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University of Calgary PRISM: University of Calgary's Digital Repository Graduate Studies The Vault: Electronic Theses and Dissertations 2012-07-19 Molecular cloning and characterization of sesquiterpene synthases from valeriana officinalis Pyle, Bryan Wilkinson Pyle, B. W. (2012). Molecular cloning and characterization of sesquiterpene synthases from valeriana officinalis (Unpublished master's thesis). University of Calgary, Calgary, AB. doi:10.11575/PRISM/26983 http://hdl.handle.net/11023/129 master thesis University of Calgary graduate students retain copyright ownership and moral rights for their thesis. You may use this material in any way that is permitted by the Copyright Act or through licensing that has been assigned to the document. For uses that are not allowable under copyright legislation or licensing, you are required to seek permission. Downloaded from PRISM: https://prism.ucalgary.ca

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Molecular cloning and characterization of sesquiterpene synthases from valeriana officinalisGraduate Studies The Vault: Electronic Theses and Dissertations
2012-07-19
Pyle, Bryan Wilkinson
Pyle, B. W. (2012). Molecular cloning and characterization of sesquiterpene synthases from
valeriana officinalis (Unpublished master's thesis). University of Calgary, Calgary, AB.
doi:10.11575/PRISM/26983
http://hdl.handle.net/11023/129
University of Calgary graduate students retain copyright ownership and moral rights for their
thesis. You may use this material in any way that is permitted by the Copyright Act or through
licensing that has been assigned to the document. For uses that are not allowable under
copyright legislation or licensing, you are required to seek permission.
Downloaded from PRISM: https://prism.ucalgary.ca
by
A THESIS
IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE
DEGREE OF MASTER OF SCIENCE
DEPARTMENT OF BIOLOGICAL SCIENCES
ii
Abstract
Valeriana officinalis (valerian) is a popular medicinal plant in North America and
Europe. Its root extract is commonly used as a mild sedative and anxiolytic. Valerenic acid, a
C15 sesquiterpenoid, has been suggested as the active ingredient responsible for the sedative
effect. Recently, medical uses of valerenic acid as anti-depressant and anti-inflammatory drugs
were suggested due to its affinity for the γ-aminobutyric acid type A (GABAA) receptor as an
agonist and its inhibition of the nuclear factor kappa-light-chain-enhancer of activated B cells
(NF-B) pathway, respectively. Despite its importance, biochemistry of valerenic acid in
valerian remains unknown. To identify the first committed enzymatic step in valerenic acid
biosynthesis, next-generation sequencing (Roche 454 titanium) was used to generate ~1 million
transcript reads from valerian root. Subsequently, three cDNAs for sesquiterpene synthases
(VoTPS1/2/3) were identified and their corresponding recombinant enzymes were purified.
Three recombinant enzymes efficiently catalyze the synthesis of valerena-4,7(11)-diene,
germacrene C/D, and drimenol, respectively, based on the spectral match in the mass
spectrometry library. Additional structural analyses using GC-MS and 13
C-NMR spectrometry
in comparison to a semi-synthesized standard confirmed the chemical identity of valerena-
4,7(11)-diene. This is the first report of valerena-4,7(11)-diene and drimenol synthases, and the
biosynthetic mechanisms of these two products from the substrate, farnesyl diphosphate, were
proposed.
iii
Acknowledgements
I would like to thank Dr. Dae-Kyun Ro for introducing me to the world of plant
metabolites. Before I started this project I had little to no understanding of the vast complexity
and unfathomable quantity of compounds produced by plants. I can now say I comprehend that
number a little more. I must also thank Dr. Ro for pushing my own expectations of myself, for
that I owe you a great debt, the skills you have given me will help me in every decision I ever
make, from now on. To all members of the Ro lab, past and present, thank you for all your help.
Dr. Hue Tran, I thank you for teaching me the basics of protein purification. I must also thank
Dr. Benjamin Pickel for our extensive discussions on women, science and beer without which
very few men can survive science. I must also thank Dr. Pickel for valerenadiene purification
and NMR analysis. Thank you to Drs. John Vederas and Zhizeng Gao for valerenadiene
chemical synthesis. Thank you to Gillian MacNevin for the semi-quantitative PCR data. Finally
I would like to thank Dr. Paul O’Maille for the pH9GW vector, a generous gift.
Last and definitely not least I must thank my family and friends. All members of my
family have helped me in some way, shape or form throughout my life and that is priceless. To
my wife Lisa, thank you, for you have contributed so much emotionally to these past two years
and I will forever be indebted to you. Our daily walks with Bodie gave me an outlet to escape
from my second love, science. You are my best friend and I am sorry for being a “difficult” grad
student for the past two years.
iv
Dedication
To my late grandfather, Byron W. Pyle, though our religious philosophies did not always agree
our educational philosophies did; everyone deserves an opportunity at education…wherever that
may go.
1.2.2 Terpenoid Biosynthesis in Plants ................................................................................... 8
1.3 Terpene Synthases .............................................................................................................. 15
1.3.2 Phylogenetic Relationships of Terpene Synthases ....................................................... 21
1.4 Metabolic Engineering of the MVA Pathway .................................................................... 22
1.5 Ligand-Receptor Binding.................................................................................................... 25
1.7 Objectives ........................................................................................................................... 30
2.1 Plant Cultivation and Metabolite Preparations ................................................................... 32
2.2 RNA preparations ............................................................................................................... 32
2.4 Plasmid Construction for Yeast Expression ....................................................................... 33
2.5 Quantitative Transcript Analysis ........................................................................................ 34
2.6 Yeast Transformation.......................................................................................................... 35
2.8 Plasmid Construction for E. coli Expression ...................................................................... 36
2.9 Heterologous Expression Trials .......................................................................................... 39
2.10 Expression in E. coli and Protein Purification .................................................................. 39
2.11 Gas-chromatography and Mass Spectroscopy Analysis ................................................... 41
2.12 Purification and NMR of Valerena-4,7(11)-diene ............................................................ 42
2.14 NMR Analysis of Valerena-4,7(11)-diene Standard ........................................................ 42
2.15 Enzyme Activity Assays ................................................................................................... 43
2.16 Enzyme Characterization .................................................................................................. 43
2.17 Phylogenetic Analysis ....................................................................................................... 44
CHAPTER 3: RESULTS ........................................................................................................... 45
3.2 Transcript Sequencing and Candidate Gene Isolation ........................................................ 47
3.3 Functional Screening of VoTPS cDNAs in Engineered Yeast ........................................... 51
3.4 Characterization of the VoTPS2 Product ............................................................................ 57
vi
3.6 Cyclization Mechanism of Valerena-4,7(11)-diene ............................................................ 65
3.7 Identification and Characterization of an Additional Sesquiterpene Synthase, VoTPS3.... 68
3.8 Phylogenetic Analysis of VoTPS1/2/3 ............................................................................... 73
CHAPTER 4: DISCUSSION ..................................................................................................... 76
Table 1. Table of primers used in cloning experiments. ............................................................. 38
Table 2. GC-MS analysis of terpenoids synthesized from VoTPS1, VoTPS2 and VoTPS3 ...... 54
Table 3. Comparison of the 13
C-NMR signals from the purified compound of peak 4 with the
published data. .............................................................................................................................. 58
Figure 2. Chemical structures of isoprene and isopentenyl diphosphate. ..................................... 5
Figure 3. Schematic depiction of the MVA pathway. ................................................................. 13
Figure 4. Schematic depiction of the DXP pathway. .................................................................. 14
Figure 5. Schematic diagram representing the carbocation mechanism of tobacco epi-
aristolochene synthase (TEAS). .................................................................................................... 20
Figure 6. Proposed biosynthetic pathway for valerenic acid production in V. officinalis. ......... 31
Figure 7. GC-MS profile of volatile metabolites from valerian root. .......................................... 46
Figure 8. Sequence alignment of deduced amino acid sequences from VoTPS1 and VoTPS2. ... 49
Figure 9. Semi-quantitative RT-PCR analysis of the VoTPS1 and VoTPS2 transcripts in V.
officinalis root and leaf. ................................................................................................................ 50
Figure 10. Unique terpene compounds synthesized from the yeast expressing VoTPS1 or
VoTPS2. ........................................................................................................................................ 53
Figure 11. Chemical structures relating to numbers from text. .................................................. 55
Figure 12. GC-MS analysis of VoTPS1 products and the terpene standards synthesized by
tomato germacrene B/C synthase. ................................................................................................. 56
Figure 13. Validation of VoTPS2 enzyme product (peak 4) as valerena-4,7(11)-diene. ............ 59
Figure 14. Expression trials of his-tagged recombinant VoTPS1 and VoTPS2. ........................ 62
Figure 15. Purification of VoTPS1/2 by Ni-NTA column using a gradient elution. .................. 63
Figure 16. In vitro enzyme assays of VoTPS1 and VoTPS2 recombinant enzyme..................... 64
Figure 17. A proposed mechanism for valerena-4,7(11)-diene formation catalyzed by VoTPS2
(valerenadiene synthase). .............................................................................................................. 67
ix
Figure 18. Expression trial of his-tagged recombinant VoTPS3 (67 kDa). ................................ 70
Figure 19. In vitro assays for VoTPS3. ....................................................................................... 71
Figure 20. A proposed mechanism of drimenol formation by VoTPS3 (drimenol synthase). ... 72
Figure 21. A phylogenetic tree representing the seven subfamilies (a-g) of terpene synthase
enzymes......................................................................................................................................... 74
Galactose
ATPase
Secondary (or specialized) metabolites encompass a vast number of low-molecular-
weight organic compounds naturally synthesized in plants and microbes. The canonical
definition describes secondary metabolites as any compound that contributes no apparent benefit
to the host organism’s growth and reproduction. However, the specialized metabolites enhance
the fitness of synthesizers in distinct ecological niches, and therefore play a central role in
evolutionary selection of plants and microbes. In contrast, primary metabolites are essential for
day-to-day function of all organisms and are normally present at higher levels. As mutations in
genes involved in primary metabolism cause fatal effects on the survival of organisms, variation
in primary metabolism is restricted and is highly conserved across the kingdoms. On the other
hand, specialized metabolism can tolerate alterations and thus display a great metabolic plasticity.
