University of Nevada, Reno
Mountain Pine Beetle (MPB; Dendroctonus ponderosae) Pheromone Component Biosynthesis, Cytochromes P450 and Monoterpene Metabolism in Bark Beetles
A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Cell and Molecular Biology
By
Minmin Song
Dr. Claus Tittiger/Dissertation Advisors
May, 2012
THE GRADUATE SCHOOL
We recommend that the dissertation prepared under our supervision by
MINMIN SONG
entitled
Mountain Pine Beetle (MPB; Dendroctonus Ponderosae) Pheromone Component Biosynthesis, Cytochromes P450 and Monoterpene Metabolism in Bark Beetles
be accepted in partial fulfillment of the requirements for the degree of
DOCTOR OF PHILOSOPHY
Claus Tittiger, Ph.D., Advisor
Gary Blomquist, Ph.D., Committee Member
Patricia Ellison, Ph.D., Committee Member
Grant Mastick, Ph.D., Committee Member
David Shintani, Ph.D., Committee Member
Grant Mastick, Ph.D., Graduate School Representative
Marsha H. Read, Ph. D., Associate Dean, Graduate School
May, 2011
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Abstract
Bark beetles successfully attack host trees by metabolizing host monterpenes
efficiently and producing aggregation pheromones. Investigating enzymes involved in
pheromone biosynthesis and monoterpene metabolism may identify unique molecular
targets and provide information for developing new means to deal with bark beetle
infestations. The three major mountain pine beetle (MPB, Dendroctonus ponderosae)
pheromone components are frontalin, exo-brevicomin and trans-verbenol. Although
these components were identified several decades ago, none of their biosynthetic
pathways are known in detail and none of the involved enzymes are identified or characterized.
This dissertation explores biochemical details of two important MPB pheromone components: exo-brevicomin and trans-verbenol, and bark beetle monoterpene
metabolism in the context of specific enzymes.
In vivo assays revealed that exo-brevicomin was predominantly produced in male
fat bodies, and not in other tissues or by females, and that fat bodies catalyzed the
conversion of decanoic acid to nonen-2-one, confirming that (6Z)-non-6-en-2-one is
likely derived from fatty acid. Fat bodies converted (6Z)-non-6-en-2-one to 6,7-
epoxynonan-2-one, the direct precursor of exo-brevicomin, and (6Z)-non-6-en-2-ol. The
epoxide was stable under physiological conditions. These results implicate a
cytochrome P450 and cyclase in the terminal steps of exo-brevicomin biosynthesis and
(6Z)-non-6-en-2-ol may be the precursor of (6Z)-non-6-en-2-one.
Two novel enzymes are implicated in exo-brevicomin production: a cytochrome
P450, CYP6CR1, and a novel short chain dehydrogenase, ZnoDH. Their mRNA profiles
were consistent with exo-brevicomin production suggests their coordinate regulation. ii
Both enzymes were expressed in Sf9 cells for enzyme assays. Recombinant ZnoDH
oxidized (6Z)-non-6-en-2-ol to the corresponding methyl-ketone, (6Z)-non-6-en-2-one, which serves as the substrate for CYP6CR1. CYP6CR1 converted (6Z)-non-6-en-2-one to 6,7-epoxynonan-2-one. These results suggest both CYP6CR1 and ZnoDH are in the exo-brevicomin biosynthetic pathway. Furthermore, two alternative pathways yielding
production of the C9 precursor from a 10:1-fatty acid are discussed in light of these new
data. While a direct decarboxylation of a β-ketoacyl-CoA intermediate has been
suggested, evidence presented here supports oxidative decarbonylation of an
unsaturated C10 aldehyle to produce 3-nonene, followed by hydroxylation to (6Z)-non-6- en-2-ol and oxidation by ZnoDH to (6Z)-non-6-en-2-one.
Other cytochromes P450 were investigated to explore the evolutionary link
between pheromone production and resin detoxification. Enzyme assays showed
CYP6DH1 and its paralog, CYP6DH2, likely have complementary roles. CYP6DH1
produced verbenol from α-pinene, but did not hydroxylate other monoterpenes and was
not induced by its substrates. Therefore, it likely functions as a pheromone (trans-
verbenol) biosynthetic enzyme. In contrast, CYP6DH2 had a broad substrate range,
suggesting a role in resin detoxification. CYP6DH1 most likely evolved from the
detoxification enzyme, CYP6DH2, by duplication followed by genetic drift.
Similarly, I. pini CYP9T2 had essentially the same substrate profile as CYP6DH2
even though CYP9T2 has been confirmed as a pheromone biosynthetic enzyme. Both
enzymes converted the same substrates into different products, suggesting that different
substrate binding regions may be involved in orienting the substrates. These results
support the evolutionary mechanism that CYP9T2 or its ancestor originally worked as a
detoxification enzyme, but it is now dedicated to pheromone production.
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Co-Authorship
Bessie Liu and Tracy Nguyen (Department of Chemistry, UNR) produced penta- deuterium labeled (6Z)-non-6-en-2-one, (6Z)-non-6-en-2-ol and epoxide (Chapter 2, 3).
Dr. C. Jeffery’s directed the syntheses.
Mory Aw performed CYP6CR1 qRT-PCR analysis for developmental groups (Chapter
3).
Patrick Delaplain contributed to ZnoDH sequencing, qRT-PCR analysis and generated the P3 viral stock (Chapter 4).
Leah Plaugher assisted performing enzyme assays (Chapter 4).
Dr. Chris Keeling (University of British Columbia) provided the unpublished CYP6DH1 cDNA clone (Chapter 5).
Leah Plaugher generated the CYP6DH1 P3 viral stock and helped conduct enzyme assays (Chapter 5).
Andrew Gorzalski and Patricia Kennel determined full-length sequences, performed the qRT-PCR, and produced CYP6DH2 and CYP9T2 viral stocks (Chapter 5). Most of that work appears in A.Gorzalski’s M.Sc. Thesis (Gorzalski, 2010).
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Table of Contents
Chapter 1. Introduction: Mountain pine beetle pheromone component biosynthesis, cytochromes P450 and monoterpene metabolism in bark beetles…………………………1
Chapter 2. exo-Brevicomin biosynthesis in the mountain pine beetle……………………34
Chapter 3. CYP6CR1: a mountain pine beetle cytochrome P450 involved in exo- brevicomin biosynthesis……………………………………………………………………….70
Chapter 4. ZnoDH is a novel dehydrogenase involved in exo-brevicomin biosynthesis in the mountain pine beetle…………………………………………………………………….104
Chapter 5. Monoterpene-metabolizing P450s in pine bark beetle pheromone biosynthesis and resin detoxification………………………………………………………..134 .
Chapter 6. Discussion and future directions…………………………………………….....183
Appendix……………………………………………………………………………………….203
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Chapter 1
Introduction: Mountain pine beetle pheromone component biosynthesis, cytochromes P450 and monoterpene metabolism in bark beetles
I. Biology of bark beetle: Dendroctonus ponderosae II. Cytochromes P450 III. Insect cytochromes P450 IV. Bark beetle cytochromes P450 V. Monoterpene metabolism and evolution in D. ponderosae and Ips pini VI. Objectives and hypotheses VII. References VIII. Figure legends
I. Biology of bark beetle: Dendroctonus ponderosae
I.1. Life cycle
The mountain pine beetle (MPB), Dendroctonus ponderosae Hopkins, is the most destructive pest of coniferous forests in Alaska, Canada, and the western US.
They destroy millions of acres of forest annually and cause significant economic damage through the devaluation of timber, soil destabilization, and increased risk of forest fires
(CBC, 2003; Drooz, 1985; Furniss & Carolin, 1977; Pitman et al., 1968; USDA, 2002;
Waters, 1985). MPBs preferentially attack lodgepole pine (Pinus contorta) and ponderosa pine (P. ponderosae), but they can attack any pine tree during outbreak conditions (Amman et al., 1990). These beetles infest the trunks of large, mature trees.
The current outbreak in North America has extended the MPB range beyond its traditional habitat boundary, and has damaged millions of acres of mature forests. Up to
80% of pine trees in western Canada are predicted to be infested and killed by 2013
(Stickney, 2007).
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The mountain pine beetle is an obligate parasite of pine trees, spending most of its life cycle beneath the bark. Egg laying, hatching and development through larvae, pupae to teneral adults all happen within the phloem of the brood tree. They emerge only for a short, pheromone-mediated flight to new host trees. Pioneer beetles chew through the outer bark to get into the inner phloem. During this process, the tree responds by producing a toxic resin to pitch out the attacking beetles (Phillips & Croteau,
1999; Steele et al., 1995). Healthy trees can produce enough volume to stop a beetle infestation, but under stressful conditions (e.g. drought), the attacked tree cannot
produce enough resin to stop the infestation (Kurz et al., 2008; Rudinsky, 1996). Resin
components produced by pine trees, such as myrcene, α and β-pinene, Δ3-carene, and
limonene, appear most toxic to beetles (Byers, 1995; Raffa et al., 1985). The beetles
also vector an associated blue-stain fungus that grows in the xylem. The extensive
damage from bark beetles and fungi reduces water and nutrient flow, and causes tree
mortality (Graham, 1967; Seybold et al., 2000).
Like many bark beetles, D. ponderosae relies on aggregation pheromones to
coordinate the “mass attack” required for successful tree colonization. The MPB
pheromone system has three main components: trans-verbenol (4,7,7-
trimethylbicyclo[3.1.1]hept-3-en-2-ol), exo-brevicomin (exo-7-Ethyl-5-methyl-6,8-
dioxabicyclo[3.2.1]octane) and frontalin ((1S,5R)-1,5-dimethyl-6,8-
dioxabicyclo[3.2.1]octane). Pioneer MPB females initially attack the host tree, and
hydroxylate α-pinene to produce trans-verbenol, which attracts both males and females
to the attacked tree (Billings et al., 1976; Libbey et al., 1985; Pitman, 1969; Pitman et al.,
1969; Pitman et al., 1968). Males emerging from a brood tree arrive at the new host tree
by responding to trans-verbenol and produce exo-brevicomin, which attracts more
females (Borden et al., 1983; Conn et al., 1983; Libbey et al., 1985). The mixture of
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pheromone components from pioneer and newly arriving beetles serves as a potent
aggregation pheromone, which results in mass attacks. During later stages of the attack, male MPBs reduce exo-brevicomin production and instead produce frontalin as an anti- aggregation signal to repel further MPB colonization (Borden et al., 1987; Miller, 2002;
Pureswaran et al., 2000; Ryker & Libbey, 1982).
I.2 Pheromone biosynthesis
Pheromones produced by the MPBs are very important for their reproductive cycle. Pheromone components may be synthesized either by modifying host precursor molecules, or by de novo synthesis. Most published studies of bark beetle pheromone component biosynthesis indicate de novo pheromone production. Ipsdienol and ipsenol, pheromone components in Ips spp., are synthesized de novo via the mevalonate pathway (Seybold et al., 1995; Tillman et al., 1998). De novo frontalin biosynthesis has also been reported for D. jeffreyi and D. ponderosae (Barkawi et al., 2003). However, in
most cases the enzymes involved in pheromone component biosynthesis have not been
identified.
The three major D. ponderosae pheromone components appear to arise from
different sources (Figure 1.1). trans-Verbenol is a bicyclic monoterpenoid alcohol. It is a female-specific aggregation pheromone component and can be induced by feeding
(Billings et al., 1976; Conn et al., 1983; Hughes, 1973; Libbey et al., 1985; Pierce et al.,
1987; Pitman et al., 1968; Pureswaran et al., 2000). trans-Verbenol is most likely
produced via cytochrome P450-mediated hydroxylation of a host tree resin component,
α-pinene (Greis et al., 1990; Hunt & Smirle, 1988; Pierce et al., 1987) although there is
evidence supporting de novo biosynthesis in D. frontalis (Renwick et al., 1973). De novo
biosynthesis via the mevalonate pathway may be possible in D. ponderosae (Blomquist
et al., 2010; C. Keeling, personal communication).
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The exo-brevicomin biosynthetic pathway is not fully understood. Teneral male
MPBs contain a detectable level of exo-brevicomin, but females do not (Borden et al.,
1987; Libbey et al., 1985; Pureswaran et al., 2000; Vanderwel et al., 1992). Feeding
decreases exo-brevicomin levels (Pureswaran et al., 2000). exo-Brevicomin is likely
synthesized de novo by epoxidation and cyclization of its precursor (6Z)-non-6-en-2-one
(Francke et al., 1996; Vanderwel et al., 1992). (6Z)-Non-6-en-2-one is believed to be
derived either via β-oxidation of long chain fatty acids, as proposed for pheromone
biosynthesis in Lepidoptera (Rafaeli & Jurenka, 2003), or directly via fatty acid
elongation to decanoic acid, followed by desaturation and a decarboxylation of decenoic
acid (Vanderwel et al., 1992). Conversion of (6Z)-non-6-en-2-one to exo-brevicomin
likely involves a cytochrome P450 mediated formation of the keto-epoxide intermediate
(Vanderwel et al., 1992) that is subsequently cyclized either enzymatically or possible
without an enzyme catalyst (Belles et al., 1969).
Frontalin, a male specific anti-aggregation pheromone component (Borden,
1985; Pureswaran et al., 2000; Ryker & Libbey, 1982), is believed to be synthesized de
novo via the mevalonate pathway in the midgut of male beetles (Barkawi et al., 2003;
Hall et al., 2002). It is likely made from its precursor, 6-methylhept-6-en-2-one, via
cytochrome P450-mediated epoxidation followed by a cyclization (Perez et al., 1996). 6-
Methylhept-6-en-2-one may be derived from monoterpenoid or longer chain precursors
via a dioxygenase-catalyzed reaction (Blomquist et al., 2010).
II. Cytochromes P450
Cytochromes P450 (abbreviated as P450s) are a large superfamily of heme thiolate proteins (Feyereisen, 1999). They were discovered over fifty years ago.
Cytochromes P450 have been named on the basis of their cellular location and
5 spectrophotometric characteristics: when the reduced heme iron forms an adduct with
CO, P450 enzymes absorb light at wavelengths near 450 nm as a characteristic peak
(Bernhardt, 2006). These enzymes have a broad substrate range, including metabolic intermediates such as lipids and steroidal hormones, as well as xenobiotic substances such as drugs and other toxic chemicals. Over 11,500 distinct CYP proteins have been identified in in animals, plants, fungi, protists, bacteria, archaea, and even viruses
(Danielson, 2002; htpp://drnelson.utmem.edu /CytochromeP450.html; Sigel et al., 2007).
In humans they play very important roles in drug and other xenobiotic metabolism, as well as cholesterol biosynthesis and steroidogenesis. Due to their diverse range of functions, cytochromes P450 have been investigated by researchers in many fields
(Bernhardt, 2006). In insects, P450 studies have focused mostly on roles in pesticide resistance (Chiu et al., 2008; Feyereisen, 2005; McCart & Ffrench-Constant, 2008) and adaption to host plants (Li et al., 2007).
Genomic studies show that the number of P450 genes differs among different species: 57 genes for human, over 100 genes for mice, more than 250 for some plants, and three genes for yeast (Saccharomyces cerevisae). Insects have approximately 100
P450 genes on average (Helvig et al., 2004; Werck-Reichhart & Feyereisen, 2000).
P450s differ in size and location in different cells. Eukaryotic P450s are usually about 500 amino acids, and are localized either in the inner mitochondrial membrane or cytoplasmic side of the endoplasmic reticulum (ER) membrane via an N-terminal hydrophobic peptide domain, whereas prokaryotic P450s are usually about 400 amino acids and soluble (Feyereisen, 2005; Werck-Reichhart & Feyereisen, 2000).
Endoplasmic reticulum P450s are usually isolated from mitochondrial and soluble proteins by differential centrifugation.
6
The monooxygenase reaction is the most common reaction catalyzed by cytochromes P450, which inserts one atom of oxygen into an organic substrate while the other oxygen atom is reduced to water (Feyereisen, 2005). P450s also can perform a large variety of reactions including epoxidation, hydroxylation, dehalogenation, N- and S-
oxidations, O-, N-, S-dealkylation, and many other uncommon reactions (Bernhardt,
2006; Feyereisen, 1999). Most P450s require a protein partner to deliver one or more
electrons to reduce the iron. Therefore, cytochromes P450 are part of P450-containing
systems of proteins. P450-containing systems include four classes. Class I is found on
the membranes of mitochondria or in bacteria, in which a flavin adenine dinucleotide
(FAD)-containing ferredoxin reductase and an iron-sulfur protein forms a short electron
chain to supply the electrons (Hanukoglu, 1996; Werck-Reichhart & Feyereisen, 2000).
Class II includes microsomal P450s that are anchored on the outer membrane of the ER,
and are made up of an FAD- and flavin mononucleotide (FMN)-containing NADPH-
dependent cytochrome P450 reductase (CPR) and a P450 (Bernhardt, 2006; Hanukoglu,
1996). Class III and IV systems are less common. Class III P450s were originally found
in Bacillus megaterium, containing the same enzymes as class II, but fused in a single
continuous polypeptide (Hanukoglu, 1996; Werck-Reichhart & Feyereisen, 2000). Class
IV P450s receive their electrons directly from NAD(P)H without any partner protein
(Werck-Reichhart & Feyereisen, 2000). Class IV P450s include thromboxane-A
synthase, a CYP5A family in humans (Baek et al., 1996; Yokoyama et al., 1991) and
CYP8A1, an enzyme involved in prostanoid biosynthesis in humans (Yokoyama et al.,
1996).
Most P450s share common properties, i.e., heme and oxygen binding, electron
transfer and oxygen activation. The differences among them lie in the properties of their
substrates. The substrates can be a variety of small and large molecules (Fennell et al.,
7
2004; Lambalot et al., 1995). Some P450s are specific for their substrates and have
narrow product profiles, whereas many P450s can accept multiple substrates and some
even catalyze multiple reactions (Bernhardt, 2006; Feyereisen, 1999; Wen et al., 2005;
Wen et al., 2003). Primary sequences of P450s are highly diverse. Cytochromes P450
in a specific family or with high sequence homology do not necessarily have similar
biological function or enzymatic activity (Feyereisen, 2005; Werck-Reichhart &
Feyereisen, 2000). Their substrate specificities do not depend on only overall sequence,
but also on hyper-variable regions (Wen et al., 2005). The contribution of these regions
for substrate selection remains poorly understood. Indeed, single amino acid differences
in regions that do not directly bind substrate can dramatically change substrate
specificities (Wilderman et al., 2011). Therefore, functional characterization of individual
P450 to assign biological roles is very important (Wen et al., 2005; Wen et al., 2003).
II. 1 Nomenclature
The root for cytochrome P450 names is CYP and families are designed by an
Arabic number indicating the gene family, followed by a capital letter indicating the
subfamily, and another numeral for the individual gene (htpp://drnelson.utmem.edu
/CytochromeP450.html). The guidelines for assigning a name suggest that members of
new CYP families share >40% amino acid identity, while members of subfamilies must
share >55% amino acid identity (htpp://drnelson.utmem.edu/CytochromeP450.html).
New P450s are typically assigned names by the P450 Nomenclature Committee,
supervised by D. Nelson (U. Tennessee).
II.2 Catalytic cycle
Although properties and reactions catalyzed by P450s differ in many aspects, there is a common P450 catalytic cycle. The catalytic cycle depends on the P450 heme group and related electron transfer chain (Hanukoglu, 1996; Oprea et al., 1997). The
8 heme group contains a protoporphyrin ring with four methyl, two vinyl, and two propionic acid side chains and a central iron atom. The iron has six ligands; four are from the four porphyrin pyrrole ring nitrogens, the fifth ligand tethers the heme to the protein via a thiolate bond from a cysteine residue on the proximal side of the heme, and the sixth ligand on the distal side of the heme coordinates with the substrate, or with water in the absence of substrate (Werck-Reichhart & Feyereisen, 2000). The first four steps of the
P450 catalytic cycle are well known, but the properties of final steps are still unclear
(Feyereisen, 1999; Poulos, 2005; Werck-Reichhart & Feyereisen, 2000). Substrate binding to the active site induces a conformation change in the active site (Meunier et al.,
2004). This change favors the transfer of an electron from NAD(P)H via P450- containing systems (Sligar et al., 1979) to reduce the ferric heme iron to the ferrous state.
When carbon monoxide (CO) binds to this reduced P450, the protein-CO complex yields the classic CO difference spectrum with a maximum absorbance at 450 nm (Feyereisen,
2005; Omura & Sato, 1964) and the catalytic cycle is disrupted (Meunier et al., 2004;
Montellano & Paul, 2005). If the reduced heme iron binds O2, the second electron transfer via the electron-transport system is induced (Bernhardt, 2006; Feyereisen,
1999). The O-O bond is cleaved, resulting in release of one water molecule and formation of a highly reactive species (Compound I) (Meunier et al., 2004; Poulos, 2005).
The oxygen atom remaining bound to the heme is then inserted in the substrate (Rittle &
Green, 2010). Finally, hydrocarbon hydroxylation is believed to involve hydrogen atom abstraction from a C-H bond of the substrate, insertion of the iron-ligated oxygen atom to the substrate, and release of the newly formed hydroxylated product (Bernhardt, 2006;
Guallar et al., 2003; Kamachi & Yoshizawa, 2003). After the product has been released from the active site, the enzyme returns to its original state.
The 450 nm peak of cytochrome P450 is different from other heme containing
9
proteins. For example, hemoglobin absorbs at 420 nm because its fifth ligand is an
imidazole from histidine instead of a thiolate provided by cysteine in P450s (Oprea et al.,
1997; Werck-Reichhart & Feyereisen, 2000). CO difference spectrum analysis is routinely used to identify functionally expressed P450 and quantify the total active P450 in a sample (Feyereisen, 2005). The heme iron in P450 protein is reduced with sodium dithionite, and then bound to CO. The difference spectrum is measured between reduced enzyme and reduced enzyme-CO complex (Omura & Sato, 1964). The
functional P450 concentration can be calculated according to Omura & Sato (1964).
III. Insect Cytochromes P450
Cytochromes P450 have been widely studied in insects to understand pesticide
resistance and the adaptation to host plants (Feyereisen, 2005; Li et al., 2007). For
example, CYP6Z1, a P450 in the mosquito (Anopheles gambiae), can metabolize DDT
(Chiu et al., 2008) and CYP6G1 is implicated in insecticide resistance in DDT-resistant
Drosophila melanogaster (McCart & Ffrench-Constant, 2008). CYP6A1 is a xenobiotic-
metabolizing P450 from the house fly (Andersen et al., 1997; Jacobsen et al., 2006;
Sabourault et al., 2001). Most insect cytochromes P450 belong to the CYP 4, 6, 9 and
12 families. CYP4 and CYP6 are the two largest families (Helvig et al., 2004; Ranson et
al., 2002). The CYP4 family is an ancient family distributed across many species
(Feyereisen, 2005; Maïbèche-Coisne et al., 2002). The CYP 6 and 9 families represent
environmental response genes (Berenbaum, 2002). The CYP12 family includes
mitochondrial P450s (Feyereisen, 1999; Le Goff et al., 2006). The average number of
P450 genes in several insects is about 100 according to genomic analyses (Feyereisen,
2006; htpp://drnelson.utmem.edu/ CytochromeP450.html; Tijet et al., 2001). However,
there is some variability among different species. For example, the honeybee, Apis
10
mellifera, has only 46 P450s, while the red flour beetle, Tribolium castaneum, has the
most, 143 P450s (Feyereisen, 2006). Insect P450s play important roles in the synthesis and degradation of endogenous compounds (Feyereisen, 2005). Currently only a few of
insect P450s have assigned functions (Feyereisen, 2005, 2006). Some studies have
shown that transcriptional responses of a P450 can be regulated by inducers such as
substrates or some other regulator, such as a hormone. Xenobiotic-metabolizing P450s
are often induced by their substrates (Le Goff et al., 2006; Poupardin et al., 2008; Wen
et al., 2009). This information offers great advantage to identify and assign P450
functions by combining expression analysis and functional assays of expressed
recombinant proteins.
