UNIVERSITY OF CALGARY Functional Genomics and Metabolite

UNIVERSITY OF CALGARY
Functional Genomics and Metabolite Profiling as Tools for Alkaloid Biosynthetic Gene
Discovery
by
Donald Reed Dinsmore
A THESIS
SUMITTED TO THE FACULTY OF GRADUATE STUDIES
IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE
DEGREE OF MASTER OF SCIENCE
GRADUATE PROGRAM IN BIOLOGICAL SCIENCES
CALGARY, ALBERTA
NOVEMBER, 2015
© Donald Reed Dinsmore 2015
ABSTRACT
The benzylisoquinoline alkaloids (BIAs) are diverse group of plant specialized
metabolites found in the families Papaveracea, Ranunculaceae, Berberidaceae and
Menispermaceae. Plants remain the only commercial source for BIAs and their biosynthesis is
poorly understood. O-methyltransferases (OMTs) are wide spread in BIA biosynthesis. Putative
OMTs were found in stem and root Next-Generation Sequencing transcriptomic databases.
Putative OMT cDNAs were isolated from Papaver somniferum and commercially synthesized.
Recombinant protoberberine 2-O-methyltransferase (2OMT) was heterologously expressed in
Escherichia coli and assayed. 2OMT demonstrated the 2-O-methylation of protoberberine
alkaloids and the 7-O-methylation of simple BIAs. The substrate range and tissue specific
expression of 2OMT suggest its in vivo role is converting (S)-cheilanthifoline to (S)-sinactine. A
LC-MS based targeted alkaloid profiling of twenty BIA producing species from the families
Papaveracea, Ranunculaceae, Berberidaceae and Menispermaceae was conducted.
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ACKNOWLEDGEMENTS
The completion of this thesis would not have been possible without the continued support
and help from my fiancé, Alexis Greene, my parents Murray and Debra Dinsmore, my coworkers Scott Farrow and Guillaume Beaudoin and finally my supervisor Dr. Peter Facchini. I
would also like to thank my committee members, Dr. Greg Moorhead and Dr. David Schriemer
for their guidance throughout the duration of my program.
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TABLE OF CONTENTS
ABSTRACT............................................................................................................................................................... I
ACKNOWLEDGEMENTS......................................................................................................................................II
TABLEOFCONTENTS ....................................................................................................................................... III
LISTOFTABLES................................................................................................................................................. VII
LISTOFFIGURES ..............................................................................................................................................VIII
LISTOFSYMBOLS,ABBREVIATIONSANDNOMENCLATURE................................................................IX
CHAPTERONE:INTRODUCTION .................................................................................................................... 1
1.1SECONDARYORSPECIALIZEDMETABOLISMINPLANTS .............................................................................................. 1
1.2ALKALOIDBIOSYNTHESISINPLANTS.............................................................................................................................. 4
1.3BENZYLISOQUINOLINEALKALOIDBIOSYNTHESIS ..................................................................................................... 12
1.3.1Structuraldiversityofbenzylisoquinolinealkaloids ............................................................................... 12
1.3.2Molecularbiologyandbiochemistryofbenzylisoquinolinealkaloids ............................................. 14
1.3.3ThecatalyticmechanismofOMTs................................................................................................................... 22
1.4APPROACHESTOGENEDISCOVERYANDFUNCTIONALCHARACTERIZATIONOFALKALOIDBIOSYNTHETIC
GENES........................................................................................................................................................................................ 24
1.4.1Traditionalapproachestogeneidentification .......................................................................................... 24
1.4.2Genomics-basedgenediscoveryapproaches .............................................................................................. 24
1.4.3Integrationoftargetedmetabolomicsandtranscriptomicsforgenediscovery ...................... 26
1.4OBJECTIVES ...................................................................................................................................................................... 27
CHAPTERTWO:MATERIALSANDMETHODS..........................................................................................29
2.1NUCLEICACIDISOLATIONANDANALYSIS .................................................................................................................. 29
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2.1.1IsolationofcDNAsencodingputativeOMTsfromopiumpoppystem............................................. 29
2.1.2IsolationofcDNAsencodingputativeOMTsfromopiumpoppyroot.............................................. 29
2.2PHYLOGENETICANALYSIS.............................................................................................................................................. 30
2.3CONSTRUCTIONOFEXPRESSIONVECTORSANDHETEROLOGOUSEXPRESSION.................................................... 32
2.3.1StemOMTgenecandidatesIzOMT1throughIzOMT6 ........................................................................... 32
2.3.2RootOMTgenecandidatesDDOMT1-4......................................................................................................... 34
2.4PURIFICATIONOFRECOMBINANTPROTEINSANDANALYSIS .................................................................................. 35
2.4.1PurificationofrecombinantIzOMT1-6andDDOMT1-4 ..................................................................... 35
2.4.2SDS-PAGE.................................................................................................................................................................... 36
2.4.3WesternBlotAnalysis ........................................................................................................................................... 36
2.5RECOMBINANTOMTCANDIDATEASSAYS ................................................................................................................ 36
2.5.1RoutineEnzymeassays......................................................................................................................................... 36
2.5.2DDOMT1TemperatureandpHOptimaassay............................................................................................ 37
2.5.3DDOMT1KmAssays ................................................................................................................................................ 37
2.6LC-MSCONDITIONSFORENZYMEASSAYANALYSIS ................................................................................................. 38
2.6.1LCconditionsforenzymeassays ...................................................................................................................... 38
2.6.2Massanalyzerconditions .................................................................................................................................... 38
2.7REAL-TIMEQUANTITATIVEPCR(QPCR).................................................................................................................. 39
2.8LC-MSBASEDTARGETEDALKALOIDPROFILINGOF20BIAPRODUCINGPLANTSSPECIES.............................. 40
2.8.1Plants............................................................................................................................................................................ 40
2.8.2ChemicalStandards ............................................................................................................................................... 40
2.8.3ExtractionofalkaloidsfromBIAproducingplantspecies.................................................................... 41
2.8.4LCconditionsfortargetedalkaloidprofiling ............................................................................................. 42
2.5.2Massanalyzerconditionsfortargetedalkaloidsprofiling ................................................................... 42
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CHAPTERTHREE:THEISOLATIONANDFUNCTIONALEXPRESSIONOFAMOLECULARCLONE
ENCODING2-PROTOBERBERINEO-METHYLTRANSFERASE...............................................................44
3.1INTRODUCTION ................................................................................................................................................................ 44
3.2RESULTS............................................................................................................................................................................ 47
3.2.1Identificationandphylogeneticanalysisof2OMTcDNA ...................................................................... 47
3.2.2Heterologousexpressionof2OMT ................................................................................................................... 50
3.2.3Enzymaticpropertiesof2OMT ......................................................................................................................... 50
3.2.4Relativetranscriptabundanceof2OMTinopiumpoppytissue ........................................................ 63
3.3DISCUSSION ...................................................................................................................................................................... 66
3.4CONCLUSIONS .................................................................................................................................................................. 76
CHAPTERFOUR:TARGETTEDALKALOIDPROFILINGOFTWENTYBENZYLISOQUINOLINE
PRODUCINGSPECIES ........................................................................................................................................78
4.1INTRODUCTION ................................................................................................................................................................ 78
4.2RESULTS............................................................................................................................................................................ 80
4.2.1TargetedalkaloidprofilingbyLC-MS/MS ................................................................................................... 80
4.3DISCUSSION ...................................................................................................................................................................... 92
4.3.1SimpleBIAs ................................................................................................................................................................ 92
4.3.2Protoberberinealkaloids ..................................................................................................................................... 94
4.3.3Protopinealkaloids ................................................................................................................................................ 96
4.3.4Benzo[c]phenanthridinealkaloids .................................................................................................................. 98
4.4.5Aporphinealkaloids ............................................................................................................................................... 99
4.4.6Morphians ................................................................................................................................................................100
4.3.6.Limitationsoftargettedalkaloidprofiling...............................................................................................101
4.4CONCLUSIONS ................................................................................................................................................................104
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CHAPTERFIVE:DISCUSSION ....................................................................................................................... 105
5.1OVERVIEW ......................................................................................................................................................................105
5.2FUTUREDIRECTIONS ....................................................................................................................................................106
5.2.1Furthercharacterizationof2OMT................................................................................................................106
5.2.2Futuretargetedmetaboliteprofiling...........................................................................................................107
5.2.3UsingthetargetedalkaloidprofilesoftwentyBIAproducingplantsinconjunctionwithNGS
..................................................................................................................................................................................................108
REFERENCES..................................................................................................................................................... 112
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LIST OF TABLES
Table 2.1 Protein abbreviations and GenBank accession numbers for sequences used in
phylogenetic analysis of SAM-dependent O-methyltransferases from selected plants........ 31
Table 2.2. Primer sequences used to amplify gene candidates from stem and root opium
poppy cDNA ......................................................................................................................... 33
Table 2.3. Real-Time quantitative PCR Primers used to study the relative gene expression of
2OMT in different tissue types from opium poppy. ............................................................. 39
Table 3.1. The products formed by assaying different chemicals with 2OMT. m/z ratios
were determined by LC-MS and reaction products were determined by CID. The
methylation pattern of the substrate is indicated by the positions N, 6, 7, 4’ and 3’.
Activities are relative to scoulerine....................................................................................... 56
Table 3.2. The ESI[+]CID pattern of substrates and the products from the substrate range
experiments determined by LC-MS/MS ............................................................................... 61
Table 4.1 Chemical standards used for the targeted alkaloid profiling of 20 BIA producing
species by LC-MS/MS. ......................................................................................................... 83
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LIST OF FIGURES
Figure 1.1. The four general groups of plant specialized metabolites and their origins in
primary metabolism. ............................................................................................................... 3
Figure 1.2. Different classes of plant alkaloids and their biosynthetic origins (Dewick 2009)..... 5
Figure 1.3. Monoterpenoid indole alkaloid (MIA) biosynthesis and diversity.............................. 7
Figure 1.4. Tropane alkaloid biosynthesis and diversity. ............................................................ 10
Figure 1.5 Purine alkaloids biosynthesis and diversity................................................................. 11
Figure 1.6. Biosynthetic pathways showing benzylisoquinoline alkaloids accumulating in
Papver somniferum and other plant species.......................................................................... 16
Figure 3.1. Neighbour-joining tree of derived from the amino acid sequences of selected
plant SAM-dependant O-methyltransferases. ....................................................................... 49
Figure 3.2. Heterologous expression of 2OMT synthetic gene in E. coli..................................... 51
Figure 3.3. The effect of pH on Papaver somniferum 2OMT activity with scoulerine................ 52
Figure 3.4. The effect of temperature on Papaver somniferum 2OMT activity with scoulerine . 53
Figure 3.5 Extracted ion chromatograms (EICs) showing the lack of O-methylation activity
observed when scoulerine was assayed with each heterologously expressed enzyme (A)
IxOMT2, (B) IzOMT3, (C) IzOMT6, (D) DDOMT2, (E) DDOMT3 and (F) DDOMT4. .. 57
Figure 3.6 Extracted ion chromatograms (EICs) showing the O-methylation activity of
2OMT on (A) cheilanthifoline, (B) Scoulerine, (C) coclaurine, (D) dopamine, (E)
Reticuline and (F) 6-O-methylnorlaudanosoline. ................................................................. 58
Figure 3.7 The O-methylation activity of 2OMT on various BIA substrates (A)
cheilanthifoline, (B) scoulerine, (C) coclaurine, (D) dopamine, (E) reticuline and (F) 6O-methylnorlaudanosoline.................................................................................................... 60
Figure 3.8 Michaelis-Menten plot for Papaver somniferum protoberberine 2OMT with (S)scoulerine with a constant concentration of SAM. ............................................................... 64
Figure 3.9 Relative abundance of 2OMT gene transcripts in Bea’s Choice opium poppy
tissues .................................................................................................................................... 65
Figure 3.10 The proposed in vivo role of 2OMT in the biosynthesis of cryptopine and
rhoeadine alkaloids. .............................................................................................................. 77
Figure 4.1 Annotated LC-MS chromatograms from 20 BIA producing species.......................... 89
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LIST OF SYMBOLS, ABBREVIATIONS AND NOMENCLATURE
Acronym
Definition
[M]+
[M+H]+
2OMT
3’OHase
3’OMT
4’OMT2
Parent Ion
Protonated parent ion
Protoberberine 2-O-methyltransferase
Uncharacterized 3’-hydroxylase
Uncharacterized 3’-O-methyltansferase
3’-Hydroxyl-N-methylcoclaurine 4’-Omethyltransferase
4-Hydroxyphenylacetaldehyde
4-Hydroxyphenylpyruvate decarboxylase
6-O-methylnorlaudanosoline-5’-Omethyltransferase
norcoclaurine-6-O-methyltransferase
Reticuline 7-O-methyltransferase
Aspartate
Berberine bridging enzyme
Benzyl isoquinoline alkaloid
Basic Local Alignment Search Tool
Canadine synthase
Collisional energy
Cheilanthifoline synthase
Collisionally-induced dissociation
Coclaurine N-methyltransferase
Codeine O-demethylase
Columbamine O-methyltransferase
Codeinone reductase
Caffeine synthase
Cetyl triethylammonium bromide
Cytochrome P450
N-Methylcanadine 1-hydroxylase
Cysteine
Dihydrobenzophenanthridine oxidase
3, 7-dimethylxanthosine N-methyltransferase
Deoxyxyulose phosphate
Extracted ion chromatogram
Electrospray ionization
Expressed sequence tag
Flavin adenine dinucleotide
FAD-dependent oxidoreductases
4HPAA
4HPPDC
5’OMT
6OMT
7OMT
Asp
BBE
BIA
BLAST
CAS
CE
CFS
CID
CNMT
CODM
CoOMT
COR
CS
CTAB
CYP
CYP82Y1
Cys
DBOX
DXMT
DXP
EIC
ESI
EST
FAD
FADOX
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FPKM
FTICR-MS
FTIR
Gln
Gly
His
HPLC
HPR
IPP
IPTG
LB
LC
Lys
MIA
MIAMET
MPO
MS
MSH
MSn
MXMT
N7OMT
NCS
NGS
NMCH
NMR
NMT
NOS
ODC
ODD
OMT
ORF
P6H
PCR
Phe
PMT
qPCR
QqQ
RT-PCR
SalAT
SalSyn
SAM
SanR
Fragments per kilobase of exon per million
fragments mapped
Fourier transform ion cyclotron resonance mass
spectrtometry
Fourier transform infrared spectroscopy
Glutamine
Glycine
Histidine
High-performance liquid chromatography
Horseradish peroxidase
Isopentenyl diphosphate
Isopropyl ß-D-1-thiogalactopyranoside
Lauria-Bertani
Liquid chromatography
Lysine
Monoterpenoid indole alkaloid
Minimum information about a metabolomics
experiment
Methylputrescine oxidase
Mass spectrometry
N-Methylstylopine14-hydroxylase
Tandem mass spectrometry
7-methylxanthosine N-methyltransferase
Norreticuline 7-O-methyltransferase
Norcoclaurine synthase
Next-generation sequencing
N-methylcoclaurine 3’-hydroxylase
Nuclear magnetic resonance
N-methyltransferase
Noscapine synthase
Ornithine decarboxylase
2-Oxoglutarate/Fe(II)-dependent dioxygenases
O-methyltransferase
Open reading frame
Protopine 6-hydroxylase
Polymerase chain reaction
Phenylalanine
Putrescine N-methyltransferase
Real-time quantitative polymerase chain reaction
Triple quadrupole
Real-time quantitative polymerase chain reaction
Salutaridinol 7-O-acetyltransferase
Salutaridine synthase
S-adenosylmethionince
Sanguinarine reductase
x
SDR
Ser
SOMT1
SPE
SPS
STOX
STR
T6ODM
TNMT
TOF
Trp
TS
TYDC
Tyr
TyrAT
VIGS
Short-chain dehydrogenase/reductase
Serine
Scoulerine-9-O-methyltransferase
Solid-phase extraction
Stylopine synthase
(S)-Tetrahydroxyprotoberberineoxidase
Strictosidine synthase
Thebaine 6-O-demethylase
Tetrahydroprotoberberine N-methytransferase
Time-of-flight
Tryptophan
Theobromine synthase
Tyrosine decarboxylase
Tyrosine
Tyrosine aminotransferase
Virus-induced gene silencing
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CHAPTER ONE: INTRODUCTION
1.1 Secondary or specialized metabolism in plants
Plants have the remarkable ability to produce a wide variety of low molecular weight
metabolites. Ancient metabolic pathways common to almost all organisms are referred to as
‘primary’ and the rest, by convention, are referred to as ‘secondary’. Although primary
metabolites comprise a relatively small subset of plant metabolites they are essential for normal
growth, development and reproduction (Pichersky and Gang 2000; Kliebenstein and Osbourn
2012). All other metabolites made by plants are known as secondary metabolites. Over 200,000
defined structures are considered secondary metabolites and are found among most plant groups
(Hartmann 2007; Pichersky and Gang 2000). Since ancient times humans have exploited the
properties of plant secondary metabolites for use as dyes, fragrances, poisons and medicines.
Even today plants remain the sole source for several compounds essential for the production of
commercial and pharmaceutical products, in addition plants can serve as natural chemical
libraries for the development of new products and drugs (Butler 2008). Despite their usefulness
to humans, the scientific community viewed secondary metabolites as “flotsam and jetsam on the
metabolic beach” (Haslam 1986). As it were, secondary metabolites were thought to be
metabolic waste or detoxification products and conferred no advantage to the plant (Hartmann
2007). It wasn’t until the right experiments were performed by plant entomologists studying
herbivore-plant relationships that a more meaningful role for plant secondary metabolites was
considered (Fraenkel 1959). After decades of experiments, it is now widely accepted that many
secondary metabolites represent the chemical adaptations of certain plants to specific ecological
conditions and serve functional roles in plants to increase their overall fitness (Hartmann 2007;
Pichersky and Lewinsohn 2011). Secondary metabolites can increase the fitness of plants in
1
several different ways. For example, secondary metabolites can act as defence compounds
against herbivores and pathogens; as attractants for pollinators and seed-dispersing animals; and
as growth suppressants for neighbouring plants who compete for resources and sunlight
(Facchini et al. 2007; Liscombe and Facchini 2008; Pichersky and Gershenzon 2002; Theis and
Lerdau 2003; Dixon 2001). The anachronistic term, secondary metabolites, has fallen out of use
in recent years in favour of the new term, specialized metabolites, which more accurately reflects
the importance of these compounds (Ferrer et al. 2008; Pichersky and Lewinsohn 2011).
Plant specialized metabolites can be arranged in four general groups based on their
chemical structures: phenolics, terpenoids, non-aromatic polyketides, cyanogenic and sulfurcontaining metabolites and alkaloids (Figure1.1) (Micheal Wink 2010). Phenolics draw their
name from the presence of a phenol moiety in their structure. All phenolics originate from the
shikimate or malonate/acetate pathways. There are approximately 9000 known phenolics.
Figure 1.1 illustrates some of the different subclasses of phenolics, flavonoids,
phenylpropanoids, polyactylenes and polyketides (Micheal Wink 2010).
The largest group of plant specialized metabolites are called terpenoids with
approximately 25 000 known compounds. The 5-carbon precursor to all terpernoids, isopentenyl
diphosphate (IPP), is generated by the cytosolic mevalonate pathway or the plastidic
deoxyxyulose phosphate (DXP) pathway. The diversity of terpenoids results from the
polymerization and cyclization of IPP monomers by terpene synthases to generate various
terpenoid backbones classified by the number of IPP monomer combined to make them (Micheal
Wink 2010). Examples of each class of terpenoid are shown in Figure 1.1.
2
Figure 1.1. The four general groups of plant specialized metabolites and their origins in
primary metabolism.
The color of each box corresponds to the category of specialized metabolites the illustrated
compounds belong to: red, phenolics; green, terpenoids; blue, alkaloids; yellow,
cyanogenic and sulfur-containing. Cys, cysteine; DXP, deoxyxyulose phosphate; IPP,
isopentyl diphosphate.
3
Cyanogenic and sulfur-containing specialized metabolites are illustrated in Figure 1.1 and
include: cyanogenic glucosides, glucosinolates and cysteine sulfoxides. Of these three groupings
only approximately 160 compounds are known (Micheal Wink 2010). Cyanogenic glucosides
are hydrolyzed by some plants for defence purposes. The enzymatic hydrolysis of cyanogenic
glucosides releases toxic by-products in response to tissue damage or wounding (Fahey,
Zalcmann, and Talalay 2001; Dewick 2009). One such example is the compound amygdalin,
found in almonds. When almond tissue containing the amygdalin is crushed a series of
enzymatic steps hydrolyze it into benzaldehyde and hydrogen cyanide (Dewick 2009). The
primary constituent of garlic flavour is a cysteine sulfoxide known as allicin. Allicin is also a
known antimicrobial (Dewick 2009). The final class of plant specialized metabolites, alkaloids
(Figure 1.1), will be discussed at length in the preceeding section.
1.2 Alkaloid biosynthesis in plants
Alkaloids are amino acid derived low molecular weight compounds of 100 to 900 Da,
often with at least one nitrogen in a heterocyclic ring. These nitrogen-containing compounds are
produced by approximately 20% of plant species (Ziegler and Facchini 2008). Although some
animals and microbes produce alkaloids the majority of known alkaloids are made by plants.
Over 12 000 alkaloids have been identified making them the second largest groups of plant
specialized metabolites after terpenoids (Ziegler and Facchini 2008). Several alkaloids have
potent pharmacological effects, including but not limited to use as stimulants, poisons, narcotics
and anti-tumour agents (Ky et al. 2001; Reynolds 2005; Beaudoin and Facchini 2014; Noble
1990). As a result of their pharmacological effects, the properties of alkaloids have been
exploited by humans for centuries (Theis and Lerdau 2003). This class of plant specialized
metabolites can be divided into subgroups shown in Figure 1.2 that represent the chemical
4
Figure 1.2. Different classes of plant alkaloids and their biosynthetic origins (Dewick
2009).
Abbreviations: Trp, tryptophan; DXP, deoxyxylose phosphate; IPP, isopentenyl
disphosphate; Glu, glutamate; Gln, glutamine; His, histidine; Asp, aspartate; Lys, lysine;
Ser, serine; Gly, glycine; Tyr, tyrosine; Phe, phenylalanine.
5
diversity of alkaloids (Dewick 2009). The biosynthesis of different alkaloid subgroups depicted
in Figure 1.2 has been proposed based on data from a variety of experiments ranging from
alkaloids extractions, biosynthetic feeding studies and biomimetic syntheses. The biosynthesis
leading to different alkaloid subgroups have not been thoroughly investigated on a molecular
biochemical basis. The best studied biosynthetic routes leading to alkaloids are the terpenoid
indole, tropane, purine and benzylisoquinoline alkaloids pathways (Ziegler and Facchini 2008).
Monoterpenoid indole alkaloids (MIAs) (Figure 1.3) are a family of more than 3 000
members found primarily in the plant families, Apocynaceae, Loganiaceae, Nissacecae and
Rubiaceae (Loyola-Vargas, Galaz-Avalos, and Ku-Cauich 2007). The most widely used model
species for studying the biochemistry and molecular biology of MIA biosynthesis have been
Catharanthus roseus and Rauvolfia serpentina (Facchini and DeLuca 2008). All MIAs are
derived from tryptophan which is decarboxylated to form tryptamine. In the first commited step
in MIA metabolism, strictosidine synthase catalyzes the Pictet-Spengler condensation of
typtamine with secologanin to form strictosidine, illustrated in Figure 1.3 (O’Connor and Maresh
2006; Facchini and DeLuca 2008; Loyola-Vargas, Galaz-Avalos, and Ku-Cauich 2007; Kutchan
et al. 1988). In addition to strictosidine synthase over 12 different cDNAs encoding MIA
biosynthetic enzymes have been discovered and functionally characterized (Jörg Ziegler and
Facchini 2008). All the unique structural subclasses of MIAs are illustrated in Figure 1.3. In
many cases enzymatic steps have been characterized using traditional biochemical methods but
the corresponding cDNA has not been cloned (Ziegler and Facchini 2008; Dethier and De Luca
1993).
6
Figure 1.3. Monoterpenoid indole alkaloid (MIA) biosynthesis and diversity.
Strictosidine synthase (STR), catalyzes the Pictet-Spengler condensation of tryptamine and
secologanin to form strictosidine in the first committed step of MIA biosynthesis. The
structural subclasses of MIAs derived from strictosidine are italicized and represented by
the different colored boxes while the names of the illustrated compounds are written below.
Quinoline, red; Sarpagan, orange; Coryanthe, purple; Aspidosperma, green; Ajmalan,
yellow; Iboga, blue. Thin arrows represent a single enzyme catalyzed reaction and thick
arrows represent multiple enzymatic and/or spontaneous chemical reactions.
7
Tropane alkaloids are an important class of anticholinergic and stimulant compounds
found primarily in the families Solanaceae, Erythroxylaceae and Convolvulaceae (Figure 1.4)
(Griffin and Lin 2000; Ziegler and Facchini 2008). The importance of some tropane alkaloids
such as nicotine, cocaine, scopolamine and hyoscyamine has led to the isolation and
characterization of several genes encoding enzymes involved in the biosynthesis of tropane
alkaloids (Ziegler and Facchini 2008). Much of the pioneering work on tropane alkaloids has
been done with tobacco, Hyoscyamus niger and Attropa belladonna, however, recent studies
performed using Eruthroxylum coca have changed paradigms surrounding tropane alkaloid
biosynthesis (Ziegler and Facchini 2008; Jirschitzka et al. 2012; Docimo et al. 2012).
Interestingly, some evidence suggests that tropane alkaloid formation in distant angiosperm
lineages has evolved independently. For example, a tropinone-reduction step which converts
methylecgonone to methylecgonine with the help of a short-chain dehydrogenase/reductase
(SDR) family reductase has been found in Eruthroxylum coca and not in the taxonomically
remote Solanaceae family (Jirschitzka et al. 2012).
Purine alkaloids are a small group of plant specialized metabolites that are well known
and widely used by human. Purine nucleotides provide the precursors for plants like Coffea
Arabica (coffee), Camellia sinensis (tea) and Theobroma cacao (cacao) to synthesize alkaloids
such as, caffeine, theobromine and methyluric acid (Figure 1.5) (Suzuki, Ashihara, and Waller
1992; Ashihara, Sano, and Crozier 2008). The major route to caffeine starts with xanthosine
which undergoes three O-methylation steps followed by the hydrolysis of the ribosyl moiety
(Ashihara, Sano, and Crozier 2008). cDNAs encoding for each methyltransferase in the pathway
has been functionally characterized. Although the enzyme responsible for the hydrolysis of 7methylxanthosine has been partially purified from tea leaves, the corresponding cDNA has never
8
been found despite some evidence to suggesting the preceding O-methylation step and the
nucleoside cleavage are catalyzed by the same enzyme (Ashihara, Sano, and Crozier 2008;
Mccarthy and Mccarthy 2014). Caffeine synthase (CS), also referred to as 3,7dimethyxanthosine N-methyltransferase (DXMT), accepts both theobromine and caffeine as
substrates and is capable of completing the last two steps of caffeine biosynthesis, while
theobromine synthase (TS) or 7-methylxanthosine N-methyltransferase (MXMT) is specific for
the conversion of 7-methylxanthine to theobromine (Figure 1.5) (Ashihara, Sano, and Crozier
2008).