Four major classes of secondary metabolites are terpenoids, alkaloids, phenylpropanoids,
and polyketides (Figure 1). Although these compounds have a small finite metabolic role in an
individual species, many of these compounds collectively have important functions as pollinator
attractants, anti-feedants, repellents, toxins, and antibiotics. The extremely large structural
diversity of specialized metabolites makes them, in an anthropocentric view, useful to humans as
food additives, fibers, bio-polymers, pharmaceuticals, and nutraceuticals. For example, Papaver
somniferum, Valeriana officinalis, Cannabis sativa, Humulus lupulus, Atropa belladona have all
been used as medicinal plants for thousands of years. Ancient documents dating over 4,600
years old listed 1,000 plant species for possible medical uses, and most of these plants are still in
use today (Newman et al., 2000).
2
The contemporary impact of specialized metabolites in our day-to-day lives may have a
much broader influence on our society than generally realized. For example, it has been
suggested that hydrocarbon terpenes and aromatic phenolics can be developed as alternative
fuels (Zhang et al., 2011). Although the estimates of oil reserves tend to vary depending on
various factors (e.g., source of information, production, consumption, and quality), undoubtedly
its quantity is finite (Owen et al., 2010). For example, synthetic rubber manufactured from
petroleum (~3.9 million tonnes per year), for example, will ultimately need to be replaced by
natural rubber, which currently accounts for 40% of total rubber production (Cornish, 2001).
This will make substantial impacts on many manufacturing industries, such as goods, medical
devices, research, and pharmaceuticals.
Medicinal plants also impact our lives as 63% of all new chemical entities from 1981-
2006 were specialized metabolites or their semi-synthetic derivatives (Newman and Cragg,
2007). For example, three generic anti-cancer drugs currently produced by partial chemical
synthesis are vinblastine, vincristine, and paclitaxel, which were first identified from the plant
species Catharanthus roseus and Taxus brevifolia. Consequently, these compounds are
produced at minute levels by their respective plants. The Pacific yew (T. brevifolia) produces
~30 mg taxol/kg of bark in one 100-year-old tree which is equivalent to a single dosage of
treatment (Horwitz, 1994; Kirby and Keasling, 2009). Currently, paclitaxel is produced either by
semi-synthesis from a naturally more abundant intermediate, 10-deacetyl baccatin III, or from
plant cell culture, significantly reducing the cost and also protecting the environment (Horwitz,
1994; Kirby and Keasling, 2009). Similar efforts have been attempted to produce vinblastine
and vincristine in C. roseus cell cultures, but it is still very challenging to meet the 3 kg/yr
worldwide demand (Verpoorte et al., 1993; Julsing et al., 2006).
3
(terpene phenolic), proanthocyanidin (phenylpropanoid), lovastatin (polyketide).
4
Terpenoids have a long etymological and biosynthetic history. The word terpenoid,
sometimes called isoprenoid (historical term), was derived from the German word “terpentin”, or
more conspicuously known as turpentine. Turpentine refers to the essential oils of conifer
species used to investigate chemical structures in the 19 th
century (Chappell, 1995). Turpentine
oils are composed of mono-, di-, and minor amounts of sesqui-terpenes which are believed to be
used as a chemical defense against pests and pathogens in conifer trees (Zulak and Bohlmann,
2010). Terpenoids contribute to primary metabolism as sterols, photosynthetic pigments, prenyl
modification of proteins, and various hormones, but they also play critical eco-physiological
roles in plant-plant, plant-pathogen, and plant-herbivore interactions. This is largely due to the
sessile nature of plants and relates to the complex evolution of specialized metabolism. In the
past 25 years, research in the field of terpene metabolism has exploded with chemical structure
estimates reaching 65,000 (Oldfield and Lin, 2012), making terpenoids, by far, the largest and
most structurally diverse class of natural products known.
The extreme chemical diversity of terpenoids attracted scientists to elucidate the structure
of camphor, a monoterpene. Otto Wallach was able to propose the structure of camphor by
proposing the isoprene rule. The ‘isoprene rule’ establishes the C5 isoprene as structural
building blocks, which are synthesized in a head-to-tail conjugation reaction. This simple
proposal could explain why many terpenes have carbon structures following the C5 x n (n = 2, 3,
4, 6, 8) rule (Ruzicka, 1953). Leopold Ruzicka further advanced the theoretical aspect of terpene
biogenesis by defining the ‘biogenetic isoprene rule’, which is based on the unique carbocation
mechanism involving the various allylic rearrangements, hydride- and methyl-shifts, and
5
deprotonation reactions. The central biological precursor of all terpenoids was then proposed to
be isopentenyl diphosphate (IPP), and not isoprene (Figure 2) (Ruzicka, 1953).
Figure 2. Chemical structures of isoprene and isopentenyl diphosphate.
1.2.1 Ecological Functions of Isoprenoids and Terpenoids
Plants are sessile organisms. This inherent stationary nature results in complex
interactions on many different trophic levels as plants must deal with many biotic and abiotic
stressors, often concurrently. Emission of mixtures of volatile compounds from floral organs
and vegetative parts after herbivore damage, and from roots into the soil are examples of
evolutionary mechanisms that plants have developed to deal with such stressors. Plant volatiles
consist mostly of terpenoids, phenylpropanoids, benzenoids, fatty acids, and amino acid
derivatives, but terpenoids are the most diverse (Dudareva et al., 2004). Normally, volatiles are
lipophilic with high vapor pressures, and hence are able to cross membranes and diffuse through
the atmosphere or soil. Consequently, these compounds are important for plant defense and
reproduction. The simplest example is C5 isoprene synthesized by enzymatic dephosphorylation
of IPP (precursor to all terpenoids) in certain plant species. In isoprene synthesizing plants, up to
6
1-2% of the carbon fixed by photosynthesis is released to the atmosphere as a volatile gas
(Vickers et al., 2009) and most of this is in the form of isoprene ~500 Tg C/yr globally (Sharkey
and Yeh, 2001; Sasaki et al., 2007). The biological implication of such massive isoprene release
is still being debated, but some physiological experiments suggest that plants release a large
amount of isoprene in response to thermal stresses (Vickers et al., 2009). Interestingly, some
plants that have lost the ability to synthesize and emit isoprene have replaced isoprene with
mono-terpenes (Harley et al., 1997). Therefore, evolutionarily it may be reasonable to assume
that plants lacking the ability to synthesize isoprene for protection against thermal stress have
replaced isoprene with mono- and sesqui-terpenes (Vickers et al., 2009), implying a significant
evolutionary consequence for ecological function of isoprene or a terpene replacement.
Strictly speaking, research into the ecological function of isoprene (C5 units) with respect
to plants has been limited to mostly abiotic and oxidative stress (Dudareva et al., 2006; Vickers
et al., 2009). However, terpenoids (C10) are much more diverse in their ecological functions
and are implicated in many plant defense, plant-plant, and reproductive interactions. Hybrid
poplar under herbivore attack by forest tent caterpillars showed local (wound site) and systemic
emission of E--ocimene (monoterpene) in addition to several other mono- and sesquiterpenes
(Arimura et al., 2004). Other examples exist wherein plants damaged by an herbivore may
induce expression of pathways involved in production of plant defense compounds such as
jasmonic acid or ethylene (Arimura et al., 2000; Arimura et al., 2002). Plants also use terpenoids
to influence the life cycle of adjacent plants (referred to as allelopathy), as it has been shown that
emission of the monoterpene 1,8-cineole from a root can inhibit germination and growth of
competing plants (Romagni et al., 2000). Recently, belowground interactions involving the
sesquiterpene E--caryophyllene in maize was identified as the first root insect-induced
7
virgifera, but only after herbivory induced emission of E--caryophyllene by maize (Rasmann et
al., 2005). In a similar interaction, transgenic Arabidopsis thaliana engineered to produce the
sesquiterpene E--farensene prevented attack of a common aphid pest (Aharoni et al., 2003) by
mimicking the common aphid alarm pheromone (Beale et al., 2006). Examples of tobacco
species attracting herbivore predators in the wild by volatile terpenoids has also been
documented (Kessler and Baldwin, 2001).
The terpenoids and -pinene, -mycrene, and -phellandrene have been implicated in
plant reproductive fitness experiments in an orchid species, Epipactis ventrifolia (Stokl et al.,
2011). Herbivorous aphids known to feed on E. ventrifolia emit a similar mixture of terpenoids
as alarm pheromones in times of distress. Consequently, the orchid species has evolved to emit
these terpenoids from its flower as a ‘generalized mimicry’, which means that the volatile
compounds emitted do not exactly mimic the aphid alarm pheromone proportions, but mimic
only the compounds present. This generalized mimicry by the orchid attracts hoverfly females
for oviposition on the orchid. Afterward, the larvae predate herbivorous aphids, grow into
adults, and become pollinators.
Consequently, as plants have developed mechanisms to deal with herbivore and pathogen
attack, herbivores have also evolved to acquire counter solutions. For example, emission of a
volatile with the intent of attracting carnivores or perhaps as a warning signal to other plants
could inadvertently attract herbivores. Therefore, many plants have evolved to emit volatiles in a
rhythmic pattern. For example, some plants may emit volatiles to attract specific carnivores
8
which are only diurnally active, whereas certain herbivores have evolved to feed nocturnally to
avoid diurnal predators (Shiojiri et al., 2006). Finally, an example of simultaneous herbivory of
aerial and root tissues results in systemic reduction in volatile emission and can cause increased
attack by herbivorous insects on adjacent unharmed plants (Rasmann and Turlings, 2007; Soler
et al., 2007). All of these cases exemplify the dynamic nature of life and the constant
evolutionary pressures that result in specialized metabolite profiles in plants.
1.2.2 Terpenoid Biosynthesis in Plants
The discovery of IPP, a biologically active precursor of terpenoids, influenced the works
of Lynen, Bloch, Cornforth, and Popjak in establishing the metabolism of cholesterol. By
combining genetic and biochemical studies, they elucidated that the mevalonic acid (MVA)
pathway is responsible for IPP biosynthesis in both human and yeast (Bloch, 1965, 1987).