IV. Bark Beetle Cytochromes P450
Successful tree colonization by bark beetles requires detoxification enzyme
systems and pheromonal enzyme systems. Bark beetles have developed detoxification
enzyme systems to tolerate poisonous alleochemicals in pine resin, including
monoterpenes (Berenbaum, 2002; Seybold et al.,1995). Monoterpene metabolism
commonly involves cytochromes P450 in several organisms. For example, the
hydroxylation of limonene with plant cytochrome P450 limonene hydroxylases has been
characterized (Karp et al., 1987; Karp et al., 1990; Kjonaas et al., 1985; Lupien et al.,
1999). Recombinant human CYP2B6, CYP2C19 and CYP2D6 hydroxylate Δ3-carene
into Δ3-carene-10-ol, while human CYP1A2 converts Δ3-carene to Δ3-carene-epoxide
(Duisken et al., 2005). Δ3-Carene metabolism by larvae of the common cutworm
(Spodoptera litura) suggested a cytochrome P450 was involved (Miyazawa & Kano,
2010). Monoterpene-metabolizing P450s have been proposed or demonstrated in bark beetles (Pierce et al., 1987; Sandstrom, 2007; White et al., 1979; White et al., 1980).
11
Hunt et al. (1988) showed P450s were involved in (-)-α-pinene and myrcene metabolism
in MPB. Blocking P450 with piperonyl butoxide, a general P450 inhibitor, resulted in
deceased levels of certain terpene alcohol pheromone components, along with a buildup
in the levels of their precursors (Hunt & Smirle, 1988). Cytochromes P450 are also good
candidates for producing pheromone components (Seybold & Tittiger, 2003). Ips pini
CYP9T2 and Ips confusus CYP9T1, as pheromonal biosynthetic enzymes, hydroxylate
myrcene to ipsdienol, confirming that P450s are involved in pheromone biosynthesis
(Sandstrom et al., 2008; Sandstrom et al., 2006). Biosynthesis of cis-verbenol (Byers,
1983; Renwick et al., 1976), trans-verbenol (Barkawi et al., 2003; Byers, 1983; Perez et
al., 1996; Pitman et al., 1969), exo-brevicomin (Perez et al., 1996; Vanderwel et al.,
1992), and frontalin (Barkawi et al., 2003; Perez et al., 1996) in MPB almost certainly
involves cytochromes P450. However enzymes involved in the biosynthesis of these
pheromone components have not been identified.
V. Monoterpene metabolism and pheromone evolution in D. ponderosae and Ips
pini
Some bark beetle pheromone components are the same as monoterpene
metabolites, suggesting an evolutionary path by which hydroxylated monoterpenes
became aggregation pheromone components (Blomquist et al., 2010). In doing so, the
enzyme systems initially developed to hydroxylate monoterpenes probably gave rise to
pheromone biosynthetic enzymes (Blomquist et al., 2010).
The link between pheromones and host tree resins was suggested as soon as the first bark beetle pheromones were identified as monoterpenoid alcohols (Hughes,
1973). Ancestral beetles successfully colonized a host by utilizing resin detoxification
mechanism(s), which mostly involve P450s (Huber et al., 2007; Hunt & Smirle, 1988;
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Pierce et al., 1987; White et al., 1979; White et al., 1980) to hydroxylate tree
alleochemicals to less toxic and more soluble forms (Bernhardt, 2006; Tillman et al.,
1999). Hydroxylated monoterpenes, such as ipsdienol and trans-verbenol, became aggregation pheromone components (Hughes, 1973; Renwick et al., 1976), suggesting a clear evolutionary path. The metabolic monoterpene products would have exerted selective evolutionary pressure on conspecifics which were able to sense the presence of the metabolite and respond to its meaning. Later, beetles would have evolved the ability to control the synthesis of monoterpene alcohols to better regulate the pheromonal signal (Blomquist et al., 2010; Seybold & Tittiger, 2003). Thus, products from detoxification reactions became aggregation pheromone components in these ancestral beetles. Evidence for this is present in modern bark beetles. For example, there is no published evidence supporting de novo trans-verbenol biosynthesis, suggesting that pheromonal trans-verbenol is derived either from host tree α-pinene or from endosymbionts (Rao et al., 2003; Renwick et al., 1976). Although ipsdienol in I. pini is a de novo product (Seybold et al., 1995), host tree precursors may join the same metabolic pool as de novo precursors, and they are indistinguishable to downstream enzymes (Hendry et al., 1980; Ivarsson & Birgersson, 1995; Lu, 1999). Lu et al. (1999) showed that exogenously supplied myrcene was converted into pheromonal ipsdienol by
I. pini. In contrast, frontalin and exo-brevicomin are both semiochemicals with no link to detoxification. They most likely represent alterations of existing metabolic pathways to form novel chemicals with informational content. In this respect, they are more similar to lipid-based pheromone components in other insects (Rafaeli & Jurenka, 2003). Indeed, exo-brevicomin is likely a lipid-based pheromone component (Perez et al., 1996;
Vanderwel et al., 1992). Frontalin and exo-brevicomin are often grouped together because of structural similarity between them or their precursors, although their
13 precursors are derived from different pathways. Frontalin is clearly synthesized from the mevalonate pathway in the midgut (Barkawi et al., 2003; Hall et al., 2002), whereas exo- brevicomin is likely derived from fatty acids (Vanderwel et al., 1992). It is possible that the same P450 catalyzes the epoxidation of both precursors. Both endo- and exo- brevicomin are produced by male MPB with a similar production pattern from their precursors, (6E)-non-6-en-2-one and (6Z)-non-6-en-2-one, respectively (Pureswaran et al., 2000; Vanderwel et al., 1992). Their production may depend on related desaturases that produce trans- or cis- desaturated fatty acid precursors. Downstream enzymes may not discriminate between trans- and cis- precursors. A change in desaturase activity can result in changes in pheromone biosynthesis in insects (Liénard et al., 2008; Roelofs
& Rooney, 2003). Symonds & Elgar (2008) suggested that minor changes to existing metabolic pathways can generate a largely diverse suite of pheromone components.
Pheromone biosynthetic enzymes may be distinguishable from detoxification enzymes based on alterations in enzyme activity and or regulation. The difference between the two may conceivably occur following simple point mutations in coding or regulatory regions or duplication. Ips pini GPPS is likely derived from its ancestral
GGPPS (Gilg et al., 2005). The primary structural differences between GPPS and
GGPPS result in differences in the substrate binding pockets: GPPS has a smaller and less selective pocket and can accept GPP as a substrate to produce myrcene (Gilg et al.,
2009). Ips pini CYP9T2 is not induced by myrcene (A. Griffith, unpublished data), and clearly is a pheromone-biosynthetic enzyme (Sandstrom et al., 2006). It also hydroxylates α-pinene (Sandstrom, 2007). These may be evidence for an evolutionary history, a detoxification enzyme becoming a pheromone biosynthetic enzyme.
VI. Objectives and Hypotheses
14
Current pest management strategies have proven ineffective to regulate pine bark beetles. Future control strategies may benefit if species-specific targets are identified. Pheromone biosynthesis is an attractive focus area because pheromone components are often species-specific. In addition, understanding how pheromone production evolved can help us understand and predict how bark beetles will respond to management efforts. Unfortunately, very few confirmed pheromone-biosynthetic enzymes have been identified in bark beetles.
The link between resin detoxification and pheromone production suggests that enzymes in the two processes should be closely related. CYP9T2 likely evolved from a detoxification enzyme, and later became a pheromone biosynthesis enzyme (Sandstrom,
2007; Sandstrom et al., 2006). trans-Verbenol could also be an original monoterpene detoxification product, and then became a pheromonal component, implicating the P450 responsible for its production may be involved with the evolution from a detoxification enzyme. Comparing the substrate and product profiles of bark beetle cytochromes P450 in D. ponderosae and I. pini should provide molecular information about the evolution of pheromone production and address the hypothesis that the pathways for monoterpene detoxification and monoterpenoid pheromone biosynthesis are evolutionarily related
(Greis et al., 1990; Seybold et al., 2000; Vanderwel & Oehlschlager, 1987). Pheromone biosynthetic P450s should be distinguishable from detoxification P450s based on activity and regulation. A detoxification P450 is likely to have a broad substrate range and be induced by those substrates, while a pheromone biosynthetic enzyme may have a strong preference for a specific precursor and perhaps be regulated by hormone and feeding (Blomquist et al., 2010).
Microarray analysis from cDNA derived from RNA isolated from 11 developmental groups demonstrated over 30 cytochromes P450 in the MPB (Aw et al.,
15
2010). Some P450 genes have expression patterns that indicate that they may be
important for detoxification or pheromone biosynthesis. For example, CYP6CR1 expression is consistent with exo-brevicomin production. In contrast, CYP6DH2 is
expressed in a pattern suggesting a role in digestion or detoxification (Aw et al., 2010).
Aw et al. (2010) showed a cluster of three genes, including CYP6CR1 and ZnoDH (for
(6Z)-non-6-en-2-one dehydrogenase; identified by its EST, DPG022F04, see below,
Chapter 4), with apparent coordinate regulation consistent with exo-brevicomin
biosynthesis. The “functional genomics followed by enzyme characterization” approach
was applied in I. pini and led to the identification of important pheromone-biosynthetic
enzymes (Figueroa-Teran et al., 2011; Keeling et al., 2004; Sandstrom et al., 2006).
Thus, we hypothesized that both CYP6CR1 and ZnoDH are responsible for exo-
brevicomin biosynthesis (Figure 1.1). However, none of these enzymes have been
biochemically characterized or assigned a function, so their putative roles remain
unconfirmed. Ips pini CYP9T2, as a pheromonal enzyme, has been studied in some
detail (Sandstrom, 2007; Sandstrom et al., 2008; Sandstrom et al., 2006), but its
potential role in detoxification remains unknown.
Using these preliminary data, I began a series of investigations into exo-
brevicomin biosynthesis, the roles of P450s in exo-brevicomin and trans-verbenol
production, and monoterpene metabolism and evolution by cytochromes P450 in D.
ponderosae and I. pini. This research was based on our understanding of the exo-
brevicomin pathway and biased towards those P450s potentially involved in pheromone
production and detoxification (Aw et al., 2010; Gorzalski, 2010; Sandstrom, 2007;
Sandstrom et al., 2008; Sandstrom et al., 2006). CYP6CR1, ZnoDH and CYP6DH2
from D. ponderosae were cloned and sequenced utilizing an expressed sequence tag
(EST) library (See Appendix Figure A.1-A.2; Gorzalski, 2010); CYP6DH1, a CYP6DH2
16 homolog with 16 amino acids different and likely involved in pheromonal biosynthesis
(Figure 1.1; Gorzalski, 2010; Appendix Figure A.3), was a gift from C. Keeling (UBC,
Canada). Ips pini CYP9T2 (Sandstrom et al., 2006) also was studied to understand its detoxification function. A molecular approach to study bark beetle cytochromes P450 should lead to a better understanding of pheromone biosynthesis, bark beetle-host tree interactions and the expected evolutionarily related pathways for monoterpene detoxification and monoterpenoid pheromone biosynthesis. It may eventually also provide valuable information for developing new means to control these pests.
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VIII. Figure legends:
Figure 1.1 Biosynthetic pathways for D. ponderosae pheromone components. exo-
Brevicomin is likely derived from a fatty acid precursor (Vanderwel et al., 1992), frontalin is synthesized de novo via the mevalonate pathway (Barkawi et al., 2003), and trans- verbenol probably arises from host tree α-pinene (Gries et al., 1990), although de novo biosynthesis is possible (dashed line; C. Keeling personal communication).
Uncharacterized steps are shown in grey, and uncharacterized enzymes are indicated in parentheses. HMG-R, 3-hydroxy-3-methylglutaryl coenzyme A reductase; HMG-S, 3- hydroxy-3-methylglutaryl coenzyme A synthase; GPP, geranyldiphosphate; GPPS, geranyl diphosphate synthase; IPPI, isopentenyl diphosphate isomerase; P, phosphate.
33
Fatty acid Pathway Mevalonate Pathway Host Tree -O O acetate Thiolase O O O- P HMG-R, OH Fatty acid biosynthetic O enzymes HMG-S glyceraldehyde-3 pyruvate IPPI -phosphate 16:0 /18:0 acid GPPS (Thioesterase II) DOX-P pathway (Desaturase) P P 16:1 / 18:1 acid GPP P P (Limited β-oxidation) (Dioxygenase) O GPP 10:0 /10:1 acid (Desaturase/ sulcatone dehydrogenase/ ZnoDH?) (Isomerase) decarboxylation α-pinene O O
(6Z)-non-6-en-2-one 6-methylhept-6-en-2-one (?) (ingestion/inhalation) (P450/CYP6CR1?) (P450) O O O O 6-methylhept-6-epoxy-2-one α-pinene (Cyclase) (Cyclase) (P450/ CYP6DH1?)
O O O HO O trans-Verbenol exo-Brevicomin Frontalin
Figure 1.1 34
Chapter 2
exo-Brevicomin biosynthesis in the mountain pine beetle
I. Abstract II. Introduction III. Materials and Methods IV. Results V. Discussion VI. Acknowledgements VII. References VIII. Figure Legends
I. Abstract
The mountain pine beetle (MPB; Dendroctonus ponderosae) relies on pheromones to complete its reproductive cycle. Investigating pheromone biosynthesis in the MPB may identify unique molecular targets and provide information for developing new means to deal with this devastating forest pest.
exo-Brevicomin is a male-specific MPB aggregation pheromone component.
Feeding results in decreased exo-brevicomin levels. Its biosynthetic pathway is likely de novo from the fatty acid-derived precursor, (6Z)-non-6-en-2-one, which is likely produced from the 10-carbon fatty acid through decarboxylation. (6Z)-Non-6-en-2-one is likely converted to 6,7-epoxynonan-2-one by a cytochrome P450, and further converted to exo-brevicomin by a cyclase. Examination of exo-brevicomin distribution in MPB coupled with GC-MS analysis confirmed that exo-brevicomin was present in unfed males, but not in females. Surprisingly, exo-brevicomin was present in male fat bodies, but not in carcasses or guts. This is the first example of a bark beetle pheromone produced in fat body, not in midgut. Unfed male fat bodies converted tri-deuterium 35
labeled decanoic acid to nonan-2-one, suggesting a decarboxylation activity in the fat
body, strongly implicating that 10:1 fatty acid is the direct precursor to (6Z)-non-6-en-2-
one and that there is no short chain fatty acid desaturase activity in the male fat body.
Homogenized male fat bodes converted (6Z)-non-6-en-2-one into the 6,7-epoxy-nonan-
2-one, the direct precursor of exo-brevicomin. A P450 inhibitor reduced the amount of epoxide produced by fat bodies. The epoxide cyclized into exo-brevicomin non- enzymatically at pH below 3.0. Taken together, these data strongly suggest that a decarboxylase, a cytochrome P450 and a cyclase in male fat body are required for the terminal steps of exo-brevicomin production. Surprisingly, homogenized male fat bodes also converted (6Z)-non-6-en-2-one into (6Z)-non-6-en-2-ol, implying that (6Z)-non-6-en-
2-ol may be the direct precursor of (6Z)-non-6-en-2-one in the beetle and that a dehydrogenase may be required for this step.
II. Introduction:
The mountain pine beetle (MPB; Dendroctonus ponderosae) is the most destructive pest of coniferous forests in Alaska, Canada, and the western US. They destroy millions of acres of forest annually and cause significant economic damage through the devaluation of timber, soil destabilization, and increased risk of forest fires
(CBC, 2003; Drooz, 1985; Furniss & Carolin, 1977; Pitman et al., 1968; USDA, 2002;
Waters, 1985). Up to 80% of pine trees in western Canada are predicted to be infested and killed by 2013 (Stickney, 2007). Infestations by these beetles could prevent the boreal forest from removing greenhouse gas (CO2) from the atmosphere and thereby
contribute to climate change (Kurz et al., 2008). MPBs preferentially attack lodgepole
(Pinus contorta) and ponderosa (P. ponderosae) pines, but they can attack any pine tree
during outbreak conditions (Amman et al., 1990). They preferentially infest the trunks of 36
large mature trees. These aggressive beetles spend most of their lives under the bark of
trees and emerge only for a short, pheromone-mediated flight to new host trees.
Because they are physically protected from sprayed pesticides, there are currently few effective tools to control the beetles.
Pheromones are very important for the beetle’s reproductive cycle (Rudinsky,
1996; Wood, 1982). Bark beetle populations may be managed if a pheromone
biosynthetic pathway is disrupted. Therefore, investigating pheromone production may
identify unique molecular targets and provide information for developing new means to
deal with bark beetle infestations, such as developing dsRNA to target specific
pheromone biosynthesis gene or producing a pheromone as a bait using recombinant
pheromone biosynthetic enzymes.
A few “pioneer” females initially attack the host tree, and feeding results in
production of the aggregation pheromone, trans-verbenol (4,7,7-
trimethylbicyclo[3.1.1]hept-3-en-2-ol), which attracts both males and females (Hughes,
1973; Libbey et al., 1985; Pitman et al., 1968; Pitman et al., 1969). Males emerging
from a brood tree produce exo-brevicomin (exo-7-ethyl-5-methyl-6,8-
dioxabicyclo[3.2.1]octane), which attracts more females to assist the attack (Borden et
al., 1983; Conn et al., 1983; Libbey et al., 1985). The mixture of pheromones from newly
arriving and pioneer beetles serves as a potent aggregation pheromone, which results in
mass attacks. During later stages of the attack, trans-verbenol and exo-brevicomin
levels decline and males instead produce frontalin [(1S,5R)-1,5-dimethyl-6,8-
dioxabicyclo[3.2.1]octane] as an anti-aggregation signal to repel further colonization (
Borden et al., 1987; Francke et al., 1996; Miller, 2002; Pureswaran et al., 2000; Ryker &
Libbey, 1982). 37
The exo-brevicomin biosynthetic pathway is not fully understood. Emerged male
MPBs contain detectable exo-brevicomin, but not females (Borden et al., 1987; Pierce et al., 1987; Pureswaran et al., 2000; Vanderwel et al., 1992). Feeding results in
decreased exo-brevicomin levels (Pureswaran et al., 2000). (6Z)-Non-6-en-2-one was proposed as a natural precursor of exo-brevicomin in MPBs and western balsam bark beetles (WBBB; Dryocoetes confusus) because it was detected in the volatiles of male
MPBs and WBBBs (Vanderwel et al., 1992). Vanderwel et al. (1992) also showed that male MPBs produced significantly more exo-brevicomin when exposed to the precursor than did females.
The biosynthetic origin of (6Z)-non-6-en-2-one is unclear. It is possible that this compound is supplied by the host trees or by associated fungi (Borden, 1985; Vanderwel
& Oehlschlager, 1987). A more likely possibility is de novo synthesis or modification of a
ω-3 mono-unsaturated 10-carbon fatty-acyl compounds (Figure 2.1) (Vanderwel et al.,
1992a). There are at least two possible routes to the 10–carbon fatty acyl intermediate:
one is from a long chain (16 or 18-carbon) fatty acyl precursor through desaturation and
limited chain shortening by β-oxidation (Figure 2.1), as proposed for pheromone
production in Lepidoptera (Rafaeli & Jurenka, 2003). Alternatively, the 10-carbon
intermediate may be directly produced by curtailing fatty acid elongation during synthesis
(Figure 2.1). Vanderwel et al. 1992) proposed that the 10-carbon intermediate, (Z)-7-
decenoyl-CoA, may be oxidized to a 3-keto-(Z)-7-decenoyl intermediate and then
decarboxylated to (6Z)-non-6-en-2-one. Experiments in drosophila show that fatty acid- derived, odd-numbered 2-ketones are likely produced by decarboxylation of the β-keto- fatty acid (Skiba & Jackson, 1994). The conversion of (6Z)-non-6-en-2-one to exo- brevicomin likely involves a cytochrome P450-mediated formation of a keto-epoxide intermediate (Vanderwel et al., 1992) (Figure 2.1), and this keto-epoxide in turn is 38
subsequently cyclized either enzymatically or possibly without an enzyme catalyst
(Silverstein et al., 1968) (Figure 2.1). However, none of these steps are confirmed.
In this study, we hypothesized that exo-brevicomin is produced in the midgut
because most bark beetle pheromones are made in the midgut (Barkawi et al., 2003;
Hall et al., 2002a; Hall et al., 2002b). We also hypothesized that a 10-carbon mono-
unsaturated fatty acid is converted into (6Z)-non-6-en-2-one by a series of
dehydrogenating and decarboxylation steps, and that the conversion of (6Z)-non-6-en-2- one to exo-brevicomin is through the epoxidation of the keto-alkene, followed by the cyclization of the formed keto-epoxide (Figure 2.1).
To confirm these hypotheses, I performed in vivo experiments using beetles or
beetle tissues coupled with gas chromatography–mass spectrometry (GC-MS) analysis
to better understand exo-brevicomin biosynthesis in detail. I confirmed that exo-
brevicomin is present in unfed males, and not in females. Surprisingly, exo-brevicomin
was produced by male fat bodies, but not by carcasses or digestive tissue. This is the
first report of a bark beetle pheromone produced in fat body, not in midgut. Isolated
intact male fat bodies converted tri-deuterated decanoic acid to tri-deuterated nonan-2- one. Homogenized male fat bodes converted penta-deuterium labeled (6Z)-non-6-en-2- one into penta-deuterium labeled keto-epoxide, the direct precursor of exo-brevicomin, and not into exo-brevicomin, and this epoxide was cyclized into exo-brevicomin in the presence of acid with pH value less than 3. These studies confirmed the hypothesized biosynthetic pathway of exo-brevicomin and strongly suggest that a decarboxylase, a cytochrome P450 and a cyclase in male fat body are required for the terminal steps of exo-brevicomin production. (6Z)-non-6-en-2-ol yielded from (6Z)-non-6-en-2-one by fat bodies implies a possible alternative pathway in the production of (6Z)-non-6-en-2-one in the beetle. 39
III. Materials and methods
Reagents and chemicals. Hink’s TNM-FH Medium 1x (Supplemented Grace’s
Medium) was from Mediatech, Inc. (Herndon, VA). Unlabeled (6Z)-non-6-en-2- one and exo-brevicomin were purchased from PheroTech Inc. (Delta, BC,
Canada). NAD+, NADP+, NADPH, piperonyl butoxide (PBO), 10,10,10-
deuterated decanoic acid, BF3/MeOH, phenylmethylsulfonyl fluoride (PMSF) and
protease inhibitor cocktail were from Sigma-Aldrich (St. Louis, MO). Deuterium
labeled penta-(6Z)-non-6-en-2-one, epoxide, and (6Z)-non-6-en-2-ol were made by C. Jeffery (Department of Chemistry, University of Nevada, Reno). 2X
NADPH regeneration system was from AAT Bioquest (Sunnyvale, CA).
Beetles. Mountain pine beetles were obtained from ponderosa pine bolts collected from the Sierra Nevada in California and Nevada, USA. The bolts were stored in a greenhouse and emerged adults were collected daily and stored for up to 1-2 weeks at 4 ºC on moist paper towels in loosely capped jars (Browne,
1972). The emerged beetles were separated by sex (Wood, 1982). Only healthy
beetles were selected for experiments, and all groups of beetles were dissected
on the same day. For unfed samples, males and females were held separately
in plastic cups with the lids allowing enough air and moisture to enter, covered
with moist paper in the dark at room temperature overnight as previously
described (Keeling et al., 2004). For tissue distribution studies, male MPBs were
starved overnight, and then dissected under a microscope in 100 mM sodium
phosphate buffer pH 7.8. Fat bodies, midguts and carcasses were isolated into
0.5 ml ice-cold cell lysate buffer (100mM sodium phosphate, pH 7.8, 1.1 mM
EDTA, 0.1 mM DTT, 0.5 mM PMSF, 1/1000 vol/vol Sigma protease inhibitor
cocktail, 20% glycerol) or culture medium (20 μg/ml gentamycin, 4/1000 vol/vol 40
Sigma protease inhibitor cocktail, 15 μg/ml tetracycline, 1 mM PMSF and 12 mM glucose in Hink’s TNM-FH medium). Fresh isolated tissues were used immediately. Ten beetles were used for carcass and midgut groups and 20
beetles for fat body groups for each replicate with four replicates in total. To
check exo-brevicomin production in males or females, whole bodies from each
group were used, and each sample contained seven insects/replicate with three
replicates for each group.
exo-Brevicomin extraction. Whole bodies of unfed males, females, or isolated unfed
male fat bodies, midguts and carcasses were homogenized in 0.5 ml ice-cold cell lysate
buffer and extracted twice with pentane: ether (1:1) spiked with 250 ng/ml n-octanol
(internal standard). The upper layer (organic phase) was collected and concentrated to
approximately 100 µl with N2 gas, then directly analyzed by GC-MS at the Nevada
Proteomics Center (UNR). A Thermo Finnigan Polaris Q ion trap was used with a molecular weight scanning range of 40–180 atomic mass unit (amu) at an ionization potential of 70 eV. A Trace gas chromatograph containing a 60 m x 0.25 mm (ID), 0.25
µm film thickness Agilent HP – INNOWAX column (P/N 19091N-136, J&W Scientific,
Palo Alto, CA) was programmed for an initial temperature of 50 ºC (1 min hold), increase to 240 ºC at 5 ºC /min (20 min hold). The injector was split at a ratio of 10:1 at a temperature of 225 ºC with a column flow of 1.2 ml He/min. The detector was set at 200
ºC. exo-Brevicomin produced by beetles was determined by comparing GC retention times and MS fragmentation patterns with an authentic standard, and was quantified according to the concentration of internal standard (n-octanol) and the ratio of areas of
GC traces of exo-brevicomin and n-octanol. The total protein in each reaction was quantified by BCA assay (Thermo Scientific, Rockford, IL) using BSA as standard and
according to the supplier’s protocol. exo-Brevicomin levels were calculated as ng exo- 41
brevicomin per mg total protein.