9
Figure 1.4. Tropane alkaloid biosynthesis and diversity.
The spontaneously derived N-methyl-Δ1- pyrrolium cation is the branch point from which
scopolamine (yellow) and cocaine (blue) biosynthesis split. Enzymes for which cognate
cDNAs have been isolates are in green. Thin arrows represent a single enzyme catalyzed
reaction and thick arrows represent multiple enzymatic and/or spontaneous chemical
reactions. ODC, ornithine decarboxylase; PMT, putrescine N-methyltransferase; MPO,
methylputrescine oxidase.
10
Figure 1.5 Purine alkaloids biosynthesis and diversity.
Enzymes for which cognate cDNAs have been isolated are in green. Thin arrows represent
a single enzyme catalyzed reaction and thick arrows represent multiple enzymatic and/or
spontaneous chemical reactions. XMT, xanthosine N-methyltransferase; MXMT, 7methylxanthosine N-methyltransferase; CS, caffeine synthase; TS, theobromine synthase;
DXMT, 3,7-dimethyxanthosine N-methyltransferase.
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1.3 Benzylisoquinoline alkaloid biosynthesis
1.3.1 Structural diversity of benzylisoquinoline alkaloids
Occurring primarily in the orders Papaveracea, Ranunculaceae, Berberidaceae and
Menispermaceae, benzylisoquinoline alkaloids (BIAs) are one of the largest and most diverse
alkaloid groups with over 2,500 unique structures identified to date (Ziegler and Facchini 2008).
BIAs have been used by humans for thousands of years. Some examples include the wellknown, analgesic morphine and the cough-suppressant codeine (Hagel and Facchini 2013).
Many other BIAs also possess potent pharmacological activities for example, noscapine is
thought to interact with microtubules to inhibit the proliferation of cancer cells (Landen, Lang,
and McMahon 2002). Papaverine and (+)-tubocurarine act as vasodilators and muscle relaxants,
while sanguinarine and berberine are known antimicrobial agent (Ziegler et al. 2009; Hagel and
Facchini 2013). The chemical complexity of BIAs, owing to the presence of at least one chiral
center, precludes the industrial synthesis of most BIAs. Therefore, in many cases plants remain
the sole source economic source of BIAs indispensible to modern medicine (Xiao, Zhang, Chen,
Lee, Barber, Chakrabarty, Desgagné-Penix, et al. 2013).
Despite the incredible structural diversity of BIAs derived from (S)-norcoclaurine there
are relatively few enzyme types associated with BIA metabolism. The only reported enzyme
types involved in BIA metabolism are S-adenosylmethionine-dependent N- methyltransferases
(NMTs) and O-methyltransferases (OMTs), cytochrome P450s (CYPs), acetyl-CoA-dependent
O-acetyltransferases, 2-oxoglutarate/Fe(II)-dependent dioxygenases (ODDs), Aldo-keto
reductases, short-chain dehydrogenases/reductases (SDRs), carboxylesterases and FADdependent oxidoreductases (FADOXs) (Beaudoin and Facchini 2014). Most evidence suggest
that the evolution of BIA biosynthesis was monophyletic (Liscombe et al. 2005) and as a result
12
many BIA biosynthetic enzymes, even between species, share considerable sequence identity.
The enzymes involved in BIA biosynthesis are thought to have arisen through the duplication of
genes recruited from primary metabolism. Random mutations in the duplicated gene allowed for
the development of novel catalytic functions without losing function of the original gene (Ober
and Hartmann 2000). The process of gene duplication and random mutation offers an
explanation as to how such a vast array of structurally unique BIAs could originate from only a
handful of enzyme types.
All BIAs are comprised of an isoquinoline group linked to a benzyl moiety which
provides the 1-benzylisoquinoline backbone. Using a limited catalytic tool kit the 1benzylisoquinoline backbone can be modified and decorated with functional groups to create
over 2,500 different BIAs. Alterations of the 1-benzylisoquinoline backbone by the formation of
intramolecular C-C and C-O bonds is catalyzed by oxidative enzymes, CYPs, FADOXs and
ODDs. Different backbone rearrangements are the criteria by which BIAs are classified into
subgroups and are known as, simple benzylisoquinolines, protoberberines, protopines,
aporphines morphinans, promorphinans, cularines, benzophenantridine and phtalideisoquinolines
(Shulgin and Perry 2002). CYPs are known to catalyze a range of different reactions from
hydroxylations and C-C and C-O couplings and are responsible for most rearrangements of the
1-benzylisoquinoline backbone (Beaudoin and Facchini 2014; Jörg Ziegler and Facchini 2008).
FADOXs are able to create C-C and C-N bonds in BIA metabolism one such examples is the
berberine bridging enzyme (BBE) which forms (S)-scoulerine from (S)-reticuline creating the
protoberberine backbone (Dittrich and Kutchan 1991; Beaudoin and Facchini 2014). Despite the
various roles of ODDs in other plant specialized metabolite biosynthetic pathways, to date,
13
ODDs involved in BIA metabolism have only demonstrated dealkylation activity (Hagel and
Facchini 2010; Farrow and Facchini 2013).
Decorative reactions of different BIA scaffolds can be performed by several enzyme
types. S-adenosylmethionine-dependent N-methyltransferases (NMTs) add methyl groups to the
nitrogen in the BIA backbone. One such enzyme is coclaurine N-methytransferase (CNMT) that
N-methylates (S)-coclaurine to form (S)-N-methylcoclaurine (Loeffler, Deus-Neumann, and
Zenk 1995). O-methylation is a common event in BIA metabolism and is often regiospecific.
Most OMTs are involved in the O-methylation of the 1-benzylisoquinoline backbone such as
norcoclaurine-6-O-methyltransferase (6OMT) which converts (S)-norcoclaurine to (S)-coclaurine
(Inui et al. 2007; Ounaroon et al. 2003) or reticuline 7-O-methyltransferase (7OMT) that
methylates (S)-reticuline to yield (S)-laudanine (Ounaroon et al. 2003). Other OMTs involved in
BIA biosynthesis O-methylate the protoberberine backbone such as scoulerine-9-Omethyltransferase (SOMT1) which converts (S)-scoulerine to (S)-tetrahydrocolumbamine (Dang
and Facchini 2012; Takeshita et al. 1995). Acetylations by enzymes like salutaridinol 7-Oacetlytranferase (SalAT) (Grothe et al. 2001) and hydroxylations by CYPs such as (S)-Nmethylcoclaurine 3’-hydroxylase (NMCH) (Pauli and Kutchan 1998) are other forms of
decorative reactions that contribute to the structural diversity of BIAs.
1.3.2 Molecular biology and biochemistry of benzylisoquinoline alkaloids
Biochemical and molecular research on BIA biosynthesis has focused on a limited
number of plant species. Most BIA biosynthetic genes that have been isolated and functionally
characterized are from only 4 species despite the prevalence of BIAs in the plant kingdom.
Coptis japonica (Japanese goldthread) and Thalictrum flavum (Yellow meadow rue) from the
Ranunculaceae family have been adopted as model organisms, particularly in the study of
14
berberine biosynthesis. The family Papaveracea has provided two model organisms as well
Eschscholzia californica (California poppy) and Papaver somniferum (Opium poppy). Both
have been used to study sanguinarine biosynthesis and morphinan alkaloid production
respectively (Ziegler and Facchini 2008).
BIA biosynthesis begins with the stereospecific condensation of two tyrosine derivatives,
dopamine and 4-hydroxyphenylacetaldehyde (4HPAA) by norcoclaurine synthase (NCS) to form
norcoclaurine (Figure 1.6). NCS is a member of the pathogenesis-related (PR)10/Bet v 1 protein
family (Liscombe et al. 2005; Samanani and Facchini 2002; Lee and Facchini 2011). 4HPAA is
derived from tyrosine in two steps: (1) the transamination of tyrosine by tyrosine
aminotransferase (TyrAT) (Lee and Facchini 2011) and (2) the subsequent decarboxylation of 4hydroxyphenylpyruvate by 4-hydroxyphenylpyruvate decarboxylase (4HPPDC) to form 4hydroxyphenylacetaldehyde (Figure 1.6). The gene encoding 4HPPDC has not been isolated but
the enzyme has been partially purified from Berberis Canadensis callus cultures (Martina
Rueffer and Zenk 1987). Dopamine is formed from tyrosine in two distinct steps, the first being
the decarboxylation of tyrosine by tyrosine decarboxylase (TYDC) to form tyramine (Facchini
and De Luca 1994) followed by the 3’-hydroxylation of tyramine to form dopamine by an
uncharacterized enzyme referred to in the literature as 3’OHase (Figure 1.6) (Beaudoin and
Facchini 2014).
15
Figure 1.6. Biosynthetic pathways showing benzylisoquinoline alkaloids accumulating in
Papver somniferum and other plant species.
Enzymes for which a corresponding gene has been found in Papaver somniferum are shown in green. If the
gene has been isolated from another BIA producing species the enzymes are shown in blue. Enzymes for
which the corresponding gene has not been found are in black. TYDC tyrosine/DOPA decarboxylase, 3OHase
tyrosine/tyramine 3-hydroxylase, 4HPPDC, 4-hydroxyphenylpyruvate decarboxylase, NCS norcoclaurine
synthase, 6OMT norcoclaurine 6-O-methyltransferase, CNMT coclaurine N-methyltransferase, NMCH Nmethylcoclaurine 3’-hydroxylase, 4’OMT2 3’-hydroxyl-N-methylcoclaurine 4’-O-methyltransferase, BBE
berberine bridge enzyme, SOMT1 scoulerine 9-O-methyltransferase, CAS canadine synthase, TNMT,
tetrahydroprotoberberine N-methytransferase, CYP82Y1 N-methylcanadine 1-hydroxylase, NOS noscapine
synthase, STOX (S)-tetrahydroxyprotoberberineoxidase, CoOMT columbamine O-methyltransferase, CFS
cheilanthifoline synthase, SPS stylopine synthase, MSH N-methylstylopine14-hydroxylase, P6H protopine 6hydroxylase, DBOX dihydrosanguinarine oxidase, SanR sanguinarine reductase, SalAT salutaridinol 7-Oacetyltransferase, T6ODM thebaine 6-O-demethylase, COR codeinone reductase, CODM codeine Odemethylase, N7OMT norreticuline 7-O-methyltransferase, 3’OHase uncharacterized 3’-hydroxylase, 3’OMT
uncharacterized 3’-O-methyltansferase, 2OMT uncharacterized protoberine 2-O-methyltransferase. Adapted
from Beaudoin and Facchini, 2014.
16
Many BIAs are derived from the branch point metabolite (S)-reticuline which is formed
from (S)-norcoclaurine. Although the order of reactions leading to (S)-reticuline was inferred by
the substrate preference of various enzymes recent evidence suggests that the pathway is not
linear and operates as lattice (Desgagné-Penix and Facchini 2012). The first conversion is a 6-Omethylation of (S)-norcoclaurine to form (S)-coclaurine by norcoclaurine-6-O-methyltransferase
(6OMT) (Figure 1.6) (Inui et al. 2007; Ounaroon et al. 2003). (S)-coclaurine is then Nmethylated by coclaurine N-methyltransferase (CNMT) to form (S)-N-methylcoclaurine (Choi,
Morishige, and Sato 2001). The hydroxylation of (S)-N-methylcoclaurine by (S)-Nmethylcoclaurine 3’-hydroxylase (NMCH) forms (S)-3’hydroxy-N-methylcoclaurine (Pauli and
Kutchan 1998) which is subsequently converted to (S)-reticuline by 3’hydroxy-Nmethylcoclaurine 4’-O-methyltransferase (4’OMT) (Morishige et al. 2000). NMCH has very
strict stereo- and substrate specificity however, most O-methylation and N-methylation steps
exhibit more promiscuity in terms of substrate and regiospecificity which contributes to the
pathway’s lattice structure.
Two metabolic routes have been proposed for the biosynthesis of papaverine. The first
starting at (S)-reticuline followed by N-demethylation of one intermediate leading to papaverine
carried out by a hypothetical enzyme (Han et al. 2010). The second, starting at (S)-norreticuline
and travelling to papaverine thus eliminating the requirement for an N-demethylation step
(Desgagné-Penix and Facchini 2012). Biochemical support for the N-desmethyl pathway comes
from the efficient incorporation of radiolabelled N-desmethyl compounds into papaverine in
tracer studies (Brochmann-Hanssen et al. 1975; Han et al. 2010) and from the isolation and
characterization of norreticuline 7-O-methyltransferase (N7OMT) which exhibits strict substrate
17
acceptance for N-desmethyl compounds converting norrecticuline to norlaudanine which could
subsequently be converted to papaverine (Pienkny et al. 2009).
Virus-induced gene silencing (VIGS) experiments knocking down expression of several
BIA biosynthetic enzymes also supported an N-desmethyl route to papaverine (Desgagné-Penix
and Facchini 2012). The VIGS mediated reduction of CNMT transcript levels in opium poppy
plants increased papaverine levels. In addition, when transcript levels of N7OMT, the Ndesmethyl specific enzyme were reduced, they corresponded to a reduction of papaverine levels
in planta. As further evidence, when 7OMT transcript levels were knocked down there was no
change in papaverine accumulation. Despite the evidence supporting an N-desmethyl route to
papaverine a recent study using mass spectrometry and a heavy stable-isotope of (S)-reticuline
found that the some of the label was incorporated into papaverine supporting a N-methylated
pathway. Most of the label however was incorporated into morphinan alkaloids (Han et al.
2010). Another recent study compared the transcriptomes of high and low papaverine producing
varieties of opium poppy. The high papaverine plants up-regulated transcripts associated with the
N-desmethyl pathway, 6OMT, CNMT, 4’OMT and most importantly N7OMT, while 7OMT was
down regulated in comparison to low papaverine producing plants (Pathak et al. 2013). Thus it
seems the major route to papaverine is via an N-desmethyl pathway and perhaps the Nmethylated route contributes in a minor way to papaverine biosynthesis (Figure 1.6).
The final step in papaverine biosynthesis involves the oxidation of the fully Omethylated, N-desmethyl alkaloid, tetrahydropapaverine by dihydrobenzophenanthridine oxidase
(DBOX) (Figure 1.6). DBOX is also responsible for the final step in the biosynthesis of the
benzo[c]phenanthridine sanguinarine. DBOX is transcribed exclusively in the roots of opium
poppy and suggest there may be transport of papaverine from the roots to the aerial organs and
18
latex (Hagel et al. 2012). The presence of DBOX in the roots presents the possibility that other
steps of papaverine biosynthesis could occur in the roots, namely the uncharacterized 3’-Omethylation step.
Morphine biosynthesis is a rare event in the plant kingdom. Only a few plants species
from the family Papaveracea are known to make it. Although other morphinans such as
salutaridine are less rare than morphine the only other plant family that makes morphinans is
Euphorbiaceae (Theuns, Theuns, and Lousberg 2014). The biosynthesis of morphine is
predicated on the ability of the plant to epimerize (S)-reticuline to (R)-reticuline. The reaction
mechanism has been postulated to occur in two distinct steps: (1) the dehydrogenation of (S)reticuline to a 1,2-dehydroreticulinium ion (Hirata et al. 2004) and (2) the reduction of the 1,2dehydroreticulinium ion to (R)-reticuline (De-Eknamkul and Zenk 1992). The enzyme
responsible for the first step was named 1,2-dehydroreticuline synthase and has been partially
purified. The enzyme for the second step, 1,2-dehydroreticuline reductase, has been purified to
homogeneity and partial characterized. The genes corresponding to both of the enzymes have
not been identified.
The morphinan backbone is generated by the C-C phenol-coupling of (R)-reticuline by
salutaridine synthase (SalSyn, CYP719B1) to form salutaridine (Figure 1.6) (Gesell et al. 2009).
Salutaridine is converted to salutaridinol by the short-chain dehydrogenase/reductase,
salutaridine reductase (SalR) (Ziegler et al. 2006), next salutaridinol is O-acetylated by
salutatidinol 7-O-acetyltansferase (SalAT) (Grothe et al. 2001; Lenz and Zenk 1995). The
cyclization of salutaridinol 7-O-acetate to form thebaine is spontaneous under basic conditions,
pH8-9. The spontaneous cyclization of salutaridinol 7-O-acetate in basic conditions suggests the
19
reaction occurs in a basic subcellular compartment or there is an uncharacterized enzyme
involved in the cyclization process (Fisinger et al. 2007).
The morphine pathway splits at thebaine (Figure 1.6). In the major route thebaine-6-Odemethylase (T6ODM) converts thebaine to neopinone which spontaneously converts to
codeinone (Hagel and Facchini 2010). Codeinone is subsequently reduced by the aldo-keto
reductase codeinone reductase (COR) to codeine (Unterlinner, Lenz, and Kutchan 1999). The
formation of morphine from codeine is catalyzed by codeine O-demethylase (CODM).
Conversely in the minor route, CODM catalyzes the first step by converting thebaine to
oripavine. Next, T6OMD catalyzes the formation of morphinone from oripavine, which is
subsequently reduced by COR to morphine.
All protoberberine, protopine, phthalideisoquinoline and benzo[c]phenanthridine
alkaloids are ultimately derived from conversion of (S)-reticuline to (S)-scoulerine by the
berberine bridge enzyme (BBE) (Figure 1.6) (Dittrich and Kutchan 1991). (S)-Scoulerine can be
converted to (S)-tetrahydrocolumbamine by scoulerine 9-O-methyltransferase (SOMT1) (Dang
and Facchini 2012; Takeshita et al. 1995). (S)-tetrahydrocolumbamine has two possible fates, it
can be converted to (S)-tetrahydropalmatine by columbamine O-methyltransferase (CoOMT)
(Takashi Morishige et al. 2002) or a methylenedioxy bridge can be added to (S)tetrahydrocolumbamine by the CYP, canadine synthase (CAS) to form (S)-canadine (Figure 1.6)
(Diaz Chavez et al. 2011; Ikezawa et al. 2003; Winzer et al. 2012). (S)-Canadine can either lead
to noscapine as described below or (S)-canadine can yield berberine through an oxidation by the
FAD-linked enzyme dihydrobenzophenanthridine oxidase (DBOX) (Hagel et al. 2012).
Recently, a gene cluster from opium poppy was found encoding for 10 different genes purported
to be involved in noscapine biosynthesis. The function of some of the genes of the cluster were
20
investigated using VIGS (Winzer et al. 2012) and some of the cDNAs have since been isolated
and functionally characterised as described below. (S)-Canadine is N-methylated by (S)tetrahydroprotoberberine N-methyltransferase (TNMT) to form (S)-N-methylcanadine. TNMT
also accepts several different alkaloids with a protoberberine backbone (Liscombe and Facchini
2007). (S)-N-methylcanadine 1-hydroxylase (CYP82Y1) is responsible for the creation of (S)-1hydroxy-N-methylcanadine from N-methylcanadine (Dang and Facchini 2014). (S)-1-hydroxyN-methylcanadine then undergoes aliphatic and aromatic ring hydroxylations and Omethylations which lead to norotinehemiacetal which is converted to noscapine by the shortchain dehydrogenase/reductase noscapine synthase (NOS) (Figure 1.6) (Chen and Facchini
2014).
Alternatively, (S)-scoulerine can be converted to (S)-cheilanlthifoline by cheilanthifoline
synthase (CFS) a CYP (Figure 1.6) (Diaz Chavez et al. 2011). O-methylation at the 2 position of
(S)-cheilanthifoline backbone to form (S)-sinactine is an uncharacterized step in BIA metabolism
(Beaudoin and Facchini 2014). (S)-Stylopine can be derived from (S)-cheilanthifoline by
addition of another methylenedioxy bridge, the enzyme responsible for this conversion is the
CYP, stylopine synthase (SPS) (Diaz Chavez et al. 2011; Ikezawa, Iwasa, and Sato 2007). The
N-methylation of (S)-stylopine by TNMT forms (S)-cis-N-methylstylopine (Liscombe and
Facchini 2007). Protopine is formed by the 14-hydroxylation of the quaternary protoberberine
alkaloid (S)-cis-N-methylstylopine by the CYP (S)-cis-N-methylstylopine 14-hydroxylase
(MSH) which leads to the subsequent ring tautomerization by breaking the C-N bond and
forming a keto moiety at C14 (Figure 1.6) (Beaudoin and Facchini 2013). T6ODM, CODM and
other ODDs have been implicated in protopine alkaloid regulation through the oxidative
21
dealkylation of methylenedioxy bridges or methoxy groups on the protopine backbone (Farrow
and Facchini 2013).
All benzo[c]phenanthridine alkaloids are derived from protopine with the exception of
chelerythrine which is derived from allocryptopine. The formation of dihydrosanguinarine from
protopine is catalyzed by protopine 6-hydroxylase (P6H) another CYP (Figure 1.6) (Takemura et
al. 2013). The 6-hydroxylation of protopine results in a spontaneous rearrangement of 6hydroxyprotopine to dihydrosanguinarine. The oxidation of dihydrosanguinarine to sanguinarine
is catalyzed by the FAD-linked enzyme dihydrobenzophenanthridine oxidase (DBOX) (Hagel et
al. 2012). The reduction of sanguinarine back to dihydrosanguinarine is catalyzed by
sanguinarine reductase (SanR) (Vogel et al. 2010; Weiss et al. 2006). The reductive process is
thought to have occurred in order to reduce the cytotoxic effect of sanguinarine for the less toxic
dihydrosanguinarine.
Rhoeadine alkaloids are only known to occur in members of Papaveracea (Shulgin and
Perry 2002). The origins of rhoeadine alkloids in planta were discovered by feeding labelled
protopine to Papaver bracteatum (Ronsch 1972). The suppression of gene transcripts encoding
CODM, responsible for the O-demethylation and/or the O,O-demethylenation of morphinan and
protopine alkaloids, by VIGS resulted in elevated levels of rhoeadine alkaloids (Farrow and
Facchini 2013). The enzymatic steps involved in the conversion of protopines to rhoeadine
alkaloids are not known.
1.3.3 The catalytic mechanism of OMTs
The widespread belief is that the ability to produce BIAs in angiosperms arose from a
monophyletic origin (Liscombe et al. 2005). One result of the monphyletic evolution of BIA
metabolism is the incredible structural diversity of BIAs is the result of remarkably few enzyme
22
types. BIA metabolism dominated by enzymes of these categories: cytochrome P450s,
reductases, oxidases, acetyltransferases, a single Pictet-Spenglerase, N-methyltransferases and,
O-methyltransferases.
SAM-dependant O-methylation which catalyze the regiospecific transfer of a methyl
group from SAM to a free hydroxyl moiety on the BIA backbone are widespread in BIA
metabolism (Beaudoin and Facchini 2014). Genes encoding O-methyltransferases involved in
BIA biosynthesis have been isolated from several plants that produce BIAs and often share
significant sequence similarity (Beaudoin and Facchini 2014). Indeed O-methylation is a
common reaction occurring in many distinct biological system (Schubert, Blumenthal, and
Cheng 2003). Yet in most plants, O-methylation is performed by SAM-dependant OMTs. SAM
is used a the methyl donor and over the course of the reaction is converted to S-adenosyl-Lhomocysteine. These enzyme are very diverse in terms of substrate preference yet all retain the
important SAM-binding domain (Zubieta et al. 2001). The mechanism of O-methylation in
plants is thought to be conserved. OMTs perform a SN2 reaction with the help of a catalytic triad
(Brandt, Manke, and Vogt 2015). The substrate binding pocket positions the substrates free
hydroxyl group near the activated methyl of SAM and the amino group of a near by His. Two
Glu residues bracket the His and ensure the proper orientation of histidine’s imidazole ring
towards the substrate through hydrogen bonding. His acts as a catalytic base and deprotonates
the hydrogen on the substrates free hydroxyl. The negatively charged oxygen that was
deprotonated acts as a nucleophile and attacks the methyl carbon on SAM. This results in the
SN2 transfer of the SAM methyl group to the substrate (Brandt, Manke, and Vogt 2015;
Schubert, Blumenthal, and Cheng 2003; Zubieta et al. 2001).
23
1.4 Approaches to gene discovery and functional characterization of alkaloid biosynthetic
genes
1.4.1 Traditional approaches to gene identification
Several BIA biosynthetic genes have been isolated and characterized using classical
biochemistry. Some such examples are the isolation of cDNA encoding for BBE from E.
califormica (Dittrich and Kutchan 1991) and SOMT, 6OMT and 4’OMT were isolated from C.
japonica (Takeshita et al. 1995; T Morishige et al. 2000). In most cases, a specific biosynthetic
enzyme was purified to homogeneity from a cell culture suspension or plant tissue. Next the
purified protein would be digested with a protease like trypsin, the amino acid sequences would
be sequenced and the sequences of the peptide fragments were used to design degenerate
oligonucleotide primers. The fragments of the corresponding gene were amplified by
polymerase chain reaction (PCR). Amplified DNA fragments from the corresponding to the
enzyme of interest were then used as probes to isolate a full-length molecular clone from a
cDNA library.
1.4.2 Genomics-based gene discovery approaches
The advent of genomic technologies has led to several new approaches for the discovery
of BIA biosynthetic genes and has aided in the efficient isolation of new genes (Goossens and
Rischer 2007). The use of Sanger sequencing to develop expressed sequence tag (EST)
collections streamlined the gene discovery process. ESTs represent the transcribed genes
derived from the large-scale random sequencing of a cDNA library from a specific plant species
and tissue type. The resulting sequences are organized and managed in a computer database and
the library can be mined for gene candidates using sequence similarity to a reference gene of
24
known function with Basic Local Alignment Search Tool (BLAST) (Altschup et al. 1990).
Candidate genes can be subsequently amplified from appropriate cDNA libraries and expressed
in bacteria or yeast.
Sanger-based EST projects have yielded over 25 000 ESTs from various BIA-producing
tissues of Papaver somniferum (Facchini et al. 2007). Similar initiatives have led to the isolation
of various BIA biosynthetic genes. For example, the isolation and characterization of 7OMT and
6OMT from opium poppy (Ounaroon et al. 2003) as well as 4’OMT from opium poppy (Ziegler
et al. 2005).
Recently high-throughput next-generation sequencing (NGS) technologies such as
Roche-454 and Illumina have changed the paradigm of functional genomics and have proven to
be fast and cost-effective methods of generating deep transcriptomic datasets (Xiao, Zhang,
Chen, Lee, Barber, Chakrabarty, Desgagné-Penix, et al. 2013). Nonetheless, NGS approaches
have the draw backs of shorter reads, higher base-call error rate and non uniform coverage when
compared to more traditional sequencing methods (C. Chen et al. 2014). Often genomic datasets
generated by NGS are mapped by comparing the dataset to a genomic dataset for a similar
species which is called a reference sequence in order to make the computational task of creating
a de novo library. In situations where there are no reference sequences available, as is the case
with many non-model species like opium poppy, RNA-Seq is used to develop a transcriptomic
dataset (Nakamura et al. 2011). NGS technologies have been used to develop RNA-Seq based
transcriptomic databases for many different plants producing specialized metabolites. This
undertaking has led to the discovery of several novel biosynthetic genes from several plant
species involved in specialized metabolism, some examples include the discovery of two unique
sesquiterpene synthases from the root of Valeriana officinalis (Pyle et al. 2012), a novel
25
sesquiterpene synthase involved the biosynthesis of the natural sweetener, hernandulcin from
Lippia dulcis (Attia, Kim, and Ro 2012) and an O-methyltransferase and a short-chain
dehydrogenase/reductase involved in noscapine biosynthesis (Dang and Facchini 2012; Chen and
Facchini 2014).