However, stable-isotope labeling patterns of IPP in bacteria did not fit the accepted prediction,
suggesting that an independent IPP pathway could be present in bacteria. Further studies
identified mevalonate-independent pathways operating in bacteria, which use pyruvate and
glyceraldehyde 3-phosphate as starting precursors. Definitive evidence for the 1-deoxy-D-
xylulose 5-phosphate (DXP) pathway was obtained from the NMR analysis of hopanoids
(cholesterol equivalent in bacteria) (Rohmer et al., 1993; Rohmer, 1999). The DXP pathway was
only fully understood in 2000, and is perhaps the last hidden metabolic pathway conserved
across various kingdoms. The eponymous DXP pathway has also been termed the non-
mevalonate, MEP (methyl erythritolphosphate pathway), or Rohmer pathway.
Through decades of work, it is now firmly established that the biosynthesis of terpenoids
occurs in almost all living organisms via two distinct metabolic pathways, the MVA and DXP
9
pathways. The MVA pathway is present in the cytosol of most eukaryotes and some
archaebacteria, but most prokaryotes do not have the MVA pathway (Rohmer, 1999; Estevez et
al., 2001). Therefore, most bacteria utilize the DXP pathway to synthesize IPP. However, plants
are the only organisms that possess both MVA and DXP pathways. The DXP pathway is present
in the plastid of the plant, and the MVA pathway in the cytosol. Since it is generally accepted
that plastids originated from bacteria by an ancient symbiotic event, presence of the DXP
pathway in plastid is not surprising. Both pathways, independent of starting materials, produce
isopentenyl diphosphate (IPP) and dimethylallyl diphosphate (DMAPP), which are key
precursors of all terpenoids. DMAPP and IPP are structural isomers of each other, and are
interchangeable by IPP isomerase. IPP isomerase converts IPP to DMAPP which acts as a
fundamental primer molecule in the synthesis of longer prenyl diphosphates, such as C10 geranyl
diphosphate, C15 farnesyl diphosphate, and C20 geranyl geranyl diphosphate. These prenyl
diphosphates are the direct biosynthetic precursors of C10 monoterpenes, C15 sesquiterpenes,
and C20 diterpenes respectively (Figures 3 and 4). Additionally, C5 hemiterpenes, C30
triterpenes, and C40 tetraterpenes can be synthesized from IPP and DMAPP.
The MVA pathway uses acetyl-CoA in the cytosol as a precursor to synthesize
cholesterol or the corresponding equivalent compounds, depending on the organisms (Figure 3).
Two carbon-carbon bonds are formed in the first two reactions of the MVA pathway by
acetoacetyl-CoA thiolase and HMG-CoA synthase, which convert two acetyl-CoA molecules to
3-hydroxy-3-methylglutaryl CoA (HMG-CoA). HMG-CoA is subsequently reduced to
mevalonate by the highly regulated HMG-CoA reductase (HMGR). Mevalonate is then
phosphorylated by two kinases and finally decarboxylated to produce IPP from mevalonate
diphosphate. IPP and its isomer DMAPP are condensed to produce various prenyl diphosphates
10
described above (Miziorko, 2011). One major metabolic fate of IPP synthesized from the MVA
pathway is cholesterol and its derivatives in animals.
The DXP pathway utilizes pyruvate and glyceraldehyde-3-phosphate (G3P) from
glycolysis (Figure 4). In the first step of the DXP pathway, pyruvate and G3P are
decarboxylated followed by two reductions and a skeletal rearrangement, catalyzing the
formation of methylerythritol phosphate (MEP). Subsequently, a cytidyl phosphate moiety is
transferred to DXP followed by phosphorylation. A unique 8-membered ring is then formed
which is facilitated by the cleavage of the nucleoside group. Ring opening and reduction are
followed by a last reduction step yielding either IPP or DMAPP. Both of the last enzymatic
steps are thought to employ a carbocation reaction (Graewert et al., 2011).
The rate-limiting enzyme for the MVA pathway is HMGR, which is competitively
inhibited by lovastatin, whereas the slowest enzyme in the DXP pathway is the reductoisomerase
(DXR), which is inhibited by fosmidomycin. Therefore, these two enzymes are the central
targets to regulate the MVA and DXP pathways. However, detailed regulation of MVA and DXP
pathways has yet to be fully understood in plants, and recent research has revealed a much more
complex feedback regulation system with multiple bottlenecks regulated at transcriptional and
post-transcriptional levels, depending on environmental and developmental cues (Rodriguez-
Concepcion, 2006).
What also seems to be unclear is the level at which metabolic crosstalk exists between the
two pathways. Based on the metabolic compartmentalization, sesquiterpenes (C15) are
synthesized in the cytosol from FPP (C15) derived from the MVA pathway, whereas
monoterpenes (C10) and diterpenes (C20) are synthesized in the plastid from GPP (C10) and
11
GGPP (C20), respectively, derived from the DXP pathway. However, some experimental
evidence suggests that the precursors for terpene synthases (TPS; prenyl diphosphates such as
FPP, GPP, and GGPP) can be transported to and from the plastid. In addition, some terpene
synthases can efficiently use physiologically non-relevant substrates. For example, Aharoni et
al. have found a cytosolic sesquiterpene synthase (FaNES1) from Fragaria ananassa (garden
strawberry) capable of synthesizing both linalool (monoterpene) and nerolidol (sesquiterpene)
from GPP and FPP, respectively (Aharoni et al., 2004). In their experiment, overexpression of
FaNES1 in the plastid surprisingly resulted in production of relatively high quantities of linalool
as well as small amounts of nerolidol (Aharoni et al., 2004). Therefore, this cytosolic enzyme has
the capability to synthesize a monoterpene from a physiologically non-relevant substrate, GPP.
Other literature evidence further implies a plastidal proton symporter which could transport
plastidic prenyl diphosphates to the cytosol, although there is no additional biochemical or
genetic data to support this report (Bick and Lange, 2003). In snapdragon, sesquiterpene
volatiles were shown to be synthesized from IPP, originating from the plastid, by labeling and
inhibitor studies (Bick and Lange, 2003). However, it is not certain if this metabolic crosstalk is
a specific case in one species or if it is a widespread phenomenon in the plant kingdom.
Nonetheless, it is evident that prenyl diphosphates can be transported from plastid to cytosol
efficiently (Dudareva et al., 2005). Similarly, cytosol to plastid transport has also been proposed
to occur in some plants (Rodriguez-Concepcion, 2006). Whether the activities of TPS enzymes
can truly catalyze these two distinctly separated reactions or if it happens inadvertently due to
promiscuous activities developed during the evolution of homologous TPSs remains to be seen
and implies a deeply complex system of subcellular regulation (Nagegowda et al., 2008).
12
Advancement in reverse genetics (e.g., RNAi and virus-induced gene silencing) has
allowed researchers the ability to investigate complex transcriptional regulation by silencing
specific transcripts, thus allowing further elucidation of crosstalk between the DXP and MVA
pathways. Additional support for the crosstalk was observed by RNAi-silencing of DXS in the
DXP pathway. Knock-down of DXS unexpectedly increased the level of sesquiterpenes in the
cytosol; subsequently, precursors from the MVA pathway were incorporated into monoterpenes
in the plastid by isotope-labeling studies (Paetzold et al., 2010). Therefore, a body of direct and
indirect experimental data strongly suggests that metabolic pools originating from both the MVA
and DXP pathways are interchangeable in plants.
The condensation of IPP to the priming molecule (DMAPP) occurs in a trans-
configuration, and until recently it has been believed that trans-GPP, FPP, and GGPP (or E,E-
prenyl diphosphates) are the only natural substrates for terpene synthases. However, a novel
pathway in wild tomato capable of producing sesquiterpenes from a cis-configured FPP (or Z,Z-
FPP) was identified and shown to be localized in the plastid (Sallaud et al., 2009). This is a
good example that TPS function is highly versatile and cannot be predicted by sequence
information alone.
Grey text indicates continuation of pathway to primary metabolite production. The MVA
pathway’s subcellular location is within the cytosol.
14
Grey text indicates continuation of pathway to production of primary metabolites. The DXP
pathway’s subcellular location is contained within chloroplasts.
15
1.3 Terpene Synthases
The exceptionally large diversity of terpenoids can be attributed to the catalytic plasticity
of the terpene synthase (TPS) enzyme family. TPSs catalyze acyclic and cyclic rearrangements
of their linear prenyl diphosphate and squalene precursors into a plethora of different terpenoids.
TPSs have a great degree of specificity towards their respective prenyl diphosphate precursors
but exhibit large variation in their catalytic mechanisms, resulting in enzymes producing single
and multiple terpene products. For example, two multi-product TPSs from Abies grandis were
shown to produce 34 and 52 different terpenes, respectively, whereas a third synthase from the
same species produces only -bisabolene (Steele et al., 1998). Evolution of the TPS family is
proposed to have occurred by duplication of general or specific metabolic genes, and subsequent
adaptive radiation of duplicated TPS genes (i.e., mutations), leading to enzymes that synthesize a
distinct product from the same substrate (Pichersky and Gang, 2000). To date, roughly 300
specific terpene skeletal structures have been reported, which most likely arose from the diverse
activities originating from gene duplications and neo-functionalization of TPSs (Bohlmann et al.,
1998).