Tissue culture assay. Isolated fat bodies were pooled from unfed males and
cultured with or without 23.3 mM (2 mg) tri-deuterated decanoic acid in 0.5 ml
culture medium in a six-well tissue culture plate at 27 ºC in a moist chamber
overnight. Two mg tri-deuterated decanoic acid in 0.5 ml culture medium without
tissue was used control. The overnight cultures were mixed with 0.5 ml
extraction solvent (CHCl3: MeOH: H2O= 1: 2: 0.8), homogenized, and centrifuged
at 10,000 x g for 2 min at 4 oC in a microcentrifuge to remove debris. The
supernatants were transferred into clean glass tubes. 0.6 ml each CHCl3 and
H2O were added to the supernatants, mixed very well, and let stand to allow
phases to separate. The upper (aqueous) layer was removed, and the lower
(organic) layer was placed in a 60 oC sand bath and dried to near dryness under
N2. One hundred µl methanolic NaOH (0.5N NaOH in methanol) was added and
the samples were incubated at room temperature for 5-10 min to dissolve lipids.
This mixture was transferred into a clean screw cap glass tube. An equal volume
of BF3/MeOH (14% Boron trifluoride in methanol solution) solution was added,
sealed and mixed very well, then heated in a 60 oC sand bath for approximately
one hour. Saturated NaCl (0.5 ml) was added and mixed very well. The mixture
was extracted with pentene twice, and the top layer was transferred into a clean
glass tube, concentrated by N2 gas, and analyzed by GC-MS. The extraction
protocol was according to previous references (Bligh &Dyer, 1959; Doss & Oette,
1965). All steps after the first centrifugation were performed in glass containers.
Twenty beetles were used for each replicate, and in total three replicates were performed. 42
Enzyme assays. Fat bodies were isolated from unfed male MPBs and homogenized as described above. Reactions with unlabeled (6Z)-non-6-en-2- one were conducted in 500 µl volumes containing 468 µl of homogenized fat bodies, 284 µM unlabeled substrate and 300 µM NADPH (final concentrations).
Reactions (n =3) were incubated for 30 min in a 30 ºC water bath. Reactions with penta-deuterium labeled (6Z)-non-6-en-2-one (284 µM final concentration) were performed in 500 µl volumes containing 250 µl of homogenized fat bodies and 250 µl 2X NADPH regeneration system with 6 hours incubation in a 30 ºC water bath (n=1). For reactions in the presence of P450 inhibitor, homogenized fat bodies were incubated with 2.5 mM PBO at room temperature for 30 minutes before performing enzyme assays. Fat bodies without PBO were also incubated at room temperature for 30 minutes. Fat bodies with or without PBO were mixed with the substrate, and reactions were initiated with the addition of NADPH or 2 X
NADPH regeneration system. Experimental conditions included reactions run without the substrate, and the reaction with the substrate in the presence and absence of P450 inhibitor. Assays then were extracted twice with pentane: ether
(1:1) spiked with 5-20 μg/ml n-octanol. The organic phase was concentrated to approximately 100 µl with N2 gas and directly analyzed by GC-MS. For assays
with unlabeled substrate, 12 beetles/sample/replicate were used.
When the epoxide was used as the substrate (n=1), enzyme assays contained
468 µl homogenized fat bodies, 600µM penta-deuterium labeled epoxide, and with or without 300 µM NAD+ or NADP+. The mixtures were incubated in a 30 ºC water bath for two hours. Experimental conditions included fat bodies only and fat bodies with NAD+ or
NADP+ in the presence of epoxide. The reactions were terminated and extracted with
pentane: ether and analyzed by GC-MS as described above. Products were identified 43
by comparing retention times and mass spectra with their controls and an authentic
standard of penta-deuterium labeled epoxide or unlabeled exo-brevicomin. The
percentage conversion of substrate to product was calculated from the areas of GC
traces by dividing the amount of product by the total amount of products and substrates
(product/(products + substrates)).
Epoxide Cyclization. The reaction mixture in pentane: ether from enzyme
assays in which fat bodies were incubated with penta-deuterium labeled (6Z)-
non-6-en-2-one as described above was treated with acetic acid at different pH
values. Briefly, 50 µl of sample in pentane: ether was mixed well with 200 µl
water, followed by incubating separately with either 200 µl glacial acetic acid, or
acetic acid to pH 3.0, 4.0, 5.0,or 6.0 for two hours at room temperature. Each
sample was extracted twice with pentane: ether, and analyzed by GC-MS as
described above. The untreated reaction mixture was used as a control.
IV. Results:
exo-Brevicomin tissue distribution. exo-Brevicomin was detected in unfed
male MPBs, but not in females (Figure 2.2A). The average exo-brevicomin
concentration in male MPBs was 87.6 ± 4.1ng/mg protein, while exo-brevicomin in females was undetectable (Figure 2.2A). Surprisingly, exo-brevicomin was detected in male fat bodies (approximately 668 ± 207 ng/mg protein), but no exo- brevicomin was detected in carcasses or midguts (Figure 2.2B). (6Z)-Non-6-en-
2-one, the exo-brevicomin precursor, was not detected in any beetles or tissues
(not shown).
Production of (6Z)-non-6-en-2-one from C10:1 fatty acid through decarboxylation.
Since exo-brevicomin is specifically produced in the fat body, the terminal steps of exo- 44
brevicomin biosynthesis (i.e. decarboxylation) are probably localized in this tissue.
Decarboxylation is the removal of the terminal carboxylic acid carbon atom from a
carbon chain. Intact unfed male fat bodies cultured with 23.3 mM (2 mg) tri-deuterated
decanoic acid (32.02 min) yielded a product with a GC retention time of 13.20 min
(Figure 2.3B). This product was not detected in extracts of fat bodies cultured without
substrate (Figure 2.3A) or in the tri-deuterated decanoic acid standard (Figure 2.3C).
The 13.2 min peak in extracts of fat bodies cultured without tri-deuterated substrate
(Figure 2.3A) had a different mass spectrum (not shown) from the 13.2 min peak
observed in Figure 2.3B. MS analysis confirmed that the product at 13.2 min in (B) is tri-
deuterated nonan-2-one because its mass spectrum (Figure 2.3D) is similar to unlabeled
nonan-2-one (Figure 2.3E) with the base peaks shifted three amu from m/z=127.0
(Figure 2.3E) to m/z=130.1 (Figure 2.3D) and from 142 (Figure 2.3E) to 145.2 (Figure
2.3D).
Production of 6,7-epoxy-nonan-2-one from (6Z)-non-6-en-2-one. Given that exo- brevicomin localized to the male fat body, and that male MPBs exposed to (6Z)-non-6- en-2-one produced exo-brevicomin (Vanderwel et al., 1992), we hypothesized that the fat body would convert (6Z)-non-6-en-2-one to epoxide or directly to exo-brevicomin, and that a cytochrome P450 would be involved in the reaction. To test this hypothesis, we
assayed the ability of homogenized fat bodies from unfed males to metabolize (6Z)-non-
6-en-2-one in the presence or absence of the P450 inhibitor, PBO. Analysis with the
base peak m/z=46 showed that male fat body homogenates converted penta-deuterium
labeled (6Z)-non-6-en-2-one (18.64 min) to a product at 27.45 min (Figure 2.4B,C) which has a similar retention time and identical MS (Figure 2.4E) as the 6,7-epoxy-nonan-2- one standard (27.47 min) (Figure 2.4D, F). This product was not detectable in fat bodies incubated without substrate (Figure 2.4A) or in the pre-incubated substrate (not shown). 45
In the presence of 2.5 mM PBO, about 11.8% labeled (6Z)-non-6-en-2-one (18.64 min) was converted to the product (27.45 min) (Figure 2.4B). In the absence of PBO, 42.8% labeled substrate (18.64 min) was converted to the product (27.45 min) (Figure 2.4C).
Deuterium-labeled exo-brevicomin, which would have appeared as an peak at approximately 16.63 min (see Figure 2.6C) and eluted before (6Z)-non-6-en-2-one, was not observed in any of these reactions (Figure 2.4B,C). Similarly, assays of fat bodies incubated with unlabeled substrate also produced a barely detectable product (not shown). MS analysis confirmed that products from unlabeled (not shown) and labeled substrates (Figure 2.4 B, C) represented the same product because the mass spectra of diagnostic fragments of the labeled product were three to five amu larger than the unlabeled product (not shown).
Surprisingly, fat bodies incubated with deuterium labeled (6Z)-non-6-en-2- one also produced a product at 22.20 min (Figure 2.5B) which had a similar retention time and identical MS (Figure 2.5E) to a penta-deuterium labeled nonen-2-ol standard (22.16 min) (Figure 2.5C, F). This product was not present in the assay with fat body only (Figure 2.5A) and penta-deuterium labeled (6Z)- non-6-en-2-one standard (Figure 2.5D). The retention time of this product is very close to the internal standard n-octanol (21.98 min) (Figure 2.5B), but their mass spectra are very different (not shown).
Epoxide cyclization. 6,7-Epoxy-nonan-2-one could cyclize into exo-brevicomin in the presence of acid at the room temperature (C. Jeffery, personal
communication). To confirm that the epoxide produced by fat bodies can be
converted to exo-brevicomin, the epoxide was incubated at different pH values.
The analysis with a base peak m/z=85 was used because it is a major fragment
for both the epoxide and exo-brevicomin. Untreated samples (e.g. Figure 2.6A) 46
retained the 27.45 min peak corresponding to 6,7-epoxy-nonan-2-one, while the sample treated with glacial acetic acid yielded a product with a retention time of
16.63 min with the complete disappearance of 27.45 min peak (Figure 2.6B).
The conversion of epoxide to exo-brevicomin did not occur with any other (pH 3.0
- 6.0) acetic acid treatments (not shown). MS analysis showed shifts in base peaks from the product (16.63 min) with m/z=161.1, 117 and 46.1 (Figure 2.6D) to the standard of unlabeled exo-brevicomin (15.37min) (Figure 2.6E) with m/z=155.9, 114 and 43 respectively.
Epoxide to exo-brevicomin in fat bodies. Because of the apparent stability of the 6,7- epoxy-nonan-2-one in physiological conditions (see above) and cyclization requiring a pH below 3, we hypothesized that a cyclase in the fat body catalyzes the cyclization of
6,7-epoxy-nonan-2-one to exo-brevicomin. To confirm this hypothesis, assays were performed with homogenized unfed male fat bodies and penta-deuterium labeled epoxide in the presence or absence of NAD+ or NADP+. Analysis with the base peak at
m/z = 46 showed that the epoxide standard (27.48 min) was contaminated with a
product at 15.21min (Figure 2.7D). The peaks at 15.21min from Figure 2.7D, B and at
15.20 min from Figure 2.7A, C had mass spectra identical (not shown) to the 16.63 min
peak in Figure 2.6B which was identified as penta-deuturium labeled exo-brevicomin
(Figure2.6D, E), and diagnostic fragments from this mass spectra were three to five amu larger (not shown) than those of the unlabeled exo-brevicomin standard (Figure
2.7E,m/z=85, MS not shown). The calculated percentage of contamination was approximately 2.4% (area of the contaminated product at 15.21 min/area of penta- deuturium labeled epoxide at 27.48) (Figure 2.7D). In the absence of NAD+ and NADP+, this percentage (areas of 15.21 peak/27.44 peak) was ~ 2.1% (Figure 2.7A). However, 47
this percentage decreased to approximately 0.85% in the presence of NAD+ (Figure
2.7B), and approximately 0.77% in the presence of NADP+ (Figure 2.7C).
V. Discussion
exo-Brevicomin was identified several decades ago as a male specific MPB aggregation pheromone component (Conn et al., 1983; Libbey et al., 1985; Pureswaran et al., 2000). However, details about exo-brevicomin biosynthesis, including the site of synthesis and enzymes in the biosynthetic pathway remain unknown. exo-Brevicomin is believed to be derived from (6Z)-non-6-en-2-one (Figure 2.1), which is naturally present in the beetles (Pureswaran et al., 2000; Vanderwel et al., 1992). The origin of (6Z)-non-
6-en-2-one is not known though it is speculated to arise from a fatty acid precursor
(Jurenka, 2003; Vanderwel et al., 1992). All published reports on the location of pheromone biosynthesis show that bark beetle pheromones are produced in midgut
(Barkawi et al., 2003; Hall et al., 2002a; Hall et al., 2002b), leading to the reasonable hypothesis that exo-brevicomin is also synthesized there.
V.1. exo-Brevicomin localization
In this study, we surveyed exo-brevicomin levels in unfed male and female
MPBs. exo-Brevicomin was detected in unfed males, not in females (Figure 2.2A), confirming previous reports (Pureswaran et al., 2000). Surprisingly, exo-brevicomin was detected in male fat bodies, but not in carcasses or guts (Figure 2.2B), strongly suggesting that this pheromone component is synthesized in the fat body, and not in the midgut. This is the first report of a bark beetle pheromone produced in fat body, not in midgut.
V.2. Fatty acid conversion to 6(Z)-non-6-en-one 48
Decarboxylation of fatty acids in insects has been proposed (Skiba & Jackson,
1994). Because (6Z)-non-6-en-2-one is a natural precursor of exo-brevicomin in MPB, and exo-brevicomin was found specifically in male fat bodies, it is likely that (6Z)-non-6- en-2-one is produced de novo through the derivatization of ω -3 fatty-acyl compounds via decarboxylation in the fat body (Vanderwel et al., 1992). Incubation of intact fat bodies from unfed males with tri-deuterated decanoic acid yielded tri-deuterated nonan-
2-one (Figure 2.3B), very likely via a decarboxylation reaction. This product was not observed in assays of fat bodies alone (without substrate) (Figure 2.3A) or in the pre- reacted substrate (Figure 2.3C), suggesting an enzyme activity in the fat bodies is responsible for tri-deuterated nonan-2-one production. Given that an unsaturated derivative of the decanoic acid precursor was not observed, there likely is no desaturase activity for short chain fatty acids in the male fat body. These data implicate that 10:1 fatty acid is the direct precursor to (6Z)-non-6-en-2-one, and that 10:1 fatty acid is likely derived from a long chain fatty acid precursor via chain shortening via β-oxidation, but not from fatty acid elongation. Incubation of intact of male fat bodies with labeled long chain fatty acids may confirm this. These data also suggest that male fat body contains decarboxylase activity that may accept both saturated and unsaturated C10 fatty acid precursors.
Surprisingly, homogenized fat bodies converted penta-deuterum labeled
(6Z)-non-6-en-2-one to a product (22.2 min) (Figure 2.5B) which had a similar retention time and identical mass spectra (Figure 2.5E) to penta-deuterum labeled nonen-2-ol standard (Figure 2.5C, F). This product was not detected in the assay with fat bodies only (Figure 2.5A) or in the substrate (Figure 5.5D), suggesting that this alcohol product results from oxidoreductase activity in the fat body. (6Z)-nonen-2-ol production was observed in male and female MPB 49
exposed to (6Z)-non-6-en-2-one vapors (Vanderwel et al., 1992). These data
imply that nonen-2-ol may be the precursor of (6Z)-non-6-en-2-one and a oxidoreductase in the fat bodies may be required for the production of (6Z)-non-
6-en-2-one. If this hypothesis is true, nonen-2-ol may be present in assays of fat bodies incubated with decanoic acid (C10) (Figure 2.3). However, nonen-2-ol was not identified in these assays. There are several possible explanations for this. One is that the large n-octanol peak may overlap that of nonan-2-ol because of their close retention times. It is also possible that the pathway to nonen-2-ol production does not accept saturated substrates. Finally, nonan-2-ol may be quickly converted to nonan-2-one. No natural nonen-2-ol was detected in fat bodies (Figure 2.3A). In the beetle, this intermediate may rapidly be catalyzed by the enzyme system to either the final product or to intermediates that require more energy for the next step (i.e. a bottleneck), such as hydroxylation or epoxidation by a P450.
V.3. 6(Z)-Non-6-en-2-one epoxidation
Homogenized male fat bodies catalyzed the conversion of (6Z)-non-en-
2-one to 6,7epoxy-nonan-2-one, the direct precursor to exo-brevicomin (Figure
2.4). This epoxide was not detected in fat bodies incubated without the substrate. In the presence of piperonyl butoxide (PBO), a non-specific P450 inhibitor, fat bodies produced less epoxide than in the absence of this P450 inhibitor, confirming that a cytochrome P450 in the fat body contributes to the conversion of (6Z)-non-6-en-2-one to epoxide (Vanderwel & Oehlschlager,
1992).
V.4. Epoxide cyclization 50
To rule out unlabeled exo-brevicomin from fat bodies, the base peak m/z=46 was used to ensure measurement of de novo-synthesized exo-
brevicomin from labeled (6Z)-non-6-en-2-one. Interestingly, we did not detect
any penta-deuterum labeled exo-brevicomin from these assays (Figure 2.4B, C).
This result implies that the epoxide does not spontaneously cyclize under
physiological conditions as proposed (Silverstein et al., 1968), and therefore a
cyclase is required for the final step of exo-brevicomin production. It is likely that
homogenization disrupted the complete pathway. It will be interesting to see if
exo-brevicomin will be produced from (6Z)-non-6-en-2-one using intact isolated
fat bodies.
The epoxide can be cyclized to exo-brevicomin in the absence of an
enzyme catalyst in the presence of acid (C. Jeffrey, personal communication).
To confirm the conversion of this produced epoxide to exo-brevicomin, epoxide
was treated with acid at different pH values. Epoxide (Figure 2.6A) treated with
glacial acetic acid was completely cyclized to exo-brevicomin (Figure 2.6B), but
not under milder conditions (pH 3.0-6.0) (not shown). These results further
support the idea that the conversation of epoxide to exo-brevicomin requires a
cyclase in the fat body.
To confirm a cyclase catalyzes the final step of exo-brevicomin
production, unfed homogenized male fat bodies were incubated with labeled
epoxide (Figure 2.7). The conversion of epoxide to exo-brevicomin does not
involve hydride transfer because both epoxide and exo-brevicomin contain the
same components of atoms. It is possible that this final step requires NAD+ or
NADP+ as cofactor during cyclization. However, neither the presence nor the
absence of NAD+ and NADP+ resulted in exo-brevicomin production by this assay 51
(Figure 2.7). The labeled epoxide substrate contains about 2.4% labeled exo- brevicomin (Figure 2.7D). Interestingly, the percentage of contaminated exo- brevicomin in the epoxide decreased to half the starting value in the presence of
NAD+ or NADP+ (Figure 2.7B, C), while in the same assay condition, this percentage of exo-brevicomin did not change in the absence of NAD+ and
NADP+ (Figure 2.7A) although only one replicate was performed and this decrease was not significant. These results suggest that NADH or NADPH or other ions may be required as coenzymes for activation of the cyclase. Fat bodies incubated with epoxide in the presence of NADH, NADPH or other ions, such as Mg2+ would confirm this hypothesis.
V.5. Summary
The data presented here clarify many aspects of the exo-brevicomin biosynthetic pathway and also raise new questions (Figure 2.8). They confirm previous suggestions that exo-brevicomin is derived from a fatty acyl-precursor (Vanderwel et al., 1992).
Biosynthesis occurs in unfed male fat bodies. A long chain ω-3-fatty acyl-CoA is likely chain-shortened to decenyl-CoA by standard β-oxidation, at which point the precursor exits β-oxidation and is decarboxylated. The decarboxylation mechanism may involve production of free β-keto decenoic acid from a β-oxidation cycle intermediate as proposed by Skiba and Jackson (1994) or proceed via the more canonical hydrocarbon- biosynthetic oxidative decarbonylase pathway (Reed et al., 1994) to (6Z)-nonene, followed by hydroxylation to (6Z)-nonen-2-ol by a cytochrome P450 (G. Blomquist, personal communication). The fact that fat body incubations with (6Z)-nonen-2-one yielded 6(Z)-nonen-2-ol (Figure 2.5) suggests an oxidoreductase activity in fat bodies carries out this reaction, which may proceed in the opposite direction under normal conditions. This reaction would support the oxidative decarbonylation/P450 52
hydroxylation pathway over decarboxylation. (6Z)-Nonen-2-one requires P450 activity to form the keto-epoxide, which appears stable under physiological conditions. Thus, a
cyclase is likely required for the last step of exo-brevicomin production.
None of the enzymes catalyzing the reactions above have been described. The
information presented here is useful to guide searches of functional genomic data in
order to isolate and characterize these enzymes and thereby confirm the proposed
pathway. Work towards this goal is described in the following two Chapters.
VI. Acknowledgments
I thank X. Liu, T. Nguyen and C. Jeffery (Department of Chemistry, UNR) for
providing penta-deuterium labeled (6Z)-non-6-en-2-one and epoxide, and C.
Jeffrey for helpful advice; D. Quilici and R. Woolsey at the Nevada Proteomics
center for GC/MS analysis; members of GJB and CT’s laboratories, especially A.
Gorzalski, P. Delaplain for assistance with collecting beetles, dissections, assays,
S. Young and R. Figueroa-Teran for helpful advice; and the Bureau Of Land
Management (BLM), US forest Service, South Tahoe District, and the Whittell
Board which oversees the UNR little valley forest for permission to collect beetle-
infested trees. This research was funded by USDA-NIFA grant #2009-05200.
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VIII. Figure Legends
Figure 2.1. Proposed exo-brevicomin biosynthetic pathway in MPB. 10:0, 16:0 and 18:0 acids represent saturated fatty acids; 10:1, 16:1 and 18:1 represent unsaturated fatty acids. P450, cytochrome P450. Enzymes catalyzing the proposed reactions have not been identified, and uncharacterized steps are shown by dashed arrows.
Figure 2.2. exo-Brevicomin distribution. (A), exo-brevicomin in unfed males and females. (B), exo-brevicomin distribution in fat bodies, carcasses and midguts of unfed males. Male MPBs starved overnight were used. The amount of exo- brevicomin was calculated according to the concentration of internal standard (n- octanol) and the ratio of areas of GC traces of exo-brevicomin and n-octanol.
The total protein from each reaction was determined by BCA assay. Values are averages ± standard error
Figure 2.3. GC-MS analysis with mass/charge (m/z= 43) of intact male fat bodies incubated with tri-deuterated decanoic acid. Culture assays: (A), fat bodies only; (B), fat bodies incubated with tri-deuterated decanoic acid; and (C), tri-deuterated decanoic acid 59
only. n-Octanol; 17.58 min in (A), 17.65 min in (B) and 18.16 min in (C). (D, E), Mass
spectra of the peak from 13.2 min in (B) and nonan-2-one from (NIST#: 114362 ID#:
7482, http://www.nist.gov/index.html), respectively. The substrate, tri-deuterated
decanoic acid, is indicated by an arrow (B, C), and the product by an arrowhead (B).