1.4.3 Integration of targeted metabolomics and transcriptomics for gene discovery
Targeted metabolomic strategies have also been used to help identity genes involved in
plant specialized metabolism (Desgagné-Penix et al. 2012; Farrow, Hagel, and Facchini 2012;
Le, Mccooeye, and Windust 2012). “Soft” ionization techniques such as electrospray ionization
tandem mass spectrometry (ESI-MSn) has been widely employed to characterize plant
specialized metabolites owing to its high sensitivity, rapid analysis time, low levels of sample
consumption and ability to provide structural information of the analytes investigated
(Desgagné-Penix et al. 2012; Farrow, Hagel, and Facchini 2012; Le, Mccooeye, and Windust
2012; Stevens, Reed, and Morré 2009; W. Wu et al. 2005). When a targeted metabolite profile is
integrated with a NGS transcriptomic database a powerful biochemical genomics tool to identify
novel genes is created (Desgagné-Penix et al. 2012). When this methodology is extended to
different varieties of the same species or of different species producing similar compounds,
choosing gene candidates is streamlined. Gene candidates can be ranked in terms of their
predicted relative expression using fragments per kilobase of exon per million fragments mapped
(FPKM) as mRNA levels are often correlated with protein abundance (C. Chen et al. 2014). For
example consider a comparative analysis of unigenes annotated as O-methyltransferases across 5
different species. Of these 5 species only one produces papaverine. Searching for OMTs based
on sequence similarity will yield a two lists of genes annotated as OMTs. One list will be OMTs
expressed in all 5 of the species and the other list will be OMTs that are expressed exclusively in
26
the papaverine producing plant. The second list will constitute strong candidate genes for an
OMT involved in papaverine biosynthesis.
1.4 Objectives
The primary objective of this study was to discover and functionally characterize a novel
gene involved in BIA biosynthesis using an integrated transcriptomic and metabolomic
approach.
Specific objectives were:
1. To isolate and characterize a molecular clone encoding a 3’-Omethyltranferase involved in papaverine biosynthesis from opium poppy.
2. To generate targeted alkaloid profiles of 20 different BIA producing
species using a triple quadrupole (QqQ) liquid chromatography-mass
spectrometry (LC-MS).
Chapter Two describes the materials and methods used for the studies described in the
subsequent chapters.
Chapter Three reports the isolation and functional expression of a molecular clone
encoding 2-protoberberine O-methyltransferase (2OMT).
The in vitro characterization and
enzyme kinetics of 2OMT. In addition, the relative expression of 2OMT was investigated across
different opium poppy tissue type by quantitative PCR (qPCR).
27
Chapter Four reports the targeted alkaloid profiles of 20 different BIA producing species
by QqQ LC-MS.
The goal was to aid in the development of an integrated
transcriptomic/metabolomic database for 20 different species to aid in the discovery of BIA
biosynthetic genes.
Chapter 5 discusses the importance of the finding from the previous chapters and suggest
ideas for future research.
28
CHAPTER TWO: MATERIALS AND METHODS
2.1 Nucleic Acid Isolation and analysis
The isolation or RNA from opium poppy cultivars Roxanne, Veronica and Bea’s Choice
root, stem, capsule, leaf and bud was performed using a previously described CTAB (cetyl
triethylammonium bromide) extraction method (Cairney, Puryear, and Chang 1993). Opium
poppy cDNA libraries were constructed using the isolated RNA in conjunction with the cDNA
library construction kit, SuperScript III First-Strand Synthesis System (Invitrogen, Carlsbad,
CA).
2.1.1 Isolation of cDNAs encoding putative OMTs from opium poppy stem
OMT gene candidates were identified using opium poppy stem Illumina Velvet Oases
(San Diego, CA) and Roche 454 MIRA (Branford, CT) Next-Generation sequencing libraries
(Citation). Known SAM-dependent OMT genes involved in BIA metabolism were used as
BLAST queries. Queries were: 6OMT from P. somniferum, C. Japonica and E. californica
(Ounaroon et al. 2003; Inui et al. 2007); 4’OMT1/2 from C. japonica and P. somniferum
(Morishige et al. 2000; Facchini and Park 2003); 7OMT from P. somniferum (Ounaroon et al.
2003); N7OMT from P. somniferum (Pienkny et al. 2009); SOMT1 from P. somniferum (Dang
and Facchini 2012). Based on their sequence similarity to known OMTs involved in BIA
biosynthesis 6 gene candidates were selected, IzOMT1-6.
2.1.2 Isolation of cDNAs encoding putative OMTs from opium poppy root
OMT gene candidates were identified using opium poppy root Illumina Velvet Oases
(San Diego, CA) and Roche 454 MIRA (Branford, CT) Next-Generation sequencing libraries.
29
Known SAM-dependent OMT genes involved in BIA metabolism were used as BLAST queries.
Queries were: 6OMT from P. somniferum, C. Japonica and E. californica (Ounaroon et al. 2003;
Inui et al. 2007); 4’OMT1/2 from C. japonica and P. somniferum (T Morishige et al. 2000;
Facchini and Park 2003); 7OMT from P. somniferum (Ounaroon et al. 2003); N7OMT from P.
somniferum (Pienkny et al. 2009); SOMT1 from P. somniferum (Dang and Facchini 2012).
Based on their sequence similarity to known OMTs involved in BIA biosynthesis and their
absence in stem transcriptomic databases, 4 gene candidates were selected, DDOMT1-4.
2.2 Phylogenetic analysis
Phylogenetic analysis based on putative amino acid sequence was performed with
ClustalW (Chenna et al. 2003) with a BLOSUM cost matrix. The phylogenetic tree was
constructed using Geneious Tree Builder (Biomatters; Newark, NJ; http://www.geneious.com),
using the Jukes-Cantor model (Jukes and Cantor 1969), neighbour-joining tree build method and
bootstrap resampling. Gap open cost: 10 and Gap extend cost: 0.1. Number of random seed was
946 361 and the number of replicates was 1000. Protein abbreviations and GeneBank accession
numbers are listed in Table 2.1 below.
30
Table 2.1 Protein abbreviations and GenBank accession numbers for sequences used in
phylogenetic analysis of SAM-dependent O-methyltransferases from selected plants.
Abbreviation
Protein name
Accession #
Ec7OMT
CjCoOMT
TtOMT
PsCaOMT
Ps7OMT
PsN7OMT
Ps6OMT
Ps4’OMT2
Ps4’OMT1
Cc4’OMT
Cj4’OMT
Tf4’OMT
Cj6OMT
Tf6OMT
VvReOMT
PtFlOMT
CjSOMT
PaCafOMT
CaCafOMT
ObEuOMT
MpFlOMT
ObCafOMT
CbEuOMT
CbCafOMT
AmCafOMT
PsSOMT1
PsSOMT2
PsSOMT3
Eschscholzia californica, reticuline 7OMT
Coptis japonica, columbamine OMT
Thalictrum tuberosum, catechol OMT
Papver somniferum, catchol OMT
Papaver somniferum, reticuline 7OMT
Papaver somniferum, norreticuline 7OMT
Papaver somniferum, norcoclaurine 6OMT
Papaver somniferum, 3’-hydroxy-N-methylcoclaurine 4’OMT2
Papaver somniferum, 3’-hydroxy-N-methylcoclaurine 4’OMT1
Coptis chinensis, 3’-hydroxy-N-methylcoclaurine 4’OMT
Coptis japonica, 3’-hydroxy-N-methylcoclaurine 4’OMT
Thalictrum flavum, 3’-hydroxy-N-methylcoclaurine 4’OMT
Coptis japonica, norcoclaurine 6OMT
Thalictrum flavum, norcoclaurine 6OMT
Vitis vinifera, resveratrol OMT
Populus trichocarpa, flavonoid OMT predicted protein
Coptis japonica, scoulerine 9OMT
Picea abies, caffeate OMT
Capsicum annuum, caffeate OMT
Ocimum basilicum, engenol OMT
Mentha X piperitta, Flavonoid 8OMT
Ocimum basilicum, caffeate OMT
Clarkia breweri, (iso)eugenol OMT
Clarkia breweri, caffeate OMT
Ammi majus caffeate OMT
Papaver somniferum, scoulerine 9OMT1
Papaver somniferum, narcotoline OMT2
Papaver somniferum, scoulerine OMT3
BAE79723.1
Q8H9A8.1
AF064697.1
AY268895.1
Q6WUC2.1
AC88562.1
AAP45315.1
AAP45314.1
AAP45314.2
ABY75613.1
Q9LEL5.1
AAU20768.1
Q9LEL6.1
AAU20765.1
CAQ76879.1
XP_002312933.1
Q39522.1
CAI30878.1
AAG43822.1
AAL30424.1
AAR09600.1
AAD38189.1
AAC01533.1
O23760.1
AAR24095.1
JN185323.1
JN185324.1
JN185325.1
31
2.3 Construction of expression vectors and heterologous expression
2.3.1 Stem OMT gene candidates IzOMT1 through IzOMT6
PCR was used to amplify open reading frames (ORFs) encoding selected proteins from P.
somniferum Bea’s choice cultivar stem cDNA using Green Taq DNA Polymerase (Genescript,
Piscataway, NJ), forward primers contained a BamHI restriction site and the reverse primers
contained a Xho1 restriction site. Primer sequences are listed in Table 2.2. PCR products were
ligated into pGEM-T Easy vectors (Promega, Madison, WI). Constructs were subsequently
transformed into XL1-Blue competent E. coli cells by heat shocking according to the
manufacturer’s protocol (New England Biolabs, Ipswich, MA). After an overnight incubation at
37 ºC on Lauria-Bertani (LB) agar plates supplemented with 100 µg/mL ampicillin, 30 µg/mL
X-gal and 0.1 mM isopropyl ß-D-1-thiogalactopyranoside (ITPG) transformed colonies were
selected by blue/white colony screening. White colonies were used to inoculate 2 mL of LB
with 100 µg/mL ampicillin and grown overnight at 37 ºC. Plasmids were purified using the
High-Speed Plasmid Mini Kit (Geneaid, New Taipei City, TW) and sent for sequencing to
confirm the insert. Upon confirmation of the gene sequence the inserts were excised using
BamHI and Xho1 (New England Biolabs, Ipswich, MA) and ligated into pET29b (Novagen,
Madison, WI) using the engineered restriction sites.
E. coli Arctic Express RP (DE3) competent cells (Agilent, Santa Clara, CA) were
transformed with expression vectors, grown at 37 ºC in LB medium supplemented with 50
µg/mL kanamycin and 34 µg/mL chloramphenicol to A600 0.8. Cultures were induced with
0.1mM IPTG for 4 h at 37 ºC and at 25 ºC as well as 24 h and 48 h at 4 ºC and harvested by
32
centrifugation at 12000 RPM for 5 min. Recombinant proteins were detected in bacterial
extracts with Coomassie brilliant blue following separation by SDS-PAGE.
Table 2.2. Primer sequences used to amplify gene candidates from stem and root opium
poppy cDNA
ORF
2OMT
Forward Primer
Reverse Primer
GGATCCAATGGATATTGCAGAAGAAAGGTTGA
CTCGAGTTCTGGAAAAGCCTCGAT
DDOMT2
GGATCCGATGACTATGAATGGGAAT
CTCGAGTTTATGAAACTCAATGAGATGAAGTC
DDOMT3
GGATCCTATGCTTGACCGTATGTTG
CTCGAGATTCTTGTGGCACTCCATAAT
DDOMT4
GGATCCCATGGATATCAAATTAGAAGATGAAGAA
CTCGAGTGGATATGCAACAATAACTGATTGAAT
IzOMT1
GATGGATCCAATGGATATCAAATTAGAAGATG
CTACTCGAGTGGATGTGCAACAATAACTG
IzOMT2
GATGGATCCAATGGAAATCAAACTAGAAGATC
CTACTCGAGAGGATATGCGACAATAACC
IzOMT3
GATGGATCCAATGATGGCTAATGACTCTCGC
GTGCTCGAGAGGATATACCTCAATAAGGGAAACG
IzOMT4
GATGGATCCAATGTATTATGAGAGGTCTTCAG
CTACTCGAGATCCTTAGATGGTACTGC
IzOMT5
GATGGATCCAATGGGTTCAGGCCATCACAGC
CTACTCGAGATTCTTGTGGCACTCCCATAATCC
IzOMT6
GATGGATCCAATGGGTTCGATCCAGAACGTAG
CTACTCGAGGTTTTTGTAGAATTCCATAACG
Primers used to amplify ORFs from cDNA for ligation to pRSETA
33
2.3.2 Root OMT gene candidates DDOMT1-4
PCR was used to amplify open reading frames (ORFs) encoding DDOMT3 and
DDOMT4 from P. somniferum Bea’s choice cultivar root cDNA using Green Taq DNA
Polymerase (Genescript, Piscataway, NJ), forward primers contained a BamHI restriction site
and the reverse primers contained a Xho1 restriction site. Primer sequences are listed in Table X.
DDOMT1 and DDOMT 2 ORFs were commercially synthesized with a BamH1 restriction site
at the 5’ end of the gene and a Xho1 restiction site at the 3’ end of the gene in the pUC57a vector
(Genscript, Piscataway, NJ). PCR products were ligated into pGEM-T Easy vectors (Promega,
Madison, WI). Constructs for the amplified gene candidates and the synthesized gene candidates
were subsequently transformed into XL1-Blue competent E. coli cells by heat shocking
according to the manufacturer’s protocol (New England Biolabs, Ipswich, MA). After an
overnight incubation at 37 ºC on Lauria-Bertani (LB) agar plates supplemented with 100 µg/mL
ampicillin, 30 µg/mL X-gal and 0.1 mM isopropyl ß-D-1-thiogalactopyranoside (ITPG)
transformed colonies were selected by blue/white colony screening. White colonies were used to
inoculate 2 mL of LB with 100 µg/mL ampicillin and grown overnight at 37 ºC. Plasmids were
purified using the High-Speed Plasmid Mini Kit (Geneaid, New Taipei City, TW) and sent for
sequencing to confirm the insert. Upon confirmation of the gene sequence the inserts were
excised using BamHI and Xho1 (New England Biolabs, Ipswich, MA) and ligated into pET29b
(Novagen, Madison, WI) using the engineered restriction sites.
E. coli Arctic Express RP (DE3) competent cells (Agilent, Santa Clara, CA) were
transformed with expression vectors, grown at 37 ºC in LB medium supplemented with 50
µg/mL kanamycin and 34 µg/mL chloramphenicol to A600 0.8. Cultures were induced with 0.1
mM IPTG for 4 h at 37 ºC and at 25 ºC as well as 24 h and 48 h at 4 ºC and harvested by
34
centrifugation at 12 000 RPM for 5 min at 4 ºC. Recombinant proteins were detected in
bacterial extracts with Coomassie brilliant blue following separation by SDS-PAGE.
2.4 Purification of recombinant proteins and analysis
2.4.1 Purification of recombinant IzOMT 1-6 and DDOMT 1-4
1 L LB cultures with 50 µg/mL kanamycin and 34 µg/mL chloramphenicol were
inoculated with E. coli Arctic Express RP cells containing pET29b constructs containing ORFs
for IzOMT 1-6 and DDOMT 1-4 and grown to A600 0.8 at 37 ºC 200 RPM. LB cultures were
induced with 0.1 mM IPTG and transferred to 4 ºC 180 RPM for 48 hours. Cultures were
centrifuged at 12 000 RPM for 5 min at 4 ºC to pellet the cells. Cells were resuspended in 10 mL
of protein extraction buffer, 50 mM NaH2PO4, pH 7.5, 10% glycerol, 750 µg/mL lysozyme from
chicken egg white (Sigma-Aldrich, St. Louis, MO), 1 mM ß-mercaptoethanol (Sigma-Aldrich,
St. Louis, MO) and 1% (v/v) Tween 20 (Sigma-Aldrich, St. Louis, MO). Cells were lysed by
sonication. Cell debris was removed by centrifugation and the supernatant was bound to Talon
cobalt affinity resin (Clonetech, Mountain View CA) by shaking on ice for 60 min, washed three
times with protein purification buffer, 50 mM NaH2PO4, pH 7.5, 10% glycerol, 750 µg/mL
lysozyme from chicken egg white (Sigma-Aldrich, St. Louis, MO), 1 mM ß-mercaptoethanol
(Sigma-Aldrich, St. Louis, MO) and 1% (v/v) Tween 20 (Sigma-Aldrich, St. Louis, MO).
Proteins were eluted with 1 mL aliquots of protein purification buffer containing increasing
concentration of imidazole (5, 50, 100 and 200 mM). Protein elutions were desalted on PD-10
columns (Amersham Biosciences, Piascataway, NJ).
35
2.4.2 SDS-PAGE
Proteins were fractionated using 12% polyacrylamide SDS-PAGE with the MiniPROTEAN Tetra Cell (Bio-Rad Laboratories, Mississauga. ON) following the instructions of the
manufacturer.
2.4.3 Western Blot Analysis
Soluble proteins were separated by SDS-PAGE and transferred to BioTrace NT
nitrocellulose membranes (Pall Life Sciences, Pensacola, FL). Protein blots were incubated with
1º antibody, anti-His tag mouse (Genscript, Piscataway, NJ) diluted to 1:1000 in blocking buffer
(137 mM NaCl, 10 mM Na2HPO4, 2.7 mM KCl, 1.8 mM KH2PO4, 0.1% (v/v) Tween 20 and 5%
(w/v) skim milk powder) for 12 hours. Following the incubation the membranes were washed in
1x PBS with 0.1% Tween 20 (137 mM NaCl, 10 mM Na2HPO4, 2.7 mM KCl, 1.8 mM KH2PO4,
0.1% (v/v) Tween 20) 4 times for 10 minutes each. Membrane was subsequently incubated with
2º antibody, goat anti-mouse antibody, horseradish peroxidase (HPR) conjugated (Bio-Rad
Laboratories, Mississauga. ON) diluted with 1:10 000 in blocking buffer for 1 hour. Membrane
was washed in 1x PBS with 0.1% Tween 20 again the same way as above. Proteins were
visualized with SuperSignal West Pico Chemiluminescent Substrate (Thermo Pierce, Rockfort,
IL).
2.5 Recombinant OMT Candidate Assays
2.5.1 Routine Enzyme assays
Standard in vitro enzyme assays were performed using 4 µg of purified recombinant
protein in a 50 µL reaction of 50 mM glycine pH 9.0, 500 µM S-adenosyl methionine, 50 µM
substrate and 50 mM ß-mercaptoethanol which were incubated for 2 hours at 30 ºC. Assays
36
were stopped with the addition of 50 µL of methanol. The reactions were then dried down in a
Speed-Vac concentrator (Savant, Ramsey, MN) and resuspended in 50 µL of 1% (v/v) formic
acid.
2.5.2 DDOMT1 Temperature and pH Optima assay
Temperature optimum assays were conducted as described in 2.4.1 with a few alterations.
Scoulerine was used as the substrate and assays were incubated at 4, 20, 25, 30, 37, 42 and 55 ºC
for 2 hours. All assays were quenched with 50 µL of methanol, dried down in a Speed-Vac
concentrator (Savant, Ramsey, MN) and resuspended in 50 µL of 1% (v/v) formic acid.
pH optimum assays were performed at pH 5 (citrate), pH 6 (KPO4-), pH 7 (KPO4-), pH 8
(Tris-HCl) and pH 9 (glycine). A 50 µM concentration of scoulerine was used in all assays.
Assays were incubated at 30 ºC for 2 hours and quenched with 50 µL of methanol. Assays were
dried down in a Speed-Vac concentrator (Savant, Ramsey, MN) and resuspended in 50 µL of 1%
(v/v) formic acid.
2.5.3 DDOMT1 Km Assays
The Michealis constant (Km) was determined by making enzymatic assays with 50 mM
glycine pH 9.0, 500 µM S-adenosyl methionine, 50 mM ß-mercaptoethanol and 0.08 µg/µL of
purified recombinant protein which were incubated for 2 hours at 30 ºC with varying
concentrations of scoulerine from 1 to 500 µM. Enzyme assays were stopped with equal
volumes of methanol and dried down in a Speed-Vac concentrator (Savant, Ramsey, MN).
Dried reaction were then resuspended in 1% formic acid to give a final total alkaloid
concentration of 1 µM across all samples.
37
2.6 LC-MS conditions for enzyme assay analysis
2.6.1 LC conditions for enzyme assays
LC analysis was performed with a 1200 liquid chromatograph coupled to a 6410 triple
quadrupole mass spectrometer (Agilent, Santa Clara, CA). Solvent A, was 1% aqueous formic
acid and solvent B was 100% acetonitrile. The chromatography column used was a Poroshell
120 SB C18 column (2.1 X 50 mm, 2.7 µm particle size; Agilent, Santa Clara, CA) with a flow
rate of 0.7 mL/min and a temperature of 55 ºC. The column was equilibrated in solvent A and
the elution conditions were as follows: Each extract was injected onto the column with a volume
of 10 µL. Compounds were eluted using a flow rate of 0.7 mL/min using a gradient of 1%
HCOOH (solvent A, HCOOH:H2O) and C2H3N (solvent B). Initial HPLC conditions were
100% solvent A changing linearly to 60% (v/v) solvent A over 6 min, and then to 1% (v/v)
solvent A by 7 min min. Mobile phase constituents remained constant for 1 min and then
returned to starting conditions at 8.1 min for a 4-min re-equilibration period. Total analysis time
was 12.1 min per sample.
2.6.2 Mass analyzer conditions
Samples were introduced to the mass analyzer via an electrospray ionization (ESI) probe
inlet with a capillary voltage of 4000 kV, a fragmentor voltage of 100V, a gas temperature of
350 ºC, the gas flow was set to 10 L/min and the nebulizer pressure was 50 psi. Mass
spectrometry data was acquired in positive ion mode.
Full scan analysis were performed with quadrupole 1 and 2 set to RF only while
quadrupole 3 scanned a variable mass range (100 – 400 m/z).
Collisionally-induced dissociation (CID) spectra were recorded using a collision
energy of 25 eV applied to quadrupole 2. MS2 fragments were analyzed in quadrupole 3 by
38
scanning from m/z 40 to m/z 2 greater than the precursor ion. Resulting spectra and retention
times were compared to standards when available and previously published spectra for
identification purposes.
2.7 Real-time quantitative PCR (qPCR)
Real-time quantitative PCR was performed on cDNA synthesized at 30 ng/µL from Bea’s
Choice variety opium poppy stem, root, capsule, leaf and bud tissues. Each 10 µL reaction
contained approximately 1.5 ng/µL of cDNA, 1X KAPA SYBR FAST qPCR kit (Kapa
Biosystems, Boston, MA) and 200 nM of forward and reverse primers specific to 2OMT.
Sequences of the primers for 2OMT and ubiquitin 10 are listed in Table 2.3. Analysis was
performed using a 7300 Real-Time PCR System (Applied Biosystems Life Technologies,
Burlington, ON). The AAC method was used to determine the relative gene expression levels
with ubiquitin as an endogenous control.
Table 2.3. Real-Time quantitative PCR Primers used to study the relative gene expression
of 2OMT in different tissue types from opium poppy.
Abbreviations: 2OMT, protoberberine 2-O-methyltranferase; UBQ 10, ubiquitin 10.
Gene
Forward Primer
Reverse Primer
2OMT
AAATGCGCTGTTGAACTTGGT
TGTTGATGATCTCCGACATAGTGA
UBQ 10
GGGAACACAAACGACACCAAA
TCGTCTTCGTGGTGGTAACTAGAG
39
2.8 LC-MS based targeted alkaloid profiling of 20 BIA producing plants species
2.8.1 Plants
Selected tissues were harvested at the outdoor Jardin Botanique de Montréal (Montréal,
Québec; http://espacepourlavie.ca) from the plants Hydrastis canadensis, Sanguinaria
canadensis, Nigella sativa, Mahonia aquifolium, Menispermum canadense, Stylophorum
diphyllum, and Xanthoriza simplicissima. Chelidonium majus, Papaver bracteatum, Argemone
mexicana, Eschscholtzia californica, Nandina domestica, Glaucium flavum, Thalictrum flavum
and Corydalis cheilanthifolia were grown from seed germinated in potted soil under standard
open air greenhouse conditions at the University of Calgary (Calgary, Alberta). Jeffersonia
diphylla and Berberis thunbergii plants were purchased from Plants Delights Nursary (Raleigh,
North Carolina; www.plantdelights.com) and Sunnyside Greenhouses (Calgary, Alberta;
www.sunnysidehomeandgarden.com), respectively. All seeds were purchases from Band T
World Sees (http://b-and-t-world-seeds.com) with the exception of T. flavum and P. bracteatum
which were purchased from Jelitto Standensamen (www.jelitto.com) and La Vie in Rose
Gardens (www.lavieenrosegardens.com) respectively. Callus cultures of Cissampelos
mucronata, Cocculus trilobus, and Tinospora cordifolia were purchased from Deutsche
Sammlung von Mikroorganismen und Zellkulturen (DSMZ, Braunschweig, Germany;
http://www.dsmz.de).
2.8.2 Chemical Standards
(S)-coclaurine was purchased from Toronto Research Chemicals (Toronto, ON;
http://www.trc-canada.com). (S)-Reticuline was a gift from Tasmanian Alkaloids (Westbury,
Australia; http://www.tasalk.com.au/). Morphine and codeine were gifts from Sanofi-Aventis
(Paris, France; http://en.sanofi-aventis.com). Allocryptopine, (S)-scoulerine and (S)-canadine
40
were purchased from Chromadex (Irvine, CA; http://www.chromadex.com). (S)-Isocorydine,
(S)-boldine, (S)-corytuberine, and (S)-glaucine were from Sequoia Research Products (St. James
Close, UK; http:// http://www.seqchem.com/). Cryptopine was purchased from MP Biomedicals
(Santa Ana, CA; http://www.mpbio.com).
Thebaine, oripavine were prepared from P. somniferum latex as described previously
(Hagel and Facchini 2010). Stylopine was produced synthetically from berberine as described
previously (Liscombe and Facchini 2007). Cheilanthifoline was produced enzymatically from
scoulerine using recombinant cheilanthifoline synthase and purified by TLC as described by
Diaz Chavez et al. 2011.
2.8.3 Extraction of alkaloids from BIA producing plant species
Tissue was harvested from plants and flash frozen in liquid nitrogen. Frozen tissue
samples were ground with a TissueLyser II (Qiagen, Venlo, NL) using 30 mL liquid nitrogen
cooled stainless steel grinding jars with 20 mm grinding balls (Retsch, Haan, DE). Samples were
lyophilized until dry.
Ground, freeze-dried tissue was mixed with Bieleski’s solution (15:1:4) (v/v), methanol,
formic acid and water, to a concentration of approximately 30 mL Bieleski’s to each gram of
dried plant tissue. Solutions were sonicated at full power for 1 min in an ice water bath. Tubes
were shaken at 200 RPM at 4 ºC for 2 hours and then left at -20 ºC for 18 hours to passively
diffuse. Debris was removed by centrifugation at 14 000 g for 10 min at 4 ºC. The resulting
supernatants were removed from the cell debris and filtered through 22 µm Millex filters (EMD
Millipore, Billerica, MA) into pre-weighed tubes. Samples were dried down in a Speed-Vac
concentrator (Savant, Ramsey, MN) and the total mass of alkaloid extract was determined with
an analytical balance. Dried total alkaloid extracts were subsequently reconstituted in 10 mM
41
ammounium acetate, 5% acetonitrile at pH 5.5 to a concentration of 5 mg/mL. The prepare the
alkaloids extract for analysis, the reconstituted alkaloid extracts were diluted 1:10 and 1:100 in
glass auto-sampler vials in 10 mM ammounium acetate, 5% acetonitrile at pH 5.5.