Currently, the crystal structures for several TPSs from plant, bacteria, and fungi have
been determined. Several plant TPS structures have been described so far, including the (+)-
bornyl diphosphate synthase from Salvia officinalis (Whittington et al., 2002), -bisabolene
synthase from Abies grandis (McAndrew et al., 2011), epi-aristolochene synthase from
Nicotiana tabacum (Starks et al., 1997), (+)--cadinene synthase from Gossypium arboreum
(Gennadios et al., 2009), and ent-copalyl synthase from A. thaliana (Koeksal et al., 2011), as
well as taxadiene synthase from Taxus brevifolia (Koeksal et al., 2011). In general, the study of
TPS-mediated reactions involves either ionization-dependent or protonation-dependent
16
carbocation formation. This is quite similar to the prenyl transferase enzymes from which the
terpene synthases are believed to have evolved. For example, the ionization-dependent terpene
synthases have an -helical fold termed the class I TPS-fold, whereas the protonation-dependent
synthases possess an unrelated -barrel fold, class II fold. However, exceptions to this rule exist
throughout the terpene synthase family. Abietadiene synthase, a diterpene synthase, possesses
both class I and II folds, in a single polypeptide and hence can catalyze both ionization- and
protonation-dependent reactions. Tobacco epi-aristolochene synthase (TEAS) also has both
structural elements, but only the class I fold is active and located in the C-terminal domain
(Christianson, 2006). The sesquiterpene synthase -bisabolene synthase from A. grandis is an
exceptional enzyme in that it has a vestigial -domain normally present in diterpene synthases
(McAndrew et al., 2011). This has interesting implications in terpene synthase evolution, as the
current theory reasons that sesquiterpene synthases evolved from diterpene synthases (Trapp and
Croteau, 2001), which could point to -bisabolene synthase from A. grandis as the most recently
diverged sesquiterpene synthase known.
1.3.1 Sesquiterpene Synthase Structure-Function Relationships
Many TPSs, from bacteria, fungi, and plant, lack similarity in primary structure but share
distinct structural domains, such as the N-terminal domain and the catalytic C-terminal domain
(Starks et al., 1997; Bohlmann et al., 1998). The structural characterization of the first TPS from
plant, TEAS, revealed that it completely consists of an -helical structure with short connecting
loops forming an -helical barrel active site (Starks et al., 1997), which is now known to be
conserved throughout bacteria, fungi, and plants termed the ‘terpene synthase fold’ (Bohlmann et
al., 1998). Other specific structural elements of terpene synthases include a conserved DDXXD
17
motif involved in binding divalent metal ions for stabilization of the diphosphate moiety upon
ionization, and variations or duplications of this ‘aspartate rich’ motif result in reduced activity.
The highly hydrophobic aromatic-rich active site in TPS accommodates the long olefin chain of
the prenyl diphosphate substrate while the two Mg 2+
ions are complexed with the aspartate-rich
motif (DDXXD), stabilizing the ionized diphosphate group. A third Mg 2+
is complexed by a
molecule (Christianson, 2006). The N-terminal domain contains two flexible regions termed the
A-C and J-K loops which help to prevent solvent-access to the hydrophobic active site when
bound to a substrate. The A-C loop contains two generally conserved arginine residues. One
helps to stabilize the lid forming action of the J-K loop when a substrate binds to the TPS and the
other helps to stabilize the negatively charged diphosphate. This action is presumably important
to prevent the regeneration of the initial FPP or its tertiary allylic isomer, nerolidyl diphosphate.
Further stabilization of the carbocation intermediates occur through conserved aromatic residues
via -cation interaction. Whether these and other aliphatic residues have active or passive
functions within the carbocation reaction mechanism is currently a topic of debate (Miller and
Allemann, 2012). For example, recent studies of patchouli alcohol synthase from the plant
species, Pogostemon cablin, putatively implicated a single leucine in active reorientation during
catalysis, effectively creating a second active site pocket (Faraldos et al., 2010). Conversely, the
skeletal structure of the terpene may rely more on the initial orientation of the substrate upon
binding the active site, implying a more passive role for the active-site residues as chaperones to
a product. For example, TEAS and Hysocyamus premnaspirodiene synthase are two
evolutionarily related enzymes that have been shown to share a carbocation intermediate but
yield different products. Studies in which various amino acids were mutated, independent of the
18
active sites and in increasing radii from the active sites, resulted in switching of their respective
products (Greenhagen et al., 2006).
Initial insight into the structure-function relationships between sesquiterpene synthases
and their substrate (FPP) came from the TEAS crystal structure. This evidence led to the
synthetic carbocation mechanism of TEAS and became the template for which most TPS
catalyzed reactions proceed. Stabilization and positioning of the electrophilic carbon (C1)
facilitates attack by the C10-C11 pi-bond creating a cyclic carbocation (Figure 5). Termination
of this carbocation, which produces a germacrene A intermediate, was initially proposed to occur
by an acidic tyrosine. However, this has also become a contentious observation. Site-directed
mutagenesis of other Tyr residues from sesquiterpene synthases from two bacterial species
Penicillium roquefortii (Felicetti and Cane, 2004) and Fusarium sporotrichioides resulted in no
change of product. This result caused researchers to conclude that the diphosphate ion may be
involved with acid/base catalysis (Shishova et al., 2007). Substrate docking of the bisabolyl
cation in modeling simulations, with two sesquiterpene synthases from Sorghum bicolor,
indicates the diphosphate ion as a proton acceptor (Garms et al., 2012). Subsequently, the
reaction from the germacrene A intermediate in TEAS proceeds by addition of a proton to C6 via
a Asp-Tyr-Asp catalytic triad where the last two residues are contained within the J-K loop
(Figure 5). Consequently, a second ring closure at C2 and C7 would occur creating the
eudesmane carbocation intermediate. Final termination of the carbocation cascade by
deprotonation of the eudesmane intermediate by the indole ring of a tryptophan would be
facilitated by the formation of an arenium cation (Figure 5). Fundamentally, the termination of
the carbocation cascade can occur by capture of water, which creates a terpene alcohol or by
proton abstraction and different TPSs apply different quenching methods.
19
Relationships between reaction mechanism and enzyme kinetics have yet to be
scientifically explored. Roughly 100 sesquiterpene synthases have been characterized as of 2008
(Degenhardt et al., 2009), and most sesquiterpene synthases have an apparent Km ranging from
0.4-10 M (Picaud et al., 2005) with the exception of -bisbolene synthase which exhibits a Km
of 49.5 M (McAndrew et al., 2011). Slower rates of catalysis are observed with enzymes
involved in sesquiterpene biosynthesis, in general, and sesquiterpene synthases show relatively
low kcat values ranging from 0.033 - 4.0x10 -3
s -1
20
Figure 5. Schematic diagram representing the carbocation mechanism of tobacco epi-
aristolochene synthase (TEAS).
1.3.2 Phylogenetic Relationships of Terpene Synthases
TPSs are believed to have originated from prenyl transferases (e.g., GPP and FPP
synthase). However, little empirical evidence exists for such conclusions. Elucidation of TEAS
3D structure revealed convincing evidence as the C-terminal backbone of TEASs tertiary
structure aligns with avian FPP synthase, despite apparent lack of primary sequence similarity
(Starks et al., 1997). Further convincing evidence from TPS phylogenetic alignments of amino
acid sequences (>40% similarity) revealed that gymnosperm monoterpene, sesquiterpene, and
diterpene synthases are more closely related to each other than to their counterparts in
angiosperm (Bohlmann et al., 1997; Bohlmann et al., 1998). This indicates convergent evolution
of specialized TPSs after the angiosperm and gymnosperm bifurcation (Bohlmann et al., 1997;
Bohlmann et al., 1998). Classification of TPSs based on the phylogenetic analysis showed that
seven TPS clades or sub-families are present in nature and fit into the following nomenclature,
TPS-a to -g (Bohlmann et al., 1998; Aubourg et al., 2002; Dudareva et al., 2003). The TPS-a
subfamily consists of casbene synthase, a diterpene synthase, and sesquiterpene synthases from
various angiosperms. TPS-b consists of monoterpene synthases from angiosperm but is distinct
from TPS-a. TPS-c and TPS-e contain diterpene synthases from primary metabolism, and
therefore have fewer representative members. The subfamily of TPS-f contains only one
presumably ancient linalool synthase, and the TPS-d subfamily contains gymnosperm TPSs.
Recently, three monoterpene synthases from Antirrhinum majus, one monoterpene synthase from
A. thaliana, and a sesquiterpene synthase, nerolidol synthase, from Fragaria ananassa comprise
a new subfamily, TPS-g, characterized by a lack of an RRx8W motif, which is present in all
characterized monoterpene synthases from angiosperm TPS-b and gymnosperm TPS-d
subfamilies (Bohlmann et al., 1997; Aubourg et al., 2002; Dudareva et al., 2003; Jones et al.,
22
2011). Function of this motif is thought to be involved in cyclization of prenyl diphosphates as
all synthases lacking this motif produce acyclic products (Dudareva et al., 2003).
1.4 Metabolic Engineering of the MVA Pathway
Many plant terpenoids have been traditionally used as aromas, flavors, pharmaceuticals,
and nutraceuticals, but the natural abundance of terpenoids is minute. Furthermore, the structural
complexity of terpenoids has prevented their chemical synthesis on a commercial scale.
Therefore, biotechnological efforts have focused on the over-production of rare but valuable
terpenoids in fast growing heterologous microbial hosts such as E. coli and yeast. E. coli and
yeast provide genetically amenable platforms for reconstitution and manipulation of complex
metabolic pathways, such as the MVA and DXP pathways for improved terpenoid production.
Reconstitution and manipulation of the MVA or DXP pathways have been attempted and
proven to be successful in E. coli and yeast. Prokaryotes, such as E. coli, do not possess the
MVA pathway, and thus reconstruction of the pathway in E. coli could create an organism
implemented with an entirely synthetic metabolic pathway. The synthetic MVA pathway in E.
coli is expected to be free from any endogenous regulatory mechanisms and hence avoids
complicated feedback regulation (Dudareva et al., 2003). Although manipulation of the
endogenous DXP pathway in E. coli has been proven to increase the level of terpenoids, the
endogenous regulatory mechanisms controlling the DXP pathway in E. coli are highly complex
and not fully understood and hence the scalable production of terpenes was not achieved
(Kajiwara et al., 1997; Farmer and Liao, 2001; Kim and Keasling, 2001). Recently, complete
reconstitution of the MVA pathway in tobacco chloroplasts was successful in producing higher
23
than normal amounts of FPP derivatives, indicating plant metabolic engineering is also feasible
(Kumar et al., 2012).