Figure 2.4 GC-MS analysis (m/z=46) of homogenized unfed male fat bodies incubated with penta-deuterium labeled (6Z)-non-6-en-2-one by (n=1). 30 beetles
were used in total. Reactions containing 2X NADPH regeneration system and:
(A), fat bodies only; (B), fat bodies with labeled (6Z)-non-6-en-2-one in the
presence of P450 inhibitor, piperonyl butoxide; (C), fat bodies with labeled (6Z)-
non-6-en-2-one in the absence of P450 inhibitor. (D), penta-deuterum labeled epoxide standard (27.47 min). The substrate, penta deuterium labeled (6Z)-non-
6-en-2-one (B, C) in arrow, products (B, C) in arrowhead. (E, F), mass spectra
from 27.45 min peak in (C) and 27.47 min peak in (D), respectively.
Figure 2.5 GC-MS analysis (m/z=67) of homogenized unfed male fat bodies
incubated with penta-deuterium labeled (6Z)-non-6-en-2-one (n=1). Thirty
beetles were used in total. Reactions containing 2X NADPH regeneration
system and: (A), fat bodies only; (B), fat bodies with labeled (6Z)-non-6-en-2-one.
(C), penta-deuterum labeled nonen-2-ol standard (22.16 min). (D), penta- deuterium labeled (6Z)-non-6-en-2-one (18.75 min). Penta-deuterium labeled
(6Z)-non-6-en-2-one in thick arrow, the product and labeled nonen-2-ol in arrowhead, and n-octanol in thin arrow. (E, F), mass spectra from 22.2 min peak in (B) and 22.16 min peak in (C), respectively.
60
Figure 2.6. GC-MS analysis of exo-brevicomin yielded from epoxide in the presence of glacial acetic acid (n=1). Analysis of base peak (m/z = 85) with reactions: (A), the sample from the mixture of Figure 2. 4 (B) and (C); (B), the sample from (A) treated with pure acetic acid; (C), unlabeled exo-brevicomin
(15.37min), penta-deuterum labeled (6Z)-non-6-en-2-one (18.78 min) standards. n-Octanol; 21.94 min in (A), 22.07 min in (B) and 21.19 min in (C). Penta- deuterum labeled (6Z)-non-6-en-2-one: 18.63 min in (A), 18.71 min in (B) and
18.78 min in (C). Penta-deuterum labeled epoxide at 27.45 min in arrow.
Product 16.63 min from epoxide in arrowhead. (D, E) Mass spectrum from 16.63
min peak in (B) and 15.37 min in (C) respectively.
Figure 2.7 GC-MS analysis (m/z=46) of homogenized male fat bodies incubated
with 6,7-epoxynonan-2-one (n=1). 25 beetles were used in total. Reactions
were conducted with fat bodies incubated with (A) deuterium labeled penta-
epoxide only, (B) penta-deuterium labeled epoxide in the presence of NAD+, or
(C) penta-deuterium labeled epoxide in the presence of NADP+. (D), penta- deuterium labeled epoxide standard. (E), unlabeled exo-brevicomin standard
(15.47min; m/z=85). Penta-deuterium labeled epoxide in thick arrow, contaminated penta-deuterium labeled exo-brevicomin in thin arrow.
Figure 2.8. Proposed revisions to the exo-brevicomin biosynthetic pathway.
16:0 and 18:0 acids represent saturated fatty acids; 10:1, 16:1 and 18:1
represent unsaturated fatty acids. P450, cytochrome P450. Unconfirmed
reactions are indicated by question marked enzymes. Enzymes catalyzing the 61
proposed reactions have not been identified, and uncharacterized steps are shown by dashed arrows.
62
Fatty Acid Pathway
O -O acetate
Fatty Acid Biosynthetic Enzymes 16:0 /18:0 acid Desaturase Thioesterase II 16:1 / 18:1 acid
β-oxidation 10:0/10:1 acid Desaturase Dehydrogenase? Decarboxylation?
O
(6Z)-non-6-en-2-one
P450?
O O
Cyclase?
O O exo-Brevicomin
Figure 2.1 63
100 (A)
s.e. 90
± 80 70 60 50 40 30 brevicomin /mg /mg protein brevicomin - 20 exo
10
ng 0 unfed male unfed female
1000 (B) s.e. 900 ± 800 700 600 500 400 300 brevicomin /mg /mg protein brevicomin - 200 exo 100 ng 0 carcass fat body midgut
Figure 2.2 64
(A) Fat body only
6 17.58 5 4 3 2 1 13.2
(B) Fat body & C10:0 fat acid
6 32.02 5 17.65 4 3 13.2 2 1 Relative Relative Abundance (C) C10:0 fat acid
20 18.16 32.0 15
10
5
12 14 16 18 20 22 24 26 28 30 32 34 Time (min)
(D) 13.2 in B 10 0 43.1 8 0 58.1 6 0
4 0 71.1 85.1 2 0 102.1 113.2 130.1 145.2 0
40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 (E) Nonan-2-one Relative Relative Abundance
m/z
Figure 2.3 65
100 (A) FB only 21.98
0 (B) FB & P450 inhibitor & labeled (6Z)-non-6-en-2-one 100 18.64 22.19 27.45
0 (C) FB & labeled (6Z)-non-6-en-2-one 100 22.19 18.64 27.45 Relative Abundance Relative
0 100 (D) Std of epoxide 27.47
0 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 Time (min)
(E) 27.45 in C
100 46.1
57.2 85.1 102.2 69.2 117.1 0 (F) 27.47 in D 100 46.1 Relative Abundance Relative
57.2 85.1 102.1 69.2 117.1 0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 m/z
Figure 2.4 66
21.98 100 (A) FB only
0 21.98 100 (B) FB & labeled (6Z)-non-6-en-2-one 22.20 18.65
0 22.16 100 (C) Labeled nonen-2-ol
Abundance Relative 21.95 0 18.75 100 (D) labeled (6Z)- non-6-en-2-one
0 16.0 17.0 18.0 19.0 20.0 21.0 22.0 23.0 24.0 Time (min) (E) 22.2 in B 100 67.1
82.2 46.1 55.2 100.2 111.1 129.1 147.2 0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 (F) 22.16 in C 100 67.0 Abundance Relative
46.2 55.0 82.0 100.0 110.9 128.9 0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 m/z
Figure 2.5 67
12 (A) untreated 21.94 14.94 18.63 27.45
0 (B) Treated with acetic acid 12 18.71 22.07
16.63
0 Relative Abundance Relative (C) Std of exo-brevicomin 12 15.37 18.78
22.19
0 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 Time (min)
(D) 16.63 in B 117.0 100 46.1
86.2
68.2 57.2 102.2 131.1 142.2 161.1 0 (E) 15.37 in C 100 43.0
Relative Abundance Relative 85.0 114.0
67.0 57.1 99.1 127.0 138.0 155.9 0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180
Figure 2.6 68
exo-Brevicomin Epoxide
(A) FB & epoxide 27.44 5 15.20
0 (B) FB & epoxide & NAD+ 27.44 5 15.21
0 (C) FB & epoxide & NADP+ 27.43 5 15.20 0 (D) Std of epoxide 27.48 Relative Relative Abundance 5 15.21
0 (E) Std of exo-brevicomin 5 15.47
0 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 Time (min)
Figure 2.7 69
O -O acetate
Fatty Acid Biosynthetic Enzymes 16:0 /18:0 acid Desaturase Thioesterase II 16:1 / 18:1 acid
β-oxidation 10:1 acyl-CoA
β-Oxidation Fatty acyl-CoA reductase?
10:1 β-keto-acyl-CoA 10:1 aldehyde Oxidative decarbonylase? Thiolase? nonene 10:1 β-keto-acid hydroxylation (P450)? nonen-2-ol Decarboxylation? Dehydrogenase? O
(6Z)-non-6-en-2-one
P450
O O
Cyclase NAD+/NADP+? NADH/NADPH?
O O exo-Brevicomin Figure 2.8 70
Chapter 3
CYP6CR1: a mountain pine beetle cytochrome P450 involved in exo-brevicomin
biosynthesis
I. Abstract II. Introduction III. Materials and Methods IV. Results V. Discussion VI. Acknowledgements VII. References VIII. Figure Legends
I. Abstract
Investigating pheromone production in mountain pine beetle (MPB;
Dendroctonus ponderosae) may identify unique molecular targets and provide information for developing specific new strategies to deal with their infestations. exo-
Brevicomin is a male-specific aggregation pheromone component of the MPB. It is specifically produced in male fat body. Male fat bodies convert (6Z)-non-6-en-2-one into the epoxide precursor of exo-brevicomin, likely via a cytochrome P450 (Chapter 2).
Microarray analysis revealed a novel cytochrome P450, contig608 (CYP6CR1), with an expression pattern consistent with exo-brevicomin production. In this chapter, quantitative real-time PCR (qRT-PCR) analysis showed that CYP6CR1 was highly expressed in the fat body of unfed males, consistent with GC-MS analyses showing that unfed male fat bodies specifically produce exo-brevicomin. This strongly implicates
CYP6CR1 in exo-brevicomin biosynthesis. To confirm this, we produced functional recombinant CYP6CR1 in Sf9 (insect) cells using a baculovirus expression system.
Enzyme assays with non-labeled and penta-deuterium labeled (6Z)-non-6-en-2-one 71
confirmed that CYP6CR1 epoxidized (6Z)-non-6-en-2-one into the direct 6,7- epoxynonan-2-one precursor of exo-brevicomin. This functional recombinant CYP6CR1
did not accept (6E)-non-6-en-2-one, 6-methylhept-6-en-2-one, or monoterpenes as substrates, indicating high selectivity for (6Z)-non-6-en-2-one as substrate, and that
CYP6CR1 is specific for exo-brevicomin biosynthesis. This is the first identified enzyme in MPB pheromone biosynthesis.
II. Introduction:
The mountain pine beetle (MPB; Dendroctonus ponderosae) is the most destructive pest of coniferous forests in Alaska, Canada, and the western US.
Investigating pheromone production in MPB may identify unique molecular targets and provide information for developing specific new strategies to deal with their infestations. exo-Brevicomin is a male-specific aggregation pheromone component. Its biosynthetic pathway is not fully understood. Emerged males, but not females, contain detectable exo-brevicomin, (Borden et al., 1987; Pierce et al., 1987; Vanderwel et al., 1992) and
feeding decreases exo-brevicomin levels (Pureswaran et al., 2000). (6Z)-Non-6-en-2-
one is believed to be a natural exo-brevicomin precursor in MPB and western balsam
bark beetles (WBB; Dryocoetes confusus) (Vanderwel et al., 1992). Male MPBs
exposed to (6Z)-non-6-en-2-one produce more exo-brevicomin than females (Vanderwel et al., 1992). (6Z)-Non-6-en-2-one is most likely derived from endogenous 10 carbon fatty-acyl compounds that contain a double bond in the ω -3 position (Chapter 2). This
10-carbon fatty acyl intermediate is likely derived from a longer-chain fatty acyl precursor
through chain shortening (β-oxidation) as has been proposed in pheromone biosynthesis
in Lepidoptera (Rafaeli & Jurenka, 2003). exo-Brevicomin biosynthesis likely involves
the formation of 6, 7-epoxynonan-2-one, which is produced by cytochrome P450 72
mediated epoxidation of (6Z)-non-6-en-2-one (Francke et al., 1996; Vanderwel et al.,
1992; Vanderwel & Oehlschlager, 1992). Once formed, 6, 7-epoxynonan-2-one is likely
cyclized to exo-brevicomin, though cyclization may occur in the presence of acid without
an enzyme, or it can be induced thermally under neutral conditions (Silverstein et al.,
1968).
Our previous study showed a clear pathway of exo-brevicomin biosynthesis:
6(Z)-decenoic acid or its derivative undergoes decarboxylation and oxidation steps, resulting eventually in (6Z)-non-6-en-2-one; the conversion of (6Z)-non-6-en-2-one to epoxide requires a cytochrome P450; and a cyclase is likely required for the final step
(Chapter 2). However, none of the enzymes involved have been identified.
In a preliminary effort to identify pheromone-biosynthetic enzymes, microarray analysis indicated that the cytochrome P450 represented by Contig608 (GenBank I.D.
EZ115564, later named “CYP6CR1”) may be involved in exo-brevicomin biosynthesis because its expression pattern (Aw et al., 2010) consistent with exo-brevicomin production (Chapter 2). Functional genomics have been successfully applied to identify pheromonal ipsdienol biosynthetic enzymes in I. pini (Figueroa-Teran et al., 2011;
Keeling et al., 2006; Keeling et al., 2004; Sandstrom et al., 2006). This method offers great advantage to identify and assign P450 functions by combining expression analysis and functional assays of expressed recombinant proteins.
We hypothesized that CYP6CR1 catalyzes the epoxidation of (6Z)-non-6-en-2- one as the penultimate biosynthetic step for exo-brevicomin (Figure 3.1), and that the epoxide product is then likely cyclized in the presence of cofactor such as NADH or
NADPH by a cyclase (Figure 3.1, Chapter 2). Since male MPBs also produce endo- brevicomin from its precursor, (6E)-non-6-ene-2-one (Vanderwel et al., 1992) with the same pattern as exo-brevicomin production (Pureswaran et al., 2000), it is possible that 73
CYP6CR1 also converts (6E)-non-6-ene-2-one to the epoxide precursor of endo- brevicomin. Frontalin, a MPB antiaggregation pheromone, is often grouped with exo- brevicomin because of the similarity of their and their precursors’ structures (Symonds &
Elgar, 2004; Symonds & Wertheim, 2005), even though their precursors are derived from different pathways and in different tissues (Barkawi et al., 2003; Hall et al., 2002;
Perez et al., 1996; Vanderwel et al., 1992). The two chemicals may be metabolically linked and may be epoxidized by the same P450. To confirm these hypotheses,
functional recombinant CYP6CR1 was produced in Sf9 cells using a baculovirus
expression system. Enzyme assays were performed with possible substrates.
Recombinant CYP6CR1 epoxidized (6Z)-non-6-en-2-one to form 6,7-epoxynonan-2-one, the direct epoxide precursor of exo-brevicomin, but exo-brevicomin was not produced.
This confirmed that the final step of exo-brevicomin production likely requires a cyclase
(Chapter 2). CYP6CR1 did not accept (6E)-non-6-en-2-one, 6-methylhept-6-en- 2-one, or monoterpenes as substrates, suggesting high selectivity for (6Z)-non-6-en-2-one and specific for exo-brevicomin biosynthesis. This is the first identification of a pheromone- biosynthetic enzyme in D. ponderosae.
III. Materials and methods
Reagents and chemicals. Hink’s TNM-FH Medium 1x (Supplemented Grace’s
Medium) and Grace’s Insect Basal Medium 1x were from Mediatech, Inc.
(Herndon, VA) and FBS was from Atlas Biologicals (Fort Collins, CO). Direct baculovirus DNA kit, SF-900 II (1X) and (1.3X) media were from Invitrogen
(Carlsbad, CA). The Sf9 cells were a gift from B. Perrino (Department of
Physiology, UNR). The housefly cytochrome P450 reductase (CPR) viral clone
(Wen et al., 2003) was kindly provided by M. Schuler (U. Illinois at Urbana- 74
Champaign). Unlabeled (6Z)-non-6-en-2-one, (6E)-non-6-en-2-one, exo- brevicomin and frontalin were purchased from PheroTech Inc. (Delta, BC,
Canada). Penta-deuterium labeled (6Z)-non-6-en-2-one and epoxide were synthesized by C. Jeffery (Department of Chemistry, (UNR)) and penta- deuterium labeled 6-methylhept-6-en- 2-one was synthesized by J. Dickschat
(Technical University, Germany). Agrose was from Bio-Express (Keysville, UT).
Phenylmethylsulfonyl fluoride (PMSF), (+)- / (-)-α-pinene, Δ-3-carene, α-
phellandrene, β-pinene, γ-terpinene, protease inhibitor cocktail, hemin, δ- aminolevulinic acid and ferric citrate were from Sigma- Aldrich (St. Louis, MO).
96-well microplates were purchased from Greiner Bio-One (Monroe, North
Carolina). 2X NADPH regeneration system was from AAT Bioquest (Sunnyvale,
CA). All oligonucleotides were from Integrated DNA Technologies (Coralville,
IA).
Beetles. Mountain pine beetles were obtained from ponderosa pine bolts collected from
the UNR little valley forest (the Sierra Nevada in Nevada, USA). The bolts were stored
in a greenhouse and emerged adults were collected daily and stored for up to 1-2 weeks
at 4 ºC in moist paper towels in loosely capped jars as described previously (Browne,
1972). Emerged beetles were separated by sex (Wood, 1982). Healthy beetles were
selected for experiments. The different groups of the beetles were dissected on the
same day. “Unfed” beetles were held in plastic cups in the dark at room temperature
with moist paper towels as previously described (Keeling et al., 2004).
Sequence analysis. Contig608 is represented by two ESTs: DPG017M13 and
DPG015M04, of 651 bp and 685 bp, respectively. DPG017M13 was selected for further
sequencing of purified plasmid template DNA (Qiagen, QIAprep Spin Miniprep Kit,
Valencia, CA) using the ABI BigDye Terminator Cycle Sequencing Ready Reaction Kit 75 v3.1 and sequence-specific primers as outlined in Table 3.1. The reactions were run on an ABI3730 DNA Analyzer at the Nevada Genomics Center (NGC) at UNR and the sequences were analyzed using Vector NTI Advance 9 software (Invitrogen).
Expression analysis. CYP6CR1 mRNA levels were determined by quantitative Real-
Time reverse transcriptase PCR (qRT-PCR). Template cDNAs from 11 biological groups were prepared previously (Aw et al., 2010). For the tissue distribution analysis, males were kept unfed overnight at room temperature in the dark and then dissected in
100 mM sodium phosphate buffer pH 7.8 under a stereo-microscope. Tissues from the head (including the prothorax), anterior midgut, posterior midgut, hindgut, fat body, and carcass of ten beetles were pooled and frozen in liquid N2 as one replicate. Four biological replicates were made for each sample. The tissues were kept at -80 ºC until
RNA was extracted using the RNeasy Plant Mini Kit (Qiagen). Twenty percent of the isolated RNA was reverse-transcribed using Superscript III reverse transcriptase and random primers (Invitrogen) according to the manufacturer’s protocol.
Primers for qRT-PCR were designed using Primer Express v 2.0 software
(Applied Biosystems, Foster City, CA) and analyzed with vector NTI Advance 9 software.
Primers with minimal potential for primer-dimer and hairpin loop formation were selected
(Table 3.1) and tested for non-specific amplification by visual inspection of melting curves. Their amplification efficiencies were determined using a relative standard curve method. Each 10 μl PCR reaction contained 1.5 μM each forward and reverse primers,
3 μl of appropriately diluted cDNA template, and 5 μl SYBR Green PCR Master Mix
(Applied Biosystems). Amplifications were performed in triplicate, for a total of four replicates per sample, at the NGC. Relative expression values for all genes were determined using the ΔΔCT method normalized to tubulin and Ubiquitin, which are stably 76
expressed in MPB (Aw et al., 2010). Reactions included no template controls to monitor
for genomic DNA contamination.
Expression cloning. The CYP6CR1 open reading frame (ORF) was amplified by PCR,
directionally cloned into the BamH I and Xho I sites of pENTR4 vector (Invitrogen)
modified to remove the Nco I site in the poly-linker (Sandstrom et al., 2006) and
transformed into DH5α cells using standard methods. The forward primer, CYP6CS1F1,
was used with two different reverse primers, CYP6CR1R1 and CYP6CR1R3 (Table 3.1),
in separate reactions to create separate constructs with or without a C-terminal
extension containing a V5 epitope and the poly-histidine tag encoded by the vector,
respectively. Each 100 µl PCR reaction contained 0.5 µM of forward and reverse
primer, 1x Pfu buffer (10 mM (NH4)2SO4, 20 mM Tris–HCl (pH 8.8), 2 mM MgSO4, 10 mM KCl, 0.1% Triton X-100 and 1 mg/ ml BSA), 0.5 mM dNTP, 5–10 ng of DPG017M13 plasmid template, and 1 U Pfu Turbo DNA Polymerase (Stratagene, La Jolla, CA). The profile for linker-ramp PCR was: 95 ºC for 1 min, two cycles of 94 ºC for 40 s, 37 ºC for 1 min, 0.3 ºC/s ramp to 72 ºC, and 72 ºC for 2.5 min, followed by 34 cycles of 94 ºC for 40 s, 60 ºC for 1 min, 72 ºC for 2.5 min, and a final extension of 72 ºC for 10 min. The recombinant plasmids, pENTR4(-Nco I)-6CR1 (untagged) and pENTR4(-Nco I)-6CR1his
(tagged) were confirmed by sequencing.
Recombinant protein production. Protocols for growth and maintenance of Sf9 cells,
recombinant baculovirus construction, and heterologous expression using the
BaculoDirectTM Expression Kit were as described by Invitrogen. Briefly, an LR
recombination reaction between each pENTR4 recombinant clone and BaculoDirect
linear DNA produced untagged and tagged recombinant baculoviral CYP6CR1 clones
that were transfected separately into Sf9 cells and grown in the presence of 100 µM
ganciclovir to select for recombinant virus. High titer P3 viral stocks for each construct 77 were produced by successive 72-hour amplifications of the initial and P2 stocks.
Approximate viral titers were determined by a plaque assay as described by Invitrogen.
The viral stocks were used to infect Sf9 cells at 1.0 x 106 cells/ml in a disposable shaking flask (Bio-Express, Keysville, UT). Hemin (5 µg/ml final concentration) or the heme precursor, δ-aminolevulinic acid (0.3 mM final conc.), and ferric citrate (0.2 mM final conc.) were added at the time of infection.
To produce recombinant CYP6CR1, CPR, or both together, Sf9 cells were infected with recombinant CYP6CR1 baculovirus and/or CPR baculovirus (Wen et al.,
2003) at multiplicities of infection (MOIs) ranging from 1 to 20 pfu/cell for tagged
CYP6CR1, 1 to 100 pfu/cell for untagged CYP6CR1, and 0.05 to 5 pfu/cell for CPR.
Various MOI ratios for recombinant CYP6CR1: CPR from 10:1 to 1:2 were also examined. Sf9 cells were cultured at 27ºC with shaking until harvest. Cells infected with epitope-tagged CYP6CR1 were harvested at day 1, 2, 3 and 4 post-infection (PI) for western blotting.
Microsomal preparation. Sf9 cells producing recombinant CYP6CR1 and CPR were harvested at day 3 PI, and microsomes were prepared by differential centrifugation essentially as per Wen et al. (2003). Briefly, cells were pelleted by centrifugation at
3000 x g at 4 ºC for 10 min. The pellets were resuspended in 1/5 cell culture volume of
100 mM ice-cold sodium phosphate buffer (pH 7.8) and repelleted twice at 3000 x g for
10 min. The pellets were resuspended in 1/30 cell culture volume of ice-cold cell lysate buffer (100 mM sodium phosphate pH 7.8, 1.1 mM EDTA, 0.1 mM DTT, 0.5 mM PMSF,
1/1000 vol/vol Sigma protease inhibitor cocktail, 20% glycerol) and lysed by sonication three times for 15 s on ice with a Branson Sonifier 450, followed by vortexing for 15 seconds. The lysate was centrifuged at 10,000 x g for 20 min at 4ºC in a micro- centrifuge and the supernatant was either further centrifuged in a TLA110 rotor at 78
120,000 x g for 2 h in a Beckman-Coulter Optima ultracentrifuge to pellet microsomes or used directly for CO spectral assay. The microsomal pellet was resuspended in 1/30 cell culture volume of ice-cold cell lysate buffer cold cell lysate buffer and used immediately or flash-frozen in N2 and stored at -80 ºC.
Recombinant protein detection. Cells infected with epitope-tagged CYP6CR1 were harvested at day 1, 2, 3 and 4 PI and sonicated as described above. Non-infected Sf9 cells were prepared similarly. The total protein concentration in each microsomal preparation was quantified by BCA assay kit (Thermo Scientific, Rockford, IL) using bovine serum albumin as a standard. Equal amounts of total protein were boiled for 5 min in an equal volume of Laemli loading buffer (Bio-Rad, Hercules, CA) and loaded into wells of a 10% SDS acrylamide gel, separated by electrophoresis, and then transferred to nitrocellulose membrane by electrophoresis. Protein production was determined by western blotting using 1:5000 mouse anti-V5 primary antibody (Invitrogen), 1:2,000 goat anti-mouse lgG secondary antibody (Biorad, Hercules, CA), and SuperSignal West Pico
Chemiluminescent Substrate (Pierce, Rockford, IL).