2.8.4 LC conditions for targeted alkaloid profiling
LC analysis was performed with a 1200 liquid chromatograph coupled to a 6410 triple
quadrupole mass spectrometer (Agilent, Santa Clara, CA). Solvent A, was 10 mM ammounium
acetate, 5% acetonitrile at pH 5.5 and solvent B was 100% acetonitrile. The chromatography
column used was a Zorbax Eclipse Plus C18 column (2.1 X 50 mm, 1.8 µm particle size (Agilent,
Santa Clara, CA) with a flow rate of 0.5 mL/min and a temperature of 45 ºC. The column was
equilibrated in solvent A and the elution conditions were as follows: Each extract was injected
onto the column with a volume of 10 µL. Compounds were eluted using a flow rate of 0.5
mL/min using a gradient of 10 mM C2H3O2NH4 (solvent A, C2H3O2NH4:C2H3N; pH5.5) and
C2H3N (solvent B). Initial HPLC conditions were 100% solvent A changing linearly to 50%
(v/v) solvent A over 10 min, and then to 1% (v/v) solvent A by 12 min. Mobile phase
constituents remained constant for 1 min and then returned to starting conditions at 13.1 min for
a 4-min re-equilibration period. Total analysis time was 18.1 min per sample.
2.5.2 Mass analyzer conditions for targeted alkaloids profiling
Samples were introduced to the mass analyzer via an electrospray ionization (ESI) probe
inlet with a capillary voltage of 4000 kV, a fragmentor voltage of 100V, a gas temperature of
350 ºC, the gas flow was set to 10 L/min and the nebulizer pressure was 50 psi. Mass
spectrometry data was acquired in positive ion mode.
42
Full scan analysis were performed with quadrupole 1 and 2 set to RF only while
quadrupole 3 scanned a variable mass range (200 – 700 m/z). From these experiments a list of
m/z found in the alkaloid extracts was compiled and were used to establish subsequent CID
experiments.
Collisionally-induced dissociation (CID) spectra were recorded using a collision energy
of 25 eV applied to quadrupole 2. MS2 fragments were analyzed in quadrupole 3 by scanning
from m/z 40 to m/z 2 greater than the precursor ion m/z. Resulting spectra and retention times for
the alkaloid standards were used to identify alkaloids from the 20 BIA producing species.
43
CHAPTER THREE: THE ISOLATION AND FUNCTIONAL EXPRESSION OF A
MOLECULAR CLONE ENCODING 2-PROTOBERBERINE OMETHYLTRANSFERASE
3.1 Introduction
Although benzylisoquinoline alkaloids accumulate in diverse taxa of angiosperms they
are most common among the orders Papaveracea, Ranunculaceae, Berberidaceae and
Menispermaceae (Ziegler and Facchini 2008). BIAs and other alkaloids are known to contribute
to the reproductive fitness of plants due to their roles in plants defence against herbivores and
pathogens (Hartmann 2007; Pichersky and Lewinsohn 2011; Wink 2003). The ability to produce
BIAs in angiosperms is thought to have arisen from a monophyletic origin (Liscombe et al.
2005). The incredible structural diversity of BIAs is the result of remarkably few enzyme types.
The structural diversity of BIAs has been explained by the duplication of genes recruited from
primary metabolism followed by random mutation of the duplicated genes (Ober and Hartmann
2000). This processes allowed for the generation of novel catalytic functions without loss of the
genes’ original functions.
Certain reaction types are widespread in BIA metabolism, one such reaction type is
SAM-dependant O-methylation which catalyze the regiospecific transfer of a methyl group from
SAM to a free hydroxyl moiety on the BIA backbone (Beaudoin and Facchini 2014). Genes
encoding O-methyltransferases involved in BIA biosynthesis have been isolated from several
plants that produce BIAs and often share significant sequence similarity (Beaudoin and Facchini
2014). 6OMT has been isolated from Coptis japonica (Inui et al. 2007) and P. somniferum
(Ounaroon et al. 2003). A molecular clone for 4’OMT has been isolated from C. japonica
(Morishige et al. 2000) while 4’OMT activity has been found in the cell cultures from several
44
plants such as, Berberis koetineana, Mahonia nervosa and Eschscholzia californica (Frenzel and
Zenk 1990). SOMT1 has been isolated from both P. somniferum (Dang and Facchini 2012) and
C. japonica (Takeshita et al. 1995). Other known OMTs involved BIA metabolism include
7OMT (Ounaroon et al. 2003) and N7OMT (Pienkny et al. 2009) both were isolated from P.
somniferum.
At least two OMTs expected to be involved in BIA metabolism in opium poppy have
never been isolated from cDNA or functionally characterized (Beaudoin and Facchini 2014).
The first is a 3’OMT involved in papaverine biosynthesis and the second a 2OMT involved in
the formation of 2-O-methylated protoberberines including (S)-tetrahydropalmatine, (S)-sinactine
and (S)-tetrahydropalmatrubine. There is a report of a 3’OMT that was partially purified and
characterized from Argemone platyceras cell suspension cultures (Rueffer et al. 1983). The 5’
and 3’ positions are equivalent on the simple BIA backbone because both sites are meta to the
isoquinoline linking carbon, as such the enzyme was named, 6-O-methylnorlaudanosoline-5’-Omethyltransferase (5’OMT). 5’OMT had a molecular weight of 47 kDa, a pH optimum of 7.5
and a temperature optimum of 35 ºC. 5’OMT was assayed with simple BIAs possessing various
O-methylation and N-methylation patterns, 6-O-methylnorlaudanosoline (6-O-methylated, Ndesmethyl), norlaudanosoline (no O-methylation, N-desmethyl) and laudanosoline (no Omethylation, N-methylated). 6-O-methylnorlaudanosoline was the only substrate accepted by
5’OMT (Rueffer et al. 1983). More recently, 3’OMT activity was found in the search for
scoulerine 9-O-methyltransferase from P. somniferum (Dang and Facchini 2012). The 9-position
of the protoberberine backbone is equivalent to the 3’-position of the simple BIA backbone. An
enzyme called SOMT1 was discovered and catalyzed the sequential O-methylation at the 9position of scoulerine followed the O-methylation of the 2-position of scoulerine which resulted
45
in the formation of the fully O-methylated protoberberine tetrahydropalmatine. An analogous
reaction was catalyzed when the simple BIA reticuline was used as a substrate, reticuline was Omethylated at the 3’-position and subsequently O-methylated at the 7-position to form the fully
O-methylated simple BIA laudanosine. Due to the nearly 300 fold reduction of catalytic
efficiency between scoulerine and reticuline, the authors postulated that SOMT1 would be
unable, on its own, to account for the high basal levels of papaverine observed in opium poppy
and suggested the presence of a more regiospecific and substrate specific 3’OMT in opium
poppy. Additional support for the presence of a dedicated 3’OMT came from the systematic
silencing of BIA biosynthetic genes from P. somniferum (Desgagné-Penix and Facchini 2012).
The VIGS-mediated knockdown of biosynthetic genes upstream of papaverine suggested that
papaverine biosynthesis proceeded via an N-desmethylated pathway which excludes reticuline.
SOMT1 preference for the N-methylated simple BIA reticuline over the N-desmethyled
norreticuline suggests that SOMT1 is unlikely to be involved in a major pathway leading to
papaverine.
Although an enzyme with protoberberine 2OMT activity has never been reported in
opium poppy, O-methylation activity at the 2-position of the protoberberine backbone as been
reported in the literature. Columbamine O-methyltransferase was first discovered in suspension
cultures of Berberis wilsoniae. CoOMT was partially purified using classical biochemistry
approaches and assayed. CoOMT catalyzed the SAM-dependent O-methylation of columbamine
at the 2-position to form palmatine and exhibited high substrate specificity (Rueffer, Amann, and
Zenk 1986). A molecular clone of CoOMT was eventually isolated from C. japonica cell
cultures and heterologously expressed in E. coli (Morishige et al. 2002). Surprisingly,
CjCoOMT had a higher degree of sequence similarity to C. japonica 6OMT, which accepts
46
simple BIAs as substrates, than to C. japonica SOMT which accepts protoberberine alkaloids as
substrates. CjCoOMT had a molecular weight of 40 kDa, a pH optimum of 8.4 and a
temperature optimum of 30 ºC. Among the substrates tested in the study columbamine was the
best substrate followed by (S)-scoulerine which indicates that the fully aromatized backbone is
preferred. (S)-tetrahydropalmatine was also accepted as a substrate however CoOMT did not
accept alkaloids with a 1-benzylisoquinoline backbone indicating a strong preference for the
protoberberine backbone.
The following chapter describes the identification and functional characterization of a
cDNA encoding protoberberine 2-O-methyltransferase (2OMT) from opium poppy roots.
2OMT is the only OMT reported from opium poppy that catalyzes the transfer of a methyl group
from SAM to a free hydroxyl group at the 2 position of protoberberine alkaloids.
3.2 Results
3.2.1 Identification and phylogenetic analysis of 2OMT cDNA
The molecular cloning of several cDNAs encoding OMTs involved in BIA metabolism
were used as query sequences to identify full-length candidate cDNAs encoding opium poppy
2OMT. Query sequences were used to mine next-generation sequence databases as described in
Chapter 2.1. Similar approaches have been used previously to identity other BIA biosynthetic
genes (Ikezawa, Iwasa, and Sato 2008; Liscombe and Facchini 2007). Six gene candidates were
amplified from Bea’s Choice Stem cDNA named, IzOMT1, through IzOMT6, and four gene
candidates were amplified from Bea’s Choice Root cDNA named DDOMT 1 through DDOMT4.
All candidates were selected in part based on their high FPKM values which are indicative of
transcript abundance. FPKM value for IzOMT1 was 60.1; IzOMT2 was 42.1; IzOMT3 was
47
1.23; IzOMT4 was 15.58; IzOMT 6 was 119.5; DDOMT1 was 117; DDOMT2 was 68.79;
DDOMT3 was 178.1 and DDOMT4 was 60.1. Of the amplified genes, IzOMT1 contained a
stop codon in the middle of the predicted ORF, IzOMT3 was too short to be a SAM-dependent
OMT, IzOMT5 contained a premature stop codon followed by an insertion, DDOMT1 and 2 also
contained a premature stop codon followed by an insert. Full length ORFs for DDOMT1 and
DDOMT2 were obtained through gene synthesis. DDOMT1 was renamed protoberberine
2OMT.
Several SAM-dependent O-methyltransferases involved in BIA metabolism were
subjected to phylogenetic analysis along with gene candidates to determine the evolutionary
relationships between candidate genes and characterized plant OMTs (Figure 3.1). One clade
forms around 7OMT and 2OMT. Other BIA biosynthetic enzymes also group together, 6OMT,
N7OMT and 4’OMT for a clade at the top of figure 3.1. SOMT1 and DDOMT2 form a clade
near the bottom of the bottom left of figure 3.1.
The amino acid sequence of 2OMT was analyzed using InterPro Protein Sequence
analysis and classification tool (Jones et al. 2014). Three domains were located. The first was a
winged helix-turn-helix DNA-binding domain with a plant methlytransferase dimerisation
domain embedded within the winged helix-turn-helix. The C-terminus of the protein contained a
SAM-dependent methyltransferase domain.
48
Figure 3.1. Neighbour-joining tree of derived from the amino acid sequences of selected
plant SAM-dependant O-methyltransferases.
Amino acid sequences were aligned using Clustal W (Chenna et al. 2003). The tree was
constructed and bootstrap analysis was performed using Geneious.
49
3.2.2 Heterologous expression of 2OMT
The identity of the isolated cDNA as protoberberine 2OMT was established by the
production of recombinant polyhistidine-tagged proteins in E. coli Arctic Express (DE3) RP cells
containing pET29b::2OMT. The recombinant enzyme had a molecular mass of ~40 kDa which
was similar to the expected due to the addition of the C-terminal His-tag (Figure 3.2 A). The
recombinant His-tagged 2OMT was only found in total protein extracts and purified protein
extracts of E. coli harbouring the pET29b::2OMT (Figure 3.2 B).
His-tagged recombinant 2OMT was purified by cobalt affinity chromatography as shown
in Figures 3.2 A and B. The protein was subsequently desalted using PD-10 columns. IzOMT2,
IzOMT3, IzOMT6, DDOMT2, DDOMT3 and DDOMT4 were expressed and purified in the
same manner as 2OMT.
3.2.3 Enzymatic properties of 2OMT
The pH optimum for 2OMT was approximately 8 and half-maximum activity occurred at
approximately pH 6.3 (Figure 3.3). Recombinant 2OMT showed maximum activity at
approximately 37 ˚C and half-maximum activity was predicted to occur at approximately 25 ˚C
and 40 ˚C (Figure 3.4).
50
Figure 3.2. Heterologous expression of 2OMT synthetic gene in E. coli
(A) SDS-PAGE analysis of protein extracts of from non-induced (-IPTG) and induced
(+IPTG) E. coli cells harbouring the pET29b 2OMT expression construct. Purified
recombinant 2OMT is also shown. (B) Western blot analysis performed on samples from A
using a polyhistidine tag monoclonal antibody shows the presence of recombinant proteins.
Number of the left of A and B show the protein molecular weight standards.
51
Figure 3.3. The effect of pH on Papaver somniferum 2OMT activity with scoulerine
Different pH ranges were achieved by using different buffers. pH 5 (citrate), pH 6-7
(potassium phosphate), pH 8(Tris-HCl), and pH 9 (glycine).
52
Figure 3.4. The effect of temperature on Papaver somniferum 2OMT activity with
scoulerine
53
Substrates from different BIA structural subgroups were used to investigate the substrate
specificity of protoberberine 2OMT and the other heterologously expressed recombinant
enzymes. Figure 3.5 shows the EICs from each purified recombinant enzyme being assayed with
scoulerine as a negative result. Figure 3.6 shows the extracted ion chromatograms for substrates
that were accepted by 2OMT. In each frame of figure 3.6 (A-F), the top two chromatogram
represent, negative controls with boiled enzyme, the first chromatogram represents the extract
ion chromatogram (EIC) corresponding to the m/z of the substrate and the second chromatogram
represents the EIC corresponding to the m/z of the product which is equivalent to the m/z of the
substrate plus the mass of a single O-methylation event (14 Da). The two lower chromatograms
represent the extracted ion chromatograms for the m/z of the substrate and the product with
native enzyme. All assays were conducted for 2 hours and incubated at 30 °C. Frame A shows
the activity of 2OMT with cheilanthifoline. The controls show that cheilanthifoline (m/z 326)
elutes at 3.2 minutes. The product, sinactine (m/z 340), elutes slightly later than cheilanthifoline
at 3.7 minutes. Frame B shows the activity of 2OMT with scoulerine. Scoulerine (m/z 328)
elutes at 3.2 minutes and the O-methylated product, tetrahydropalatrubine (m/z 342) elutes at 3.5
minutes. Frame C shows the activity of 2OMT with coclaurine. Coclaurine (m/z 286) elutes at
2.8 minutes and the O-methylated product of the reaction, norarmepavine (m/z 300) elutes at 3.2
minutes. Frame D shows the activity of 20MT with dopamine. Dopamine (m/z 154) elutes at 0.4
minutes and the O-methylated product, 4-methoxytyramine (m/z 168) elutes at 0.8 minutes.
Frame E shows the activity of 2OMT with reticuline. Reticuline (m/z 330), retention time, 3.1
minutes, is O-methylated to form laudanidine (m/z 344) with a retention time of 3.2 minutes.
Frame F shows the activity of 6-O-methylnorlaudanosoline with 2OMT. The substrate, 6-Omethylnorlaudanosoline (m/z 302) has a retention time of 2.7 minutes. The product is 6,7-O54
dimethylnorlaudanosoline (m/z 316) elutes at 3.4 minutes. IzOMT2, IzOMT3, IzOMT6,
DDOMT2, DDOMT3 and DDOMT4 were assayed with the same substrates and no activity
could be detected.
Table 3.1 lists all the substrates assayed with 2OMT, the methylation pattern of substrate,
the relative activity of each substrate relative to scoulerine and the product formed. The relative
activities were calculated by comparing the integrations of the EICs and normalizing them to
scoulerine. The best substrate for 2OMT under the conditions tested was the protoberberine (S)cheilanthifoline which was methylated at the 2 position to make (S)-sinactine at % 1010 relative
activity followed by (S)-scoulerine which was methylated at the 2 position to form (S)tetrahydropalmatrubine with a relative of activity % 100. (S)-Coclaurine was converted to (S)norarmepavine by 2OMT with an O-methylation at the 7 position with a relative activity of %
41.9. The phenylethylamine and precursor to all BIAs, dopamine was also accepted by 2OMT
which is thought to form 4-methyoxytryamine with a relative activity of % 21.1. (S)-Reticuline
and (S)-6-O-methylnorlaudanosoline were also accepted by 2OMT in trace amounts to form (S)laudanidine and (S)-6,7-O,O-methylnorlaudanosoline respectively.
Figure 3.7 shows the chemical conversions of the substrates to the reaction products by
2OMT. The products of each reaction were determined by CID at 25 eV. In all cases chemical
standards were not available for the products of reaction and the chemical structures were
inferred by comparing the fragmentation mass spectra of the substrate and using fragmentation
schemes for protoberberine alkaloids and simple BIAs in the literature (Schmidt et al. 2005;
Schmidt et al. 2007). Table 3.2 shows the CID fragmentation patterns of the substrates and the
enzymatic products of 2OMT. Across all samples a characteristic gain of 14 Da was observed
on the isoquinoline moiety of the product.
55
Table 3.1. The products formed by assaying different chemicals with 2OMT. m/z ratios
were determined by LC-MS and reaction products were determined by CID. The
methylation pattern of the substrate is indicated by the positions N, 6, 7, 4’ and 3’.
Activities are relative to scoulerine
as a substrate.
Substrate
Substrate
m/z
Product
m/z
N
6
7
4'
3'
Relative
Activity (%)
Products
Dopamine
154.2
168.2
H
OH
OH
-
-
21.1
4-Methoxytyramine
(S)-Coclaurine
286.4
300.4
H
OMe
OH
OH
-
41.9
(S)-Norarmepavine
(S)-Norlaudanosoline
288.2
302.2
H
OH
OH
OH
OH
0
-
(S)-6-O-Methylnorlaudanosoline
302.2
316.2
H
OMe
OH
OH
OH
0.2
(S)-6,7-O,O-Dimethylnorlaudanosoline
Norreticuline
316.2
330.2
H
OMe
OH
OMe
OH
0
-
(S)-Reticuline
330.4
344.4
Me
OMe
OH
OMe
OH
2.61
(S)-Laudanidine
Boldine
328.4
342.4
Me
OH
OMe
OMe
OH
0
-
Isocorydine
342.2
356.2
Me
OMe OMe
OMe
OH
0
-
(S)-Scoulerine
328.2
342.2
H
OMe
OH
OMe
OH
100
(S)-Tetrahydropalmatrubine
(S)-Cheilanthifoline
326.4
340.4
H
OMe
OH
MDO MDO
1.01E+03
(S)-Sinactine
Oripavine
298.2
312.2
Me
OH
OMe
0
-
Morphine
286.2
300.2
Me
OH
OMe
0
-
56
Figure 3.5 Extracted ion chromatograms (EICs) showing the lack of O-methylation activity
observed when scoulerine was assayed with each heterologously expressed enzyme (A)
IxOMT2, (B) IzOMT3, (C) IzOMT6, (D) DDOMT2, (E) DDOMT3 and (F) DDOMT4.
For each substrate the top two EICs (controls) represent boiled enzyme negative controls
and the bottom two lines represent purified native recombinant enzyme.
57
Figure 3.6 Extracted ion chromatograms (EICs) showing the O-methylation activity of
2OMT on (A) cheilanthifoline, (B) Scoulerine, (C) coclaurine, (D) dopamine, (E) Reticuline
and (F) 6-O-methylnorlaudanosoline.
58
For each substrate the top two EICs (controls) represent boiled enzyme negative controls
and the bottom two lines represent purified native recombinant 2OMT. The incubation of
native 2OMT with cheilanthifoline (m/z 326) yielded sinactine (m/z 340) based on its CID
spectrum. The incubation of 2OMT with scoulerine (m/z 328) formed
tetrahydropalmatrubine (m/z 342) based on its CID spectrum. The incubation of 2OMT
with coclaurine (m/z 286) formed norarmepavine (m/z 300) based on its CID spectrum. The
incubation of 2OMT with dopamine (m/z 154) formed 4-methyoxytyramine (m/z 168) based
on its CID spectrum. The incubation of 2OMT with reticuline (m/z 330) formed
laudanidine (m/z 344) based on its CID spectrum. The incubation of 2OMT with 6-Omethylnorlaudanosoline (m/z 302) formed 6,7-O,O-dimethylnorlaudanosoline (m/z 316)
based on its CID spectrum.
59
Figure 3.7 The O-methylation activity of 2OMT on various BIA substrates (A)
cheilanthifoline, (B) scoulerine, (C) coclaurine, (D) dopamine, (E) reticuline and (F) 6-Omethylnorlaudanosoline.
The incubation of native 2OMT with cheilanthifoline (m/z 326) yielded sinactine (m/z 340)
based on its CID spectrum. The incubation of 2OMT with scoulerine (m/z 328) formed
tetrahydropalmatrubine (m/z 342) based on its CID spectrum. The incubation of 2OMT
with coclaurine (m/z 286) formed norarmepavine (m/z 300) based on its CID spectrum. The
incubation of 2OMT with dopamine (m/z 154) formed 4-methyoxytyramine (m/z 168) based
on its CID spectrum. The incubation of 2OMT with reticuline (m/z 330) formed
laudanidine (m/z 344) based on its CID spectrum. The incubation of 2OMT with 6-Omethylnorlaudanosoline (m/z 302) formed 6,7-O,O-dimethylnorlaudanosoline (m/z 342)
based on its CID spectrum.
60
Table 3.2. The ESI[+]CID pattern of substrates and the products from the substrate range
experiments determined by LC-MS/MS
The common name of each alkaloid in indicated as well as the m/z ratio of the molecular
ion or the protonated parent ion [M]+ or [M+H]+. The identity of the products was
inferred by comparing the CID patterns of the products to those of substrates and by
studying the fragmentation patterns for protoberberine alkaloids and simple BIAs in the
literature. The retention time is indicated by RT and is expressed in minutes. CE indicates
the collision energy in eV used in CID experiments. ESI-CID spectrum m/z indicates the
m/z of the fragments resulting from the CID of a given compound and the number in
brackets represents the relative intensity of the ion in the spectrum. The structure is also
illustrated in the table
or [M]+
RT
(min)
CE
(eV)
ESI-CID Spectrum
m/z (Relative
Intensity)
(S)-Cheilanthifoline
326
3.2
25
326 (17), 178 (100),
176 (11), 151 (23),
149 (6), 91 (17)
2
(S)-Sinactine
340
3.7
25
340 (7.2), 192 (100),
176 (10), 165 (46)
3
(S)-Scoulerine
328
3.2
25
328 (9.02), 178 (100),
151 (10.77)
4
(S)Tetrahydropalmatrubine
342
3.5
25
342 (14), 192 (100),
165 (46), 151 (8), 150
(7)
5
(S)-Coclaurine
286
2.8
25
286 (18), 269 (65),
178 (5), 175 (3), 143
(12), 107 (100)
6
(S)-Norarmepavine
300
3.2
25
300 (14), 283 (92),
192 (8), 190 (6), 175
(5), 107 (100)
7
Dopamine
154
0.5
25
154 (1), 137 (100),
119 (12), 91 (9)
8
4-Methoxytyramine
168
0.8
25
168 (1), 151 (100),
133 (8)
9
(S)-Reticuline
330
3.1
25
192 (100), 177 (8.09),
175 (19.32), 151
(5.45), 143 (25), 137
(41.49)
No.
Compound
1
[M+H]+
61
Structure
10
(S)-Laudanidine
344
11
(S)-6-Omethylnorlaudanosoline
302
12
(S)-6,7-O,Odimethylnorlaudanosoline
316
25
344(1), 206 (100),
189 (7), 143 (52), 137
(38)
2.7
25
302 (1), 253 (5), 207
(9), 178 (83), 175
(27), 163 (12), 143
(58), 137 (12), 123
(100), 115 (12)
3.4
25
316 (1), 192 (85), 189
(22), 143 (25), 123
(100)
3.2
62
Varying concentrations of (S)-scoulerine from 5 to 200 µM and a constant concentration
of SAM at 500 µM, produced a typical Michaelis-Menten substrate saturation kinetics. The
apparent Km value for (S)-scoulerine was 51.8 ± 13.0 µM and the apparent Vmax was calculated as
1.51 x 106 Counts/min/µg. Both the apparent Km and apparent Vmax, shown in Figure 3.9, were
calculated by regression of the Michaelis-Menten curve using SigmaPlot.
3.2.4 Relative transcript abundance of 2OMT in opium poppy tissue
The relative transcript abundance of 2OMT in different opium poppy tissue types was
determined by qPCR. Primers were designed to anneal to a unique region of 2OMT’s 3’UTR.
Of the five tissue types analyzed, root had by far the greatest relative transcript abundance while,
stem, capsule, leaf and bud had almost no relative transcript abundance (Figure 3.10).
63
Figure 3.8 Michaelis-Menten plot for Papaver somniferum protoberberine 2OMT with (S)scoulerine with a constant concentration of SAM.
The Km and Vmax were calculated by regression using SigmaPlot.
64
Figure 3.9 Relative abundance of 2OMT gene transcripts in Bea’s Choice opium poppy
tissues
RT-PCR was performed with cDNA synthesized using total RNA from each organ. Error
bars represent the standard error of the mean for three independent measurements.
65
3.3 Discussion
Many studies have described the cloning and functional characterization of OMTs
involved in BIA biosynthesis (Dang and Facchini 2012; Morishige et al. 2000; Ounaroon et al.
2003; Pienkny et al. 2009). To date, all OMTs involved in BIA biosynthesis utilize SAM as the
methyl donor and transfer a methyl group with a certain degree of regiospecificity (Beaudoin and
Facchini 2014). When this study began at least two OMTs involved in BIA biosynthesis
remained unknown. The first was a 3’OMT involved in the production of papaverine and the
second a protoberberine 2OMT involved in the production of sinactine.
The goal of this study was to isolate the 3’OMT gene and functionally characterize it. To
this end, 6 candidates were found in 454 and Illumina NGS stem transcriptomic databases and 4
more gene candidates were selected from 454 and Illumina NGS root transcriptomic databases.
Candidates were selected based on their sequence similarity to characterized OMTs involved in
BIA biosynthesis. Candidate selection was prioritized based on each contig’s FPKM value
provided by the Illumina databases. Expression levels of a given gene can be predicted by their
FPKM values in the database because FPKM values are related the number of reads used to
assemble any given gene. The assumption was made that mRNA abundance, expressed by the
FPKM value, was correlated to the protein abundance. Due to the high accumulation of
papaverine in opium poppy latex, only the most highly expressed putative OMT genes, as
indicated by their FPKM values, were selected as gene candidates. A recent study described an
enzyme, DBOX, responsible for the conversion of tetrahydropapaverine to papaverine was
expressed in root tissue (Hagel et al. 2012). The presence of DBOX in the roots opened the
66
possibility that other steps of papaverine biosynthesis also occurred in the roots and led to
prioritization of gene candidates exclusively from roots over gene candidates found in the stem.