Metabolic engineering of yeast relies heavily on modifications of the endogenous MVA
pathway, and the best example for increased C15 sesquiterpene production involved increasing
FPP abundance (Ro et al., 2006; Shiba et al., 2007; Ro et al., 2008). Four central points of
importance in achieving enhanced carbon flux for de novo terpene synthesis are: i) to increase
the pool of acetyl-CoA that serves as a precursor to the MVA pathway, ii) to increase cellular
activity of the rate-limiting enzyme, 3-hydroxy-3-methylglutaryl-coenzyme A reductase
(HMGR) and deregulate it from feedback inhibition, iii) to re-route FPP from ergosterol (yeast
sterol) to sesquiterpene biosynthesis, and iv) to overexpress the transcription factor activating
the steroid (i.e., MVA) biosynthetic pathway.
Firstly, implementing the pyruvate dehydrogenase bypass in yeast can alleviate the
bottleneck created by pathway precursor supply of acetyl-CoA to the MVA pathway. The
pyruvate dehydrogenase bypass converts pyruvate into acetyl-CoA in three steps by pyruvate
decarboxylase, acetaldehyde dehydrogenase, and acetyl-CoA synthetase. By overexpressing the
endogenous acetaldehyde dehydrogenase and heterologously expressing a Salmonella enterica
acetyl-CoA synthetase variant, Shiba et al. were able to increase acetate production in
engineered Saccharomyces cerevisiae (Shiba et al., 2007). Secondly, the major metabolic
bottleneck of the MVA pathway is caused by the rate-limiting enzyme HMGR, and thus
overexpression of a deregulated version (N-terminal truncated) of HMGR could significantly
enhance the flux. HMGR is regulated by several intermediate products of the MVA pathway
including FPP, and its membrane bound N-terminal domain appears to mediate the feedback
24
inhibitory effect. N-terminal truncation of tHMGR was shown to abolish inhibitory activity and
increase squalene production in yeast (Donald et al., 1997; Polakowski et al., 1998). Thirdly,
squalene synthase condenses two C15 FPP molecules to synthesize C30 squalene, however this
synthase can be down-regulated to increase the availability of FPP. Sterol biosynthesis in yeast
is an essential pathway, and the biosynthesis of sterol in S. cerevisiae involves over 20 distinct
reactions from the precursor acetyl-CoA, proceeding through FPP, which is a branch point for
sterol and sesquiterpene production (Shiba et al., 2007). Squalene synthase is the first committed
step in sterol biosynthesis. Therefore, down-regulating the expression of squalene synthase
(ERG9) can have a marked impact on increasing FPP (Ro et al., 2006). Lastly, a point-mutant
version of UPC2 transcription factor (upc2-1) can constitutively up-regulate several genes in the
MVA pathway.
Additional studies have employed more drastic methods to block squalene synthesis from
FPP by the complete knockout of squalene synthase. Complete aberrant removal of this gene
would result in a lethal mutant (sue) (Takahashi et al., 2007), but the phenotype can be rescued
by an external supply of ergosterol (yeast cholesterol), producing an abundant level of FPP.
However, cytotoxicity becomes a significant problem with engineering overproduction of FPP,
and consequently the yeast dephosphorylate FPP by diacylglycerol pyrophosphate phosphatase
(DPP1) to form farnesol (FOH), which is less toxic. Therefore, knocking out dpp1 is a rational
step in committing carbon to the production of sesquiterpenes (Faulkner et al., 1999). Similar
efforts to improve flux of FPP towards terpene hydrocarbon production, such as overexpression
of the FPP synthase, have been used but have little additive effect (Jackson et al., 2003; Ro et al.,
2006).
25
Another potentially significant problem exists with the consequence of high-level
production of terpenoids as there may be innate toxicity related to the terpene being produced
(Ro et al., 2008). Consequences of such toxicity have been observed in yeast engineered to
produce high levels of arteminisic acid. Ro et al. found that yeast engineered to produce large
quantities of artemisinic acid, an anti-malarial drug precursor to artemisinin, resulted in the
induction of multiple pleiotropic drug resistance genes (Ro et al., 2008). Global transcription
analysis by yeast microarray, as well as quantitative PCR, identified genes from the major
facilitator superfamily, in addition to ATP-binding cassette transporters, in response to the
overproduction of the weak acid, artemisinic acid.
1.5 Ligand-Receptor Binding
Several terpenoids have been shown to bind pharmacologically important receptors with
high specificity, and therefore have relevance as anti-cancer anti-psychotic and anti-malarial
drugs (Eckstein-Ludwig et al., 2003; Jordan and Wilson, 2004; Yan et al., 2005; Winther et al.,
2010). For example, salvinorin A is a lipophilic neutral small molecule, which selectively binds
a G-protein coupled receptor (GPCR) (Yan et al., 2005). Another diterpene anti-cancer drug,
paclitaxel, has been shown to be a potent mitotic inhibitor (Jordan et al., 1996). Other examples
of terpenoids that have selective biological targets are artemisinin and thapsigargin which both
attenuate activity of Ca 2+
ion pumps in Plasmodium falciparum ATP6 and its mammalian
homolog sarcoplasmic endoplasmic reticulum Ca 2+
ATPase (SERCA), respectively (Eckstein-
Ludwig et al., 2003; Winther et al., 2010).
Salvinorin A is a hallucinogenic diterpene produced by the sage Salvia divinorum and has
historically been used by the Mazatec people of Oaxaca, Mexico in shamanic rituals. The
26
hallucinogenic properties of salvinorin A lie in its ability to selectively bind the -opioid receptor
(Yan et al., 2005). Salvinorin A was the first non-alkaloid opioid subtype-selective drug and
exhibits no affinity for the traditional target of most natural hallucinogenic compounds, such as
N,N-dimethyltryptamine, psilocybin, and mescaline, and it rivals the potency of synthetic
hallucinogens, such as lysergic acid diethylamide (LSD) (Roth et al., 2002). Stabilization of the
compound in the binding pocket is through unusual and generally unconserved binding residues
isoleucine, glutamate and tyrosine (Yan et al., 2005).
Inhibition of mitosis represents a powerful approach in controlling cancer cell
proliferation. Microtubules play an extremely important role in the proliferation of metastatic
tumors as these generally advance through mitosis rapidly. For example, during prometaphase,
microtubules must rapidly extend and retract in an effort to adhere to the kinetochores. This
highly dynamic nature of microtubule formation is the basis for ‘microtubule binding agents’.
Paclitaxel is a microtubule stabilizing agent (MSA) which promotes polymerization whereas
vinblastine or vincristine bind and inhibit polymerization. The consequence to the cell is loss of
the dynamic nature needed for advancement to anaphase, and consequently the cell enters
apoptosis. Paclitaxel binds the -subunit of tubulin and is located on the inner surface of
microtubule structures (Nogales et al., 1995). Elucidation of the actual binding site has
identified an arginine residue as the specific amino acid involved (Rao et al., 1999). The
mechanism by which paclitaxel promotes polymerization is unknown, and there is only one
binding site on every molecule of tubulin (heterodimer of and subunits). Initially it was
believed that taxanes and other MSAs diffused through fenestrations in the microtubule wall.
However, kinetic studies have revealed that binding of paclitaxel is too fast to occur in this
manner, leading scientists to propose a second mechanism (Diaz et al., 2003). Recent modeling
27
evidence supports a proposed second binding site whereby taxanes bind first to the outer-
microtubule surface before moving to the -subunit binding site on the inner surface of the
tubulin structure (Magnani et al., 2009).
Calcium balance within the endoplasmic reticulum is an important process as Ca 2+
is an
important second messenger in cell signaling processes. Consequently, Ca 2+
flux is tightly
governed by Ca 2+
ion channels, and disruption of such processes can lead to pro-apoptotic
cascades, indirectly inducing cytochrome c release, caspases, and finally cell death (Scorrano et
al., 2003; Deng et al., 2009). Therefore, inhibition of normal SERCA function by thapsigargin
regardless of the proliferative state of the cell could make this sesquiterpene lactone a potent
anti-cancer drug (Winther et al., 2010). The lipophilic nature of thapsigargin allows it to
selectively bind the E2 form of SERCA (Toyoshima and Nomura, 2002). Unfortunately,
selective targeting of non-proliferative cells is not feasible with most anti-cancer drugs, and
hence a pro-drug mechanism has been designed for thapsigargin where a short H-S-S-L-Q-L
amino acid sequence attached to a short linker at O-8 allows for specific recognition by a
prostate-specific antigen protease (Denmeade et al., 2003). In a similar mechanism, artemisinin
inhibits PfATP6 Ca 2+
levels in P. falciparum, and mutagenesis studies have identified a single
amino acid which can abolish the inhibitory activity of artemisinin (Uhlemann et al., 2005). The
impetus for which malarial parasites develop resistance to artemisinin may impinge on
elucidation of this mutation in natural settings (Krishna et al., 2010).
1.6 Sesquiterpene Biosynthesis in Valeriana officinalis
Valeriana officinalis is a medicinal plant native to Asia and Europe where it has been
used for centuries as a potent sedative, although contemporary uses are more common to Europe
28
and the United States. In fact, valerian made the top-ten list of top selling herbal remedies in the
United States in 2002 (Anderson et al., 2005). The first biological activity relating to valerian
root extract was observed over 50 years ago (Stoll et al., 1957). Various metabolites of the
essential oil extracts from dried root show hundreds of specialized metabolites that include but
are not limited to chlorogenic acid, monoterpene alkaloids, terpenoids, valepotriates, furanofuran
lignans, and phenylpropanoids (Torssell and Wahlberg, 1966, 1967; Houghton, 1999; Navarrete
et al., 2006). Major terpene compounds identified from valerian root extracts are
sesquiterpenoids, such as valeranone, valerenal, valerenic acid, and several valerenic acid
derivatives (Stoll et al., 1957; Houghton, 1988, 1999). The precise compound exhibiting activity
has been contentious, but recent studies implicated the sesquiterpene, valerenic acid, as an
inhibitor of the nuclear factor kappa-light-chain-enhancer of activated B cells (NF-B) pathway
(i.e., anti-inflammatory) and agonist of the -aminobutyric acid type A (GABAA) receptor (i.e.,
sedative) (Figure 7) (Jacobo-Herrera et al., 2006; Benke et al., 2009).