CO microplate assay. Functional P450 concentrations from recombinant CYP6CR1,
CPR, and Sf9 only microsomes were determined by carbon monoxide (CO)-difference spectrum analysis (Omura &Sato, 1964) using a 96-well microplate and SpectraMax M5
Microplate Reader coupled with SoftMax® Pro software (Molecular Devices, Inc.,
Sunnyvale, CA) essentially as per Choi, et al. (2003). Briefly, 200 µl of microsome solution from reach sample was loaded into replicate wells for reference and CO treatment samples. The reference wells were tightly sealed with regular paper tape, and the plate was placed in a plastic chamber. CO gas was perfused into the top of the chamber and out from the bottom at 0.5 l/min for 3 min. All of samples were reduced by adding 10 µl fresh 0.5 M sodium hydrosulfite to 25 mM final concentration (Choi et al., 79
2003). The absorbances of the samples from 400 nm to 500 nm were measured with
SpectraMax M5 Microplate Reader. The P450 concentration was calculated using the
following formula:
[P450] (mM) =1.9* (∆450-∆490)/91, (Equation 3.1)
where ∆450 and ∆490 are the absorbance differences between the CO sample and
reference sample at 450 nm and 490 nm respectively (Omura & Sato, 1964), and 1.9
represents the molar extinction coefficient based on the depth of 200 µl with a 6.96 mm
diameter well relative to 1cm light pathway.
Enzyme assays. In vitro assays were conducted in 500 µl reactions containing 468 µl
or 250 µl of a 4:1 mixture of microsomes bearing recombinant CYP6CR1: CPR, either of
284 µM unlabeled (6Z)-non-6-en-2-one, 600 µM penta-deuterium labeled (6Z)-non-6-en-
2-one, 200-300 µM monoterpenes or 183 µM penta-deuterium labeled 6-methylhept-6- en- 2-one in pentane, and 300 µM NADPH or 250 µl 2X NADPH regenerating system.
Reactions were initiated with the addition of NADPH or NADPH regenerating system, incubated in a 30 ºC water bath for 30 min, 2 hours or 6.5 hours, and then extracted twice with pentane:ether (1:1) spiked with 5-100 µg/ml n-octanol (internal standard). The organic phase was concentrated to approximately 100 µl with N2 gas and directly
analyzed by coupled GC-MS at the Nevada Proteomics Center (UNR). Negative
controls included reactions run with the recombinant CPR alone, recombinant CYP9T2,
recombinant CYP6DH2 (See Chapter 5) or uninfected Sf9 cells. A Thermo Finnigan
Polaris Q ion trap was used with a molecular weight scanning range of 40–180 atomic
mass unit (amu) at an ionization potential of 70 eV. A trace gas chromatograph
containing a 60 m x 0.25 mm (ID), 0.25 µm film thickness DB-5 capillary column (J&W 80
Scientific, Palo Alto, CA) was programmed for an initial temperature of 50 ºC (1 min hold), increase to 200 ºC at 5 ºC/min, 10 ºC/min to 320 ºC (20 min hold). The injector was split at a ratio of 100:1 at a temperature of 280 ºC with a column flow of 1.5 ml
He/min. The detector was set at 200 ºC. The product was identified by comparing retention times and mass spectra with negative controls and an authentic standard. The percentage of substrate conversion was calculated according to areas of product and substrate by dividing the amount of product by the total amount of products and substrates (product/(products + substrates)).
IV. Results
Sequencing. The full-length cDNA from clone# DPG017F13 was completely sequenced and designated CYP6CR1 by the P450 Nomenclature Committee
(http://drnelson.utmem.edu/ CytochromeP450.html). The 1683 bp cDNA contained a
1539 nucleotide (nt) open reading frame (ORF) encoding a 513 amino acid (a.a.) protein flanked by 129 nt and 12 nt 5’ and 3’ UTRs, respectively (Figure 3.2). The predicted translation product has a molecular mass of 59 kDa and an isoelectric point of 8.8
(Gasteiger et al., 2003). The primary structure of the predicted translation product contains an N-terminal trans-membrane domain and typical P450 conserved motifs, including the WxxxR, ExxR, and PxxFxPERF (PERF) motifs, and the canonical heme- binding domain (PFxxGxRxCxG) surrounding the heme-cysteine ligand (Cys457) (Figure
3.2)(Sandstrom et al., 2006). BLAST searches (Altschul et al., 1997) indicated that
CYP6CR1 was most similar to Tribolium castaneum CYP6 family P450s, including 41% a.a. to CYP6BQ7 and 40% to CYP6BQ5 (not shown).
CYP6CR1 expression. CYP6CR1 mRNA levels were highest in unfed males; feeding decreased the level of CPY6CR1 mRNA to approximately one of third that of unfed 81
males (Figure 3.3A). In contrast, mRNA levels in larvae, pupae, pre-emerged and
mature females were nearly 100-fold lower and barely detectable (Figure 3.3A).
CYP6CR1 mRNA was distributed in all tissues examined from unfed males, but
predominately localized to the fat body and head (Figure 3.3B).
Functional expression. A baculoviral system was used to produce recombinant
CYP6CR1 for functional analysis. The V5-epitope tagged version of CYP6CR1 was
used to confirm recombinant protein production because an antibody for CYP6CR1 is
unavailable. Western blot analysis indicated an approximately 59 kDa tagged protein
was most abundant at 3 days PI with MOI of 5 and 10, but no recombinant protein was
detected at 1 day PI and in non-infected Sf9 cells (not shown). CO-difference spectra of
microsomes or cell lysate supernatants from Sf9 cells producing recombinant (untagged)
CYP6CR1 harvested at various days PI and various MOI produced a good characteristic
450 nm peak at day 3 PI with a MOI of 10 (Figure 3.4), while microsomes from
uninfected Sf9 cells or from Sf9 cells producing recombinant CPR or the epitope-tagged
version of CYP6CR1 did not produce a 450 nm peak (not shown). Microsomes from Sf9
cells co-expressing recombinant CYP6CR1 and housefly CPR at various ratios did not
produce the characteristic CO-difference peak at a 450 nm (not shown). More functional
CYP6CR1 was produced with heme precursors than with hemin (not shown).
Enzyme Assays. Because co-expression experiments with recombinant (untagged)
CYP6CR1 and housefly CPR did not yield the characteristic CO difference peak at 450
nm, we expressed each protein separately and combined the microsomes for enzyme
assays. All assays were performed with a 4:1 mixture of microsomes bearing
recombinant CYP6CR1 and CPR, with final concentration of 1.5 ~ 3.1 10-4 mM functional
P450 (Table 3.2). Over six replicates, recombinant CYP6CR1 reliably converted (6Z)- non-6-en-2-one to 6,7-epoxynonan-2-one (Table 3.2). GC-MS analysis showed that 82
recombinant CYP6CR1 incubated with penta-deuterium labeled (6Z)-non-6-en-2-one
(18.68 min) yielded a product at 27.47 min (Figure 3.5B) with a retention time and mass spectrum identical (Figure 3.6B) to penta-deuterium labeled epoxide standard (Figure
3.5C, 3.6C). The reaction incubated with unlabeled (6Z)-non-6-en-2-one yielded a product at 27.62 min (Figure 3.5A) with diagnostic MS peaks (Figure 3.6A) 3 to 5 amu smaller than those from the penta-deuterium labeled epoxide standard (Figure 3.6C).
The product was not detected in reactions run with microsomes from uninfected Sf9 cells incubated with unlabeled (6Z)-non-6-en-2-one (Figure 3.5D), or microsomes from Sf9 cells producing housefly CPR incubated with deuterium-labeled (6Z)-non-6-en-2-one
(Figure 3.5E). Reactions of microsomes from Sf9 cells producing recombinant
CYP6CR1 incubated with unlabeled (6E)-non-6-en-2-one, penta-deuterium labeled 6- methylhept-6-en- 2-one (n=1) or monoterpenes did not yield an observable product (n=2, not shown). Assays using the NADPH regeneration system and a 6.5-hour incubation showed recombinant CYP6CR1 and CYP9T2 converted about 33.9% and 8.5% (6Z)- nonen-2-one to the product, respectively, while with a 2-hour incubation, only 8.1% and
3.5% substrate was converted to the product by CYP6CR1 and CYP9T2, respectively
(Table3.2). The products from reactions catalyzed by CYP9T2 and CYP6DH2 had retention times and mass spectra identical to the epoxide standard (not shown). In an unreplicated experiment, CYP6CR1 converted 25.7% substrate to the product in the presence of 1.2 mM each CaCl2 and MgCl2, while assays in the absence of additional
CaCl2 and MgCl2, but using a higher concentration of CYP6CR1 only converted 8.1%
substrate into the product (Table 3.2).
V. Discussion 83
Previous studies showed that exo-brevicomin is specifically located in the male fat body and that a cytochrome P450 is likely involved in its synthesis (Chapter 2). A cytochrome P450 was predicted to epoxidize the known exo-brevicomin precursor, (6Z)- non-6-en-2-one, to keto-epoxide, which subsequently would be cyclized to exo- brevicomin (Figure 3.1)(Vanderwel et al., 1992). Microarray analysis showed a novel cytochrome P450, CYP6CR1, was unique among 30 putative P450s because its expression pattern (Aw et al., 2010) is consistent with exo-brevicomin production
(Pureswaran et al., 2000). In order to test the hypothesis that CYP6CR1 catalyzes the conversion of (6Z)-non-6-en-2-one to the direct epoxide precursor of exo-brevicomin, we undertook a molecular and biochemical characterization. The full sequence of
CYP6CR1 was determined; the predicted translation product has a hydrophobic N- terminal target sequence and many motifs, including the classic heme-binding domain, common to P450s (Figure 3.2). qRT-PCR studies showed CYP6CR1 mRNA levels were
100-fold higher in unfed males relative to females or other developmental stages and localized to the fat body and head of unfed males (Figure 3.3). This expression profile is consistent with both the production of exo-brevicomin (Pureswaran et al., 2000) and GC-
MS analyses showing that exo-brevicomin was specifically produced by unfed male fat bodies (Chapter 2). These data therefore strongly implicate CYP6CR1 in exo- brevicomin biosynthesis.
Functional recombinant CYP6CR1 was expressed in Sf9 cells. The 450 nm peak in the CO difference spectrum is characteristic of correctly-folded functional cytochrome
P450 (Bernhardt, 2006), indicating that enzyme assays could be done using the microsomal preparation. The significant peak at 420 nm suggests the protein was not completely reduced and/or a fraction of recombinant proteins did not properly fold
(Lambalot et al., 1995; Wen et al., 2003). 84
In vitro P450 assays are typically supplemented with additional CPR since over- expressed P450 in baculovirus systems can exhaust endogenous reductase activities
(Wen et al., 2003). Because functional recombinant CYP6CR1 could not be produced when Sf9 cells were co-infected with CYP6CR1- and CPR-encoding baculoviruses, we elected to produce both enzymes separately and mix the microsomes together for assays. The ratio of P450 to P450 reductase can be very important for catalytic activity
(Murataliev et al., 2008; Wen et al., 2003). Electron utilization may be most efficient when the concentration of P450 is greater than that of P450 reductase (Mao et al., 2008;
Murataliev et al., 2008). Therefore, in our enzyme assays, a 4:1 mixture of microsomes bearing recombinant CYP6CR1: CPR was used.
Sf9 microsomes containing recombinant CYP6CR1 readily converted (6Z)-non-6- en-2-one to a product, which was confirmed as (6,7)-epoxy-nonan-2-one, the direct precursor of exo-brevicomin, by comparison with the labeled epoxide standard (Figure
3.5). However, microsomes from uninfected Sf9 cells or cells infected with CPR alone did not epoxidize (6Z)-non-6-en-2-one (Figure 3.5D,E). Thus, epoxide production was due to CYP6CR1, and not to an endogenous activity of Sf9 cells. Increasing reaction times showed increasing product levels (Table 3.2), confirming that the epoxide is an enzymatic product.
Male MPBs also produce endo-brevicomin, which is not an aggregation pheromone component (Pureswaran et al., 2000; Rudinsky et al., 1974), from its
precursor, (6E)-non-6-en-2-one (Vanderwel et al., 1992) with the same pattern as exo-
brevicomin production (Pureswaran et al., 2000). Endo- and exo-isomers may depend
on related desaturases that produce trans- or cis- unsaturated fatty acid precursors and
these precursors may not be discriminated by downstream enzymes, CYP6CR1.
However, recombinant CYP6CR1 did not take (6E)-non-6-en-2-one as substrate, 85 suggesting that a different P450 catalyzes the conversion of (6E)-non-6-en-2-one to endo-brevicomin. (+)-exo-Brevicomin is the natural enantiomer produced by male MPB, with a 98% enantiomeric excess of (+)- to (-)-exo-brevicomin in the blend (Pureswaran et al., 2000). It is likely derived from the cis-epoxide (Silverstein et al., 1968). MPBs exposed to (6Z)-non-6-en-2-one produced (+)-exo-brevicomin, and not (-)-exo- brevicomin (Vanderwel et al., 1992). Non-enzymatic, acid hydrolysis of the cis- and trans-epoxides yields exo- and endo-bicylic products, respectively (Silverstein et al.,
1968). The chirality of the epoxide produced by CYP6CR1 is unclear and may include (-
)- and (+)-cis-epoxide. Identifying these enantiomers can be done by simply reanalyzing these samples with a chiral column.
Frontalin and exo-brevicomin are often grouped together because of the similarity of their and their precursors’ structures (Symonds & Elgar, 2004; Symonds &
Wertheim, 2005). Therefore, we hypothesized that the two chemicals may be epoxidized by the same P450. However, a single enzyme assay conducted with a low concentration of functional CYP6CR1 showed that no frontalin or its intermediates was produced from 6-methylhept-6-en- 2-one (not shown). More replicates with high concentration of functional CYP6CR1 should be examined to confirm this result, however these data suggest that a different P450 is responsible for frontalin production.
Either CaCl2 or MgCl2 or both likely accelerated the conversion of (6Z)-non-6-en-
2-one to epoxide because a higher percentage of substrate (25.7%) was converted to the product in their presence compared to 8.1% conversion in their absence. This unreplicated result is preliminary and should be replicated. Enzymes often require these ions as activators or stabilizers to improve catalytic efficiency (de Bolster, 1997; Fisher et al., 2005). The NADPH regeneration system likely yielded more products (Table 3.2).
Mazur et al., (2009) showed that using NADPH rather than an NADPH regeneration 86
system can underestimate the kinetic rates of intraluminal carbonyl reduction because
NADPH and NADP+ are relatively impermeable and have limited access to the lumen of
ER. These studies should be replicated for confirmation.
In general, P450s exhibit broad substrate ranges to metabolize a large number of endogenous and exogenous compounds, and it is common for different P450s to metabolize same chemical at different positions in the molecule (Lewis, 2000). Musca domestica CYP6A1 is relatively unspecific, metabolizing pesticides in addition to terpenoids (Le Goff et al., 2006). CYP9T2 accepts both myrcene and (+)-α-pinene as substrates (Sandstrom, 2007; Sandstrom et al., 2006). However, many P450s show high specificity toward a particular substrate and product. For example, Diploptera punctata (cockroach) CYP15A1 catalyzes the stereo selective epoxidation of methyl farnesoate to JH III (Helvig et al., 2004). CYP6CR1 appears capable of similarly robust substrate discrimination because no activity was observed with monoterpene substrates,
6-methylhept-6-en- 2-one or with (6E)-non-6-en-2-one (not shown). Recombinant
CYP9T2 and CYP6DH2, which are known monoterpene-hydroxylating P450s
(Sandstrom et al, 2006; Chapter 5) served as negative controls. They nevertheless appeared to have low activity on (6Z)-non-6-en-2-one (Table 3.2), suggesting their ability to accept a broad range of substrates (See Chapter 5). Their catalytic efficiencies in these assays appear much lower than CYP6CR1. In the same conditions, CYP6CR1 converted 33.9% or 25.7% (6Z)-non-6-en-2-one to epoxide while only 8.5% and 7.6% substrate was converted to product by CYP9T2 and CYP6DH2, respectively (Table 3.2).
Thus, CYP6CR1 appears highly specific for (6Z)-non-6-en-2-one and appears to function solely in pheromone biosynthesis. This is the third example of a pheromone- biosynthetic P450 from a bark beetle, and the first from D. ponderosae. Two pheromone-biosynthetic myrcene hydroxylase genes (CYP9T1 and CYP9T2) from Ips 87
spp. have tightly controlled expression patterns regulated in part by juvenile hormone III
(Sandstrom et al., 2008; Sandstrom et al., 2006). CYP6CR1 appears similarly highly
regulated, though the mechanism(s) controlling its expression remain unknown.
VI. Acknowledgments
I thank X. Liu, T. Nguyen from C. Jeffery’s lab (Department of Chemistry,
University of Nevada, Reno) for making penta-deuterium labeled (6Z)-non-6-en-
2-one and epoxide; J. Dickschat (Technical University, Germany) for synthesizing penta-deuterium labeled 6-methylhept-6-en- 2-one; D. Quilici and R.
Woolsey at the Nevada Proteomics for GC/MS analysis; B. Perrino (Department of Physiology, University of Nevada, Reno) for Sf9 cells; M. Schuler (U. Illinois at
Urbana-Champaign) for kindly providing the housefly reductase baculoviral clone; D. Schooley (Department of Biochemistry, University of Nevada, Reno) for generously offering Maxsoft Diode-Array Spectrophotometer for assays; M.
Aw (GJB’s lab) for doing qRT-PCR on developmental groups; members of GJB and CT’s laboratories, especially A. Gorzalski and P. Delaplain, for assistance with collecting beetles, dissection, assays, and S. Young, R. Figueroa Teran for helpful advice; and the Bureau of Land Management (BLM), US forest Service,
South Tahoe District, and the Whittell Board which oversees the UNR little valley forest, for permission to collect beetle-infested trees.
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VIII. Figure legends:
Figure 3.1. Proposed terminal steps in exo-brevicomin biosynthesis. (6Z)-Non-6-en-2- one is derived from fatty acid precursor(s) and converted to epoxide by CYP6CR1. The epoxide is cyclized to exo-brevicomin by a cyclase that may require NADH or NADPH as cofactor.
Figure 3.2. Sequence and translation of CYP6CR1 cDNA. The encoded a.a. sequence of CYP6CR1 is shown above the cDNA sequence, nt position is indicated to the left. 94
The N-terminal transmembrane sequence and conserved WxxxR, ExxR, and
PxxFxPERF motifs are underlined. The heme-binding domain, PFxxGxRxCxG, is boxed.
Figure 3.3. CYP6CR1 mRNA profile. (A) Relative CYP6CR1 mRNA levels in developmental groups, with levels in pre-merged male set at one. (B) Distribution in unfed males. Levels in the posterior midgut were set at one. H, head/pro-thorax; AM, anterior midgut; PM, posterior midgut; HG, hindgut; FB, fat body; C, carcass. The expression of CYP6CR1 is normalized to the expressions of Ubiquitin and tubulin.
Values are means ± standard error, n = 3 for developmental groups and n=4 for the tissue distribution study.
Figure 3.4. CO-difference absorption spectrum of microsomes from Sf9 cells infected with CYP6CR1 recombinant baculovirus. The characteristic absorbance peak at 450 nm indicated functional CYP6CR1 produced. The concentration of functional P450 in this example was approximately 6.3 X10-4 mM.
Figure 3.5. Functional assays. GC traces are shown for reactions including
recombinant CYP6CR1 incubated with (A), unlabeled (6Z)-non-6-en-2-one (18.79 min);
or (B), penta-deuterium labeled (6Z)-non-6-en-2-one (18.68 min). (C), penta-deuterium labeled epoxide standard (27.47 min). (D), microsomes from uninfected Sf9 cells incubated with unlabeled (6Z)-non-6-en-2-one (18.80 min); (E), microsomes from Sf9 cells producing housefly CPR incubated with deuterium labeled penta-(6Z)-non-6-en-2- one (18.69 min). All reactions were performed with the NADPH regeneration system at
30 °C for at least 2 hours. The base peak m/z = 43 was used to analyze reactions 95 conducted with unlabeled substrate, and base peak m/z = 46 was used for labeled substrate. Substrates are indicated by an arrow, and products by an arrowhead.
Figure 3.6 Mass spectra of products and penta-deuterium-labeled epoxide standard.
(A), (B), and (C) mass spectra of the peaks from 27.62 min in Figure 3.5A, 27.47 min in
Figure 3.5B and 27.47 min in Figure 3.5C, respectively. The mass spectrum of the product from B is identical to that of the epoxide standard. The mass to charge of the unlabeled product in A was 3-5 amu less than the standard in C.
96
Fatty acid precursor(s)
O
(6Z)-non-6-en-2-one
CYP6CR1
O O
6,7-epoxynonan-2-one
Cyclase + + NAD /NADP ? NADH/NADPH?
O O
exo-Brevicomin
Figure 3.1 97
1 ACTTCACACTAAGATTGAAGACCACACAATATTGAAGTAGTAGGTGCCAAAAACATCCGA 61 AACAAGTAAACCAGAAATTGTTGCACCAGAGAACACAAAATTGATGAGCAACACATTGAA M L P V P A A L M I I I I I L L L 121 GAAGAAAAGATGTTACCAGTTCCAGCTGCTCTCATGATTATCATCATAATTTTACTCTTA I I F S I V I T F S T Y T F S Y W K K R 181 ATAATTTTCTCTATTGTTATAACCTTCTCTACGTACACTTTTTCTTATTGGAAGAAACGA K F N F L E P T I P F G N A Q S F F L G 241 AAATTTAATTTCCTGGAGCCAACAATCCCATTTGGAAATGCTCAGAGTTTCTTTCTTGGA K K G L G E L Y S D W Y L E M K A K G W 301 AAAAAGGGTTTAGGAGAACTTTATAGCGATTGGTATCTGGAAATGAAAGCCAAGGGTTGG D M G G A Y F G S K P V F I P I D N K L 361 GATATGGGAGGTGCCTATTTCGGCAGCAAACCCGTTTTCATACCAATCGATAATAAGCTG I K T I L V K D F S N F Q N H G F Y I N 421 ATTAAAACGATATTGGTTAAAGATTTTTCGAATTTTCAAAATCACGGCTTTTACATCAAC E K I D P L S G H I Y N L E S S K W K N 481 GAAAAAATCGATCCACTGTCTGGGCATATCTACAATTTGGAAAGCAGCAAGTGGAAGAAC L R S K I L P A F S S S K L R N H I V V 541 CTGCGCTCCAAAATCCTCCCAGCTTTCTCTTCAAGCAAATTGAGGAACCACATAGTTGTC M D S L T K V L V N R L R N M A Q S Q L 601 ATGGATAGTCTCACCAAAGTATTAGTCAACAGACTAAGAAATATGGCACAGTCTCAATTA P I D I K S S L D R F T L D V T S A S L 661 CCAATTGACATTAAGAGTAGTTTGGATAGATTTACCTTGGACGTAACATCCGCCTCCCTT L G I E T E C L K D K N A E L M K Q T R 721 TTGGGAATCGAAACTGAATGTCTGAAAGATAAAAATGCTGAATTAATGAAGCAGACCAGA A F F D I Q L C R L F N T L V L L I P R 781 GCATTCTTTGACATACAATTATGTCGACTTTTTAACACCTTGGTACTGCTAATTCCCAGG N I L I F F N F K V Y P T Q V T N Y F I 841 AACATCCTTATATTTTTCAACTTCAAGGTGTATCCAACACAGGTAACAAATTATTTCATT N F F G N L K A Q R S V E K I R R N D L 901 AACTTCTTTGGCAATCTTAAAGCTCAAAGATCAGTGGAAAAAATAAGGAGAAACGATTTG T D I L I D L C D K T K I V S G D S G N 961 ACCGACATATTAATTGATCTATGCGACAAAACTAAAATTGTTTCCGGGGATAGTGGTAAT G L T E P L T I K E F A A Q M H L F L D 1021 GGTCTAACAGAACCCCTAACAATCAAAGAGTTTGCTGCACAAATGCACCTGTTTCTTGAC T Y E T S S A T E T F A L C E L A A Y P 1081 ACCTACGAAACTTCTTCCGCAACAGAAACTTTTGCTCTGTGTGAACTGGCTGCATATCCA D M Q T R L R N E I N T V L S R F N G V 1141 GATATGCAAACTAGATTGCGAAATGAAATCAATACTGTTTTAAGCAGATTTAATGGTGTT V E Y D A I T E M N Y L D Q V V N E T L 1201 GTTGAGTACGATGCAATCACCGAAATGAACTATTTGGATCAAGTTGTTAATGAAACATTA R K Y P V L P V L P R V C E S D Y P I P 1261 CGGAAGTATCCTGTACTACCCGTTCTTCCAAGAGTATGTGAAAGCGATTATCCAATACCG D S K L T I E K G T L V M V T N M G I H 1321 GATAGTAAACTTACCATTGAGAAAGGAACATTGGTAATGGTGACAAACATGGGAATTCAT Y D P E Y Y P D P M R F D P E R F T S E 1381 TACGATCCTGAATATTATCCAGATCCAATGCGTTTTGATCCAGAACGATTTACGTCAGAA N I A K R P F S S F V P F G E G P R S C 1441 AATATAGCGAAAAGACCATTTTCCTCATTTGTTCCGTTTGGTGAAGGTCCAAGAAGTTGT V G K R L G M L Q A K V G L I A I L R N 1501 GTCGGGAAGCGTCTTGGAATGTTGCAGGCAAAAGTTGGTTTGATTGCAATCCTTCGGAAT F K I T L S E K T K M P I Q F E K S G L 1561 TTCAAGATAACATTAAGCGAGAAGACTAAGATGCCGATCCAGTTTGAAAAATCTGGACTT F L N P E G K I W I N L E T I E * 1621 TTTTTGAACCCTGAAGGAAAAATATGGATAAACCTTGAAACAATTGAGTGATTATGACCA 1681 TTT
Figure 3.2 98
Table 3.1
Primer name Sequence Amplicon length (bp)
Sequencing DPG017M13 (CYP6CR1) DPG017M13F1 CGATCCACTGTCTGGGCA n.a. CYP6CS1F3 GCGATTATCCAATACCGG n.a. CYP6CS1R3 CCGGTATTGGATAATCGC n.a. CYP6CS1F4 CCTTGGTACTGCTAATTCCC n.a. CYP6CS1R4 GGGAATTAGCAGTACCAAGG n.a. CYP6CR1F1 GGTCCAAGAAGTTGTGTCG n.a. 6CR1F2 AACACATTGAAGAAGAAAAG n.a. 6CR1R2 GCCCAGACAGTGGATCG n.a.