Some gene candidates were dropped due their small size or the presence of introns within the
predicted ORF of the gene. Such introns could be the result of a simple assembly error due to
certain biases of the Velvet assembly software or by something more systematic like differences
in the developmental stage of the plants used for sequencing and for other experiments. Genes
for which full length cDNAs were amplified and heterologously expressed in E.coil were
IzOMT2, IzOMT3, IzOMT6, DDOMT3 and DDOMT4.
We report the characterization of a full-length gene encoding protoberberine 2OMT
found in opium poppy roots. This novel SAM-dependent OMT was identified by tBLASTn
analysis of Illumina HiSeq and Roche-454 pyrosequencing transcriptomic databases of opium
poppy root. Amplification of protoberberine 2OMT from P. somniferum variety “Bea’s Choice”
roots produced a gene containing an intron in the ORF, as a result, the gene was synthesized.
The intron could the be result of there being several genomic copies of 2OMT or the
developmentally dependant splicing of 2OMT. Similarly it is possible that there was an
assembly error resulting in a mis-call substitution in the NGS dataset. Unlike, IzOMT1 and
IzOMT5, 2OMT and DDOMT2 were not abandoned, the reasoning was due to their high FPKM
values and their presence in roots and absence in stem. Functional expression of the 2OMT
synthesized gene produced a polyhistidine-tagged enzyme in E. coli allowed 2OMT to be
purified to homogeneity by colbalt-affinity chromatography (Figure 3.2). Purified 2OMT
catalyzed the conversion of selected protoberberine alkaloids and simple BIAs to their respective
2-O-methylated and 7-O-methylated derivatives (Figure 3.7). When the protoberberine ring is
67
formed by BBE, the 7-position of the simple BIA backbone becomes the 2-position of the
protoberberine backbone (Dittrich and Kutchan 1991).
The phylogenetics of protoberberine 2OMT were studied by comparing the amino acid
sequences of 2OMT to the sequences of characterized plant SAM-dependent OMTs in a
neighbour-joining tree (Figure 3.1). 2OMT forms a clade with 7OMTs from Eschscholzia
californica and Papaver somniferum which is relatively distant from other characterized OMTs
involved in BIA metabolism suggesting they have arisen from a recent common ancestor through
a process of gene duplication and subsequent mutation. 7OMT and 2OMT both catalyze a
similar reaction: the O-methylation of the isoquinoline moiety at the same position, which
corresponds to the 7 position on the simple BIA backbone or the 2 position of the protoberberine
backbone. CjCoOMT also methylates at the 2-position of the protoberberine backbone
specifically the substrates, columbamine, tetrahydrocolumbamine and scoulerine. CjCoOMT
forms a separate and distant clad from 7OMT and 2OMT suggesting it arose independently from
7OMT and 2OMT. Other OMTs involved in BIA metabolism, 6OMT, 4’OMT and N7OMT also
formed a distinct clade from 7OMT and 2OMT which is interesting because N7OMT accepts the
N-desmethyled substrate norreticuline and forms norlaudanine by an O-methylation at the 7position of norreticuline (Pienkny et al. 2009). As a general rule, 6OMT and 4’OMT clustered
closest together based on function (O-methylation at the 6- or 4’ positions) and independently of
species. PsSOMT1, which catalyzes the O-methylation of scoulerine at the 9-position (3’position of the simple BIA backbone), is highly similar to DDOMT2. The clade formed with
SOMT and DDOMT2 led us to believe that DDOMT2 may have been the elusive 3’OMT
involved in biosynthesis of papaverine however when it was assayed with simple BIAs and
protoberberines no activity was detected.
68
The catalytic properties of recombinant opium poppy protoberberine 2OMT are generally
in agreement with those of other purified or partially purified OMTs from opium poppy and
other BIA producing plants. The molecular weight of opium poppy 2OMT is approximately
40kDa, determined by SDS-Page (Figure 3.2) which is comparable to the molecular weights of
other OMTs purified from opium poppy and related plants. For example, SOMT1 from opium
had mass of 43 kDa (Dang and Facchini 2012), 7OMT from opium poppy was 40 kDA
(Ounaroon et al. 2003) and CoOMT from C. japonica had a mass of 40 kDa (Morishige et al.
2002). Gel filtration chromatography of some OMTs has revealed native molecular masses that
are approximately double the size (Frick and Kutchan 1999), suggesting the possibility that
2OMT may exist as a dimer in vivo. The possibility of 2OMT existing as a dimer was
strengthened by the detection of a plant methyltransferase dimerisation domain by the InterPro
sequence analysis tool. The temperature and pH optima for recombinant 2OMT (Figure 3.3 and
3.4) fall in the same range as other characterized OMTs. Opium poppy SOMT1 exhibited a pH
optimum of 9.0 and a temperature optimum of 37 °C (Dang and Facchini 2012), opium poppy
7OMT has a pH optimum of 8.0 and a temperature optimum of 37 °C (Ounaroon et al. 2003),
and CoOMT from Coptis japonica demonstrated optimal activity at a pH 8.4 and 30 °C
(Morishige et al. 2002). The apparent Km value of 2OMT for (S)-scoulerine of 51.8 ± 13.0 µM
falls on the high end of Km values reported for other OMTs and could be lower if the protein had
been purified to homogeneity. SOMT1 has an apparent Km of 28.5 ± 6.8 µM for (S)-scoulerine
(Dang and Facchini 2012), while 7OMT from opium poppy is reported as having a Km of 16 µM
for (S)-reticuline (Ounaroon et al. 2003). CoOMT and 4’OMT from Coptis japonica have a Km
of 66 ± 18 µM for columbamine and 42 µM for 6-O-methylnorlaudanosoline respectively
(Morishige et al. 2002; Morishige et al. 2000). Unfortunately, due to limited availability of the
69
chemical, a Km for (S)-cheilanthifoline could not be determined for in this study. However, given
the apparent substrate preference of 2OMT for (S)-cheilanthifoline suggests that a Km for (S)cheilanthifoline would be lower than that for (S)-scoulerine. The substrate specificity of opium
poppy protoberberine 2OMT is somewhat wide in that it accepts the phenylethylamine
dopamine, which serves as a precursor to all BIAs, as well is simple BIAs and protoberberine
type alkaloids (Table 3.1). Despite their phylogenetic distance, the reactions catalyzed by
protoberberine 2OMT are similar to the reactions catalyzed by C. japonica CoOMT in that they
O-methylate at 2-position of the protoberberine backbone. CoOMT has been reported to Omethylate columbamine, tetrahydrocolumbamine and scoulerine. However, unlike 2OMT,
CoOMT does not accept simple BIAs. Protoberberine 2OMT is unique as it is the only enzyme
characterized from opium poppy that catalyzes the 2-O-methylation of the protoberberine
backbone.
As discussed above, 2OMT shows homology with other characterized OMTs involved in
BIA metabolism but is most closely related to opium poppy 7OMT (figure 3.1). Like many plant
O-methyltransferases, reticuline 7OMT was reported to have a somewhat broad substrate
specificity which often makes assigning an in vivo role difficult. Reticuline 7OMT accepted the
phenolics, guaiacol and isovanillic acid in addition to the BIAs, reticuline, orientaline,
protosinomenine, isoorientaline (Ounaroon et al. 2003). 7OMT was assayed with scoulerine and
showed no catalytic activity. Guaiuacol demonstrated the highest catalytic efficiency (kcat/Km).
However, the authors suggested that because guaiacol does not accumulate in P. somniferum,
reticuline, with the second highest catalytic efficiency, was suggested to be the endogenous
substrate of 7OMT. Reticuline 7OMT was also quite promiscuous in its ability to methylate
positions other than the 7-position. Protosinomenine and isoorientaline each have free hydroxyls
70
at C-6 and are O-methylated at the C-7, yet both were O-methylated by 7OMT at C-6. In some
cases, 7OMT could also methylate at the C-4’, when C-3’ was O-methylated, as a secondary
reaction. Despite the interesting methylation products of 7OMT, it showed the highest catalytic
efficiency at the 7-position. Unfortunately, we did not have access to many of the substrates that
7OMT accepted as substrates (orientaline, protosinomenine or isoorientaline). Lacking these
substrates or their equivalent protoberberine alkaloids, it was impossible to test whether or not
2OMT is capable of methylting at different positions on the BIA backbone. Most of the
substrates investigated in this study did not have free hydroxyls at C-6 or C-4’. The notable
exceptions to this would be coclaurine which has a free hydroxyl at C-4’ and dopamine with two
free hydroxyls. 2OMTs apparent inability to methylate at different positions and its inability to
doubly O-methylate substrates could be the result of differences in the binding pocket of 2OMT
to accommodate protoberberine alkaloids when compared to 7OMT, however, an investigation
with more substrates and using NMR as the method of identifying the products would be needed
to determine the real reasons for the differences between 7OMT and 2OMT.
The fragmentation patterns of protoberberine alkaloids and simple BIAs have been
studied at length (Schmidt et al. 2005; Schmidt et al. 2007). Published fragmentation schemes,
the fragmentation of the chemical standards and the retention times of the standards relative to
the enzymatic products, served as the foundation for interpreting the fragmentation mass spectra
of the products formed by 2OMT. Simple BIAs fragment in a very distinctive and reproducible
fashion, first the bond linking the isoquinoline and the benzyl moieties is broken which forms
two charged species, one is derived from the isoquinoline moiety and the other from the benzyl
moiety. This information allowed us to determine whether the methylation event occurred on the
isoquinoline or benzyl moiety. However, this on its own was not enough information to position
71
the methylation event on a given hydroxyl and did not preclude the possibility that the
methylation event occurred on the amine group. An alternate fragmentation route of simple
BIAs involved the loss of an amine group from the parent ion followed by the recyclization of
the isoquinoline moiety without fragmentation of the bond linking the isoquinoline group to the
benzyl group. The removal of the nitrogen as a methylation site left only the free hydroxyls on
the isoquinoline moiety as possible methylation sites. Further fragmentation of the ion without
the amine group resulted in the cleavage of the bond linking the benzyl group and the
isoquinoline moiety which leads to the formation of a charges species corresponding to the
isoquinoline moiety (Schmidt et al. 2005). Therefore the strategy to identify enzymatic products
in this experiment consisted of looking for ions in the fragmentation mass spectra corresponding
to the isoquinoline moiety and were 14 Da heavier than the equivalent ions from the
fragmentation mass spectra of the substrate. Coclaurine, 6-O-methylnorlaudanosoline and
reticuline, which are all 6-O-methylated, only provide a single possible site to for methylation on
the isoquinoline ring. The structural limitation of a single methylation site on the isoquinoline
moiety provided a strong degree of confidence that 2OMT catalyzed the 7-O-methylation on
these substrates. To be certain, NMR should be employed to identify the products.
The fragmentation patterns of protoberberine alkaloids have been studied in considerable
depth and share some characteristics with the fragmentation of simple BIAs (Schmidt et al.
2007). As with simple BIAs, protoberberines are broken into two separate ions, one representing
the isoquinoline moiety and an other representing the benzyl moiety. The CID of scoulerine
results in the formation of two major ions from the parent ion. The scoulerine parent ion is
represented by m/z 328, while the singly O-methylated isoquinoline moiety and the singly Omethylated benzylic ion are represented by m/z 178 and 151 respectively. Scoulerine has another
72
fragmentation route that splits the isoquinoline moiety from the parent ion resulting in an ion
without the amine at m/z 151. The fragmentation mass spectrum cheilanthifoline displays
equivalent ions. The parent ion of cheilanthifoline is m/z 326. The two distinct ions representing
the singly O-methylated isoquinoline moiety of cheilanthifoline are m/z 178 and 151. The ion
representing the benzyl moiety with a methylendioxy bridge is m/z 149. The strategy to interpret
the fragmentation mass spectra of the enzymatic products was similar to the strategy used for
simple BIAs, search for ions that represent the isoquinoline moiety with an additional 14 Da
when compared to the substrate and a retention time later than that of the substrate. When
assayed with 2OMT, scoulerine was converted to m/z 342. Upon fragmentation of m/z 342 the
ions m/z 342, 192, 165, 151 and 150 were observed in the mass spectrum. Ions with m/z 192
and 165 were derived from the isoquinoline moiety and are exactly 14 Da heavier than the
equivalent ions in the fragmentation mass spectrum of scoulerine, m/z 178 and 151. This data,
when taken together, suggested that a 2-O-methylation occurred on the free hydroxyl group of
the isoquinoline moiety of scoulerine. The ion representing the benzylic moiety was detected
with m/z 151, indicating no reaction occurred on the benzylic moiety of scoulerine. The product
of the enzymatic reaction of 2OMT with scoulerine was identified as (S)-tetrahydropalmatrubine.
Cheilanthifoline (m/z 326) was converted by 2OMT to m/z 340. The fragmentation mass
spectrum of m/z 340 displayed the ions m/z 340, 192, 176 and 165. Again ions m/z 192 and 165
represent the isoquinoline moiety with an additional methyl group on the free hydroxyl when
compared to the fragmentation mass spectrum of cheilanthifoline. The expected ion at m/z 149
could not be detected. Ion m/z 176 was present in the fragmentation patterns of cheilanthifoline
and m/z 340. I believe that m/z 176 is derived from the parent ion moiety and is the benzylic
moiety compliment to ion m/z 151 from cheilanthifoline or ion m/z 165 form the enzymatic
73
product m/z 340. If this is were the case, ion m/z 176 would contain an additional nitrogen and
carbon when compared to benzylic ion described above. The presence of m/z 176 in the
fragmentation mass spectra of cheilanthifoline and m/z 340 supports the idea that 2OMT did not
catalyze any reaction on the benzylic moiety. The O-methylated product of the reaction between
2OMT and cheilanthifoline was putatively identified as (S)-sinactine. These experiments should
be repeated using different setting on the MS to increase confidence in the dataset by reducing
background noise and to clean up the chromatograms. Selected-reaction monitoring should be
used in future experiments to increase the sensitivity and improve the quantitative reliability of
the data for both the substrate and the product.
Some OMTs involved in BIA metabolism are know to accept catechols and 2OMT seems
no different (Ounaroon et al. 2003). The reaction of 2OMT with dopamine was much more
difficult to interpret and the identification of the product is by no means certain. Although it is
clear that dopamine (m/z 154) was methylated to form the product m/z 168, dopamine had two
sites that can be O-methylated and it was impossible to distinguish where the O-methylation
event took place by CID. The loss of 17 Da from the fragmentation mass spectra of both
dopamine and the enzymatic product corresponds to the loss of –NH3 and preclude the Nmethylation of dopamine. Therefore the most likely site of O-methylation is either the 3-position
or the 4-position of dopamine. To draw conclusions regarding the specific site of O-methylation
for dopamine without supporting mass fragmentation data of a standard or an NMR study is not
possible.
While the study of substrates with differing methylation patterns and CID studies
provided support for the identification of the enzymatic products, without chemical standards it
is not possible to definitively identify the compounds in accordance with the minimum
74
information about a metabolomics experiment guidelines (MIAMET) (Katajamaa and Oresic
2007). Ultimately to be certain of the methylation site and the chemical structure of the
enzymatic products, NMR would have to be employed for a final identification.
RT-PCR was employed to determine the relative transcript abundance of 2OMT in
different tissue types from Bea’s choice variety opium poppy (Figure 3.9). The RT-PCR shows
a considerably higher abundance of 2OMT transcripts in root which is, to a certain degree, in
agreement with the FPKM values obtained through RNAseq, which showed high levels of
2OMT expression in roots compared to stem. The implication is that 2OMT would be expressed
and active in root tissue. RNA gel blot analysis of 7OMT revealed the highest level transcript
abundance in bud followed by stem, with almost no expression in capsule, leaf or root
(Ounaroon et al. 2003). The transcript abundance of 2OMT in roots correlates with reports
suggesting cryptopine and rhoeadine alkaloids accumulate in the roots of opium poppy (Farrow
and Facchini 2013). Figure 3.10 shows the proposed role of 2OMT in vivo from scoulerine.
Cheilanthifoline is made from scoulerine by CFS. Sinactine the product of 2OMTs Omethylation of cheilanthifoline. Cheilanthifoline is subsequently N-methylated by TNMT to
form cis-N-methylsinactine. The protopine backbone is formed by the 14-hydroxylation of cisN-methylsinactine to form cryptopine. Cryptopine would then be converted to the rhoeadine
alkaloids N-methylprophyroxine and glaudine by unknown enzymes. Future studies should
investigate the enzymatic steps involved in the conversion of cryptopine to rhoeadine alkaloids.
A good first step would be to do a comparative transcriptomic analysis of plant species that do
and do not produce rhoeadine alkaloids but produce cryptopine. The study would look in
particular for transcripts that code for oxidative enzymes present in the plants producing
rhoeadine alkaloids and are absent in non-rhoeadine producing species. This evidence described
75
in this chapter reinforces the in vivo role of 2OMT as the missing step in cryptopine metabolism
although virus-induced gene silencing of 2OMT would be necessary to establish the true in vivo
role of 2OMT.
3.4 Conclusions
Next-generation transcriptomic databases are valuable tools for homology based
discovery of novel genes involved in plant specialized metabolism. This platform was used to
identify and functionally characterize a novel OMT from opium poppy providing insight into
BIA chemical diversity.
76
Figure 3.10 The proposed in vivo role of 2OMT in the biosynthesis of cryptopine and
rhoeadine alkaloids.
Enzymes for which the corresponding genes have been isolated are labelled in green.
Enzymes that remain unknown are represented by question marks. Abbreviations used
are as follows: CFS, cheilanthifoline synthase; 2OMT, protoberberine 2-Omethyltransferase; TNMT, tetrahydroprotoberberine N-methyltransferase; MSH, Nmethylstylopine 14-hydroxylase.
77
CHAPTER FOUR: TARGETTED ALKALOID PROFILING OF TWENTY
BENZYLISOQUINOLINE PRODUCING SPECIES
4.1 Introduction
Benzylisoquinoline alkaloids are made almost exclusively by the plant families
Papaveracea, Ranunculaceae, Berberidaceae and Menispermaceae. Despite their restriction to
only a small number of plant families BIAs are one of the largest and most diverse groups of
alkaloids. The knowledge of BIA diversity across different species is the result of limited
metabolite profiling of cell culture and plant tissue employing various analytical techniques
including UV-Visible spectroscopy, Fourier transform infrared spectroscopy (FTIR), NMR,
HPLC, MS, circular dichroism and various combinations of the aforementioned techniques
(Chintalwar et al. 2003; Farrow, Hagel, and Facchini 2012; Hook, Sheridan, and Wilson 1988;
Ikuta and Itokawa 1988; Iwasa et al. 2008; Liscombe et al. 2009).
Electrospray ionization (ESI) coupled with MS has been adopted as one of the most
powerful techniques for the analysis of BIAs due to its high sensitivity and selectivity. Triple
quadrupole (QqQ), ion traps and time-of-flight (TOF) mass analyzers have all been used to
identify BIAs (Fabre et al. 2000; Le et al. 2014; Li et al. 2010; Ren et al. 2007; Zhang et al.
2006). These studies have provided indispensible data regarding the accurate mass and
fragmentation patterns of several different BIAs.
The monophyletic evolution of BIA metabolism (Liscombe et al. 2005) provides strong
support for the idea that the routes to different BIAs are achieved through similar pathways
across species. This assumption has led to the homology-dependent cloning of biosynthetic gene
orthologs from different species (Farrow et al. 2012). Although several BIA biosynthetic genes
have been identified, the enzymes responsible for the formation of most BIAs remain largely
78
uncharacterized. The biochemical snapshots obtained through multi-stage mass spectrometry
can offer substantial clues, when integrated with transcriptomic datasets, it can aid in the
selection and interpretation of new gene candidates for the discovery of new BIA biosynthetic
genes. Similar initiatives have already been undertaken, for example, the transcript and
metabolite profiling of cell cultures for 18 BIA producing species (Farrow et al. 2012). In this
study cell cultures representing BIA producing plant families Papaveracea, Ranunculaceae,
Berberidaceae and Menispermaceae were used to construct and annotate Sanger-based EST
libraries. The cell cultures were also subjected to targeted metabolite profiling by liquid
chromatography coupled to an ESI source coupled with a QqQ mass analyzer. The two datasets
were integrated by creating BIA metabolic networks based on identified or annotated compounds
that fit into previously proposed biosynthetic routes. Next biosynthetic gene candidates from
each species were used to fill in known and unknown biosynthetic steps by annotations assigned
by tBLASTn analysis. An integrated framework of this nature can provide guidance for the
functional characterization of homologous gene candidates and unknown gene candidates.
Additional benefits of an integrated transcriptomic and metabolic approach across species that
share some aspects of their metabolism is that it facilitates the isolation of genes that catalyze the
same reaction across different plant species. The availability of enzyme variants catalyzing the
same reaction is of particular interest to the emerging field of synthetic biology and could aid in
the assembly of BIA pathways in microorganism (Hawkins and Smolke 2008; Minami et al.
2008; Nakagawa et al. 2011).
This chapter describes the use of targeted alkaloid profiles for twenty BIA producing
species representing four different families of BIA producing plants, Papaveracea,
Ranunculaceae, Berberidaceae and Menispermaceae. Alkaloids were extracted from plant
79
tissues and analyzed using LC-ESI/MS/MS. The resulting retention times and CID patterns were
compared to a series of BIA standards for identification in accordance with guidelines set-out by
MIAMET (Katajamaa and Oresic 2007). The data generated by these experiments represent a
piece of a larger initiative which will integrate the aforementioned targeted metabolite profiles
with high resolution FTICR-MS data along with Roche 454 and Illumina next-generation
transcriptomic database for each species.
4.2 Results
4.2.1 Targeted alkaloid profiling by LC-MS/MS
Initial analysis of alkaloid extracts from the twenty species was performed in Full Scan
mode. Alkaloid extracts were made by grinding and subsequently lyophilizing plant tissues snap
frozen in liquid nitrogen. Alkaloids were extracted from the freeze-dried ground plant tissue
using Bieleski solution. The resulting chromatograms and mass spectra were generated by
scanning between m/z 200-700. The chromatogram and mass spectrum for each plant were used
to select compounds for further structural study. Compounds were selected for MS/MS by
extracting the mass spectrum of major peaks found on the chromatogram. The study focused on
fragmenting compounds with m/z that matched the m/z of the standards. In addition, the
constituent mass-to-charge ratios of major peaks were also selected for further analysis by CID.
CID was used to generate fragmentation data for compounds found in the full scan of the
20 plant species. The fragmentation pattern and retention times were compared the
fragmentation pattern and retention times of chemical standards for identification listed in table
4.1. The annotated chromatograms for each species of BIA producing plant is shown in figure
4.1. Argemone mexicana contains three BIA from the list of standard compounds which include
(10) berberine, (16) protopine and (19) allocryptopine. No compounds were identified in
80
Chelidonium majus. Papaver bracteatum contains (4) thebaine and (6) boldine. Stylophorum
diphyllum contains (5) stylopine and (7) scoulerine. (9) sanguinarine, (15) chelerythrine, (16)
protopine, (19) allocryptopine and (21) cryptopine were all found in Sanguinaria canadensis.
Eschscholzia californica produces (8) reticuline, (16) protopine and (19) allocryptopine. A rich
array of alkaloids were detected in Glaucium flavum including, (6) boldine, (7) scoulerine, (9)
sanguinarine, (13) isocorydine, (15) chelerythrine, (16) protopine, (17) glaucine and (19)
allocryptopine. Corydalis cheilanthifolia produces (5) stylopine, (6) boldine and (16) protopine.
No alkaloids could be positively identified in Hydrastis canadensis or Nigella sativa however,
(10) berberine and (16) protopine were identified in Thalictrum flavum samples and (10)
berberine was found in Xanthorhiza simplicissima. Mahonia aquifolium produces (9) reticuline,
(10) berberine and (13) isocorydine. Only (10) berberine could be identified from Berberis
thunbergii. No alkaloids could be identified in Jeffersonia diphylla. Several alkaloids including
(6) boldine, (8) reticuline, (10) berberine, (13) isocorydine and (16) protopine were positively
identified in Nandina domestica. No alkaloids could be identified from any of the species
surveyed from the family Menispermaceae.
Alkaloid distribution is to some degree, different across families of plants. For example,
the family Papaveracea seems to have the widest distribution of different BIAs of all the families
surveyed, however it is worth noting that 8 species from the family Papaveracea were
investigated while only 4 were species were investigated from each Ranunculaceae,
Berberidaceae and Menispermaceae. Papaveracea was the only family studied that contained
benzo[c]phenanthridine and morphinan alkaloids. The alkaloid subclasses, aporphine,
protoberberine and protopine were observed in all of the plant families with the exception of
Menispermaceae. The alkaloids (S)-boldine, berberine and protopine were the most widespread
81
alkaloids in this study and were found in 4, 6 and 7 plant species respectively and were the only
alkaloids to be positively identified across the families Papaveracea, Ranunculaceae and
Berberidaceae. No alkaloids were identified in the family Menispermaceae.
Some trends also develop if the distribution of alkaloids are considered as a function of
tissue/organ type. For example, the benzo[c]phenanthridine alkaloids sanguinarine and
chelerythrine are only observed in subterranean tissues such as rhizomes and roots. The
morphinan thebaine is only found in stem tissue and the 3’-O-methylated aporphine, glaucine
was only detected in root tissue. Protoberberine alkaloid types were identified in all of the
tissue/organ types except callus. The some alkaloids were observed in multiple tissue types, for
example, boldine, protopine and allocryptopine were found in stem, rhizome and root. No
alkaloids were identified in callus tissue tested in this study.
82
Table 4.1 Chemical standards used for the targeted alkaloid profiling of 20 BIA producing
species by LC-MS/MS.
The number of the compound is there for illustrative purposes in figure 4.1. The common
name of each alkaloid in indicated as well as the m/z ratio of the molecular ion or the
protonated parent ion [M]+ or [M+H]+. The retention time is indicated by RT and is
expressed in minutes. CE indicates the collision energy in eV used in CID experiments.
ESI-CID spectrum m/z indicates the m/z of the fragments resulting from the CID of a given
compound and the number in brackets represents the relative intensity of the ion in the
spectrum. The structure is also illustrated in the table.
83
No.