Mediation of neuronal excitability in the human brain is highly reliant on -aminobutyric
acid, which potently inhibits GABAA receptors (Khom et al., 2010). GABAA receptors are the
targets of the drug class benzodiazepines due to the important role they play in mediating the
balance between excitation and inhibition of the central nervous system. Consequently,
benzodiazepines have potentially serious side-effects. Therefore, discovery of drugs with similar
efficacy but benign manifestation of side-effects has led to extracts from plants, such as valerenic
acid from V. officinalis. Similarly, in vivo and in vitro experiments have reported valerenic acid
to be an allosteric inhibitor of the GABAA receptor, constituting it as a potential anxiolytic drug
with little toxicity (Benke et al., 2009; Khom et al., 2010). The hydrocarbon precursor to
valerenic acid, valerena-4,7(11)-diene, has also been implicated as an anxiolytic compound
29
(Takemoto et al., 2009). Ligand-receptor binding and mutagenesis studies of valerenic acid and
valerenic acid derivatives with GABAA receptors implicates Gln265 on the -3 subunit as
absolutely necessary for interaction (Benke et al., 2009).
Valerenic acid itself has been shown to have nM level binding constants with respect to
GABAA receptors (Benke et al., 2009). The unique structure of valerenic acid may be important
for activity as other derivatives have been shown to have similar and sometimes higher potency
(Khom et al., 2010; Kopp et al., 2010). Interestingly, because valerena-4,7(11)-diene and
valerenal (possible aldehyde precursor to valerenic acid) have also been determined to potentiate
GABAA receptors, therefore several of the possible pathway intermediates in valerenic acid
metabolism may have significance as sedatives and there may be a synergistic effect occuring.
30
1.7 Objectives
The goal of this project is to identify a novel sesquiterpene synthase, which catalyzes the
first committed step to valerenic acid from a medicinal plant Valeriana officinalis. Structural
analysis of valerenic acid suggests that valerena-4,7(11)-diene is the product from the
unidentified sesquiterpene synthase (Figure 6). Importantly, this sesquiterpene skeleton is
unique, and its synthase from FPP substrate has yet to be identified. Ultimately, I aim to
demonstrate the enzymatic synthesis of the medically important terpene, valerena-4,7(11)-diene.
Integrative approaches involving genomics, chemistry, and metabolic engineering tools will be
included in this project.
Four specific objectives to achieve this goal are as follows.
Specific Objectives
1. Utilize genomics resources to identify TPS genes from Valeriana officinalis.
2. Functional activity evaluation of the encoded recombinant enzymes in yeast and E. coli
systems.
4. Propose the mechanism for valerena-4,7(11)-diene synthesis based on TPS product profile.
31
Figure 6. Proposed biosynthetic pathway for valerenic acid production in V. officinalis.
32
2.1 Plant Cultivation and Metabolite Preparations
V. officinalis seeds were obtained from B & T world seeds (France). Seeds were
germinated at 20 °C, and seedlings were grown in the University of Calgary greenhouse.
Valerian root was ground by mortar and pestle with liquid N2, and 100 mg of the ground tissue
was extracted using 1 mL ethyl acetate. The organic layer was partitioned by centrifugation,
diluted 10 times, and analyzed by GC-MS under the conditions described below.
2.2 RNA preparations
Total RNAs were isolated according to a modified version of the published protocol
(Meisel et al., 2005). Valerian root and leaf were ground under liquid N2 and 1.5-2 g were
extracted with 5 mL/g tissue of extraction buffer (1% (w/v) Cetyl Trimethyl Ammonium
Bromide (CTAB), 0.5 M TRIS HCl pH 8.0, 0.25 M EDTA pH 8.0, 4% (w/v) NaCl, 0.5% (w/v)
polyvinylpyrrolidone (PVP) in diethyl pyrocarbonate (DEPC) treated H2O preheated to 65ºC.
100 L of fresh -mercaptoethanol and 50 L spermidine trihydrochloride (SPD) were added to
5 mL extraction buffer. After the extraction slurry was vortexed for 30 sec. an equivalent
volume of 24:1 (chloroform:isoamyl alcohol) was added followed by vortexing and
centrifugation at 12,000 x g for 20 min. at room temperature. The resulting aqueous phase was
decanted and extracted a second time with 24:1 (chloroform:isoamyl alcohol). The aqueous
phase was then decanted and LiCl was added to a concentration of 2 M. After an overnight
incubation at 4ºC the solution was centrifuged at 12,000 x g for 35 min at room temperature and
the supernatant removed and the pellet dried but avoiding complete dessication. The pellet was
then resuspended in 0.5 mL of DEPC-treated H2O and extracted once more with 24:1
(choloroform:isoamyl alcohol). After vortexing, the centrifugation step was performed at 14,000
33
x g for 30 min at 4ºC. Aqueous phase was extracted, 1 mL of 100% ethanol was added and the
solution incubated on ice before vortexing and precipitating for 30 min at -80ºC. The ethanol
solution was then centrifuged at 14,000 x g for 20 min at 4ºC. The supernatant was removed and
the pellet dried avoiding complete desiccation. The resulting pellet was washed with 75%
ethanol, centrifuged at 14,000 x g for 10 min at 4ºC and dried once more. Total RNA was
dissolved in 50 L DEPC-treated H2O followed by concentration and purity measurements by a
Nanodrop 1000.
2.3 cDNA Library Preparation from Total RNA
7 µg of double-stranded cDNA from root tissue were prepared by the supplier’s protocol
(Invitrogen) using Superscript II Reverse Transcriptase (Invitrogen). The 454 GS FLX Titanium
was used to sequence valerian cDNA, and the raw reads were assembled by the University of
Calgary Bioinformatics Center through the Magpie informatics platform.
2.4 Plasmid Construction for Yeast Expression
Full length sesqui-TPS cDNAs (VoTPS1/2/3) were obtained by in silico analysis of the V.
officinalis database from the PhytoMetaSyn project at the University of Calgary. VoTPS1/2/3
were amplified from valerian root cDNA by a forward primer and a reverse primer with a
restriction enzyme digestion site integrated into the primer (Table 1). General PCR conditions
were as follows: 1 cycle of 30 sec at 98C; 29 cycles of 10 sec at 98C, 30 sec. at 60C (Table
1), 1 min 45 sec at 72C; followed by 1 cycle for 10 min. at 72C. The amplified PCR product
was ligated into a pGEM vector using a TA-cloning kit (Promega). The resulting pGEM clone
harbouring one of VoTPS1/2/3 was then transformed into Top10 cells and grown overnight at
37C on plates containing 100 g/mL ampicillin. Colony PCR was then performed to confirm
34
the presence of inserts and a single positive colony was selected for growth overnight at 37C in
3 mL LB broth containing 100 g/mL ampicillin. Isolation of pGEM clones harbouring one of
VoTPS1/2/3 was done by kits (Gene-All, Korea) and subsequently, restriction mapped and
sequenced. pGEM clones containing one of VoTPS1/2/3 were then digested with their respective
restriction enzymes (Table 1) and ligated into a linearized pESC-Leu2d vector and transformed
into Top10 cells. Colonies from plates were then verified to contain the insert by colony PCR.
Positive colonies were grown at 37C in 3 mL LB broth containing 100 g/mL ampicillin and
clones isolated using a kit (Gene-All, Korea). Clones were resequenced to confirm the insert
was present in the desired vector as ampicillin was the selection marker for both pGEM and
pESC-Leu2d cloning.
2.5 Quantitative Transcript Analysis
Semi-quantitative RT-PCR analyses for VoTPS1/2 were performed using 250 ng cDNA
from V. officinalis root or aerial tissue for 30 cycles with an annealing temperature of 55°C. For
VoTPS1, the primers used were a forward primer, 5’-CTGTTTACGAACAAGACAAGTCATG
CAAC-3’, and a reverse primer, 5’-AAGTCACAAAGCGCACCAAATTCAGAACT-3’. For
VoTPS2, the primers used were a forward primer, 5’-TATCGTCGAACGATACATTATTAGC
ATCAG-3’, and a reverse primer, 5’- CTTTGTAGAATACATTCATAAAGCATG-3’. The
restriction enzyme mapping of the resulting 921-bp (VoTPS1) and 1032-bp (VoTPS2) amplicons
were performed using EcoRV and HindIII separately to confirm their sequence identities.
Identical conditions and primers were used with 250 ng of RNA from V. officinalis root or aerial
tissues as a negative control to rule out possible genomic DNA contamination. Elongation factor
1α (EF1) was used as an internal control with a forward primer, 5’-GACTGTCACACTTCTCA
CATTGCC-3’, and a reverse primer, 5’-TCTCGACCACCATAGGTTTGGT-3’, using 5 ng of
35
cDNA from V. officinalis root or aerial tissues by the same PCR conditions mentioned above.
Amplified VoTPS1/2 and EF1 fragments were mixed and run in the same lane for visualization.
Quantitative PCR of VoTPS1 was performed with a forward primer, 5’- TGGTCAAAGCATC
AACAATTATCGCT-3’, and a reverse primer, 5’-CTTCTTCTTTTGTGGCACCATGTTGT-3’.
Ten ng of cDNA from V. officinalis root or aerial tissues were used with an annealing
temperature of 58°C. The above mentioned EF1 primers were also used as the reference gene.