Cloning CYP6CS1F1 GCGGATCCAACACATTGAAGAAG CYP6CR1R1(tagged) GCCTCGAGTCAAACTGGATCGG 1514 CYP6CR1R3(untagged) GCCTCGAGAATGGTCATAATCAC 1589 qRT-PCR MPBcon608-RT-F1 GGAGCCAACAATCCCATT TG MPBcon608-RT-R1 CCAGATACCAATCGCTATAAAGTTCTC 86 n.a , not applicable
99
140 (A) 120
100 Female
80 Male 60
40
) 20 s.e.
± 0 ddCT -
14 (B)
12
10 Relative Expression (2 Expression Relative 8
6
4
2
0 H C FB MG PM HG
Figure 3.3 100
0.11
0.1
0.09
0.08
0.07
0.06 Absorbance
Δ 0.05
0.04
0.03
0.02 400 420 440 460 480 500 Wavelength (nm)
Figure 3.4 101
(A) CYP6CR1 & unlabeled (6Z)-non-6-en-2-one 18.79
27.62
(B) CYP6CR1 & labeled (6Z)-non-6-en-2-one 18.68
27.47
(C) Std of labeled epoxide 27.47 Relative Abundance Relative (D) Sf9 cells & unlabeled (6Z)-non-6-en-2-one
18.80
(E) CPR & labeled (6Z)-non-6-en-2-one
18.69
17 18 19 20 21 22 23 24 25 26 27 28 Time (min )
Figure 3.5 102
[ (A)] 27.62 min in Figure 3.5A 0
1005
0
5
0
5 43
0
5
0
5
0
5
0
5 0 O O 5 55 83 0 98 5 67.1 0 114 5 0 0 (B) 27.47 min in Figure 3.5B
[ ] 0 1005 0
5
0
5 46.1
0
5
0
5
0 5 O O 0
5 0 D2 D 5 3 0 57.2 85.1 102.2 5
0 69.2
Relative Abundance Relative 117.1 5 0 0 (C) 27.47 min in Figure 3.5C
0
5
1000 5 46.1 0
5
0
5
0
5
0
5 O O
0
5 0 D2 D3 5 0 57.2 85.1 102.1 5
0 69.2 117.1 5 161.1 0 0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 m/z
Figure 3.6 103
Table 3.2: Preliminary studies of CYP6CR1 activity
incubation NADPH – MgCl Product/(Product+ [P450] NADPH 2 time reg* ,CaCl2 Substrate) (min) X 10-4 mM (300 μM) 1X 1.2 mM CYP6CR1 Neg. Control
Exp. 1 30 1.35 + - - 5.8% Sf9 0 Exp. 2 30 2.1 + - - 5.6% CPR 0 Exp. 3 2 2 + - - 5% CPR 0 Exp. 4 2 3.1 - + - 8.1% 9T2 3.4% Exp. 5 6.5 3.1 - + - 33.9% 9T2 8.5% Exp. 6 2 1.4 - + + 25.7% 6DH2 7.6% NADPH–reg*: NADNPH regeneration system 104
Chapter 4
ZnoDH is a novel dehydrogenase involved in exo-brevicomin biosynthesis in the
mountain pine beetle
I. Abstract II. Introduction III. Materials and Methods IV. Results V. Discussion VI. Acknowledgements VII. References VIII. Figure Legends
I. Abstract
Investigating pheromone production in mountain pine beetles (MPB;
Dendroctonus ponderosae) may identify unique molecular targets and provide information for developing specific new strategies to deal with their infestations. exo-
Brevicomin is a male-specific aggregation pheromone component produced in the fat body. Male fat bodies incubated with (6Z)-non-6-en-2-one produced nonen-2-ol
(Chapter 2), and unfed male fat bodies converted tri-deuterated decanoic acid to nonan-
2-one via decarboxylation, strongly implicating that 10:1 fatty acid is the precursor to
(6Z)-non-6-en-2-one (Chapter 2). Microarray analysis from the MPB showed a unique dehydrogenase, DPG022F04, (renamed as ZnoDH) has an expression pattern consistent with exo-brevicomin production. We hypothesized that ZnoDH could function as either a decarboxylation or dehydration enzyme to produce (6Z)-non-6-en-2-one, or a cyclase to convert epoxide to exo-brevicomin. In this study, we show that ZnoDH was highly expressed in male fat bodies, and that the recombinant ZnoDH converted penta- deuterium labeled nonen-2-ol to (6Z)-non-6-en-2-one, but did not catalyze either 105
decarboxylation or the cyclization of epoxide to exo-brevicomin. This finding leads to a
new hypothesized pathway to (6Z)-non-6-en-2-one based on the hydrocarbon
biosynthetic pathway in insects.
II. Introduction:
exo-Brevicomin (exo-7-ethyl-5-methyl-6,8-dioxabicyclo[3.2.1]octane) is a male
specific mountain pine beetle (MPB; Dendroctonus ponderosae) pheromone component
(Borden et al., 1987; Pierce et al., 1987; Pureswaran et al., 2000; Vanderwel et al.,
1992). Studying the exo-brevicomin biosynthesis pathway may identify unique
molecular targets and provide information for developing specific new strategies to deal
with pine bark beetle infestations. Mature male MPBs contain exo-brevicomin when they
exit the brood tree; feeding results in decreased exo-brevicomin levels (Pureswaran, et
al., 2000). (6Z)-Non-6-en-2-one is believed to be a natural precursor of exo-brevicomin
(Vanderwel, et al., 1992). MPB beetles exposed to (6Z)-non-6-en-2-one produce exo-
brevicomin and nonen-2-ol (Vanderwel, et al., 1992). Our previous studies showed that homogenized male fat bodies converted (6Z)-non-6-en-2-one into the direct epoxide precursor of exo-brevicomin via CYP6CR1 (Chapter 2, 3). These incubations also produced nonen-2-ol (Chapter 2). The epoxide is very stable in physiological conditions and only cyclizes into exo-brevicomin at pH < 3 (Chapter 2), suggesting a cyclase is required in the final step of exo-brevicomin biosynthesis.
(6Z)-Non-6-en-2-one is believed to be derived from fatty acyl precursor(s)
(Vanderwel, et al., 1992; Vanderwel & Oehlschlager, 1992); indeed, we showed that
(6Z)-non-6-en-2-one is likely derived from C10 fatty-acyl compounds that contain a double bond in the ω -3 position through de novo synthesis because male fat bodies
converted tri-deuterated decanoic acid (10 carbon) to nonan-2-one (9 carbon) (Chapter 106
2). These studies confirm the general pathway that incorporates β-oxidation to shorten
long chain fatty acids, such as C16:1 or C18:1, to C10:1 (Figure 4.1). However, the
mechanism used to convert C10:1 fatty acyl-CoA to (6Z)-non-6-en-2-one remains unknown.
(6Z)-Non-6-en-2-one production from a C10 fatty acyl-CoA precursor requires a
chain-shortening step to remove the carboxylic carbon and yield a C9 product. While β-
oxidation is not appropriate because it removes two carbons per cycle, direct
decarboxylation would remove just the carboxylic carbon. (Z)-10-Heptadecen-2-one, an
aggregation pheromone component of Drosophila mulleri, is derived from the fatty acid
elongation pathway, likely via decarboxylation (Skiba & Jackson, 1994). In this system,
the β-keto-acid intermediate from fatty acid elongation is decarboxylated to the corresponding 2-ketone (Skiba & Jackson, 1994). Decarboxylation also occurs via a different mechanism during hydrocarbon biosynthesis in insects as named decarbonylation (Reed et al., 1994; Rule & Roelofs, 1989; Vaz et al., 1988). More universally, there are several dehydrogenases that catalyze decarboxylation, including pyruvate dehydrogenase, isocitrate dehydrogenase and α-ketoglutarate dehydrogenase in the tricarboxylic acid cycle (Denton et al., 1975; Krebs & Weitzman, 1987; Lowenstein,
1969). It is likely that a dehydrogenase catalyzes the reaction from an unsatured C10
fatty acyl-CoA precursor to (6Z)-non-6-en-2-one in the MPB (Figure 4.1) although the β- keto acid is relatively unstable and may spontaneously decarboxylate (Skiba and
Jackson, 1992).
Nonen-2-ol production in fat bodies incubated with (6Z)-non-6-en-2-one raises an alternative possible pathway for (6Z)-non-6-en-2-one production, in which C10: 1 fatty acid is decarboxylated to 3-nonene via fatty acyl-CoA reductase and oxidative decarbonylase – the standard metabolic pathway from fatty acid to hydrocarbon in 107
insects (Reed, et al., 1994). 3-Nonene could then be hydroxylated to (6Z)-non-6-en-2-ol by a P450, and then oxidized to (6Z)-non-6-en-2-one (G. Blomquist, personal communication) (Figure 4.1). Short-chain hydrocarbon hydroxylation is precedent in D. jeffreyi, a sibling species of MPB that readily hydroxylates heptane to heptan-2-ol and heptan-1-ol (Paine et al., 1999).
A “functional genomics followed by enzyme characterization” approach in I. pini led to the identification of important pheromone-biosynthetic enzymes in that species
(Figueroa-Teran et al., 2011; Keeling et al., 2006; Keeling et al., 2004; Sandstrom et al.,
2006). This strategy to combine functional genomics, molecular biology, and biochemistry was also successfully used in identifying the function of CYP6CR1 in exo- brevicomin biosynthesis (Chapter 3). Clustering analysis from MPB microarray data revealed a novel dehydrogenase, represented by EST DPG022F04 (GenBank ID#
GO494274), later renamed “ZnoDH’ (for (6Z)-non-6-en-2-ol Dehydrogenase), that clustered with CYP6CR1 (Aw et al., 2010). Its expression pattern is consistent to the production of exo-brevicomin: it is highly expressed in unfed male and feeding decreases its expression level (Aw et al., 2010). In this study, we hypothesized that
ZnoDH is involved in exo-brevicomin biosynthesis either as cyclase acting downstream of CYP6CR1, or a dehydrogenase dehydrating nonen-2-ol to (6Z)-non-6-en-2-one, and possibly as an oxidoreductase drawing carbon away from β-oxidation at decanoic acid
(Figure 4.1). In the latter case, the nine-carbon intermediate could exit β-oxidation as a secondary alcohol, (6Z)-non6-en-2-ol, or as a ketone, (6Z)-non-6-en-2-one, either of which could be interconverted by a dehydrogenase (Figure 4.1).
To confirm these hypotheses, qRT-PCR and recombinant ZnoDH production in
Sf9 cells with a baculovirus expression system were used to confirm gene expression profiles and to perform enzyme assays to find possible substrates. ZnoDH, like 108
CYP6CR1, is highly expressed in male fat bodies. Recombinant ZnoDH readily
converted nonen-2-ol to (6Z)-non-6-en-2-one in the presence of NAD+ or NADP+, with a
preference of NAD+. Sf9 cells infected with ZnoDH recombinant DNA cultured with tri-
deuterium-labeled decanoic acid did not yield any products, and recombinant ZnoDH did
not cyclize epoxide into exo-brevicomin. These data suggest (6Z)-non-6-en-2-ol is an
intermediate in the pathway and support the hypothesis that (6Z)-non-6-en-2-one biosynthesis uses components for insect hydrocarbon biosynthesis (Reed et al.,1994).
III. Materials and methods
Reagents and chemicals. Hink’s TNM-FH Medium 1x (Supplemented Grace’s
Medium) and Grace’s Insect Basal Medium 1x were from Mediatech, Inc.
(Herndon, VA) and FBS was from Atlas Biologicals (Fort Collins, CO). Direct
baculovirus DNA kit, SF-900 II (1X and 1.3X) media were from Invitrogen
(Carlsbad, CA). The Sf9 cells were a gift from B. Perrino (Department of
Physiology, University of Nevada, Reno (UNR)). Housefly cytochrome P450
reductase (CPR) (Wen et al., 2003) was kindly provided by M. Schuler (U. Illinois
at Urbana-Champaign). Unlabeled (6Z)-non-6-en-2-one was purchased from
Phero Tech Inc. (Delta, BC, Canada). Penta-deuterium labeled (6Z)-non-6-en-2- one and nonen-2-ol (6Z)-non-6-en-2-ol) were synthesized by C. Jeffery
(Department of Chemistry, UNR). Agrose was from Bio-Express (Keysville, UT).
Phenylmethylsulfonyl fluoride (PMSF), protease Inhibitor cocktail, NAD+ and
NADP+ were from Sigma- Aldrich (St. Louis, MO). All oligonucleotides were from
Integrated DNA Technologies (Coralville, IA).
Beetles. Mountain pine beetles were obtained from ponderosa pine bolts collected
from the UNR little valley forest (the Sierra Nevada in Nevada, USA). The bolts were 109
stored in a greenhouse and emerged adults were collected daily and stored for up to 1-2
weeks at 4 ºC in moist paper towels in loosely capped jars as described previously
(Browne, 1972). Emerged beetles were separated by sex (Wood, 1982). Healthy beetles were selected for experiments. The different groups of the beetles were dissected on the same day. Unfed beetles were held in plastic cups in the dark at room temperature with moist paper towels to prevent desiccation as previously described
(Keeling, et al., 2004).
ZnoDH cDNA sequencing. The ZnoDH cDNA clone (represented by DPG022F04) was grown from a frozen permanent stock and completely sequenced by primer walking
(Table 4.1) of purified plasmid template DNA (Qiagen, QIAprep Spin Miniprep Kit,
Valencia, CA) using the ABI BigDye Terminator Cycle Sequencing Ready Reaction Kit v3.1. Sequencing reactions were run on an ABI3730 DNA Analyzer at the Nevada
Genomics Center (UNR) and the sequences were analyzed using Vector NTI. Advance
9 software (Invitrogen).
Expression analysis. ZnoDH mRNA levels were determined by Real-Time reverse transcriptase PCR (qRT-PCR). Template cDNAs from 11 biological groups were prepared previously (Aw, et al., 2010). For the tissue distribution analysis, males were kept unfed overnight at room temperature in the dark and then dissected in 100 mM sodium phosphate buffer pH 7.8 under a stereo-microscope. Tissues from the head
(including the prothorax), anterior midgut, posterior midgut, hindgut, fat body, and carcass of ten beetles were pooled and frozen in liquid N2. Four biological replicates
were made for each sample. The tissues were kept at -80 ºC until RNA was extracted
using the RNeasy Plant Mini Kit (Qiagen). Twenty percent of the isolated RNA was
reverse-transcribed using Superscript III reverse transcriptase and random primers
(Invitrogen) according to the manufacturer’s protocol. 110
Primers for qRT-PCR were designed using Primer Express v 2.0 software
(Applied Biosystems, Foster City, CA) and analyzed with vector NTI Advance 9 software.
Primers with minimal potential for primer-dimer and hairpin loop formation were selected
(Table 1) and tested for non-specific amplification by visual inspection of melting curves.
Their amplification efficiencies were determined using a relative standard curve method.
Each 10 μl PCR reaction contained 1.5 μM each forward and reverse primers, 3 μl of appropriately diluted cDNA template, and 5 μl SYBR Green PCR Master Mix (Applied
Biosystems). Amplifications were performed in triplicate, for a total of four replicates per sample, at the NGC. Relative expression values for all genes were determined using the ΔΔCT method normalized to tubulin and Ubiquitin, which are stably expressed in
MPB (Aw, et al., 2010). Reactions included no template controls to monitor for genomic
DNA contamination.
Antibody production. Molecular Technologies (Piscataway, NJ) was contracted to produce a rabbit anti-22F04 polyclonal antibody based on the antigenic peptide
(CNPSPDYFEERKGWR) designed and selected by the contracted company. The synthesized peptide was injected into the two rabbits separately. Four immunizations per rabbit were performed. Serum was collected, and the antibody was purified using peptide-conjugated resin. The quality of the antibody was examined by ELISA. The antibody was used to confirm recombinant protein produced in infected Sf9 cells and in beetle tissues.
Recombinant baculoviral protein expression. Protocols for growth and maintenance of Sf9 cells, recombinant baculovirus construction, and heterologous expression using the BaculoDirectTM Expression Kit were as described by Invitrogen. Sf9 cells were cultured at 27ºC in a shaking flask for all procedures. ZnoDH in pDONR was transferred into BaculoDirect C-Term Linear DNA by recombination using LR Clonase™ II Enzyme 111
Mix (Invitrogen). The recombinant baculoviral clone was transfected into Sf9 cells and
grown in the presence of 100 µM ganciclovir to select for recombinant virus. High titer
P3 viral stocks for each construct were produced by successive 72h amplifications of the
initial and P2 stocks. Approximate viral titers were determined by a plaque assay as
described by Invitrogen. The viral stocks were used to infect Sf9 cells grown to a density
of 1.0 x 106 cells/ml in a disposable shaking flask (Bio-Express, Keysville, UT).
For recombinant ZnoDH and CPR (Wen et al., 2003) expression, Sf9 cells were infected with recombinant baculovirus at multiplicities of infection (MOIs, pfu/cell) ranging from 1 to 20 for ZnoDH, and 0.1 for CPR (negative control). To determine the optimum expression conditions for ZnoDH, cells were harvested at day 1, 2, 3 and 4 post infection
(PI) and at various MOI and analyzed by SDS-PAGE and western blot. The best condition was chosen for further experiments.
Recombinant protein preparation. Sf9 cells producing recombinant ZnoDH or CPR were harvested on day 3 PI and recombinant proteins were prepared by differential centrifugation. Briefly, cells were pelleted by centrifugation at 3000 x g at 4 ºC for 10 min. The pellets were washed twice in 1/5 cell culture volume of ice-cold 100 mM sodium phosphate buffer (pH 7.8) and repelled at 3000 x g for 10 min. The pellets were resuspended in 1/30 cell culture volume of ice-cold cell lysate buffer (100 mM sodium phosphate pH 7.8, 1.1 mM EDTA, 0.1 mM DTT, 0.5 mM PMSF, 1/1000 vol/vol Sigma protease inhibitor cocktail) and lysed by sonication three times for 15 s on ice with a
Branson Sonifier 450. The lysate was centrifuged at 10,000 x g for 20 min at 4 ºC in a microcentrifuge tube and the supernatant (cell lysate supernatant) was used immediately for enzyme assays. For sub-cellular localization studies, the cell lysate supernatant was further centrifuged in a TLA110 rotor at 120,000 x g for 2 h in a Beckman-Coulter Optima ultracentrifuge to pellet microsomes. The microsomal pellet was resuspended in 1/30 112
cell culture volume of ice-cold cell lysate buffer, and 50µl solution from each step was
taken.
Western blotting. Head, fat body, anterior midgut, midgut, posterior midgut, malpighian
tubule, sex organ and carcass were dissected from male MPBs that had been unfed
overnight. Each sample contained tissues from 10 beetles. Protein concentrations were
measured at the Nevada Proteomics Center (NPC). Equal amounts of protein were
loaded into each lane of a 12% SDS-PAGE gel, separated by electrophoresis, and
transferred to nitrocellulose membrane. The membrane with transferred protein was
stained with 1X ponceau for examining the total protein loaded, then removed ponceau
with TBST buffer. For recombinant ZnoDH, the same volume of the microsomal fraction
and the supernatant were loaded to the SDS gel. The nitrocellulose membrane
containing proteins was blocked in 5% milk /TBST(20mM Tris pH 7.4, 0.15M NaCl, 0.1%
Tween), followed by the incubation of 1:1000 rabbit anti-22F04 antibody (GenScript
Corp, Piscataway,NJ,USA) containing 1% milk. After washing 5 times with TBST buffer,
the membrane was incubated in 1:2,000 goat anti-rabbit lgG secondary antibody
(BioRad, Hercules, CA) containing 1% milk. The membrane was incubated with
SuperSignal West Pico Chemiluminescent Substrate (Pierce, Rockford, IL) and imaged
with Kodak X-ray film (Carestream health, Rochester NY).
Enzyme assays. In vitro assays were conducted in 500 µl volumes containing 468 µl
of lysates from recombinant ZnoDH, boiled recombinant ZnoDH, recombinant CPR, or
uninfected Sf9 cells, and 300 µM penta-deuterium-labeled nonen-2-ol or 496 µM penta-
deuterium labeled epoxide. Reactions were initiated with the addition of 300 µM NADP+ or NAD+, and incubated in a 30 ºC water bath for 0, 25 min, 50 min, 2h or 4h for assays using nonen-2-ol as substrate. When using epoxide as substrate, NADP+, NAD+, NADH and NADPH were used as cofactors in separate reactions with 2h incubations at 30 ºC. 113
Reactions were terminated and extracted twice with pentane: ether (1:1) spiked with 5
µg/ml n-octanol (internal standard). To examine decarboxylation activity, Sf9 cells infected with recombinant ZnoDH viral DNA at MOI=10 were cultured in 2ml volumes for
72h in 6-well culture plates. Before harvesting, 2 mg tri-deuterium decanoic acid was added to the cultures, and the cultures were incubated at 27 ºC. Samples were removed at different times (30min, 1, 2, 3 and 4h) to allow Sf9 cells to make C10 fatty acyl-CoA derivatives and then extracted with pentane: ether. Samples were concentrated to approximately 100 µl with N2 gas and directly analyzed by coupled GC-
MS at the NPC. A Thermo Finnigan Polaris Q ion trap was used with a molecular weight scanning range of 40–180 atomic mass units (amu) at an ionization potential of 70 eV.
A trace gas chromatograph containing a 60 m x 0.25 mm (ID), 0.25 µm film thickness
DB-5 capillary column (J&W Scientific, Palo Alto, CA) was programmed for an initial temperature of 50 ºC (1 min hold), increased to 200 ºC at 5 ºC/min, 10 ºC/min to 320 ºC
(20 min hold). The injector was split at a ratio of 100:1 at a temperature of 280 ºC with a column flow of 1.5 ml He/min. The detector was set at 200 ºC. The product was identified by comparing retention times and mass spectra with negative controls and an authentic standard. The percentage of substrate conversion was calculated according to areas of product and substrate by dividing the amount of product by the total amount of products and substrates (product/(products + substrates)).