1
Compound
Morphine
[M+H]+
or [M]+
286.2
RT
(min)
1.01
CE
(eV)
ESI-CID Spectrum
m/z (Relative
Intensity)
25
286 (100), 229
(11.01), 211 (11.97),
209 (8.65), 201
(27.03), 193 (6.91),
185 (13.14), 183
(11.34), 180.9 (8.42),
173 (12.69), 165
(12.95), 157 (5.02),
155 (11.35), 147
(6.32), 145 (5.78), 58
(11.42), 44 (6.78)
298
(1),
234
(1),
(2), 283 (1), 267
249 (3), 237 (1),
(4), 223 (1), 221
218 (8), 196 (5),
58 (100)
300 (100), 282 (5.51),
243 (9.69), 241
(5.95), 225 (16.09),
215 (26.76), 209
(5.54), 199 (16.42),
193 (7.43), 187
(10.45), 183 (15.64),
181 (7.1), 165
(11.75), 161 (8.53),
155 (7.47), 58
(16.33), 44 (6.68)
312 (2), 281 (2), 266
(4), 255 (1), 251 (11),
249 (2), 237 (1), 234
(2), 223 (2), 221 (7),
218 (4), 195 (2), 177
(1), 58 (100)
2
Oripavine
296.2
3.5
25
3
Codeine
300.2
3.52
25
4
Thebaine
312.2
5.43
25
5
(S)-Stylopine
324
11.25
25
324 (20.4), 176 (100),
149 (39.94), 119
(6.39)
25
297 (14.39), 282
(16.54), 267 (6.43),
266 (9.19), 265
(90.58), 237 (100),
233 (14.66), 222
(8.36), 205 (32.45),
177 (8.96), 44 (8.79)
6
(R,S)-Boldine
328.2
4.98
84
Structure
7
(S)-Scoulerine
328
6.21
25
8
(S)-Reticuline
330
4.76
25
9
Sanguinarine
332.2
8.26
25
10
Berberine
336
8.02
25
11
(R,S)-Canadine
340
10.28
25
12
Papaverine
340
8.47
25
13
(S)-Isocorydine
342.2
6.97
25
14
(R,S)Tetrahydropapaverine
344
6.23
25
15
Chelerythrine
348.2
8.24
25
85
328 (9.02), 178 (100),
151 (10.77)
192 (100), 177 (8.09),
175 (19.32), 151
(5.45), 143 (25), 137
(41.49)
332 (100), 330 (6.14),
317 (15.91), 304
(22.86), 302 (7.52),
274 (14.07)
336 (45.63), 321
(56.34), 320 (100),
306 (22.08), 304
(16.43), 292 (83.38),
278 (5.47), 275 (5.7)
340 (9.82), 176 (100),
149 (9.5)
340 (70.91), 325
(7.91), 324 (74.12),
296 (11.54), 202
(100), 171 (15.27)
311 (13.26), 296
(36.67), 281 (30.93),
280 (34.69), 279
(100), 267 (5.76), 265
(39.91), 264 (84.05),
251 (24.03), 248
(64.06), 247 (18.41),
236 (40.49), 235
(5.61), 219 (9.84),
191 (8.47)
296 (6.8), 192 (100),
189 (33.2), 174
(16.15), 158 (11.77),
151 (51.84)
348 (45.51), 333
(37.27), 332 (100),
318 (31.94), 316
(8.2), 315 (8.48), 304
(56), 290 (8.73)
354 (59.25), 336
(9.18), 271 (16.84),
265 (6.09), 247 (9.8),
206 (17.05), 189
(79.71), 188 (100),
177 (6.22), 175
(5.93), 165 (14.85),
149 (46.29), 135
(6.12)
325 (8.04), 310
(37.78), 295 (30.25),
294 (100), 279
(27.57)
16
Protopine
354.2
6.81
25
17
(S)-Glaucine
356.2
7.64
25
18
(R,S)Tetrahydropalmatine
356
9.58
25
356 (10.45), 192
(100), 165 (22.38),
150 (5.5)
19
Allocryptopine
370.2
6.99
25
370 (36.41), 352
(18.72), 337 (5.03),
336 (7.29), 321
(5.84), 306 (7.59),
290 (31.63), 206
(27.8), 191 (6.21),
190 (8.31), 189
(34.38), 188 (100),
181 (18.47), 166
(6.37), 165 (13.62),
151 (9.34), 149 (9.71)
20
(S)-Canadaline
370.2
6.85
25
290 (12.64), 190
(100)
25
370 (68.18), 352
(10.46), 339 (5.46),
321 (9.4), 311 (6.87),
291 (15.92), 290
(5.28), 283 (5.57),
263 (10.91), 222
(16.66), 206 (8.09),
205 (69.54), 204
(100), 194 (8.21), 193
(11.48), 190 (32.49),
175 (10.43), 165
(89.48), 151 (5.35),
150 (12.28), 149
(23.01), 135 (5.56)
21
Cryptopine
370.2
6.65
86
22
Hydrastine
384.2
7.68
25
369 (7.93), 366
(10.08), 354 (17.54),
351 (65.15), 336
(100), 333 (5.55), 308
(6)
23
Noscapine
414.2
10.67
25
414 (5), 365 (18.39),
323 (5.23), 220 (100),
206 (5), 179 (6.43)
87
88
89
90
Figure 4.1 Annotated LC-MS chromatograms from 20 BIA producing species.
The numbers on some chromatograms indicate compounds identified by comparing the
retention times and CID patterns to the standards in figure 4.1. (A) Argenome mexicana;
(B) Chelidonium majus; (C) Papaver bracteatum; (D) Stylophorum diphyllum; (E)
Sanguinaria Canadensis; (F) Eschscholzia californica; (G) Glaucium flavum; (H) Corydalis
cheilanthifolia; (I) Hydrastis canadensis; (J) Nigella sativa; (K) Thalictrum flavum; (L)
Xanthorhiza simplicissima; (M) Mahonia aquifolium; (N) Berberis thunbergii; (O)
Jeffersonia diphylla; (P) Nandina domestica; (Q) Menispermum canadense; (R) Coculus
trilobus; (S) Tinospora cordifolia; (T) Cissampelos mucronata.
91
4.3 Discussion
Metabolomics can be used as a method for the discovery of novel BIA biosynthetic genes
particularly when used in conjunction with next-generation transcript profiles. Alkaloid extracts
from 20 different BIA producing species from the families Papaveracea, Ranunculaceae,
Berberidaceae and Menispermaceae were analyzed by LC-MS/MS to generate targeted alkaloid
profiles. This work helps to reveal the diversity of BIA metabolic networks in these unique plant
systems. The positive identification of alkaloids in any given plant implies the existence of
upstream enzymes and metabolites from a specific BIA metabolic branch. Such information,
used on its own or when integrated with transcriptomic information, can guide the search for
novel BIA biosynthetic genes and produce higher quality gene candidates. The QqQ based
targeted metabolite profiling discussed in this chapter will ultimately be combined with a
metabolite profiling by FTMS and NMR as well as next-generation transcriptomic datasets
generated by Roche-454 and Illumina (Xiao et al. 2013). The integration of these complimentary
datasets will facilitate the mapping of an extended collection of chemical and genetic
components of the combined BIA metabolic network of 20 different plant species. The ultimate
goal is to demonstrate how precise metabolic profiles can be used to predict enzyme function, to
generate a catalogue of orthologous genes with slightly different catalytic specificities and
enzymatic properties to aid in synthetic biology pathway engineering and discover novel genes.
4.3.1 Simple BIAs
The fragmentation patterns of alklaoids with the 1-benzylisoquinoline backbone are
relatively well studied and several characteristic fragmentation mechanisms have been proposed
(Schmidt et al. 2005; Schmidt et al. 2007). Typically the fragmentation of simple BIAs occurs at
the α-carbon which results in the formation of separate isoquinoline and benzyl ion moieties
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(Schmidt et al. 2005; Schmidt et al. 2007). Fragmentation at the α-carbon gives clues to the Omethylation pattern of the alkaloid. Fragmentation of the 1-benzylisoquinoline backbone also
generates naphthalene-type ions resulting from loss of an ammonia or methylamine from the
isoquinoline nitrogen of the alkaloids (Schmidt et al. 2005; Schmidt et al. 2007). The loss of
ammonia or methylamine indicates the N-methylation state of the alkaloid.
The CID spectrum generated for (S)-reticuline follows the fragmentation pattern
described in the literature (Schmidt et al. 2005). The reticuline parent ion [M+H]+ has a m/z of
330 and is fragmented at the α-carbon with a collision energy of 25 eV to form an ion with a m/z
of 192 and 137. The 192 ion represents the N-methylated and O-methylated isoquinoline moiety
of reticuline and the 137 ion represents the O-methylated benzyl ion. The m/z 175 and m/z 143
ions represent the naphthalene-types ions formed upon the loss of a methylamine, subsequent
fragmentation at the α-carbon and rearrangement of the isoquinoline moiety.
In this study, reticuline was the only simple BIA surveyed for and was detected in only
three species, E. californica, M. aquifolium and Nandina domestica. (S)-Reticuline is considered
a branch-point metabolite in BIA metabolism and gives rise to protoberberines, protopines,
benzo[c]phenanthridines, phthalideisoquinolines, aporphines, rhoeadines and morphinans
(Beaudoin and Facchini 2014). The limited detection of reticuline in this study and the apparent
low basal levels of reticuline among these 20 species could be the result of the considerable
metabolic flux of reticuline to other alkaloid types (Beaudoin and Facchini 2014). It is also
important to note that the undetectable levels of reticuline in this study do not indicate reticuline
is not present, simply that reticuline was not detected. It is possible that reticuline was present
below the limit of detection of the QqQ in several species in which it was not detected.
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4.3.2 Protoberberine alkaloids
Several protoberberine alkaloids were detected in the families Papaveracea,
Ranunculaceae and Berberidaceae. The fragmentation of protoberberine alkaloids has been
investigated in considerable detail (Schmidt et al. 2007). (S)-Scoulerine is the first
protoberberine alkaloid made by the FAD-dependent C-C coupling the branch-point intermediate
(S)-reticuline by BBE (Ziegler and Facchini 2008). The CID mass spectrum of scoulerine is
characterized by the key ions at m/z 178 representing the singly O-methylated isoquinoline
moiety and another ion at m/z 151, represents the complementary singly O-methylated benzylic
portion of the molecule (Schmidt et al. 2007). The fragmentation of the protoberberines,
tetrahydropalmatine and stylopine are similar to that of scoulerine. The CID mass spectrum for
tetrahydropalmatine show a fragment of m/z 192, which represents the dimethylated isoquinoline
moiety, and of m/z 165 representing the dimethylated benzylic ion. Stylopine is very similar in
structure to tetrahydropalamatine however the double O-methylation of the isoquinoline and
benzylic ions are replaced with methylenedioxy bridges. Owing to stylopine’s methylenedioxy
bridges the ions observed in the CID mass spectrum derived from the isoquinoline and benzylic
moieties are m/z 176 and 149 respectively. The CID mass spectrum of canadine is somewhat
different from the CID patterns of other protoberberine alkaloids described above. The expected
m/z 176 ion is derived from the isoquinoline moiety containing a methylenedioxy bridge.
However, no ion with m/z 165 is present, which would in most cases indicate the doubly Omethylated benzylic moiety. In its place there is an ion with m/z 149, which implies the loss of
16 mass units. Although the mechanism for canadine’s fragmentation is unknown the loss of 16
could be explained by the loss of –CH2 or of -O.
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Berberine is structurally unique from the other protoberberine alkaloids examined in this
study in that the ring adjacent to the benzylic moiety is aromatic. This additional aromatic ring
results in a unique CID mass spectrum relative to scoulerine, tetrahyrdopalamatine and canadine.
A handful of studies have reported MS/MS information for berberine and other similar alkaloids
(Ren et al. 2007; W. Wu et al. 2005). Although no fragmentation scheme has been proposed for
berberine it seems the data collected in this study is in agreement with the literature. Rather than
a cleavage separating the isoquinoline and benzyl moieties, as is the case with most
protoberberines, there is a loss of functional groups from the backbone of the parent ion. For
example, berberine with a m/z of 336 generated fragment ions with m/z 321 and 320 which can
be explained by the loss of –CH3 and –CH4 respectively, perhaps through the cleavage of the
methylenedioxy bridge. The fragment m/z 306 can be generated from the parent ion by the
cleavage of two CH3 groups. While the fragment ion m/z 292 can be generated by the loss –
C2H4O from the parent ion (Wu et al. 2005).
Scoulerine was detected in only two plants from the family Papavercacea, S. diphyllum
and G. flavum. Since the presence of many other downstream BIA subclasses such as,
protoberberine, protopine, benzo[c]phenanthridine and phthlideisoquinoline, are predicated on
the ability of a plants to make scoulerine, it is entirely possible that scoulerine is made by many
other plants however the flux through scoulerine metabolism and accumulation of downstream
alkaloids made from scoulerine resulted in extremely low basal levels of scoulerine in most
plants. Stylopine was detected in the families Papaveracea and Ranunculacea in the species S.
diphyllum and H. canadensis. Recent alkaloid characterizations of H. canadensis roots were
performed using Orbitrap LS-MSn and ultra performance liquid chromatography quadrupole
time-of-flight mass spectrometery (UPLC-QTOF-MSn) (Le, Mccooeye, and Windust 2012; Le,
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Mccooeye, and Windust 2014). Both studies reported several BIAs that were not detected using
the approach described in this work, conversely we detected some alkaloids that they did not.
For example, in this work stylopine was detected but not in the Orbitrap or QTOF studies. The
most obvious reason for this discrepancy would be the analysis of different tissue types. In both
the Orbitrap and QTOF studies, alkaloids were extracted from root while in this study alkaloids
were extracted from rhizome. Berberine was the most widespread protoberberine alkaloid
detected in this study. Berberine was found in the A. mexicana in the family Papaveracea.
Berberine was also found in the species T. flavum and X. simplicissima in the family
Ranunculaceae. Berberine seems to accumulate in the family Berberidaceae. The species M.
aquifolium, B. thunbergii and N. domestica all accumulate berberine. There seems to be a
certain correlation between tissue type and berberine accumulation. Excluding A. mexicana and
M. aquifolium, berberine seems to be commonly found in roots. This type of information could
guide future work on berberine biosynthesis and the search for gene orthologues involved in
berberine biosynthesis in other plants species.
4.3.3 Protopine alkaloids
In general, the fragmentation behavior of protopine alkaloids share similarities with the
fragmentation behavior of other BIAs. Protopines main fragmentation routes create ions derived
from the isoquinoline moiety and the benzyl moiety. Typically protopines will have three
isoquinoline derived ions, one of lower mass consisting only the amine, another of higher mass
containing the amine and an alcohol group, which is subsequently lost as H2O to make the third
isoquinoline derived ion (Schmidt et al. 2007). In addition, there will be at least one ion derived
from the benzyl moiety. For example, the alkaloid protopine, m/z 354, will fragment to create an
ion at m/z 189 corresponding to the isoquinoline moiety without the alcohol group. Ions at m/z
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206 and 188 will also be made corresponding to the isoquinoline ion with the alcohol group and
the dehydrated isoquinoline ion. Ion m/z 149 represents the benzylic ion. Protopine and
cryptopine are unique in that they make an additional benzylic ion, that is not observed in the
mass spectrum of allocryptopine, with an alcohol group at m/z 165 owing perhaps the presence
of the benzylic methylenedioxy bridge. The differences between the CID mass spectra of
protopine, allocryptopine and cryptopine are the result of differing O-methylation patterns as is
the case with protoberberine alkaloids.
Protopine alkaloids were detected in the families Papaveraceae, Ranunculaceae and
Berberidaceae. Protopine alkaloids were found in the stem, root and rhizome of various plants.
The biosynthesis of protopine is catalyzed by the 14-hydroxylation of (S)-cis-N-methylstylopine
by MSH, which leads to ring tautomerization by cleavage of the C-N bond and the formation of a
C14 keto moiety (Martina Rueffer and Zenk 1987; Beaudoin and Facchini 2013). This
cytochrome P450 catalyzed reaction is prerequisite to the biosynthesis of benzo[c]phenanthridine
alkaloids like sanguinarine (Beaudoin and Facchini 2014). Allocryptopine and cryptopine are
also enzymatic products of MSH that are made in parallel to protopine by the 14-hydroxylation
of (S)-cis-N-methylcanadine and (S)-cis-N-methylsinactine respectively (Beaudoin and Facchini
2013). Protopine was the most widespread of all the targetted protopine alkaloids. In the family
Papaveraceae, protopine was detected in A. mexicana, S. canadensis, E. califorinica and G.
flavum. Interestingly, in the family Papaveraceae, the plants which produce protopine also
produce allocryptopine. This shared metabolic feature suggests some degree of shared
metabolism. Of course the ability of a given plant to produce protopine and allocryptopine is
tied to the expression of MSH, but it seems it is also tied to the ability of the plant to express
TNMT which N-methylated the protoberberine backbone (Liscombe and Facchini 2007).
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Perhaps genes invovled in protopine alkaloid metabolism are in some way linked. There is new
evidence supporting gene clusters involved in BIA metabolism like the 10-gene cluster involved
noscapine biosynthesis in P. somniferum (Winzer et al. 2012). It seems however that the link
between protopine and allocryptopine metabolism does not extend into the other families
observed in this study (allocryptopine was not detected in the families Ranunculaceae,
Berberidaceae and Menisperaceae), which may mean that the possible linking of genes
regulating protopine alkaloid metbaolism are only present in the Papaveracea lineage.
Cryptopine was only detected in S. canadensis of the family Papaveracea. Cryptopine is made
from scoulerine by the formation of a methylenedioxy bridge by cheilanthifoline synthase to
make cheilanthifoline (Figure 3.11). Cheilanthifoline is subsequently 2-O-methylated by
protoberberine 2OMT to make sinactine which is then N-methylated by TNMT to make (S)-cisN-methylsinactine. (S)-cis-N-methylsinactine is then 14-hydroxylated to make cryptopine by
MSH. Cryptopine is considered to the the source of rhoaedine alkaloids (Farrow and Facchini
2013). The involvement of 2OMT in cryptopine metabolism suggest that S. canadensis may be a
good candidate in which to search for opium poppy 2OMT enzyme orthologs due to the presence
of cryptopine.
Protopine was also detected in H. canadensis and T. flavum of the family Ranunculaceae.
Protopine was also found in N. domestica of the family Berberidaceae. No protopine alkaloids
were detected in the family Menispermaceae.
4.3.4 Benzo[c]phenanthridine alkaloids
Little is known of the fragmentation properties of benzo[c]phenanthridine alkaloids. One
characteristic fragment of benzo[c]phenanthridine alkaloids is the loss of m/z 58 from the parent
ion. It is thought that the loss of m/z 58 is the result of losing -CH2-CO from the parent ion
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(Frick et al. 2005; Schmidt et al. 2007). Both sanguinarine and chelerythrine demonstrate the
characteristic loss of m/z 58. The alkaloids sanguinarine and chelerythrine were only detected in
the family Papaveracea in the species S. canadensis and G. flavum. In both cases the there is
evidence of the upstream metabolism required for the biosynthesis of benzo[c]phenanthridine
alkaloids. S. canadensis and G. flavum both produce protopine, which is upstream of
sanguinarine, and allocryptopine, which is upstream of chelerythrine (Beaudoin and Facchini
2013; Takao, Kamigauchi, and Okada 1983).
4.4.5 Aporphine alkaloids
The fragmentation of aporphine alkaloids has been well studied and fragmentation
schemes for their dissociation have been suggested (Schmidt et al. 2007; Stévigny et al. 2004;
Wu and Huang 2006). The aporphine alkaloids fragmented in this study share certain structural
characteristics which lead to similar fragmentation patterns. Boldine, isocorydine and glaucine
all contain methylated amino groups, therefore all parent ions loose -CH2NH2 (loss of 31 Da).
Subsequent losses from [M+H-RNH2]+ are dependant on the substitution patterns found on the
aromatic rings and are commonly losses of CH3OH (31 Da), CO (28 Da) or radical losses of CH3. (15 Da) of -OCH3. (31 Da). For example, in the case of isocorydine, m/z 342, the first loss
is of the –CH2NH2 creating [M+H-RNH2]+ at m/z 311. From this point the fragmentation route
appears to split, one route involves the loss of two -CH3. creating ions at m/z 296 and 281.
Alternatively, m/z 311 can lose –CH3OH to form m/z 279. The m/z 279 ion then looses, -CO, OCH3. and -CH3. to make ions m/z 251, 248 and 264 respectively. The ion m/z 264 is further
fragmented in the same manner as described above to complete the mass spectrum.
Aporphine alkaloids are found in the families Papaveraceae, Ranunculaceae and
Berberidaceae. Among the family Papavercareae, boldine is found in P. bracteatum and G.
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flavum. Isocorydine and Glaucine are also found in G. flavum. Only one aporphine, boldine, was
found in the family Ranunculaceae in the species H. canadensis. From the family
Berberidaceae, two species contained aporphines. Isocorydine was detected in M. aquifolium,
while in N. domestica, boldine and isocorydine were detected. No aporphines were detected in
the family Menispermaceae.
Aporphine metabolism is poorly understood and much of what is known comes from
tracer studies and suggests glaucine is ultimately derived from reticuline in three steps.
Although the order is unknown, an oxidation forming the aporphine bridge between carbon 8 of
the isoquinoloine moiety and the 5’ carbon of the benzyl moiety is required, in addition to two
O-methylation steps (Bhakuni and Jain 1988). The fully O-methylated glaucine is of
considerable interest because at some point in its biosynthesis it was methylated at what is
equivalent to the 3’ position of the simple BIA backbone suggesting the existance of an enzyme
with 3’-O-methyltransferase activity.
4.4.6 Morphians
The fragmentation patterns of morphinans have been described in the literature to some
extent (Raith et al. 2003; Schmidt et al. 2005). In most cases, the morphinan parent ion loses an
ion at m/z 58 corresponding to the CH3 CHNHCH3+ which is derived from the N-methyl group of
the alkaloid. Through the same process the parent ion can also lose 57 Da which produces an
ion, in the case of thebaine, at m/z 255. Further fragmentation of the thebaine derived ion, m/z
255, produces a loss of 32 Da, –CH3OH, which creates and ion with m/z 223. The ion m/z 223
looses 28 Da, -CO, to make an ion at m/z 195 which subsequently looses –H20, 18 Da, to
produce an ion with m/z 177. Variations of this fragmentation pattern for different morphinan
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species like codeine or morphine are largely due to differing oxidation states of the alcohol
groups found on the compounds.
Morphinan alkaloids are not widespread in the plant kingdom and are restricted to the
family Papavercacea with the exception of some reports in the family Euphorbiaceae (Beaudoin
and Facchini 2014; Theuns et al. 1986). In this study, the only morphinan detected was thebaine
in P. bracteatum. The identification of thebaine in P. bracteatum coroborates previous reports
which have detected thebaine in P. bracteatum (Ziegler et al. 2009). The presence of thebaine in
P. bracteatum suggest the presence of upstream morphinan metabolites and the genes
responsible for its biosynthesis. Based on what we know of morphinan biosynthesis it is likely
that the morphinans salutaridinol and salutaridine are made in P. brateatum as well as the simple
BIA (R)-reticuline.
4.3.6. Limitations of targetted alkaloid profiling
As with any technique, the LC-MS/MS approach used in this study presents certain
limitations. Targeted alkaloid profiling by LC-MS/MS is only capable of identifying alkaloids
for which a standard is available (Katajamaa and Oresic 2007). Often in the case of secondary
metabolites, chemical standards are difficult to synthesize and/or are incredibly expensive which
often becomes a limiting factor in the identification of alkaloids from a given sample. The
scarcity of chemcial standards also presents issues with quantification, in that generating
standard curves can become prohibitively expensive. Other techniques, such as NMR, are
capable of identifying chemicals without the use of a standard. Despite this obvious strength of
NMR as a technique for metabolite identification, it suffers from a lack of sensitivity when
compared to mass spectrometry based techniques (Verpoorte et al. 2008). In many cases the
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concentration of secondary metabolites in planta are below the limit of detection for most NMR
instruments.
As with most metabolomic initiatives, other methodological decisions and limitations
have no doubt effected the results of this study. One such area of concern in this study revolves
around harvesting of the plants or cell cultures. Firstly, most of the plants use in this analysis
were grown under a wide variety of conditions. Due to the narrow distribution of some of these
plants in nature and in greenhouses across Canada we had very little control to ensure that the
plants were grown under uniform conditions. It is entirely possible that variations in growing
conditions resulted in changes to the alkaloid profile of the plants analyzed in this study (Kim
and Verpoorte 2010). In addition to the growing conditions, many plants exhibit diurnal changes
in primary metabolites as well as secondary metabolites levels (Urbanczyk-Wochniak et al.
2005). Due to limited access to the plants studied in this work, diurnal effects could not be
controlled for. Plants were snap frozen in liquid nitrogen immediately following their harvesting
to limit oxidation of metabolites, enzymatic reactions and plant wounding responses. The
possibility also exists that analyzing a single tissue type from each plant resulted in false
negatives in the analysis. It is well established that BIAs are biosynthesized in a tissue specific
manner (Facchini et al. 2007). The reach of this study could have been extended by analyzing
different tissue types for each plant. At least two studies in the literature examine the alkaloids
of H. canadensis and support the possibility that there are differences in the alkaloid profiles of
tissues from H. canadensis (Le, Mccooeye, and Windust 2012; Le, Mccooeye, and Windust
2014). Both of the aforementioned studies extracted alkaloids from root compared to this study
in which we analyzed rhizome. Although many of the same alkaloids were detected in both sets
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of studies, root tissue appears to contain some additional alkaloids such as, hydrastine and
berberine.
Only cell cultures were available for three out of four species in the Menispermaceae
family. As a result we were limited to using plant cell culture in some cases. In opium poppy
there is a distinction between the alkaloid profiles of tissues harvested from the plant and cell
culture and in most studies alkaloid production is induced using a fungal elicitor. Therefore the
absence of detectable alkaloids in the family Menisperaceae could be the result of low basal
alkaloid levels in callus versus normal plant tissues.
Extraction of alkaloids from the plant matrix can also introduce error into experiments.
Although we chose a solvent system with hydrophillic and hydrophobic properties, it is probable
that we introduced extraction bias to our results. Added to the issue of using a single solvent
system for extraction is that the extraction of alkaloids from the biological matrix does not
adhere to the basic principles of chemical solvation because the mechanisms of metabolite
extractions from biological samples can be influenced by interactions with the chemical milieu
of the plant (Kim and Verpoorte 2010). In this study a range of different tissues from different
plants were analyzed each with its own biological matrix. In an attempt to improve our
extraction efficiency from distinct biological matrices all plant tissues were ground with a tissue
lyser and sonicated during the extraction process. Due to cost restraints, solid-phase extraction
(SPE) a method often used in MS based metabolomic initiatives, was not used. SPE often works
to remove compounds that may interfere with analysis, such as, salts which dampen MS signals
(Kim and Verpoorte 2010). It is possible that in some samples there were compounds present
that interfered with the analysis but without specific comparative studies is impossible to be sure
to what degree other compounds may have effected the results of this study.
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Following harvesting other considerations can effect the outcome of targeted alkaloid
profiles. The differences in ionization of different chemical species is well established and in my
personal experience I have noticed a difference in the ionization of simple BIAs compared to
other BIA alkaloid subclasses. In this study we used a mobile phase A comprised of ammonium
acetate and acetonitrile and it seems to reduce the signal of simple BIAs relative to the formic
acid mobile phase used for the enzyme assay analysis. Perhaps our choice of mobile phase had
the effect of reducing the signals of reticuline below the limit of detection in the plants samples.
4.4 Conclusions
Conclusions drawn in the targeted metabolic snapshot discussed in this chapter must be
drawn with care and the dynamic nature of the biological systems studied must be considered.
The targeted alkaloids profiling of 20 plants species that produce BIAs was able to identify
several alkaloids in all the families studied except Menispermaceae. The absence of detected
alkaloids by no means implies that it is impossible for a given plant to produce a given alkaloid,
this is due to the limitations in the collection of standards used, that only a single tissue type was
collected from each plant at a single time point in addition to other methodological decisions.
