factors
3.2. Cyclins
3.3. Regulatory complexes
3.4. Genes involved in hormonal signaling
4. Genes controlling later steps of glandular trichome development
4.1. Cytoskeleton regulators
4.2. Cuticle deposition regulators
5. Evolution of glandular trichome development regulators
5.1. Evidence supporting the conservation of glandular trichome development regulators in the Solanaceae
5.2. Some regulators appear to be conserved in distant plant families, while the others are likely to have evolved independently
5.3. Different scenarios for the evolution of glandular trichome development regulators
6. Perspectives
References