IV. Results
Sequencing. The full-length cDNA was completely sequenced from ZnoDH. The 1097 bp cDNA contained a 987 nucleotide (nt) open reading frame (ORF) encoding a 329 amino acid (a.a.) protein flanked by 60 nt and 47 nt 5’ and 3’ UTRs, respectively (Figure
4.2). The predicted translation product has a molecular mass of 36 kDa and an 114
isoelectric point of 8.6 (Gasteiger et al., 2003). Blast searches (Altschul et al., 1997)
indicate ZnoDH is most similar to retinol dehydrogenases, light dependent
protochlorophyllide oxidoreductases and classic short chain dehydrogenase/reductases
(SDR) from all domains of life. The best hit to a characterized protein was to human
WW domain-containing oxidoreductase (Bednarek et al., 2000) with 23.8% a.a. identity
(not including WW domain) and E=5.21e-25. The primary structure has several
conserved sequence motifs of classic SDRs. These include the cofactor binding site,
GXXXGXG, and substrate binding site, YXXXK (Bednarek, et al., 2000).
ZnoDH expression. ZnoDH mRNA levels were assayed by qRT-PCR. ZnoDH mRNA
levels were highest in unfed males, decreasing to approximately one fourth that level in
fed males (Figure 4.3A). In contrast, mRNA levels in larvae, pre-emerged adults, and
females were all approximately 30-fold lower than those in unfed males (Figure 4.3A).
ZnoDH mRNA was predominately localized to the fat body of unfed males (Figure 4.3B),
almost 10-fold higher than levels observed in guts and 2-fold higher than levels in heads and carcasses (Figure 4.3B).
ZnoDH localization. We expressed ZnoDH at various MOI and harvested samples
from 1 to 4 days PI to determine optimal expression conditions. An approximately 36
kDa protein was most abundant at 3 days PI in cells infected with MOI = 5 (not shown).
These conditions were used to express recombinant ZnoDH for sub-cellular localization
studies and enzyme assays. Western blot analysis confirmed recombinant ZnoDH was
present at 3 days PI at MOI = 5, but not in non-infected Sf9 cells (Figure 4.4A) and sub-
cellular fractionation assays showed most ZnoDH localized in the microsomal pellet
(Figure 4.4A). An approximately 36 kDa protein band was predominantly present in the
fat body tissues compared with other tissues (Figure 4.4C). Ponceau staining with the 115
same gel showed that total protein loaded in fat body line was much less than other
samples except for anterior midgut and midgut (Figure 4.4B).
Enzyme Assays. Recombinant ZnoDH converted penta-deuterium labeled nonen-2-ol
to a product with the retention time of 18.28 min in the presence of NAD+ (Figure 4.5C)
and 18.27 min in the presence of NADP+ (Figure 4.6C), while recombinant CPR (Figure
4.5B, 4.6B) and uninfected Sf9 cell lysates (not shown) incubated under the same conditions produced a small peak of product with the same retention time. MS analysis
(m/z= 46) showed that these products had an m/z spectrum (Figure 4.5F, 4.6F) identical to that of the penta-deuterium labeled (6Z)-non-6-en-2-one standard (Figure 4.5G, 4.
6G), although the standard had a retention time of 18.36 min (Figure 4.5D, 4.6D). In the presence of NAD+, about 80% of substrate was converted to the product (Figure 4.5C),
while in the presence of NADP+ about 37.9% substrate was converted to the product
(Figure 4.6C). For assays incubated for less than 4h, twice as much substrate was
converted to the product in the presence of NAD+ compared to the amount converted by
incubations with NADP+. When incubation times were extended to 4h, almost the same
percentage of substrate was converted to the product with both cofactors (Figure 4.7).
The percentage of substrate conversion did not change in incubations with recombinant
CPR regardless of the cofactor present (Figure 4.7).
V. Discussion
exo-Brevicomin, an male MPB aggregation pheromone component (Borden, et al., 1987; Pierce, et al., 1987; Pureswaran, et al., 2000; Vanderwel, et al., 1992), is specifically synthesized in the male fat body (Chapter 2). It has been proposed that it is likely derived from known precursor (6Z)-non-6-en-2-one via a P450 mediating epoxidation, followed by cyclization of epoxide and 6Z)-non-6-en-2-one is derived from 116 fatty acid via decarboxylation (Vanderwel, et al., 1992). Previous studies confirmed that
(6Z)-non-6-en-2-one is likely derived from C10 fat acid via decarboxylation or decarbonylation (Chapter 2) and male fat bodies converted (6Z)-non-6-en-2-one to the direct epoxide precursor of exo-brevicomin via CYP6CR1 (Chapter 2, 3) and (6Z)-non-6- en-2-ol (Chapter 2). Clustering analysis of microarray data in MPB suggested ZnoDH may play a role in the exo-brevicomin biosynthesis pathway because its expression pattern (Aw, et al., 2010) is consistent with exo-brevicomin production (Pureswaran, et al., 2000). We hypothesized that ZnoDH may work as a cyclase downstream of
CYP6CR1, or a dehydrogenase dehydrating nonen-2-ol to (6Z)-non-6-en-2-one, and possibly as a decarboxylase catalyzing the decarboxylation form C10 to C9 (Figure 4.1).
To identify the role of ZnoDH in this hypothesized pathway, we performed a series of molecular and biochemistry experiments.
The predicted translation product of ZnoDH has all SDR family critical sequence motifs including the nucleotide binding and substrate binding motifs (Figure 4.2), making
ZnoDH a classic short chain dehydrogenase, similar to retinol dehydrogenases. These enzymes show similar mechanism, but have low sequence similarity, sharing only conserved sequence motifs including a classic Rossmann-fold for nucleotide binding
(Kavanagh, et al., 2008). The best BLASTP (Altschul, et al., 1997) hit was to human
WW domain-containing oxidoreductase. The human enzyme appears to have a role in cancer tumor genesis (Bednarek et al., 2000). Finding closely-related SDRs with very different functions in different taxa is not unusual. Figueroa-Teran et al (2010) noted a pheromone-biosynthetic SDR from I. pini that was most similar to a human SDR. This type of comparison illustrates how this enzyme family has broadly diverged over time to assume different roles, while maintaining a common reaction mechanism. 117
qRT-PCR showed that ZnoDH mRNA levels were higher in unfed males relative to females or other developmental groups and specifically localized to the fat body of unfed male MPBs (Figure 4.3). This expression pattern is consistent to microarray data
(Aw et al., 2010) and with the expression pattern of CYP6CR1 (Chapter 3) and the production of exo-brevicomin (Chapter 2). These data strongly implicate ZnoDH in exo- brevicomin biosynthesis.
Recombinant ZnoDH production by Sf9 cells was confirmed by western blotting.
This recombinant protein mostly localized in microsomes (Figure 4.4). In vivo, ZnoDH was predominantly localized in the fat body (Figure 4.4) although accurate comparison is complicated by the fact that the total amount of protein differed for each sample, despite attempts to quantitate prior to loading. The denaturing detergent in the protein samples may have affected quantitation.
Sf9 cells infected with recombinant ZnoDH did not convert tri-deuterium-labeled decanoic acid into any products under several culture conditions (not shown), suggesting that ZnoDH may not be involved in decarboxylation from C10 fatty-acyl compounds to a C9 compound, and decarboxylation may be conducted upstream of
ZnoDH. Alternatively, it is possible that Sf9 cells were not able to convert C10 fat acid to the precursor of ZnoDH for decarboxylation. To further confirm functions of ZnoDH, C10
fatty-acyl derived compounds, such as hydroxyacyl CoA or 3-ketoacyl CoA could be used as substrates.
Furthermore, recombinant ZnoDH did not cyclize epoxide into exo-brevicomin
(not shown), indicating that ZnoDH does not catalyze the terminal step in exo-brevicomin biosynthesis. Interestingly, enzyme assays showed that recombinant ZnoDH readily converted penta-deuturium labeled nonen-2-ol to products using both NAD+ and NADP+
as its cofactor (Figure 4.5, 4.6). These two products are confirmed as penta-deuturium 118
labeled (6Z)-non-6-en-2-one by labeled (6Z)-non-6-en-2-one standard. Cell lysate
supernatants from uninfected Sf9 cells (not shown) or Sf9 cells producing recombinant
CPR converted about 2.5% substrate to (6Z)-non-6-en-2-one in the presence of NAD+
(Figure 4.5B) and 1.7% in the presence of NADP+ (Figure 4.6B), but this product was not observed in the boiled ZnoDH (Figure 4.5 A, 4.6A). These activities may be due to an endogenous dehydrogenase in the Sf9 cells. In the presence of NAD+, more substrate
was converted by recombinant ZnoDH to the product than in the presence of NADP+
(Figure 4.6, 4.7), suggesting a preference to use NAD+ as a cofactor. Increasing the
incubation time resulted in more products from ZnoDH with, but not from CPR,
regardless of which cofactor was present (Figure 4.7), further supporting that the
observed product was due to ZnoDH activity, but not to an endogenous activity of Sf9
cells. More products produced by ZnoDH in the presence of NAD+ than in the presence
of NADP+ indicates that the recombinant ZnoDH prefers NAD+ as its cofactor.
These results suggest ZnoDH is likely involved in exo-brevicomin biosynthesis by
specifically catalyzing (6Z)-non-6-en-2-one production. However, these data also
present an intriguing mystery regarding the source of nonen-2-ol. Standard
decarboxylation of a C10 precursor should yield the methylketone (Skiba & Jackson,
1994), so the requirement for ZnoDH is unclear. It is more likely that C10: 1 fatty acid is
decarbonylated to 3-nonene via fatty acyl-CoA reductase and oxidative decarbonylase –
the standard metabolic pathway from fatty acid to hydrocarbon in insects (Reed, et al.,
1994). 3-Nonene could then be hydroxylated to (6Z)-non-6-en-2-ol by a different P450
(G. Blomquist, personal communication). This hypothesis is supported by the observation that male fat bodies converted (6Z)-non-6-en-2-one to (6Z)-non-6-en-2-ol
(Chapter 2) and MPBs exposed to (6Z)-non-6-en-2-one produced (6Z)-non-6-en-2-ol
(Vanderwel, et al., 1992). Furthermore, a sibling species, D. jeffreyi, hydroxylates 119
heptane (Paine et al., 1999), suggesting the presence of an appropriate P450 to
hydroxylate 3-nonene in D. ponderosae. Examining if fat bodies convert 3-nonene to
nonen-2-ol may answer this hypothesis. The possibility that ZnoDH acts as a
decarboxylase should be also examined because decarboxylations are commonly
catalyzed by dehydrogenases (Rule & Roelofs, 1989; Skiba & Jackson, 1994; Vaz, et
al., 1988). However, a preliminary assay of ZnoDH-recombinant Sf9 cells cultured with
C10:0 fatty acid did not produce a 9-carbon product, suggesting ZnoDH does not
function as a decarboxylase under the conditions tested.
VI. Acknowledgments
I thank X. Liu, T. Nguyen from C. Jeffery’s lab (Department of Chemistry,
University of Nevada, Reno) for making penta-deuterium labeled nonen-2-ol and
(6Z)-non-6-en-2-one; D. Quilici and R. Woolsey at the Nevada Proteomics for
GC/MS analysis; B. Perrino (Department of Physiology, University of Nevada,
Reno) for Sf9 cells; M. Schuler (U. Illinois at Urbana-Champaign) for kindly
providing the housefly reductase baculoviral clone; P. Delaplain for doing qRT-
PCR, sequencing and making the P3 viral stock; L. Plaugher for doing functional assays; members of GJB and CT’s laboratories, especially A. Gorzalski for assistance with collecting beetles, dissection and S. Young, R. Figueroa-Teran, for helpful advice; and the Bureau of Land Management(BLM), US forest
Service, South Tahoe District, and the Whittell Board which oversees the UNR little valley forest, for permission to collect beetle-infested trees.
VII. References 120
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Borden, J. H., Ryker, L. C., Chong, L. J., Pierce, H. D., Johnston, B. D., & and
Oehlschlager, A. C. (1987). Response of the mountain pine beetle, dendroctonus
ponderosae Hopkins (coleoptera: colytidae), to five semichemicals in British
Columbia lodgepole pine Forests. Can. J. Forest Res., 17, 118-128.
Browne, L.E., 1972. An emergence cage and refrigerated collector for wood-boring
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Denton, R. M., Randle, P. J., Bridges, B. J., Cooper, R. H., Kerbey, A. L., Pask, H. T.,
Whitehouse, S. (1975). Regulation of mammalian pyruvate dehydrogenase. Mol
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Figueroa-Teran, R., Welch, W., Blomquist, G., & Tittiger, C. (2011). Ipsdienol
dehydrogenase (IDOLDH): A novel oxidoreductase important for Ips pini
pheromone production. Insect Biochem Mol Biol. 42(2):81-90. 121
Gasteiger, E., Gattiker, A., Hoogland, C., Ivanyi, I., Appel, R. D., & Bairoch, A. (2003).
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Nucleic Acids Res, 31(13), 3784-3788.
Kavanagh, K. L., Jörnvall, H., Persson, B., & Oppermann, U. (2008). Medium- and short-
chain dehydrogenase/reductase gene and protein families: the SDR superfamily:
functional and structural diversity within a family of metabolic and regulatory
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Keeling, C. I., Bearfield, J. C., Young, S., Blomquist, G. J., & Tittiger, C. (2006). Effects
of juvenile hormone on gene expression in the pheromone-producing midgut of
the pine engraver beetle, Ips pini. Insect Mol Biol, 15(2), 207-216. doi: IMB629
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pheromone biosynthesis in the pine engraver beetle, Ips pini (Coleoptera:
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Krebs, H., & Weitzman, P. (1987). Krebs' citric acid cycle: half a century and still turning.
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Lowenstein, J. (1969). Methods in Enzymology (Vol. 13). Boston: Academic Press. 728
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Paine, T., Millar, J., Hanlon, C., & Hwang, J. (1999). Identification of semiochemicals
associated with Jeffrey pine beetle, Dendroctonus jeffreyi. J. Chem. Ecol., 25,
433-453. 122
Pierce, H. D., Conn, J. E., Oehlschlager, A. C., & Borden, J. H. (1987). Monoterpene
metabolism in female mountain pine beetles, Dendroctonus ponderosae
Hopkins, attacking ponderosa pine. J. Chem. Ecol., 13(6), 1455-1480.
Pureswaran, D. S., Gries, R., Borden, J. H., & and Pierce, H. D. (2000). Dynamics of
pheromone production and communication in the mountain pine beetle,
Dendroctonus ponderosae Hopkins, and the pine engraver, Ips pini (Say)
(Coleoptera: Scolytidae). Chemoecology, 10, 153-168.
Reed, J. R., Vanderwel, D., Choi, S., Pomonis, J. G., Reitz, R. C., & Blomquist, G. J.
(1994). Unusual mechanism of hydrocarbon formation in the housefly:
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tricosene and CO2. Proc Natl Acad Sci USA, 91(21), 10000-10004.
Rule, G., & Roelofs, W. (1989). Biosynthesis of sex pheromone components from
linolenic acid in arctiid moths. Archives of Insect Biochemistry and Physiology,
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expression of a bark beetle cytochrome P450 that hydroxylates myrcene to
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Vanderwel, D., Gries, G., Singh, S. M., Borden, J. H., & and Oehlschlager, A. C. (1992).
(E)- and (Z)-6-nonen-one biosynthetic precursors of endo- and exo-brevicomin in
two bark beetles(Coleoptera: Scolytidae). J. Chem. Ecol., 18(8), 1389-1404.
Vanderwel, D., & Oehlschlager, A. C. (1992). Mechanism of brevicomin biosynthesis
from (Z)-6-nonen-2-one in a bark beetle. Journal of the American Chemical
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Vaz, A. H., Jurenka, R. A., Blomquist, G. J., & Reitz, R. C. (1988). Tissue and chain
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411-446.
VIII. Figure legends:
Figure 4.1. Possible roles for ZnoDH in exo-brevicomin biosynthesis. 10:1, 16:1 and 18:1 represent unsaturated fatty acids. Enzymes catalyzing the proposed reactions have not been identified, and uncharacterized steps are shown by dashed arrows.
124
Figure 4.2. Sequence and translation of ZnoDH cDNA. The encoded amino acid sequence of ZnoDH is shown above the cDNA sequence, and the position cDNA is indicated to the left. The conserved residues YXXXK, characteristic of a substrate binding site, and GXXXGXG, typical of a coenzyme binding site, are underlined.
Figure 4.3. qRT-PCR analysis of ZnoDH mRNA levels. (A) Relative ZnoDH mRNA levels in developmental groups, with levels in pre-merged males set at one. (B) Relative
ZnoDH mRNA levels in different tissues of unfed males, with levels in the posterior midgut set at one. The expression of ZnoDH is normalized to the expressions of
Ubiquitin and tubulin. Values are means ± standard error, n = 3 for developmental groups and n=4 for the tissue distribution study.
Figure 4.4. Western blot analysis of ZnoDH. (A) Recombinant production in Sf9 cells. 1: cell lysate supernatant from recombinant ZnoDH and 2: Sf9 cells without recombinant
DNA; 3: microsome pellet and 4: the supernatant after spinning 120,000 x g for 2 hours.
(B, C) Tissue distribution of ZnoDH in vivo. Ponceau staining (B), and western blot (C) of the same gel. H, head; FB, fat body; AM, anterior midgut; PM, posterior midgut; MT, malpighian tubule; SO, sex organ; C, carcass. ZnoDH localizes predominantly in the fat body.
Figure 4.5. GC-MS analysis at mass to charge (m/z=46) of (6Z)-non-6-en-2-one formed from penta-deuterium labeled nonen-2-ol by recombinant ZnoDH in the presence of
NAD+. Reactions contained NAD+ and penta-deuterium labeled nonen-2-ol with cell lysate supernatant (A) boiled recombinant ZnoDH, (B) recombinant CPR and (C) recombinant ZnoDH. (D), the standard of penta-deuturium labeled (6Z)-non-6-en-2-one 125
(18.36min); (E), the standard of penta-deuterium labeled nonen-2-ol at 21.79 min; (F),
the mass spectrum from the 18.28 min peak in C; (G), the mass spectrum of the 18.36
min peak in D. Substrate in arrow and product in arrowhead.
Figure 4.6. GC-MS analysis (m/z=46) of (6Z)-non-6-en-2-one formed from penta- deuterium labeled nonen-2-ol in the presence of NADP+. Reaction contains NADP+ and
penta-deuterium labeled nonen-2-ol with cell lysate supernatant (A) boiled recombinant
ZnoDH, (B) recombinant CPR, (C) recombinant ZnoDH. (D), the standard of penta-
deuturium labeled (6Z)-non-6-en-2-one (18.36min); (E), the standard of penta-deuterium
labeled nonen-2-ol (21.79 min); (F), the mass spectrum from 18.27 min peak in C; (G)
the mass spectrum from 18.36 min peak in D. Substrate in arrow and product in
arrowhead.
Figure 4.7. The percentage of substrate converted to the product by recombinant
ZnoDH and CPR during different incubation times in the presence of NAD+ (A) or NADP+
(B). There were two replicates at the120 min time point.
126
C16:1/C18:1
β-oxidation
Decarboxylation? C10:1
ZnoDH? P450? OH ZnoDH?
O ZnoDH?
(6Z)-non-6-en-2-one
CYP6CR1
O O
ZnoDH?
O O
exo-Brevicomin
Figure 4.1 127
Table 4.1
Primer name Sequence Amplicon length (bp)
Sequencing DPG022F04 MPB22F4-F1 GCGGATAATGACTGGTGATG n.a. MPB22F4-F2 TTCTCGATACAGAGCCCAGG n.a. MPB22F4-F3 TTCACCTGGCACACTCCG n.a.
qRT-PCR MPB22F4-RTF1 GTGGTTTGTTTCCTGCTTGGT MPB22F4-RTR1 TTGCCGTTTTGCCCACTA 95 n.a, not applicable 128
1 TTCCTTATAATTGCTCAACAGTCTTCTTTCTAACATCATCTTTTAATTTTTTCTCTTAAT M H F I L W F V S C L V L L K V V N A L 61 ATGCATTTTATTTTGTGGTTTGTTTCCTGCTTGGTTTTGTTGAAAGTGGTAAATGCATTG T C F W C K S K V C L V G K T A I I T G 121 ACGTGTTTCTGGTGCAAAAGCAAAGTGTGTTTAGTGGGCAAAACGGCAATCATTACAGGA G A A G I G F Q T A L A L A S K G C R V 181 GGGGCCGCAGGTATTGGTTTCCAGACCGCACTAGCTCTAGCTTCCAAAGGATGTCGGGTG I I A D I S N L T K A V D E L K N V S K 241 ATTATAGCTGATATTTCCAACTTAACTAAAGCAGTGGATGAATTGAAAAATGTTTCGAAA N Q N I I G V E V D L A S F R S V R E F 301 AATCAAAATATCATTGGTGTAGAAGTTGATCTGGCATCTTTTAGATCCGTTAGGGAATTT A K K I L D T E P R L D I L V C N A G I 361 GCGAAAAAAATTCTCGATACAGAGCCCAGGCTCGATATATTGGTTTGTAATGCAGGTATC G E H K M R I M T G D G V E K T M Q I N 421 GGTGAACACAAAATGCGGATAATGACTGGTGATGGGGTAGAAAAAACTATGCAAATTAAT Y Y S N F L M I H L L L D L L K K S A P 481 TACTACAGTAATTTCTTAATGATTCATTTGCTGTTGGATTTGTTGAAAAAATCGGCACCA S R I A I T S S V L A Y V S D L T V K E 541 AGTAGAATAGCCATAACAAGCTCCGTACTAGCATACGTTTCCGACTTGACTGTAAAGGAA L N P S P D Y F E E R K G W R T F N C G 601 CTAAATCCTAGTCCTGATTATTTCGAAGAGAGGAAAGGGTGGAGGACGTTTAACTGCGGA T Y S A S K L C L A G L T K T L A S K L 661 ACATACAGTGCGTCCAAATTATGTCTGGCAGGCTTAACTAAGACGCTCGCCTCAAAACTC Q G T E V I V N V C H P N G V R T S I Y 721 CAAGGAACGGAAGTGATCGTGAATGTTTGTCATCCCAATGGCGTGAGAACTTCAATTTAT A P V V R Q A I G L S Y F V S S L Y C L 781 GCTCCAGTAGTACGACAAGCGATAGGACTTTCGTATTTTGTATCGAGTTTGTATTGCCTT V C L F G K T P E E G A Q T L I H L A H 841 GTCTGTCTCTTTGGAAAAACCCCAGAAGAAGGAGCTCAAACATTAATTCACCTGGCACAC S E E V K L I N G R S F Y E G M V V H P 901 TCCGAAGAAGTGAAACTAATAAATGGTCGATCATTTTACGAGGGAATGGTTGTTCATCCA P F Q L T D Y F C E Q L W K A S I E Y T 961 CCTTTCCAACTAACTGATTATTTCTGCGAACAGCTGTGGAAGGCTTCCATTGAATACACC K L T P E E I K C * 1021 AAGCTTACTCCTGAGGAAATTAAGTGTTAATTAGATATGAAAATGAACTCAATTGAATAC 1081 CTAAAGTAAATACTTTA
Figure 4.2 129
50 45 40 35 30 25 20 15 10
) 5 0 s.e.