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CHAPTER FIVE: DISCUSSION
5.1 Overview
A biochemical transcriptomics approach has been employed to isolate and functionally
characterize a novel gene involved in BIA biosynthesis. The isolation and functionally
characterization of protoberberine 2OMT to provides insight into the chemical diversity of BIAs
and into the evolution of BIA biosynthesis. Gene candidate mining in Roche 454 and Illumina
NGS transcriptomic libraries of opium poppy stem and root tissue using sequence similarity and
high levels of expression, inferred by the FPKM value of each gene, yielded 10 unique OMT
gene candidates. Genes were synthesized or amplified from cDNA, heterologously expressed in
E. coli and purified by cobalt-affinity chromatography. The biochemical characterization of one
synthesized gene candidate, sharing considerable sequence similarity with opium poppy
reticuline 7OMT, demonstrated O-methylation activity and was named protoberberine 2OMT.
2OMT catalyzed the SAM-dependant 2-O-methylation of protoberberine alkaloids and the 7-Omethylation of simple BIAs. The accumulation of 2OMT transcripts in root agreed with the
FPKM values obtained from the Illumina NGS stem and root databases. The ability of 2OMT to
catalyze the 2-O-methylation of cheilanthifoline and it’s presence in roots suggest that 2OMT
may be involved with cryptopine and rhoeadine alkaloids metabolism.
Alkaloids were extracted from plant tissues and analyzed using LC-MS/MS to generate
targeted alkaloid profiles for 20 different BIA producing species. The plants used in this study
were selected from four different families, Papaveracea, Ranunculaceae, Berberidaceae and
Menispermaceae. A specific tissue type was selected from each plant species and at a single
time point. Alkaloids in the plant extracts were identified by matching their retention times and
CID patterns to the retention times and CID patterns of 23 BIA authentic standards. Ultimately
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the targeted alkaloid profiling described in this study will be combined with high resolution
FTICR-MS data and with NGS data for each plant as part of the Genome Canada PhytoMetaSyn
Project (Facchini et al. 2012; Xiao et al. 2013).
In this chapter, the significance and implications of the result of this study will be
discussed and directions for future research are suggest.
5.2 Future directions
5.2.1 Further characterization of 2OMT
There are several additional experiments that may help to establish the in vivo role of
2OMT. To begin, the amplification of 2OMT by PCR from root tissue would add support to
2OMT being an enzyme relevant to BIA metabolism. Perhaps the reason we could not isolate
the gene from root tissue was due to the developmental stage of the plant. Future attempts to
isolate 2OMT could be carried out at distinct developmental points to increase the chance of
isolating a full length gene.
To increase our understanding of the substrate specificity and methylation sites of 2OMT,
it should be tested with a broader suite of substrates. Assaying 2OMT with the simple BIAs
orientaline, protosinomenine and isoorientalinen would help determine if 2OMT shares the
ability of 7OMT to methylate at different sites on the BIA backbone. Assaying 2OMT with
tetrahydrocolumbamine and other protoberberine could also help determine the regiospecificity
of 2OMT. NMR should be employed to definitively identify the enzymatic products in the
absence of chemical standards.
Due to the limited availability of cheilanthifoline, the kinetic properties of 2OMT for
cheilanthifoline could not be determined experimentally. To add support to the idea that 2OMT
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is involved in the biosynthesis of cryptopine and ultimately rhoeadine alkaloids it will be crucial
to determine the catalytic parameters of 2OMT for cheilanthifoline.
To determine the role of 2OMT in vivo, VIGS should be conducted. Changes in 2OMT
transcript abundance can be monitored by qPCR and changes in the alkaloid profile can be
determined by LC-MS/MS. If indeed 2OMT is involved in the biosynthesis of cryptopine, the
reduction of 2OMT transcript abundance should lead to increased levels of upstream alkaloids
such as scoulerine and cheilanthifoline along with a reduction of downstream alkaloids such as
sinactine, cryptopine and rhoeadine alkaloids, like N-methylporphyroxine and glaudine, which
have been suggested to the downstream products of sinactine (Farrow and Facchini 2013). It
would also be interesting to see if the silencing of 2OMT results in a significant increase in the
abundance of simple BIAs.
5.2.2 Future targeted metabolite profiling
Future targeted metabolite profiling could add value and information to the metabolic
picture described in this study. Expanding the catalogue of BIA standards would allow us to
increase the number of alkaloids annotated in any given LC-MS based study. Additionally, it
would be valuable to develop an understanding the effect of developmental stage on alkaloid
production. Time course experiments that would allow us to extract alkaloids from distinct
developmental stages would increase our understanding of what each plant makes, when it
makes it and allow us to optimize the isolation specific biosynthetic genes based on the
abundance of related alkaloids in a given developmental stage. For example, if a certain
developmental stages shows high abundance of a certain alkaloid , it is plausible there would
higher transcript abundance of genes involved in the production of that alkaloid.
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Additional characterization of the existing samples by an alternate analytical method
could provide an interesting comparative analysis of the benefits of using one analytical method
over another. For example, analysis of the same samples using an LTQ Orbitrap could provide a
large increase in resolution, increased mass accuracy and increased dynamic range and could
thus provide a deeper metabolite analysis when compared to the QqQ approach used in this study
(Hu et al. 2005).
5.2.3 Using the targeted alkaloid profiles of twenty BIA producing plants in conjunction with
NGS
Care must be taken when drawing conclusions from any metabolomic initiative. In
essence, the targeted alkaloid profiling of twenty BIA producing plants represents a metabolic
snap shot of a single plant tissue at a single developmental time point. The presence of a given
alkaloid only indicates that a given alkaloid was presence at that time in the plant, while the
absence only means that the alkaloid could not be detected at that time in the plant. Absence
does not mean that the plant cannot make an alkaloid that was not detected. However, with that
being said, the presence of a certain alkaloid in a given plant implies the existence of the
metabolic machinery required to form that alkaloid.
For example, the presence of the alkaloid sanguinarine in S. canadensis implies the
existence of the enzymes responsible for sanguinarine biosynthesis. One would expect to find in
the NGS transcriptomic databases for S. canadensis (Xiao et al. 2013), contigs orthologous to the
opium poppy enzymes, TNMT, MSH, P6H, and DBOX.
Provided the similar orientation of the benzyl moeity in both glaucine and boldine it is
possible the same oxidative enzyme is responsible for the formation of the aporphine bridge in
the biosynthesis of both compounds. The combination of the targeted alkaloid profile described
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in this study could be used in conjunction with NGS transcriptomic data to find the unknown
oxidative enzyme involved in forming the aporphine bridge in glaucin and boldine. Recently, a
P450 was isolated from C. japonica cell culture that catalyzes the formation of corytuberine, an
aporphine, from reticuline (Ikezawa, Iwasa, and Sato 2008). CYP80G2, as it is known,
introduces a C-C bond between carbon 8 of the isoquinoline moiety and the 2’ carbon of the
benzyl moeity of reticuline. The result is a distinct orientation of the benzyl moeity in
corytuberine compared to to glaucine and isocorydine which have the aporphine bridge between
C8 and C5’. CYP80G2 was not reported to form the aporphine bridge between C8 and C5’
suggesting the possibility that the oxidative enzyme involved in the formation of glaucine and
isocorydine is different. Of course it is also a possible that the substrate specificity and binding
pocket of CYP80G2 is different in other species allowing it to catalyze the different aporphine
isoforms. This question could be answered by comparing the alkaloid profiles and
transcriptomic databases of plants that make aporphines. For example, boldine could be detected
in many plants examined in this study, P. bracteatum, G. flavum, H. canadensis, and N.
domestica. Boldine shares the same C8 C2’ aporphine bridge as corytuberine. Aporphines
containing the C8 and C5’ aporphine bridge (isocorydine and glaucine) were only detected in P.
bracteatum, M. aquifolium, and N. domestica. If one is looking for the oxidative enzyme
responsible for the formation of the aporphine bridge between C8 and C5’, CYP80G2 could be
used as a BLAST query sequence to search for similar P450s. All the P450 with a certain degree
of sequence similarity from each aporphine producing species could be compiled in a list and
analyzed in a phylogenetic tree. Amino acid sequences that share a high degree of similarity
with CYP80G2 would cluster close together and would likely not be responsible for the
formation of the aporphine bridge between C8 and C5’. Putative P450 amino acid sequences on
109
the tree that cluster away from CYP80G2 and are not found in the species that produce boldine
exclusively but are present in species that produce isocorydine or glaucine would consititute high
quality gene candidates. The genes could then be isolated from the plant, heterologously
expressed and enzymatically characterized to confirm their function.
Future work could use G. flavum as a model species to isolate a molecular clone encoding
for a 3’-O-methyltranferase. A similar methodology as described above could be employed to
find a 3’OMT. G. flavum contains the aporphine alkaloid glaucine, which is O-methylated at
C3’-position of the simple BIA backbone. In the NGS transcriptomic database of G. flavum, one
would expect to find a gene with 3’-O-methylation activity. By comparing the lists of candidate
OMTs from G. flavum and a plant that does not appear to produce 3’-O-methylated compounds
like E. californica, it would be possible to reduce the number of possible 3’OMT candidates by
eliminating genes annotated as OMTs present in both G. flavum and E. californica.
As described in Chapter Four, the presence of thebaine in P. bracteatum suggest the
presence of upstream morphinan metabolites and the genes responsible for its biosynthesis.
Based on what we know of morphinan biosynthesis it is likely that the downstream morphinans
salutaridinol and salutaridine are made in P. brateatum. Similarily, (R)-reticuline would also be
expected and the enzyme(s) responsible for the epimerization of (S)-reticuline to (R)-reticuline
should be present. At this times, the genes involved in the epimerization of reticuline are
unknown, however an enzyme thought to be involved in the epimerization of reticuline, 1,2dehydroreticuline reductase (DRR) has been purified and partially characterized from opium
poppy (Beaudoin and Facchini 2014; De-Eknamkul and Zenk 1992). The search for candidate
genes encoding enzymes involved in the epimerization of reticuline could be narrowed by a
comparative transcriptomic analysis grouping opium poppy plants with P. bracteatum and
110
comparing them to the transcriptoms of another species from the family Papveracea that does not
produce morphinan alkaloids such as S. diphyllum or S. canadensis. Uncharacterized genes
annotated as reductive enzymes and oxidative enzymes found in opium poppy and P. bracteatum
but not in S. diphyllum or S. canadensis would be strong gene candidates for genes.
Certainly these targeted alkaloid profiles could also be used to find 2OMT orthologs in
other BIA producing species. Assuming, 2OMT is involved in cryptopine metabolism one could
find plants which produce cryptopine, like S. canadensis and expect to find 2OMT homologs.
This could be achieved by searching for a 2OMT in the S. canadensis NGS trancriptomic
database using the opium poppy 2OMT sequence as a query. The list of candidate genes
generated would be a targeted list and likely to contain a 2OMT ortholog.
111
REFERENCES
Altschup, Stephen F, Warren Gish, The Pennsylvania, and Universily Park. 1990. “Basic Local
Alignment Search Tool 2Department of Computer Science.” Journal of molecular biology
215 (3): 403-410. doi:10.1016/S0022-2836(05)80360-2.
Ashihara, Hiroshi, Hiroshi Sano, and Alan Crozier. 2008. “Caffeine and related purine alkaloids :
Biosynthesis , catabolism , function and genetic engineering.” Phytochemistry 69 (4): 841856. doi:10.1016/j.phytochem.2007.10.029.
Attia, Mohamed, Soo-un Kim, and Dae-kyun Ro. 2012. “Molecular cloning and characterization
of (+)-epi-[alpha]-bisabolol synthase, catalyzing the first step in the biosynthesis of the
natural sweetener, hernandulcin, in Lippia dulcis.” Archives of Biochemistry and Biophysics
527 (1): 37-44. doi:10.1016/j.abb.2012.07.010. http://dx.doi.org/10.1016/j.abb.2012.07.010.
Beaudoin, Guillaume A W, and Peter J Facchini. 2013. “Biochemical and Biophysical Research
Communications Isolation and characterization of a cDNA encoding (S) -cis-Nmethylstylopine 14-hydroxylase from opium poppy , a key enzyme in sanguinarine
biosynthesis.” Biochemical and Biophysical Research Communications 431 (3): 597-603.
doi:10.1016/j.bbrc.2012.12.129. http://dx.doi.org/10.1016/j.bbrc.2012.12.129.
———. 2014. “Benzylisoquinoline alkaloid biosynthesis in opium poppy.” Planta (240): 19-32.
doi:10.1007/s00425-014-2056-8.
Bhakuni, Dewan, and Sudha Jain. 1988. “The Biosynthesis of Glaucine in Litsea glutinosa.”
Journal of the Chemical Society 1 (8): 1447-1449. doi:10.1039/P19880001447.
Brochmann-Hanssen, Einar, Chung-Hsiung Chen, Rolland C. Chen, Hsuch-Ching Chiang,
Albert Y. Leung, and Kenneth McMurtrey. 1975. “The Biosynthesis of 1 Benzylisoquinolines in Papaver somniferum. Preferred and Secondary Pathways; Stereochemical Aspects.” The Journal of the Chemical Society, Perkin Transactions 1 (16The
Biosynthesis of 1 -Benzylisoquinolines in Papaver somniferum. Preferred and Secondary
Pathways ; Stereo- chemical Aspects): 1531-1537. doi:10.1039/P19750001531.
Butler, Mark S. 2008. “Natural products to drugs : natural product-derived compounds in clinical
trials.” Natural Product Reports 25 (3): 475-516. doi:10.1039/b514294f.
Cairney, John, Jeff Puryear, and Shujun Chang. 1993. “A Simple and Efficient Method for
Isolating RNA from Pine Trees.” Plant Molecular Biology Reporter 11 (2): 113-116.
Chen, Xue, and Peter J Facchini. 2014. “Short-chain dehydrogenase / reductase catalyzing the
final step of noscapine biosynthesis is localized to laticifers in opium poppy.” The Plant
Journal 450 (2): 173-184. doi:10.1111/tpj.12379.
112
Chenna, Ramu, Hideaki Sugawara, Tadashi Koike, Rodrigo Lopez, Toby J Gibson, Desmond G
Higgins, and Julie D Thompson. 2003. “Multiple sequence alignment with the Clustal series
of programs.” Nucleic Acids Research 31 (13): 3497-3500. doi:10.1093/nar/gkg500.
Chintalwar, G J, S Gupta, G Roja, and V A Bapat. 2003. “Protoberberine Alkaloids From Callus
and Cell Suspension Cultures of Tinospora cordifolia.” Pharmaceutical biology 41 (2): 8186. doi:1388-0209/03/4102-08.
Choi, KB, Takashi Morishige, and Fumihiko Sato. 2001. “Purification and characterization of
coclaurine N-methyltransferase from cultured Coptis japonica cells.” Phytochemistry 56:
649-655. http://www.sciencedirect.com/science/article/pii/S0031942200004817.
Dang, Thu-Thuy T, and Peter J Facchini. 2012. “Characterization of three O-methyltransferases
involved in noscapine biosynthesis in opium poppy.” Plant physiology 159 (2) (June): 61831. doi:10.1104/pp.112.194886. http://www.ncbi.nlm.nih.gov/pubmed/22535422.
Dang, Thuy, and Peter J. Facchini. 2014. “CYP82Y1 Is N-Methylcanadine 1-Hydroxylase, a Key
Noscapine Biosynthetic Enzyme in Opium Poppy.” The Journal of biological chemistry 289
(4): 2013-2026. doi:10.1074/jbc.M113.505099.
De-Eknamkul, Wanchai, and Meinhart H. Zenk. 1992. “PURIFICATION AND PROPERTIES
OF 1,2-DEHYDRORETICULINE REDUCTASE FROM PAPAVER SOMNIFERUM
SEEDLINGS.” Phytochemistry 31 (3): 813-821. doi:10.1016/0031-9422(92)80020-F.
Desgagné-Penix, Isabel, and Peter J Facchini. 2012. “Systematic silencing of benzylisoquinoline
alkaloid biosynthetic genes reveals the major route to papaverine in opium poppy.” The
Plant Journal 72 (2): 331-344. doi:10.1111/j.1365-313X.2012.05084.x.
Desgagné-Penix, Isabel, Scott C Farrow, Dustin Cram, Jacek Nowak, and Peter J Facchini. 2012.
“Integration of deep transcript and targeted metabolite profiles for eight cultivars of opium
poppy.” Plant molecular biology 79 (3) (June): 295-313. doi:10.1007/s11103-012-9913-2.
http://www.ncbi.nlm.nih.gov/pubmed/22527754.
Dethier, Monique, and Vincenzo De Luca. 1993. “PARTIAL PURIFICATION OF AN NMETHYLTRANSFERASE INVOLVED IN VINDOLINE BIOSYNTHESIS IN
CATHARANTHUS.” Phytochemistry 32 (3): 673-678.
Dewick, P.M. 2009. Medicinal natural products : a biosynthetic approach. 3rd ed. Chichester,
U.K.: Wiley.
Diaz Chavez, Maria Luisa, Megan Rolf, Andreas Gesell, and Toni M Kutchan. 2011.
“Characterization of two methylenedioxy bridge-forming cytochrome P450-dependent
enzymes of alkaloid formation in the Mexican prickly poppy Argemone mexicana q.”
Archives of Biochemistry and Biophysics 507 (1): 186-193. doi:10.1016/j.abb.2010.11.016.
113
Dittrich, Heinz, and Toni M Kutchan. 1991. “Molecular cloning , expression , and induction of
berberine bridge enzyme , an enzyme essential to the formation of benzophenanthridine
alkaloids in the response of plants to pathogenic attack.” Proceedings of the National
Academy of Sciences of the United States of America 88 (22): 9969-9973.
Dixon, Richard A. 2001. “Natural products and plant disease resistance.” Nature 411 (June):
843-847.
Docimo, Teresa, Michael Reichelt, Bernd Schneider, and John C D Auria. 2012. “The first step
in the biosynthesis of cocaine in Erythroxylum coca : the characterization of arginine and
ornithine decarboxylases.” Plant molecular biology 78 (6): 599-615. doi:10.1007/s11103012-9886-1.
Fabre, N, C Claparols, S Richelme, M L Angelin, I Fourasté, and C Moulis. 2000. “Direct
characterization of isoquinoline alkaloids in a crude plant extract by ion-pair liquid
chromatography-electrospray ionization tandem mass spectrometry: example of
Eschscholtzia californica.” Journal of chromatography. A 904 (1) (December 22): 35-46.
http://www.ncbi.nlm.nih.gov/pubmed/11209899.
Facchini, Peter J, Joerg Bohlmann, Patrick S Covello, Vincenzo De Luca, Radhakrishnan
Mahadevan, Jonathan E Page, Dae-Kyun Ro, Christoph W Sensen, Reginald Storms, and
Vincent J J Martin. 2012. “Synthetic biosystems for the production of high-value plant
metabolites.” Trends in biotechnology 30 (3) (March): 127-31.
doi:10.1016/j.tibtech.2011.10.001. http://www.ncbi.nlm.nih.gov/pubmed/22209518.
Facchini, Peter J., Jillian M. Hagel, David K. Liscombe, Natalia Loukanina, Benjamin P.
MacLeod, Nailish Samanani, and Katherine G. Zulak. 2007. “Opium poppy: blueprint for
an alkaloid factory.” Phytochemistry Reviews 6 (1) (March 1): 97-124. doi:10.1007/s11101006-9042-0. http://www.springerlink.com/index/10.1007/s11101-006-9042-0.
Facchini, Peter J., and Vincenzo De Luca. 1994. “Differential and Tissue-specific Expression of
a Gene Family for Tyrosine/Dopa Decarboxylase in Opium Poppy.” The Journal of
biological chemistry 269 (43): 26684-26690.
Facchini, Peter J., and Sang-Un Park. 2003. “Developmental and inducible accumulation of gene
transcripts involved in alkaloid biosynthesis in opium poppy.” Phytochemistry 64 (1)
(September): 177-186. doi:10.1016/S0031-9422(03)00292-9.
http://linkinghub.elsevier.com/retrieve/pii/S0031942203002929.
Facchini, Peter J. and DeLuca, Vincenzo. 2008. “Opium poppy and Madagascar periwinkle :
model non-model systems to investigate alkaloid biosynthesis in plants.” The Plant Journal
54 (4): 763-784. doi:10.1111/j.1365-313X.2008.03438.x.
114
Fahey, Jed W, Amy T Zalcmann, and Paul Talalay. 2001. “The chemical diversity and
distribution of glucosinolates and isothiocyanates among plants.” Phytochemistry 56 (1): 551. doi:10.1016/S0031-9422(00)00316-2.
Farrow, Scott C, and Peter J Facchini. 2013. “O , O-Demethylenation with Widespread Roles in
Benzylisoquinoline Alkaloid Metabolism in Opium Poppy * □.” The Journal of biological
chemistry 288 (40): 28997-29012. doi:10.1074/jbc.M113.488585.
Farrow, Scott C, Jillian M Hagel, and Peter J Facchini. 2012. “Transcript and metabolite
profiling in cell cultures of 18 plant species that produce benzylisoquinoline alkaloids.”
Phytochemistry 77 (March 14): 79-88. doi:10.1016/j.phytochem.2012.02.014.
http://www.ncbi.nlm.nih.gov/pubmed/22424601.
Ferrer, J, M B Austin, C Stewart Jr, and J P Noel. 2008. “Structure and function of enzymes
involved in the biosynthesis of phenylpropanoids.” Plant Physiol Biochem. 46 (3): 356-370.
Fisinger U, Grobe N, Zenk MH (2007). 2007. “Thebaine synthase: a new enzyme in the
morphine pathway in P. somniferum.” Natural Product Communications 2: 249–253.
Fraenkel, Gottfried S. 1959. “The Raison d ’Etre of Secondary Plant Substances.” Science 129
(3361): 1466-1470.
Frenzel, Thomas, and Meinhart H. Zenk. 1990. “S-ADENOSYL-L-METHIONINE : 3’-hydroxyN-methyl-(S)-coclaurine-4'-O-methyl TRANSFERASE , A REGIO- AND
STEREOSELECTIVE ENZYME OF THE ( S ) -RETICULINE PATHWAY.”
Phytochemistry 29 (11): 3505-3511.
Frick, S, and T M Kutchan. 1999. “Molecular cloning and functional expression of Omethyltransferases common to isoquinoline alkaloid and phenylpropanoid biosynthesis.”
The Plant journal : for cell and molecular biology 17 (4) (March): 329-39.
http://www.ncbi.nlm.nih.gov/pubmed/10205892.
Frick, Susanne, Robert Kramell, Anthony J Fist, and Toni M Kutchan. 2005. “Comparative
Qualitative and Quantitative Determination of Alkaloids in Narcotic and Condiment
Papaver somniferum Cultivars.” Journal of natural products 63 (2): 666-673.
doi:10.1021/np0496643.
Gesell, Andreas, Megan Rolf, Jörg Ziegler, María Luisa, Díaz Chávez, Fong-chin Huang, Toni
M Kutchan, Andreas Gesell, Megan Rolf, and Toni M Kutchan. 2009. “CYP719B1 Is
Salutaridine Synthase , the C-C Phenol-coupling Enzyme of Morphine Biosynthesis in
Opium Poppy * □.” The Journal of biological chemistry 284 (36): 24432-24442.
doi:10.1074/jbc.M109.033373.
Goossens, Alain, and Heiko Rischer. 2007. “Implementation of functional genomics for gene
discovery in alkaloid producing plants.” Phytochemistry Reviews 6 (1) (February 27): 35115
49. doi:10.1007/s11101-006-9018-0. http://www.springerlink.com/index/10.1007/s11101006-9018-0.
Griffin, William J, and G David Lin. 2000. “Chemotaxonomy and geographical distribution of
tropane alkaloids.” Phytochemistry 53 (6): 623-637.
Grothe, Torsten, Rainer Lenz, M Toni, J Biol Chem, Torsten Grothe, Rainer Lenz, and Toni M
Kutchan. 2001. “Molecular Characterization of the Salutaridinol 7- O -Acetyltransferase
Involved in Morphine Biosynthesis in Opium Poppy Papaver somniferum.” The Journal of
biological chemistry 276 (33): 30717-300723. doi:10.1074/jbc.M102688200.
Hagel, Jillian M, Guillaume A W Beaudoin, Elena Fossati, Andrew Ekins, J J Vincent, Peter J
Facchini, Jillian M Hagel, et al. 2012. “Characterization of a Flavoprotein Oxidase from
Opium Poppy Catalyzing the Final Steps in Sanguinarine and.” The Journal of biological
chemistry 287 (51): 42972-42983. doi:10.1074/jbc.M112.420414.
Hagel, Jillian M, and Peter J Facchini. 2010. “Dioxygenases catalyze the O-demethylation steps
of morphine biosynthesis in opium poppy.” Nature chemical biology 6 (4) (April): 273-5.
doi:10.1038/nchembio.317. http://www.ncbi.nlm.nih.gov/pubmed/20228795.
———. 2013. “Benzylisoquinoline Alkaloid Metabolism : A Century of Discovery and a Brave
New World Special Focus Issue – Review.” Plant & cell physiology 54 (5): 647-672.
doi:10.1093/pcp/pct020.
Han, Xu, Marc Lamshöft, Nadja Grobe, Xuan Ren, Anthony J Fist, Toni M Kutchan, Michael
Spiteller, and Meinhart H Zenk. 2010. “The biosynthesis of papaverine proceeds via (S)reticuline.” Phytochemistry 71 (11-12) (August): 1305-12.
doi:10.1016/j.phytochem.2010.04.022.
http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=2900451&tool=pmcentrez&ren
dertype=abstract.
Hartmann, Thomas. 2007. “From waste products to ecochemicals : Fifty years research of plant
secondary metabolism.” Phytochemistry 68 (1): 2831-2846.
doi:10.1016/j.phytochem.2007.09.017.
Haslam, E. 1986. “Secondary Metabolism - Fact and Fiction.” Natural Product Reports 3 (1):
217-249. doi:10.1039/NP9860300217.
Hawkins, Kristy M, and Christina D Smolke. 2008. “Production of benzylisoquinoline alkaloids
in Saccharomyces cerevisiae.” Nature chemical biology 4 (9) (September): 564-73.
doi:10.1038/nchembio.105.
http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=2830865&tool=pmcentrez&ren
dertype=abstract.
116
Hirata, Kazumasa, Chotima Poeaknapo, Juergen Schmidt, and Meinhart H Zenk. 2004. “1,2Dehydroreticuline synthase, the branch point enzyme opening the morphinan biosynthetic
pathway.” Phytochemistry 65 (8): 1039-1046. doi:10.1016/j.phytochem.2004.02.015.
Hook, Ingrid, Helen Sheridan, and Graham Wilson. 1988. “ALKALOIDS OF CELL
CULTURES DERIVED.” Phytochemistry 27 (7): 2137-2141. doi:0031-9422/88.
Hu, Qizhi, Robert J Noll, Hongyan Li, Alexander Makarov, Mark Hardman, and R Graham
Cooks. 2005. “The Orbitrap: a new mass spectrometer.” Journal of mass spectrometry :
JMS 40 (4) (April): 430-43. doi:10.1002/jms.856.
http://www.ncbi.nlm.nih.gov/pubmed/15838939.
Ikezawa, Nobuhiro, Kinuko Iwasa, and Fumihiko Sato. 2007. “Molecular cloning and
characterization of methylenedioxy bridge-forming enzymes involved in stylopine
biosynthesis in Eschscholzia californica.” FEBS Journal 274 (4): 1019-1035.
doi:10.1111/j.1742-4658.2007.05652.x.