2.6 Yeast Transformation
All yeast transformations were done with the EPY300 strain according to the protocol
described by (Gietz and Schiestl, 2007). A single colony was selected for growth overnight at
30C in 2 mL SC (500 mL of media containing 0.695 g of a mixture of amino acids containing
various amounts of the following: L-Ala, L-Arg, L-Asn, L-Asp, L-Lys, L-Glu, L-Ile, L-Lys, L-
Phe, L-Pro, L-Ser, L-Thr, L-Tyr, L-Val, L-Trp, Gly, uracil and adenine; 3.35 g yeast nitrogen
base) media omitting His and Met with 2% (v/v) glucose and shaken at 200 rpm. The overnight
culture was diluted 25-fold to a 50 mL SC medium of the same components, at the same
concentrations and grown at 30C for 4-6 hrs shaking at 200 rpm, followed by two wash steps
with sterile ddH2O, pelleted for 5 min. at 4,150 rpm. This was followed by an additional two
wash steps with sterile ddH2O, centrifuged at 14,000 rpm for 30 sec. After the last wash the cells
were resuspended in 50% polyethyleneglycol, 1 M lithium acetate and single-stranded salmon
testes DNA (Sigma Aldrich). 0.5-1.0 g plasmid DNA was used for each respective
transformation and incubated at 42C for 40 min. After incubation transformations were left on
ice for 2-5 min. before plating on SC-agar media omitting His, Met and Leu supplemented with
2% (v/v) glucose and grown for 3 days at 30C..
36
2.7 In vivo Production of Terpenoids in Yeast
Transgenic yeasts were inoculated in 2 mL Synthetic Complete (SC) media omitting the
amino acids His, Met and Leu with 2% glucose, and the sub-cultures were cultivated overnight at
30C and 200 rpm. The overnight culture was diluted 25-fold to a 50 mL SC media omitting His
and Leu with 2% (v/v) galactose, 0.2% (v/v) glucose, and 2 mM Met. Five mL of dodecane
(10% of the culture volume) was overlaid to the culture medium to trap volatile terpenoids
released during culture. The 50 mL yeast was cultured at 30C for 200 rpm for 3 days. The
yeast cultures were then centrifuged at 4,000 rpm for 5 min, and 1 mL of dodecane was extracted
and diluted in hexane (100-fold dilution) for GC-MS analysis.
2.8 Plasmid Construction for E. coli Expression
The Gateway Cloning (Invitrogen) system was used for construction of the bacterial
expression clone. VoTPS1/2/3 genes were initially cloned into the pDONR207 vector using gene
specific primers with attB1 specific 5’ tails (Table 1). According to the Gateway manual a PCR
reaction was performed using the following conditions: 1 cycle for 2 min. at 95°C ; 10 cycles for
15 sec. at 94 °C, 30 sec. at 60°C; 1 min. 45 sec. at 68°C. 10 L of the previous reaction were
immediately added to 40 L of a second reaction using the following conditions: 1 cycle for 1
min. at 95°C; followed by 5 cycles for 15 sec. at 94°C, 30 sec. at 45°C, 1 min. 45 sec. at 68°C;
followed by 15 cycles for 15 sec. at 94°C, 30 sec. at 55°C, and 1 min. 45 sec. at 68°C using
primers with 3’ tails specific to the respective genes and their 5’ portions specific to attB1 sites
(Table 1). Homologous recombination of the PCR product and the pDONR207 vector were
done using conditions suggested in the Gateway manual and resulted in an entry clone
harbouring one of the genes VoTPS1/2/3. The entry clone was then transformed into Top10 cells
and grown overnight on a plate containing 30 g/mL gentamicin. After colony PCR a positive
37
single colony was selected and grown in 3 mL LB broth containing 30 g/mL gentamicin and
subsequently isolated using a kit (Gene-All, South Korea). Isolated entry clone was then
restriction mapped to confirm integration of VoTPS1/2/3. Similarly, a second recombination
reaction was performed using the entry clone harbouring VoTPS1/2/3 with the expression vector
pH9GW (provided by Dr. Paul O’Maille, John-Innes Centre, UK) according to the Gateway
manual. 1 L of the reaction product was then used to transform Top10 cells and grown
overnight on plates containing 50 g/mL kanamycin. Colony PCR was performed to verify
integration of VoTPS1/2/3 into pH9GW. A single positive colony was then selected for growth
overnight at 37C in 3 mL LB broth containing 50 g/mL kanamycin and the resulting
expression clone was isolated by a kit (Gene-All, Korea). Purified expression clones were then
restriction mapped and sequenced.
Amplicon Cloning
5′-GGGGACCACTTTGTACAAGAAAGCTGGGT-3′R attB2
2.9 Heterologous Expression Trials
VoTPS1/2/3 genes were cloned into the expression vector pH9GW (provided by Dr. Paul
O’Maille, John-Innes Centre, UK), which contains an in-frame N-terminal 9x hisitidine tag. E.
coli (BL21AI) with 50 g/mL kanamycin expressing either VoTPS1 or VoTPS2 were cultured at
37 °C until an A600 of 0.3-0.6 was reached and subsequently induced. For each clone an
uninduced and induced culture were grown at temperatures of 15 and 37°C and time points of 2,
4, and 6 hrs were sampled for protein expression by pelleting followed by lysis and visualized on
an SDS-PAGE gel. Expression of VoTPS3 was tested in a similar way but at a single
temperature of 37°C and using the Rosetta cell line.
2.10 Expression in E. coli and Protein Purification
VoTPS1 or VoTPS2 were cultured at 37 °C until an A600 of 0.3-0.6 was reached,
incubated for 30 min at 4°C, and induced for 24 hr at 15°C with 0.2% (v/v) arabinose. The
cultures were centrifuged (4,000 rpm for 30 min at 4°C), and pellets were resuspended in 25 mL
of extraction buffer (25 mM Tris-HCl pH 7.5, 100 mM NaCl, 10 mM imidazole, 10% (v/v)
glycerol, 1 mM PMSF, and 1 mM DTT), frozen in liquid N2, and stored at -80°C. After thawing
cells harbouring VoTPS2 in a 42°C water bath cells were lysed by sonication, the total lysate
centrifuged (30 min, 10,000 rpm at 4°C) and the supernatant incubated for 1 hr end-over-end at
4°C with 1 ml Ni-NTA affinity resin (Novagen). The sample was then loaded into an empty 10
mL BioRad Econo column. The column was washed with 35 column volumes of 50 mM Tris-
HCl pH 7.5, 750 mM KCl, 40 mM imidazole, 10% (v/v) glycerol, 1 mM DTT, and 0.1 % (v/v)
Triton X-100, and then washed with 5 column volumes of the same buffer without Triton X-100.
The column was eluted with 10 column volumes of 50 mM Tris-HCl pH 7.5, 100 mM KCl, 500
40
mM imidazole, 10% (v/v) glycerol, and 1 mM DTT, and 1 mL fractions were collected.
Fractions containing VoTPS2 were pooled and concentrated to 250 µL with an Amicon Ultra-4
centrifugation filter unit (10-kDa cutoff). Alternatively, the cleared lysate after centrifugation
was filtered by a 0.2 m filter, and the recombinant VoTPS1 or VoTPS2 enzymes were purified
through a Bio-scale Mini Profinity IMAC cartridge (1 mL bed volume; Bio-Rad) installed on a
Bio-Rad FPLC. Before loading the protein extract, the Ni-NTA column was equilibrated with
extraction buffer (50 mM TRIS-HCl, 1 mM PMSF, 1 mM DTT, pH 7.5). A single wash step (10
mL of 50 mM TRIS-HCl, 750 mM KCl, 10% glycerol (v/v), 40 mM imidazole) followed
equilibration. Sample loading was performed at a 1 mL min -1
rate, while wash and elution steps
were performed at 2 mL min -1
. VoTPS1 was eluted by 5 mL of 275 mM imidazole, followed by
a gradient to 500 mM imidazole (50 mM TRIS-HCl, 100 mM KCl, 10% glycerol, 500 mM
imidazole, 1 mM DTT; against the same buffer without imidazole) over a volume of 5 mL. An
additional 5 mL of buffer containing 500 mM imidazole were passed through the column to elute
any residual protein. VoTPS2 was also eluted by a linear gradient over a 10 mL volume from 0-
500 mM imidazole with the same buffers as above. Fractions of 1 mL were collected over the
entire elution and run on a 10% SDS-PAGE gel. Fractions containing either VoTPS1 or
VoTPS2 were pooled and concentrated on an Amicon concentrator (>30 kDa exclusion size).
Protein was subsequently quantified by the Bradford method (Bio-Rad).
Similarly, VoTPS3 was cloned into the expression vector pH9GW. Constructs carrying
VoTPS3 were expressed in E. coli Rosetta (DE3) pLysS cells (Novagen) and cultured with LB
broth during sub-culture stages. All cultures were incubated with 30 g/mL chloramphenicol
and 15 g/mL kanamycin. Rosetta cells carrying the VoTPS3 construct were grown in 2
41
Fernbach flasks containing 1L of rich media (TB) at 37C until an OD600 of 0.6-0.8 was reached.
After cooling cultures for 20 min. at 4C 1 mM IPTG (Inalco, Italy) was added for production of
recombinant VoTPS3 by growth at 15C for 20 hrs. Cultures were pelleted at 4,000 rpm for 30
min. at 4C. Cell pellets were weighed and resuspended in 1.5 mL/gpellet in extraction buffer (50
mM TRIS-HCl, 300 mM NaCl, 10 mM imidazole, 10% glycerol). The resuspended cells were
frozen at -80C until the day of purification.
Thawed cells were lysed by sonication (see above) and centrifuged at 10,000 rpm for 40
min at 4C. followed by decanting the supernatant and a second centrifugation step of 10,000
rpm for 30 min at 4C. The cleared lysate was then incubated at 4C overnight end-over-end
with 200 L Ni-NTA resin (Novagen). The slurry was then loaded onto a 1 mL Econo-column
(Bio-Rad) and washed with 50 mM TRIS-HCl, 500 mM KCl, 10% glycerol (v/v), 20 mM
imidazole. The column was eluted with 10 column volumes of 50 mM Tris-HCl pH 7.5, 100
mM KCl, 500 mM imidazole, and 10% (v/v) glycerol.
2.11 Gas-chromatography and Mass Spectroscopy Analysis
Organic extracts of EPY300 yeast expressing TPS1 or TPS2 were analyzed by total ion
scan and single ion mode (m/z 204) for product identification. Analysis was conducted on an
Agilent 6890N gas chromatography system coupled to an Agilent 5975B mass spectrometer.
Peaks pertaining to the expected parental mass of sesquiterpenes (m/z 204), specifically
germacrene D, valerenic acid and valerena-4,7(11)-diene, were analyzed by authentic standard.
All other sesquiterpenes identified were by the NIST5/Wiley7 mass spectra library, Massfinder
4, and/or by literature. Retention indices were calculated by using alkane standard (C10-C40)
and compared with the values in the literature and Massfinder 4 database. One L samples were
42
injected at an inlet temperature of 250C with a flow rate of 1 mL min -1
helium on a DB1-UI-MS
and DB-Wax column (30 m X 250 m i.d. X 0.25 m film thickness). The initial temperature of
the program was set to 40C followed by a linear increase of 10 C min -1
to a temperature of
220C. The Cyclodex B chiral column (30 m x 250 µm inner diameter x 0.25 µm film thickness)
was also used to compare retention behavior of authentic valerena-4,7(11)-diene and the terpene
product from VoTPS2. The program used for chiral analysis was: initially at 50 °C (hold for 5
min) followed by 5 °C min -1
linear increase to 70 °C and final ramp of 2.5 °C min -1
to 200 °C.
2.12 Purification and NMR of Valerena-4,7(11)-diene
100 mL of SC medium without His, Met, and Leu supplemented with 1.8 % (v/v)
galactose and 0.2 % (v/v) glucose was inoculated with 1.5 mL overnight culture of the EPY300
expressing VoTPS2. After 6 h, the culture was supplied with 1 mM methionine and 5 mg of
Amberlite™ XAD-4 (Sigma-Aldrich), which was washed with MeOH prior to use. After
cultivating 3 days at 30 °C and 180 rpm, the Amberlite resin™ was recovered by filtration,
washed with distilled water and submerged in MeOH. The suspension was extracted three times
with 10 mL hexane, and the combined supernatants were dried over Na2SO4 and evaporated by a
gentle N2 stream to 0.2 ml. The concentrate was separated by silica column chromatography in a
Pasteur pipette and eluted with 10 ml of n-hexane. The collected 0.5 ml fractions were analyzed
by GC-MS and valerena-4,7(11)-diene-containing fractions were pooled and evaporated to
dryness by a gentle N2 stream. For NMR analysis, the dried residue (app. 0.7 mg) was dissolved
in CDCl3, and spectra were recorded on an UltrashieldPlus 600 MHz spectrometer (Bruker) in
CDCl3 at -20°C. Chemical shifts were reported as parts per million relative to CDCl3.
2.14 NMR Analysis of Valerena-4,7(11)-diene Standard
43
The natural product, valerenic acid, was purchased from Extrasynthese (France).
Valerena-4,7(11)-diene was synthesized from valerenic acid by John Vederas’ laboratory
(University of Alberta), and a detailed synthesis procedure is given in the Appendix I. Nuclear
magnetic resonance (NMR) spectra were obtained on Varian Inova 500 MHz and 600 MHz
spectrometers. 1 H NMR chemical shifts are reported in parts per million (ppm) using the residual
proton resonance of solvents as reference: CDCl3 δ 7.26, and CD2Cl2, δ 5.32. 13
C NMR chemical
shifts are reported relative to CDCl3 δ 77.0, and CD2Cl2 δ 53.8. Infrared spectra (IR) were
recorded on a Nicolet Magna 750 or a 20SX FT-IR spectrometer. Film Cast refers to the
evaporation of a solution on a NaCl plate. Mass spectra were recorded on a Kratos IMS-50 (high
resolution, electron impact ionization (EI)), or a ZabSpec IsoMass VG (high resolution
Electrospray (ES)).
2.15 Enzyme Activity Assays
Verification of enzyme activity was done according to a modified protocol originally
described by (O'Maille et al., 2004). In 1.5 mL glass GC vials; 50 mM TRIS HCl pH 7.5, 10
mM MgCl2, 100 M FPP and 50 g protein in 500 L ddH2O were gently overlaid with 500 L
pentane and incubated at 30°C for 1 hr. The reaction was terminated by vortexing for 1 min. and
centrifuging at 4,150 rpm. Initially 300 L of pentane was extracted and concentrated with a
gentle N2 (g) stream to a volume of ~50 L. A second volume of 500 L pentane was added to
the assay vial, vortexed and centrifuged. An additional 300 L was extracted and concentrated.
Negative controls of a boiled enzyme, enzyme with 100 mM EDTA and no enzyme in addition
to the above buffer system were run in parallel with each enzyme assay.
2.16 Enzyme Characterization
44
Appropriate assay incubation time and enzyme amount were determined to ensure that
the initial velocity of the reaction was linear in the given conditions. 100 M FPP substrate was
spiked with [1- 3 H]-FPP (Perkin Elmer, Boston, USA, 23 Ci mmol
-1 ) and used as a stock solution
for serial dilution. Biochemical properties were determined in the substrate concentrations
ranging from 0.25 to 50 M in triplicates for each concentration. Assays were carried out in 100
L volumes with 1.5 g of purified protein in each assay. The reactions were overlaid with 900
L hexane and incubated for 15 min at 30C. Reactions were terminated by adding 100 L of
0.5 M EDTA and 4 M NaOH, followed by 1 min of vortexing. Reactions were then centrifuged,
and 500 L of hexane was mixed with 3.5 mL of scintillation cocktail. Total activity of the
radioisotope labeled product was analyzed by liquid scintillation counting (LS 6500 Multi-
Purpose Scintillation Counter, Beckman Coulter). Apparent Vmax and Km values were calculated
using the Enzyme Kinetics Module Sigmaplot 12.0.
2.17 Phylogenetic Analysis
All TPS sequences were extracted from the public domain. Any mono- or di-TPSs were
analyzed for a chloroplast targeting peptide by ChloroP (Expasy), which was subsequently
removed due to lack of homology of such sequences between species. All sequences were
aligned by CLC Main Workbench, saved as clustal (aln) files and analyzed by the website
www.phylogeny.fr with the following settings in á la carte mode: bootstrap value 100, neighbor
joining method and treedyn for tree rendering (Dereeper et al., 2008; Dereeper et al., 2010).
45
3.1 Metabolite Profiling of Valerian Root
Valerenic acid is known to accumulate in the root of the valerian plant (Valeriana
officinalis) (Bos et al., 1997). To ensure the presence of valerenic acid in the V. officinalis root
prior to 454 pyrosequencing, the volatile metabolites from V. officinalis root were analyzed by
gas-chromatography mass-spectrometry (GC-MS). The metabolites from root were identified by
spectral match to the mass spectra library and to an authentic valerenic acid standard. The root
sample presented a complex mixture of volatile compounds, but the two most abundant volatiles
were identified as bornyl acetate and valerenal (Figure 7A, D). Although the metabolite
composition of valerian varies depending on their ecotypes, these two compounds have been
reported as major constituents in valerian root (Bos et al., 1997; Letchamo et al., 2004). In our
initial analysis, valerenic acid was not detected, but the valerenic acid standard also could not be
measured at concentrations lower than 100 M likely due to its low volatility. To increase the
volatility of valerenic acid, the valerian root extract and valerenic acid standard were derivatized
(i.e., methylated) and re-analyzed by GC-MS. After derivatization, several new peaks appeared,
and the retention index (RI) and mass fragmentation of one later-eluting compound coincided
with the methylated valerenic acid (Figure 7A, B, C). The valerenic acid content from
greenhouse grown valerian was quantified to be 0.56 ± 0.02 mg (n=4) per g fresh weight, but no
valerenic acid was detected in aerial parts (stem and leaves) of the plant. Metabolite analysis
therefore confirmed that valerenal and valerenic acid are major terpenoid constituents of V.
officinalis root. This result also suggested valerena-4,7(11)-diene sesqui-TPS transcripts are
specific to root and likely to be abundant.
46
metabolites from valerian root.
be detected after methylation, and it
shows an identical retention index and
mass fragmentation pattern to those of
the authentic standard (B and C). D)
Mass fragmentation of valerenal is
shown (Dae-Kyun Ro).
3.2 Transcript Sequencing and Candidate Gene Isolation
From the same valerian root sample analyzed by GC-MS, cDNA was prepared and
subjected to 454 pyrosequencing. This deep transcript sequencing yielded a total of 949,214
reads with an average read length of 347-bp. After removing repetitive, AT-rich, and low quality
sequences, 759,335 high quality reads were collected and assembled via the Magpie
bioinformatics platform (The Bioinformatics Center, University of Calgary) using the MIRA
algorithm (Chevreux et al., 2004; Meisel et al., 2005). The MIRA assembly of the 454 reads
generated 55,093 unigenes, which covers 42.3 M-bp of the total transcripts. From this sequence
data set, transcripts homologous to the previously reported sesqui-TPS (e.g., amorpha-4,11-
diene, 5-epi-aristolochene, and germacrene A synthases) were retrieved by BLASTX homology
search (Wallaart et al., 2001; Ro et al., 2008).
Two full-length valerian sesqui-TPS cDNAs were distinctly identified owing to their
abundance in the database, and they are referred to as VoTPS1 and VoTPS2. The read numbers
for VoTPS1 and VoTPS2 transcripts constitute 0.03% (ranked 200th) and 0.04% (ranked 259th)
of all reads, respectively. The amino acid sequences deduced from the ORFs of VoTPS1 and
VoTPS2 share 75% identity, encoding 563 and 562 amino acids, respectively (Figure 8). These
two sesqui-TPS clones appear to be unique to valerian because the BLAST analysis shows that
the closest terpene synthase to VoTPS1 and VoTPS2 was germacrene D synthase from Vitis
vinifera with only 53% amino acid identity. The proteins encoded by VoTPS1 and VoTPS2 did
not possess any motif for plastid targeting, implying that they are not di-terpene or mono-terpene
synthases, which are known to be localized to the plastid. Semi-quantitative RT-PCR analyses
of these two transcripts in valerian root and aerial tissues (stem and leaves) showed predominant
expression patterns of VoTPS1/2 in valerian root (Figure 9). Although a marginal level of
48
VoTPS1 expression was detected in aerial tissues, the level of VoTPS1 transcripts in aerial tissues
was quantified to be 178 ± 6 fold (n=4) lower than that