———. 2008. “Molecular Cloning and Characterization of CYP80G2, a Cytochrome P450 That
Catalyzes an Intramolecular C – C Phenol Coupling of (S)-Reticuline in Magnoflorine
Biosynthesis , from Cultured Coptis japonica Cells.” The Journal of biological chemistry
283 (14): 8810-8821. doi:10.1074/jbc.M705082200.
Ikezawa, Nobuhiro, Masaru Tanaka, Raku Shinkyo, Toshiyuki Sakaki, Kuniyo Inouye, and
Fumihiko Sato. 2003. “Metabolism and Bioenergetics : Molecular Cloning and
Characterization of Enzyme That Belongs to a Novel P450 Family , from cultured Coptis
japonica Cells Molecular Cloning and Characterization of CYP719 , a Methylenedioxy
Bridge-forming Enzyme That Belong.” The Journal of biological chemistry 278 (40):
38557-38565. doi:10.1074/jbc.M302470200.
Ikuta, Akira, and Hideji Itokawa. 1988. “ALKALOIDS OF TISSUE CULTURES OF
NANDINA DOMESTICA.” Phytochemistry 27 (7): 2143-2145. doi:003l 9422/88.
Inui, Takayuki, Ken-Ichi Tamura, Nanae Fujii, Takashi Morishige, and Fumihiko Sato. 2007.
“Overexpression of Coptis japonica norcoclaurine 6-O-methyltransferase overcomes the
rate-limiting step in Benzylisoquinoline alkaloid biosynthesis in cultured Eschscholzia
californica.” Plant & cell physiology 48 (2) (February): 252-62. doi:10.1093/pcp/pcl062.
http://www.ncbi.nlm.nih.gov/pubmed/17189286.
Iwasa, Kinuko, Teturo Takahashi, Yumi Nishiyama, Masataka Moriyasu, Makiko Sugiura, and
Atsuko Takeuchi. 2008. “Online Structural Elucidation of Alkaloids and Other Constituents
in Crude Extracts and Cultured Cells of Nandina domestica by Combination of LC-MS /
MS , LC-NMR , and LC-CD.” Journal of natural products 71 (8): 1376-1385.
doi:10.1021/np8001496.
117
Jirschitzka, Jan, Gregor W Schmidt, Michael Reichelt, Bernd Schneider, and Jonathan
Gershenzon. 2012. “Plant tropane alkaloid biosynthesis evolved independently in the
Solanaceae and Erythroxylaceae.” Proceedings of the National Academy of Sciences of the
United States of America 109 (26): 10304–10309. doi:10.1073/pnas.1200473109.
Jones, Philip, David Binns, Hsin-yu Chang, Matthew Fraser, Weizhong Li, Craig Mcanulla,
Hamish Mcwilliam, et al. 2014. “Sequence analysis InterProScan 5 : genome-scale protein
function classification.” Bioinformatics: 1-5. doi:10.1093/bioinformatics/btu031.
Jukes, TH, and CR Cantor. 1969. Evolution of Protein Molecules. New York: Academic Press.
Katajamaa, Mikko, and Matej Oresic. 2007. “Data processing for mass spectrometry-based
metabolomics.” Journal of chromatography. A 1158 (1-2) (July 27): 318-28.
doi:10.1016/j.chroma.2007.04.021. http://www.ncbi.nlm.nih.gov/pubmed/17466315.
Kim, Hye Kyong, and Rob Verpoorte. 2010. “Sample preparation for plant metabolomics.”
Phytochemical analysis : PCA 21 (1) (January): 4-13. doi:10.1002/pca.1188.
http://www.ncbi.nlm.nih.gov/pubmed/19904733.
Kliebenstein, Daniel J, and Anne Osbourn. 2012. “Making new molecules – evolution of
pathways for novel metabolites in plants.” Current opinion in plant biology 15 (4): 415-423.
doi:10.1016/j.pbi.2012.05.005. http://dx.doi.org/10.1016/j.pbi.2012.05.005.
Kutchan, T M, N Hampp, F Lottspeich, and M H Zenk. 1988. “The cDNA clone for strictosidine
synthase from Rauvolfia and expression in Escherichia coli.” FEBS letters 237 (1-2): 40-44.
doi:10.1016/0014-5793(88)80167-4.
Ky, C.-L, J Louarn, S Dussert, B Guyot, S Hamon, and M Noirot. 2001. “Caffeine, trigonelline,
chlorogenic acids and sucrose diversity in wild Coffea arabica L. and C. canephora P.
accessions.” Food Chemistry 75 (2) (November): 223-230. doi:10.1016/S03088146(01)00204-7. http://linkinghub.elsevier.com/retrieve/pii/S0308814601002047.
Landen, JW, Roland Lang, and SJ McMahon. 2002. “Noscapine Alters Microtubule Dynamics in
Living Cells and Inhibits the Progression of Melanoma.” Cancer research 62: 4109-4114.
http://cancerres.aacrjournals.org/content/62/14/4109.short.
Le, Phuong Mai, Margaret Mccooeye, and Anthony Windust. 2012. “Characterization of the
alkaloids in goldenseal ( Hydrastis canadensis ) root by high resolution Orbitrap LC-MS n.”
Analytical and bioanalytical chemistry 405 (13): 4487-4498. doi:10.1007/s00216-0126539-9.
———. 2014. “Application of UPLC-QTOF-MS in MS E mode for the rapid and precise
identification of alkaloids in goldenseal (Hydrastis canadensis).” Analytical and
Bioanalytical Chemistry 406 (6): 1739-1749. doi:10.1007/s00216-013-7558-x.
118
Lee, Eun-Jeong, and Peter J Facchini. 2011. “Tyrosine aminotransferase contributes to
benzylisoquinoline alkaloid biosynthesis in opium poppy.” Plant physiology 157 (3)
(November): 1067-78. doi:10.1104/pp.111.185512.
http://www.ncbi.nlm.nih.gov/pubmed/21949209.
Lenz, Rainer, and Meinhart H. Zenk. 1995. “Acetyl Coenzyme A: Salutaridinol-7-OAcetyltransferase from Papaver somniferum Plant Cell Cultures.” The Journal of biological
chemistry 270 (52): 0-6. doi:10.1074/jbc.270.52.31091.
Li, Yubo, Tiejun Zhnag, Haiyu Xu, and Changxiao Lui. 2010. “Chemical fingerprint analysis of
Phellodendri Amurensis Cortex by ultra performance LC / Q-TOF-MS methods combined
with chemometrics.” Journal of separation science 33 (21): 3347-3353.
doi:10.1002/jssc.201000426.
Liscombe, David K, and Peter J Facchini. 2007. “Molecular Cloning and Characterization of
Tetrahydroprotoberberine cis-N-Methyltransferase , an Enzyme Involved in Alkaloid
Biosynthesis in Opium Poppy * □.” The Journal of biological chemistry 282 (20): 1474114751. doi:10.1074/jbc.M611908200.
———. 2008. “Evolutionary and cellular webs in benzylisoquinoline alkaloid biosynthesis.”
Current opinion in biotechnology 19 (2) (April): 173-80. doi:10.1016/j.copbio.2008.02.012.
http://www.ncbi.nlm.nih.gov/pubmed/18396034.
Liscombe, David K., Benjamin P. MacLeod, Natalia Loukanina, Owi I. Nandi, and Peter J.
Facchini. 2005. “Evidence for the monophyletic evolution of benzylisoquinoline alkaloid
biosynthesis in angiosperms.” Phytochemistry 66 (20) (October): 2500-2520.
doi:10.1016/j.phytochem.2005.04.044.
http://linkinghub.elsevier.com/retrieve/pii/S0031942205004255.
Liscombe, David K., Jörg Ziegler, Jürgen Schmidt, Christian Ammer, and Peter J. Facchini.
2009. “Targeted metabolite and transcript profiling for elucidating enzyme function :
isolation of novel N-methyltransferases from three benzylisoquinoline alkaloid-producing
species.” The Plant Journal 60 (4): 729-743. doi:10.1111/j.1365-313X.2009.03980.x.
Loeffler, Susanne, Brigitte Deus-Neumann, and Meinhart H. Zenk. 1995. “S-ADENOSYL-LMETHIONINE:(S)-COCLAURINE-N-METHYL- TRANSFERASE FROM TINOSPORA
CORDIFOLIA.” Phytochemistry 38 (6): 1387-1395.
Loyola-Vargas, Victor Manuel, Rosa M. Galaz-Avalos, and Roberto Ku-Cauich. 2007.
“Catharanthus biosynthetic enzymes : the road ahead.” Phytochemistry Reviews 6 (2-3):
307-339. doi:10.1007/s11101-007-9064-2.
Mccarthy, Andrew A, and James G Mccarthy. 2014. “The Structure of Two NMethyltransferases from the Caffeine Biosynthetic Pathway.” Plant physiology 144 (2):
879-889. doi:10.1104/pp.106.094854.
119
Minami, Hiromichi, Ju-Sung Kim, Nobuhiro Ikezawa, Tomoya Takemura, Takane Katayama,
Hidehiko Kumagai, and Fumihiko Sato. 2008. “Microbial production of plant
benzylisoquinoline alkaloids.” Proceedings of the National Academy of Sciences of the
United States of America 105 (21) (May 27): 7393-8. doi:10.1073/pnas.0802981105.
http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=2396723&tool=pmcentrez&ren
dertype=abstract.
Morishige, T, T Tsujita, Y Yamada, and F Sato. 2000. “Molecular characterization of the Sadenosyl-L-methionine:3’-hydroxy-N-methylcoclaurine 4'-O-methyltransferase involved in
isoquinoline alkaloid biosynthesis in Coptis japonica.” The Journal of biological chemistry
275 (30) (July 28): 23398-405. doi:10.1074/jbc.M002439200.
http://www.ncbi.nlm.nih.gov/pubmed/10811648.
Morishige, Takashi, Emilyn Dubouzet, Kum-Boo Choi, Kazufumi Yazaki, and Fumihiko Sato.
2002. “Molecular cloning of columbamine O-methyltransferase from cultured Coptis
japonica cells.” European Journal of Biochemistry 269 (22) (November 28): 5659-5667.
doi:10.1046/j.1432-1033.2002.03275.x. http://doi.wiley.com/10.1046/j.14321033.2002.03275.x.
Nakagawa, Akira, Hiromichi Minami, Ju-Sung Kim, Takashi Koyanagi, Takane Katayama,
Fumihiko Sato, and Hidehiko Kumagai. 2011. “A bacterial platform for fermentative
production of plant alkaloids.” Nature communications 2 (May) (January): 326.
doi:10.1038/ncomms1327.
http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=3112539&tool=pmcentrez&ren
dertype=abstract.
Noble, R L. 1990. “The discovery of the vinca alkaloids--chemotherapeutic agents against
cancer.” Biochemistry and cell biology = Biochimie et biologie cellulaire 68 (12)
(December): 1344-51. http://www.ncbi.nlm.nih.gov/pubmed/2085431.
Ober, D, and T Hartmann. 2000. “Phylogenetic origin of a secondary pathway: the case of
pyrrolizidine alkaloids.” Plant molecular biology 44 (4) (November): 445-50.
http://www.ncbi.nlm.nih.gov/pubmed/11197320.
Ounaroon, Anan, Gabriele Decker, Jürgen Schmidt, Friedrich Lottspeich, and Toni M. Kutchan.
2003. “( R,S )-Reticuline 7- O -methyltransferase and ( R,S )-norcoclaurine 6- O methyltransferase of Papaver somniferum - cDNA cloning and characterization of methyl
transfer enzymes of alkaloid biosynthesis in opium poppy.” The Plant Journal 36 (6)
(December): 808-819. doi:10.1046/j.1365-313X.2003.01928.x.
http://doi.wiley.com/10.1046/j.1365-313X.2003.01928.x.
O’Connor, Sarah E, and Justin J Maresh. 2006. “Chemistry and biology of monoterpene indole
alkaloid biosynthesis.” Natural Product Reports 4 (23): 532-547. doi:10.1039/b512615k.
120
Pathak, Sumya, Deepika Lakhwani, Parul Gupta, Brij Kishore Mishra, Sudhir Shukla, and Mehar
Hasan Asif. 2013. “Comparative Transcriptome Analysis Using High Papaverine Mutant of
Papaver somniferum Reveals Pathway and Uncharacterized Steps of Papaverine
Biosynthesis.” PloS one 8 (5): e65622. doi:10.1371/journal.pone.0065622.
Pauli, H H, and T M Kutchan. 1998. “Molecular cloning and functional heterologous expression
of two alleles encoding (S)-N-methylcoclaurine 3’-hydroxylase (CYP80B1), a new methyl
jasmonate-inducible cytochrome P-450-dependent mono-oxygenase of benzylisoquinoline
alkaloid biosynthesis.” The Plant journal : for cell and molecular biology 13 (6) (March):
793-801. http://www.ncbi.nlm.nih.gov/pubmed/9681018.
Pichersky, Eran, and David R Gang. 2000. “Genetics and biochemistry of secondary metabolites
in plants : an evolutionary perspective.” Trends in plant science 5 (10): 439-445.
Pichersky, Eran, and Jonathan Gershenzon. 2002. “The formation and function of plant volatiles:
perfumes for pollinator attraction and defense.” Current opinion in plant biology 5 (3)
(June): 237-43. http://www.ncbi.nlm.nih.gov/pubmed/11960742.
Pichersky, Eran, and Efraim Lewinsohn. 2011. “Convergent Evolution in Plant Specialized
Metabolism.” Annual review of plant biology 62: 549-566. doi:10.1146/annurev-arplant042110-103814.
Pienkny, Silke, Wolfgang Brandt, Jürgen Schmidt, Robert Kramell, and Jörg Ziegler. 2009.
“Functional characterization of a novel benzylisoquinoline O-methyltransferase suggests its
involvement in papaverine biosynthesis in opium poppy (Papaver somniferum L).” The
Plant journal : for cell and molecular biology 60 (1) (October): 56-67. doi:10.1111/j.1365313X.2009.03937.x. http://www.ncbi.nlm.nih.gov/pubmed/19500305.
Pyle, Bryan W, Hue T Tran, Benjamin Pickel, Tegan M Haslam, Zhizeng Gao, John C Vederas,
Soo-un Kim, and Dae-kyun Ro. 2012. “Enzymatic synthesis of valerena-4,7(11)-diene by a
unique sesquiterpene synthase from the valerian plant (Valeriana officinalis).” FEBS
Journal 279 (11): 3136-3146. doi:10.1111/j.1742-4658.2012.08692.x.
Raith, Klaus, Reinhard Neubert, Chotima Poeaknapo, Christian Boettcher, Meinhart H Zenk, and
Jürgen Schmidt. 2003. “Electrospray tandem mass spectrometric investigations of
morphinans.” Journal of the American Society for Mass Spectrometry 14 (11) (November):
1262-9. doi:10.1016/S1044-0305(03)00539-7.
http://www.ncbi.nlm.nih.gov/pubmed/14597116.
Ren, Lingling, Xingya Xue, Feifang Zhang, Qing Xu, and Xinmiao Liang. 2007. “High
performance liquid chromatography-mass spectrometry analysis of protoberberine alkaloids
in medicine herbs.” Journal of separation science 30 (6): 833-842.
doi:10.1002/jssc.200600246.
121
Reynolds, Tom. 2005. “Hemlock alkaloids from Socrates to poison aloes.” Phytochemistry 66
(12t) (June): 1399-1406. doi:10.1016/j.phytochem.2005.04.039.
http://www.ncbi.nlm.nih.gov/pubmed/15955542.
Ronsch, Hasso. 1972. “Zur Biosynthese der Alkaloide vom Rhoeadin-Typ.” European Journal
of Biochemistry 28: 123-126.
Rueffer, M, M Amann, and M H Zenk. 1986. “S-Adenosyl-L-methionine: Columbamine-Omethyl transferase, a compartmentalized enzyme in protoberberine biosynthesis.” Plant cell
reports 5 (3): 182-185.
Rueffer, Martina, N. Nagakura, and Meinhart H. Zenk. 1983. “A Highly Specific Omethyltransferase for Nororientaline Synthesis from Argemone platyceras Cell Cultures.”
Journal of Medicinal Plant Research 49: 196-198.
Rueffer, Martina, and Meinhart H. Zenk. 1987. “Distant Precursors of Benzylisoquinoline
Alkaloids and their Enzymatic Formation.” European Journal of Biosciences 42c: 319-332.
Samanani, Nailish, and Peter J Facchini. 2002. “Purification and characterization of
norcoclaurine synthase. The first committed enzyme in benzylisoquinoline alkaloid
biosynthesis in plants.” The Journal of biological chemistry 277 (37) (September 13):
33878-83. doi:10.1074/jbc.M203051200. http://www.ncbi.nlm.nih.gov/pubmed/12107162.
Schmidt, J, K Raith, C Boettcher, and Meinhart H. Zenk. 2005. “Analysis of benzylisoquinolinetype alkaloids by electrospray tandem mass spectrometry and atmospheric pressure
photoionization.” European Journal of Mass Spectrometry 11 (3): 325-333.
http://www.impublications.com/content/abstract?code=E11_0325.
Schmidt, Jürgen, Chotima Boettcher, Christine Kuhnt, Toni M Kutchan, and Meinhart H Zenk.
2007. “Poppy alkaloid profiling by electrospray tandem mass spectrometry and electrospray
FT-ICR mass spectrometry after [ring-13C6]-tyramine feeding.” Phytochemistry 68 (2)
(January): 189-202. doi:10.1016/j.phytochem.2006.10.003.
http://www.ncbi.nlm.nih.gov/pubmed/17113612.
Schubert, Heidi L, Robert M Blumenthal, and Xiaodong Cheng. 2003. “Many paths to
methyltransfer : a chronicle of convergence.” Trends in Biochem Sci. 28 (6): 329-335.
doi:10.1016/S0968-0004(03)00090-2.Many.
Shulgin, Alexander T., and Wendy E. Perry. 2002. The Simple Plant Isoquinolines. First Edit.
Berkeley, CA, USA: Transform Press.
Stevens, Jan F, Ralph L Reed, and Jeffrey T Morré. 2009. “Characterization of Phytoecdysteroid
Glycosides in Meadowfoam (Limnanthes alba) Seed Meal by Positive and Negative Ion
LC-MS/ MS.” Journal of Agricultural and Food Chemistry 56 (11): 3945-3952.
doi:10.1021/jf800211k.Characterization.
122
Stévigny, Caroline, Jean-Louis Habib Jiwan, Raoul Rozenberg, Edmond de Hoffmann, and
Joëlle Quetin-Leclercq. 2004. “Key fragmentation patterns of aporphine alkaloids by
electrospray ionization with multistage mass spectrometry.” Rapid communications in mass
spectrometry : RCM 18 (5) (January): 523-8. doi:10.1002/rcm.1343.
http://www.ncbi.nlm.nih.gov/pubmed/14978796.
Suzuki, Takeo, Hiroshi Ashihara, and George R. Waller. 1992. “PURINE AND PURINE
ALKALOID METABOLISM IN CAMELLIA AND COFFEA PLANTS.” Phytochemistry
31 (8): 2575-2584.
Takao, Narao, Miyoko Kamigauchi, and Momoyo Okada. 1983. “Biosynthesis of Benzo
[c]phenanthridine Alkaloids Sanguinarine , Chelirubine and Macarpine.” Helvetica Chimica
Acta 66 (40): 473-484.
Takemura, Tomoya, Nobuhiro Ikezawa, Kinuko Iwasa, and Fumihiko Sato. 2013.
“Phytochemistry Molecular cloning and characterization of a cytochrome P450 in
sanguinarine biosynthesis from Eschscholzia californica cells.” Phytochemistry 91: 100108. doi:10.1016/j.phytochem.2012.02.013.
http://dx.doi.org/10.1016/j.phytochem.2012.02.013.
Takeshita, N, H Fujiwara, H Mimura, J H Fitchen, Y Yamada, and F Sato. 1995. “Molecular
cloning and characterization of S-adenosyl-L-methionine:scoulerine-9-O-methyltransferase
from cultured cells of Coptis japonica.” Plant & cell physiology 36 (1) (January): 29-36.
http://www.ncbi.nlm.nih.gov/pubmed/7719631.
Theis, N, and M Lerdau. 2003. “The Evolution of Function in Plant Secondary Metabolites.”
International Journal of Plant Sciences 164 (May 2003): S93-S102.
http://www.jstor.org/stable/10.1086/374190.
Theuns, Hubert G, H Leo Theuns, and Robert J J Ch Lousberg. 2014. “Search for New Natural
Sources of Morphinans.” Economic Botany 40 (4): 485-497.
Unterlinner, Bernhard, Rainer Lenz, and Toni M Kutchan. 1999. “Molecular cloning and
functional expression of codeinone reductase : the penultimate enzyme in morphine
biosynthesis in the opium poppy Papaver somniferum.” The Plant Journal 18 (5): 465-475.
doi:10.1046/j.1365-313X.1999.00470.x.
Urbanczyk-Wochniak, Eza, Charles Baxter, Anna Kolbe, Joachim Kopka, Lee J. Sweetlove, and
Alisdair R. Fernie. 2005. “Profiling of diurnal patterns of metabolite and transcript
abundance in potato ( Solanum tuberosum ) leaves.” Planta 221 (6): 891-903.
doi:10.1007/s00425-005-1483-y.
Verpoorte, R., Y. H. Choi, N. R. Mustafa, and H. K. Kim. 2008. “Metabolomics: back to basics.”
Phytochemistry Reviews 7 (3) (March 11): 525-537. doi:10.1007/s11101-008-9091-7.
http://www.springerlink.com/index/10.1007/s11101-008-9091-7.
123
Vogel, Matthias, Michael Lawson, Udo Conrad, Werner Roos, Matthias Vogel, Michael Lawson,
Wolfgang Sippl, Udo Conrad, and Werner Roos. 2010. “Structure and Mechanism of
Sanguinarine Reductase, an Enzyme of Alkaloid Detoxification.” The Journal of biological
chemistry 285 (24): 18397-18406. doi:10.1074/jbc.M109.088989.
Weiss, Dagmar, Alfred Baumert, Matthias Vogel, and Werner Roos. 2006. “Sanguinarine
reductase , a key enzyme of benzophenanthridine detoxification.” Plant, Cell and
Environment 29 (2): 291-302. doi:10.1111/j.1365-3040.2005.01421.x.
Wink, Michael. 2003. “Evolution of secondary metabolites from an ecological and molecular
phylogenetic perspective.” Phytochemistry 64 (1) (September): 3-19. doi:10.1016/S00319422(03)00300-5. http://linkinghub.elsevier.com/retrieve/pii/S0031942203003005.
Wink, Micheal. 2010. Annual Plant Review, Biochemistry of Plant Secondary Metabolism. Ed.
J.K.C Roberts, J.A., Evans, D., Imaseki, H., McManus, M.T., Rose. 2nd ed. Hoboken, NJ,
USA: Wiley-Blackwell.
Winzer, Thilo, Thilo Winzer, Valeria Gazda, Zhesi He, Filip Kaminski, Marcelo Kern, Tony R
Larson, et al. 2012. “A Papaver somniferum 10-Gene Cluster for Synthesis of the
Anticancer Alkaloid Noscapine.” Science 336 (2012): 1704-1708.
doi:10.1126/science.1220757.
Wu, Wei, Fengrui Song, Cunyu Yan, Zhiqiang Liu, and Shuying Liu. 2005. “Structural analyses
of protoberberine alkaloids in medicine herbs by using ESI – FT-ICR-MS and HPLC – ESI
– MS n.” Journal of Pharmaceutical and Biomedical Analysis 37 (3): 437-446.
doi:10.1016/j.jpba.2004.11.026.
Wu, Wu-nan, and C Huang. 2006. “Structural elucidation of isoquinoline, isoquinolone,
benzylisoquinoline, aporphine, and phenanthrene alkaloids using API-ionspray tandem
mass spectrometry.” CHINESE PHARMACEUTICAL JOURNAL- … 58 (1): 41-55.
http://web2.tmu.edu.tw/pharmtw/cjp/issue/58-1/J005.pdf.
Xiao, Mei, Ye Zhang, Xue Chen, Eun-Jeong Lee, Carla J S Barber, Romit Chakrabarty, Isabel
Desgagné-Penix, et al. 2013. “Transcriptome analysis based on next-generation sequencing
of non-model plants producing specialized metabolites of biotechnological interest.”
Journal of biotechnology 166 (3) (July 10): 122-34. doi:10.1016/j.jbiotec.2013.04.004.
http://www.ncbi.nlm.nih.gov/pubmed/23602801.
Xiao, Mei, Ye Zhang, Xue Chen, Eun-jeong Lee, Carla J S Barber, Romit Chakrabarty, Isabel
Desgagné-penix, et al. 2013. “Transcriptome analysis based on next-generation sequencing
of non-model plants producing specialized metabolites of biotechnological interest.”
Journal of Biotechnology 166 (3): 122-134. doi:10.1016/j.jbiotec.2013.04.004.
http://dx.doi.org/10.1016/j.jbiotec.2013.04.004.
124
Zhang, Yufeng, Qirong Shi, Peiying Shi, Weidong Zhang, and Yiyu Cheng. 2006.
“Characterization of isoquinoline alkaloids , diterpenoids and steroids in the Chinese herb
Jin-Guo-Lan (Tinospora sagittata and Tinospora capillipes) by high-performance liquid
chromatography/electrospray ionization with multistage mass spectrometry.” Rapid
communications in mass spectrometry 20 (15): 2328-2342. doi:10.1002/rcm.
Ziegler, Jorg, Susan Voigtlander, Jurgen Schmidt, Robert Kramell, Otto Miersch, Christian
Ammer, Andreas Gesell, and Toni M Kutchan. 2006. “Comparative transcript and alkaloid
profiling in Papaver species identifies a short chain dehydrogenase/reductase involved in
morphine biosynthesis.” The Plant Journal 48 (2): 177-192. doi:10.1111/j.1365313X.2006.02860.x.
Ziegler, Jörg, María Luisa Diaz-Chávez, Robert Kramell, Christian Ammer, and Toni M
Kutchan. 2005. “Comparative macroarray analysis of morphine containing Papaver
somniferum and eight morphine free Papaver species identifies an O-methyltransferase
involved in benzylisoquinoline biosynthesis.” Planta 222 (3) (October): 458-71.
doi:10.1007/s00425-005-1550-4. http://www.ncbi.nlm.nih.gov/pubmed/16034588.
Ziegler, Jörg, and Peter J Facchini. 2008. “Alkaloid biosynthesis: metabolism and trafficking.”
Annual review of plant biology 59 (January): 735-69.
doi:10.1146/annurev.arplant.59.032607.092730.
http://www.ncbi.nlm.nih.gov/pubmed/18251710.
Ziegler, Jörg, Peter J Facchini, René Geissler, Jürgen Schmidt, Christian Ammer, Robert
Kramell, Susan Voigtländer, Andreas Gesell, Silke Pienkny, and Wolfgang Brandt. 2009.
“Evolution of morphine biosynthesis in opium poppy.” Phytochemistry 70 (15-16): 1696707. doi:10.1016/j.phytochem.2009.07.006.
http://www.ncbi.nlm.nih.gov/pubmed/19665152.
Zubieta, Chloe, Xian-zhi He, Richard A Dixon, Joseph P Noel, Chalcone O-methyltransferase
Chomt, and O-methyltransferase Iomt S-adenosyl- L. 2001. “Structures of two natural
product methyltransferases reveal the basis for substrate specificity in plant.” Nature
Structural Biology 8 (3): 271-279.
125