± - ddCT
16 14 12 10 Relative Expression Relative(2 Expression 8 6 4 2 0 head carcass fat body anterior midgut posterior midgut hindgut
Figure 4.3 130
(A) 1 2 3 4
36 kDa
kDa H FB AM MG PM MT SO C (B)
75 50
25
(C)
36
Figure 4.4 131
Product Substrate
21.79 (A) Boiled DH & NAD+ & Substrate
(B) CPR & NAD+ & Substrate 21.79
18.27 18.28 (C) DH & NAD+ & Substrate 21.78
(D) Std of (6Z)-non 18.36
Relative Abundance -6-en-2-one
21.79 (E) Std of nonen-2-ol
16.0 17.0 18.0 19.0 20.0 21.0 22.0 23.0 24.0 Time (min)
100 67.0 (F) 18.28
46.2
75.0 89.0 99.0 116.0 129.9 55.1 145.9 0
100 67.0 (G) 18.36
Relative Abundance 46.2
55.1 75.0 89.0 99.0 116.0 130 145.9 0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 m/z
Figure 4.5 132
Product Substrate
100 (A) Boiled DH & NADP+ & Substrate 21.79
0 100 (B) CPR & NADP+ & Substrate 21.80
18.28 0 18.27 100 (C) DH & NADP+ 21.80 & Substrate
0 100 18.36 (D) Std of (6Z)-non-
Relative Abundance 6-en-2-one 0 100 21.79 (E) Std of nonen-2-ol
0
16.0 17.0 18.0 19.0 20.0 21.0 22.0 23.0 24.0 Time (min)
100 67.0 (F) 18.27
46.2
55.1 75.0 89.0 99.0 116.0 130.0 145.0 0 100 67.0 (G) 18.36
46.2 Relative Abundance
55.1 75.0 89.0 99.0 116.0 130.0 145.9 0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 m/z
Figure 4.6 133
100 + 90 (A) NAD 80 70 60
50 ZnoDH 40 CPR 30 20 10 0 0 60 120 180 240
80
+ 70 (B) NADP
60
50
40 Percentage of substrate converted to product to converted substrate of Percentage ZnoDH 30 CPR
20
10
0 0 60 120 180 240 Time (min)
Figure 4.7 134
Chapter 5
Monoterpene-metabolizing P450s in pine bark beetle pheromone biosynthesis and
resin detoxification
I. Abstract II. Introduction III. Materials and Methods IV. Results V. Discussion VI. Acknowledgements VII. References VIII. Figure Legends
I. Abstract
A bark beetle’s successful attack is partially contributed by rapidly metabolizing toxic monoterpenes from the host tree and using some of these metabolic products as pheromones. In doing so, some detoxification enzymes may have evolved into pheromone biosynthetic enzymes. Evidence for this evolutionary relationship may be found by comparing activities of monoterpene-hydroxylating enzymes. Microarray analysis from the MPB showed that a cytochrome P450, CYP6DH2, is highly expressed in both males and females and induced by monoterpenes, and is thus likely a detoxification gene. Its paralog, CYP6DH1, having 16 amino acids different from
CYP6DH2, is highly expressed in female hindguts and induced by feeding, but not monoterpenes. CYP6DH1 likely acts as a pheromone biosynthetic gene. Functional recombinant CYP6DH1, CYP6DH2 and CYP9T2 were produced in Sf9 (insect) cells using a baculovirus expression system. Enzyme assays with various monoterpenes showed that CYP6DH1 accepted (+) and (-)-α-pinene as substrates to produce verbenol, and that CYP6DH2 and CYP9T2 accepted the same substrates: (+)-, (-)-α-pinene, Δ-3- 135
carene and Limonene, and in addition CYP9T2 accepted myrcene. These data support
that CYP6DH1 may be a pheromonal trans-verbenol biosynthetic enzyme sharing a
common ancestor with CYP6HD2, and that both CYP6DH2 and CYP9T2 are
detoxification enzymes because of their broad substrate ranges. CYP9T2 preferred
myrcene over (+)-α-pinene and Δ-3-carene, supporting that CYP9T2 originally worked as detoxification enzyme and became a pheromone biosynthetic enzyme over time.
Both CYP6DH2 and CYP9T2 converted the same substrates into different products, suggesting a similarity and difference of their structures.
II. Introduction:
The mountain pine beetle (MPB), Dendroctonus ponderosae Hopkins, is the
most lethal pest of coniferous forests in the North West America. Its ability to
successfully colonize and kill trees is evidently due in part to the ability of the pioneer
female beetles to metabolize host monoterpenes efficiently and to produce aggregation
pheromones. Pioneer females initially attack the host tree, and metabolize the tree α-
pinene ((1S, 5S)-2,6,6-trimethyl bicyclo[3.1.1]hept-2-ene) to produce trans-verbenol
(4,7,7-trimethylbicyclo[3.1.1]hept-3-en-2-ol) (Hughes, 1973) (Figure 5.1), which attracts
both males and females (Billings et al., 1976; Libbey et al., 1985; Pitman et al., 1968;
Pitman et al., 1969). During the early phases of attack, pine trees produce a defensive
resin containing toxic mono-, sesqui- and diterpenoid chemicals to pitch out beetles
(Phillips & Croteau, 1999; Steele et al., 1995). A healthy tree can produce sufficient
resins to kill the beetles. However, under stressful conditions (e.g. drought), trees may
not produce enough resins to stop the infestation (Kurz et al., 2008; Rudinsky, 1996).
The predominant components of resins from host tree are monoterpenes. To survive in
this toxic monoterpene environment (Smith, 1964, 1966), these beetles have developed 136 enzyme systems that metabolize a wide range of toxic host substances rapidly and efficiently to compounds that are less toxic and/or more easily excretable (Bernhardt,
2006; Brattsten et al., 1977; Hunt & Smirle, 1988) by non-specific oxidation (Hughes,
1973; Hughes, 1973; Renwick & Hughes, 1975) or oxidize monoterpenes to pheromones by highly specific processes (Byers, 1983; Klimetzek & Ffrancke, 1980;
Renwick et al., 1976).
Current pest management strategies have proven ineffective to regulate pine bark beetles. Future control strategies may benefit from species-specific targets.
Pheromone biosynthesis is an attractive focus area because pheromone components are often species-specific. In addition, understanding how pheromone production evolved can help us understand and predict how bark beetles will respond to management efforts. Unfortunately, very few confirmed pheromone-biosynthetic and monoterpene metabolic enzymes have been identified in bark beetles. Bark beetle pheromone biosynthesis likely evolved from existing resin detoxification reactions
(Byers, 1983; P. Hughes, 1973; Pierce et al., 1987; Tillman et al., 1999). Some pheromone components are the same as monoterpene metabolites, suggesting the evolutionary path by which hydroxylated monoterpenes became aggregation pheromone components (Blomquist et al., 2010). For example, pheromonal verbenol is derived from
α-pinene, which is likely ingested from the host tree (Rao et al., 2003; Renwick et al.,
1976). Ipsdienol, an aggregation pheromone component in I. pini, is derived from myrcene (Fish et al., 1984; Seybold & Tittiger, 2003), which is both ingested and produced de novo (Blomquist et al., 2010; Lu, 1999). The enzyme systems that detoxify monoterpenes may have evolved a secondary biological role as “pheromone biosynthetic” enzymes that hydroxylate specific monoterpenes to produce pheromones
(Blomquist et al., 2010). It is likely that monoterpenoid alcohols provided evolutionary 137
pressure for these beetles to sense and respond to their presence and meaning, and
later, beetles would have developed the ability to control and regulate the synthesis of
pheromonal monoterpenoid alcohols (Blomquist et al., 2010; Seybold & Tittiger, 2003).
Pheromone biosynthetic enzymes should be distinguishable from detoxification
enzymes based on enzyme activity and regulation. A detoxification enzyme may have a
broad substrate range and be induced by its substrates, while a pheromone biosynthetic
enzyme may have a strong preference for the precursor and perhaps be regulated by
hormones or feeding. For example, I. pini CYP9T2, a pheromone-biosynthetic enzyme
(Sandstrom et al., 2006), which is not induced by its substrate, myrcene (A. Grifffith,
unpublished data), also hydroxylates α-pinene to verbenol (Sandstrom, 2007). This may
be evidence for this evolutionary scheme. The two enzyme systems can be closely
related through mutation or duplication. Alternatively, detoxification and pheromone
production may be accomplished rapidly and simultaneously using the same enzyme. It
may be more energy efficient for beetles to use the metabolic products from
detoxification as pheromone than to produce it de novo. However, there would be less
control over the amount of pheromone produced.
Monoterpene metabolizing cytochromes P450 have been studied in several
organisms. For example, recombinant human CYP2B6, CYP2C19 and CYP2D6
hydroxylate Δ-3-carene into Δ-3-carene-10-ol, while human CYP1A2 converts Δ-3- carene to Δ-3-carene-epoxide (Duisken et al., 2005). The hydroxylation of limonene with several plant cytochrome P450 limonene hydroxylases has been characterized (Karp et
al., 1987; Karp et al., 1990; Kjonaas et al., 1985; Lupien et al., 1999). Δ-3-Carene
metabolism by larvae of the common cutworm (Spodoptera litura) suggested a
cytochrome P450 was involved (Miyazawa & Kano, 2010). Cytochromes P450 involved
in monoterpene metabolism in the bark beetles have been proposed (Pierce et al., 1987; 138
Sandstrom, 2007; White et al., 1979; White et al., 1980), but relatively little is known about these enzymes in bark beetles. Hunt et al. (1988) demonstrated that cytochromes
P450 are involved in α-pinene and myrcene metabolism in MPB. Blocking P450 activity with piperonyl buxtoxide resulted in deceased levels of certain terpene alcohols, along with a buildup in the levels of their monoterpene precursors (Hunt & Smirle, 1988). The only other bark beetle P450s with known function are CYP9T1 and CYP9T2, from I. confusus and I. pini, respectively. Both enzymes hydroxylate myrcene to ipsdienol
(Sandstrom et al., 2006, 2008). CYP9T2 also hydroxylates α-pinene to verbenol
(Sandstrom, 2007).
P450s with detoxification roles in D. ponderosae and I. pini remain unknown.
Comparing the substrate profiles of bark beetle cytochromes P450s in D. ponderosae and I. pini should provide molecular information about the evolution of pheromone production and address the hypothesis that the pathways for monoterpene detoxification and monoterpenoid pheromone biosynthesis are evolutionarily related (Greis et al.,
1990; Seybold et al., 2000; Vanderwel & Oehlschlager, 1987). In this study, we investigated the role of P450s in pheromonal trans-verbenol biosynthesis in MPB and detoxification in MPB as well as I. pini.
trans-Verbenol is an aggregation pheromone produced by female MPBs
(Hughes, 1973; Hughes, 1973; Libbey et al., 1985; Pitman et al., 1968). Feeding results in increased trans-verbenol levels (Pierce et al., 1987; Pureswaran et al., 2000). It is most likely produced via cytochrome P450-mediated hydroxylation of host tree α-pinene even though there is evidence supporting de novo biosynthesis in D. frontalis (Renwick et al., 1973). De novo biosynthesis via the mevalonate pathway may occur in D. ponderosae as well (C. Keeling, personal communication) although there is no published evidence supporting this idea. 139
While the observation that cytochromes P450 are not necessarily important for
the enantiomeric ratios of pheromonal ipsdienol (Sandstrom et al., 2008; Sandstrom et
al., 2006) contradicts earlier predictions (Renwick et al., 1976; Seybold et al., 2000),
there is strong indirect evidence supporting stereo-selective P450-mediated reactions in
the case of α-pinene hydroxylation in Dendroctonus spp. (Greis et al., 1990). Female D.
ponderosae appear to have two distinct α-pinene hydroxylating pathways. The first
pathway does not discriminate between α-pinene enantiomers and likely plays a
detoxification role, while second pathway appears specific for (-)-α-pinene and therefore
produces the bulk of the pheromone component, (-)-trans-verbenol (Pierce et al., 1987).
The P450s in the two pathways may be closely related. (-)-trans-Verbenol, not the (+)-
isomer, is the major female-aggregation pheromone in D. ponderosae (Borden et al.,
1987; Libbey et al., 1985). It is derived from (-)-α-pinene in the MPB (Pierce et al.,
1987), while (+)-trans-verbenol is likely produced from (+)-α-pinene because female
MPBs produced trans-verbenol when exposed (-)- and (+)-α-pinene (Greis et al., 1990) and male and female D. brevicomis produced (+)-trans-verbenol and (-)-trans-verbenol respectively when exposed to (+)-α-pinene and (-)-α-pinene (Byers, 1983). cis-Verbenol is also considered as an aggregation pheromone in the MPB (Miller & Lafontaine, 1991).
It is possibly derived from both (+) and (-)-α–pinene because male and female D. brevicomis produced cis-verbenol from both enantiomers when exposed to (+)-α-pinene and (-)-α-pinene (Byers, 1983). The P450s involved in trans-verbenol biosynthesis in
MPB are still not identified.
Functional genomics have been successfully applied to identify pheromonal ipsdienol biosynthetic enzymes in I. pini (Figueroa-Teran et al., 2011; Keeling et al.,
2006; Keeling et al., 2004; Sandstrom et al., 2006). Monoterpenes such as α-pinene and myrcene are potent and rapid inducers of P450 in certain insects (Brattsten et al., 140
1977). Such induction may also occur in D. ponderosae and facilitate the rapid detoxification of host monoterpenes. Therefore, screening P450 gene expression by microarray analysis in the MPB may reveal candidates for detoxification and/or trans- verbenol biosynthesis. This information offers great advantage to identify and assign
P450 functions by combining expression analysis and functional assays of expressed recombinant proteins.
Aw et al. (2010) showed that CYP6DH2 is highly expressed in both males and females, and Gorzalski (2010) showed CYP6DH2 is induced when adults are exposed to a monoterpene-saturated environment, suggesting its detoxification role. Its paralog,
CYP6DH1, with 16 amino acids different, is highly expressed in female hindguts and induced by feeding, but not by monoterpenes (Gorzalski, 2010), implying that CYP6DH1 likely acts as a pheromone biosynthesis gene. We hypothesized that CYP6DH1 is involved with trans-verbenol biosynthetic pathway (Figure 5.1), CYP6DH2 is responsible for resin detoxification, and both may share the same substrate, such as (-)-α-pinene, but CYP6DH2 may have a broad substrate range (Figure 5.1). Comparison of monoterpene substrate profiles from CYP6DH2 and CYP6DH1 as well as CYP9T2 can help understand the evolutionary relationship between detoxification and pheromone biosynthesis, and can also help understand structural elements that affect substrate docking.
Recombinant CYP6DH1, CYP6DH2 and CYP9T2 were produced with a baculovirus expression system in Sf9 cells. Enzyme assays with various monoterpenes coupled with GC-MS analysis showed that CYP6DH1 only accepted (+)- and (-)-α – pinene and produced verbenols, and CYP6DH2 accepted (+)-, (-)-α -pinene, Δ-3-carene and R-(+)-limonene, but not other tested monoterpenes. CYP9T2 had an identical substrate profile to CYP6DH2, but with the addition of myrcene as a substrate. 141
Interestingly, the two P450 families converted the same substrates into different products, implicating the similarity and difference of their structures. These data can readily be interpreted to indicate that CYP6DH1 plays a pheromone biosynthetic role, while CYP6DH2 is involved in detoxification. CYP6DH1 and CYP9T2 likely evolved from common ancestor.
III. Materials and methods
Reagents and chemicals. Hink’s 1x TNM-FH Medium (Supplemented Grace’s
Medium) and Grace’s 1x Insect Basal Medium were from Mediatech, Inc. (Herndon, VA) and FBS was from Atlas Biologicals (Fort Collins, CO). Direct baculovirus DNA kit, SF-
900 II (1X) and (1.3X) media were from Invitrogen (Carlsbad, CA). Sf9 cells were a gift from B. Perrino (Department of Physiology, University of Nevada, Reno). Housefly cytochrome P450 reductase viral clone (Wen et al., 2003) was kindly provided by M.
Schuler (U. Illinois at Urbana-Champaign). The CYP6DH1 plasmid clone was provided by C. Keeling (University British Columbia, BC, Canada). Agarose was from Bio- express (Keysville, UT). Phenylmethylsulfonyl fluoride (PMSF), (+)-, (-)-α-pinene, Δ-3- carene, α-phellandrene, β-pinene, γ-terpinene, myrcene, terpinolene, R-(+)- limonene, protease Inhibitor cocktail, δ-aminolevulinic acid and ferric citrate were from Sigma-
Aldrich (St. Louis, MO). 96-well microplates were purchased from Greiner Bio-One
(Monroe, North Carolina).
Recombinant protein production. Protocols for growth and maintenance of Sf9 cells, recombinant baculovirus construction, and heterologous expression using the
BaculoDirectTM Expression Kit were as described by Invitrogen. CYP6DH1, CYP6DH2 and CYP9T2 baculoviral clones were produced by L. Plaugher, A. Gorzalski (Gorzalski,
2010) and P. Sandstrom (Sandstrom, 2007) respectively. Briefly, an LR recombination 142 reaction between each pENTR4 recombinant clone and BaculoDirect Linear DNA produced recombinant baculoviral P450 clones that were transfected separately into Sf9 cells and grown in the presence of 100 µM gancyclovir to select for recombinant virus.
High titer P3 viral stock was produced by successive 72-hour amplifications of the initial and P2 stocks. Approximate viral titer was determined by a plaque assay as described by Invitrogen. The viral stock was used to infect Sf9 cells at 1.0 x 106 cells/ml in a disposable shaking flask (Bio-Express, Keysville, UT). The heme precursor, δ- aminolevulinic acid (0.3 mM final conc.) and ferric citrate (0.2 mM final conc.) were added at the time of infection. To produce recombinant CYP6DH1, CYP6DH2, CYP9T2 and housefly reductase (CPR), Sf9 cells were infected with recombinant baculovirus at multiplicities of infection (MOI) = 0.5 for CYP6DH1, 0.025 for 6DH2, 0.4 for CYP9T2, and
0.1 for (CPR) as described previously (Chapter 3). All recombinant proteins were produced separately. Sf9 cells were cultured at 27ºC with shaking flask and all samples were harvested on day 3 post infection (PI).
Microsome preparation. Sf9 cells and Sf9 cells producing recombinant CYP6DH1,
CYP6DH2, CYP9T2 and CPR were harvested at day 3 PI, and microsomes were prepared by differential centrifugation essentially as per Wen et al. (2003). Functional
CYP6CR1 (See above, Chapter 3) was produced as negative control (not shown).
Briefly, cells were pelleted by centrifugation at 3000 x g at 4 ºC for 10 min. The pellets were resuspended in 1/5 cell culture volume of 100 mM ice-cold sodium phosphate buffer (pH 7.8) and repelled twice at 3000 x g for 10 min. The pellets were resuspended in 1/30 cell culture volume of ice-cold cell lysate buffer (100 mM sodium phosphate pH
7.8, 1.1 mM EDTA, 0.1 mM DTT, 0.5 mM PMSF, 1/1000 vol/vol Sigma protease inhibitor cocktail, 20% glycerol) and lysed by sonication three times for 15 s on ice with a
Branson Sonifier 450, followed by vortexing for 15 seconds. The lysate was centrifuged 143 at 10,000 x g for 20 min at 4ºC in a micro-centrifuge and the supernatant was either further centrifuged in a TLA110 rotor at 120,000 x g for 2 h in a Beckman-Coulter Optima ultracentrifuge to pellet microsomes or used directly for CO spectral assay. The microsomal pellet was resuspended in 1/30 cell culture volume of ice-cold cell lysate buffer cold cell lysate buffer and used immediately.
CO microplate assay. Functional P450 concentrations from recombinant CYP6DH1,
CYP6DH2 and CYP9T2 Sf9 microsomes were determined by carbon monoxide (CO)- difference spectrum analysis (Omura & Sato, 1964) using a 96-well microplate and
SpectraMax M5 Microplate Reader coupled with SoftMax® Pro software (Molecular
Devices, Inc., Sunnyvale, CA) essentially as per Choi et al., (2003). Briefly, 200 µl of microsome solution from reach group was loaded into replicate wells for reference and
CO treatment samples. The reference wells were tightly sealed with paper tape, and the plate was placed in a plastic chamber. CO gas was perfused into the top of the chamber and out from the bottom at 0.5 l/min for 3 min. All of samples were reduced by adding
10 µl fresh 0.5 M sodium hydrosulfite to 25 mM final concentration (Choi et al., 2003).
The absorbances of the samples from 400 nm to 500 nm were measured with
SpectraMax M5 Microplate Reader. The P450 concentration was calculated using the following formula:
[P450] (mM) =1.9* (∆450-∆490)/91, (Equation 5.1)
where ∆450 and ∆490 are the absorbance differences between the CO sample and reference sample at 450 nm and 490 nm respectively (Omura & Sato, 1964), and 1.9 represents the molar extinction coefficient based on the depth of 200 µl with a 6.96 mm diameter well relative to 1cm light pathway. The total protein concentration in each 144
microsome sample was quantified by BCA assay kit as recommended by the protocol
(Thermo Scientific, Rockford, IL).
Enzyme assays. Enzyme assays were conducted in 500 µl reactions containing 468 µl
or 250 µl of a 4:1 mixture of microsomes bearing recombinant P450 or CPR as
described previously (Chapter 3), 200-300 µM monoterepene in pentane solvent, and
300 µM NADPH or 250 µl 2X NADPH regenerating system. Reactions were initiated
with the addition of NADPH or NADPH regeneration system, incubated in a 30 ºC water
bath for 30 min to 6 hours, and then extracted twice with pentane: ether (1:1) spiked with
5-100 µg/ml n-octanol (internal standard). The organic phase was concentrated to
approximately 100 µl with N2 gas and directly analyzed by coupled GC-MS at the
Nevada Proteomics Center (UNR). Negative controls included reactions run with
microsomes prepared from cells infected with recombinant CPR baculovirus only, or
microsomes from cells with recombinant CYP6CR1 and CPR baculoviruses (Chapter 3).
A Thermo Finnigan Polaris Q ion trap was used with a molecular weight scanning range
of 40–180 atomic mass unit (amu) at an ionization potential of 70 eV. A trace gas chromatograph containing a 60 m x 0.25 mm (ID), 0.25 µm film thickness DB-5 capillary column (J&W Scientific, Palo Alto, CA) was programmed for an initial temperature of 50
ºC (1 min hold), increase to 200 ºC at 5 ºC/min, 10 ºC/min to 320 ºC (20 min hold). The injector was split at a ratio of 100:1 at a temperature of 280 ºC with a column flow of 1.5 ml He/min. The detector was set at 200 ºC. The product was identified by comparing retention times and mass spectra with negative controls and an authentic standard.
For the competition experiments, a mixture of (+)-α-pinene (256 µM) and myrcene (264 µM) or Δ-3-carene (256 µM) and myrcene (264 µM) in final concentration respectively were added to recombinant CYP9T2-CPR microsomes. The 2X NADPH regeneration system was added to initiate the reactions. The enzyme assays were 145 incubated in a 30ºC water bath for three or six hours and extracted as described above.
A mixture of 4:1 Sf9 cells microsomes: Sf9 cells microsomes bearing CPR was used as a negative control. The percentage of substrate conversion was calculated according to areas of product and substrate by dividing the amount of product by the total amount of products and substrates (product/(products + substrates)).
IV. Results
Functional expression. CO-difference absorption spectra of microsomes from Sf9 cells producing recombinant CYP6DH1, CYP6DH2 and CYP9T2 harvested on 3 days PI produced characteristic 450 nm peaks (Figure 5.2A, B, and C) respectively, as did
CYP6CR1 (not shown), but not microsomes from uninfected Sf9 cells or from Sf9 cells producing recombinant CPR (not shown). The concentration of functional recombinant
P450 was approximately 6.4 X10-4 mM for CYP6DH1 (Figure 5.2A), 3.173 X10-4 mM for
CYP6DH2 (Figure 5.2B) and 4.73 X10-4 mM for CYP9T2 (Figure 5.2C) according to
Equation 5.1. The 420 nm peak is much bigger from recombinant CYP6DH1 and much smaller from both CYP6DH2 and CYP9T2 compared to the 450 nm peak (Figure 5.2).
Enzyme Assays. P450s and CPR were expressed separately. All assays were performed with a mixture of four parts P450 and one part of CPR Sf9 cell microsomes.
The substrates and products from various reactions are summarized in Table 5.1.
Recombinant CYP6DH1 converted (-)-α-pinene to a product at 24.6 min (Figure 5.3B).
GC-MS analyses showed that this product had a retention time and mass spectrum
(Figure 5.3D) identical to the standard of trans-verbenol at 24.6 min (Figure 5.3C, E).
The product was not detected in reactions containing recombinant CPR alone (Figure
5.3A). Similarly, a product at 24.61 min was produced from the incubation of CYP6DH1 146
with (+)-α-pinene (Figure 5.4B) with a similar retention time to the standard of cis-
verbenol at 24.55min (Figure 5.4C), or trans-verbenol (Figure 5.3C). MS analysis
showed that the product (Figure 5.4D) had a mass spectrum identical to the standard of cis-verbenol and trans-verbenol (Figure 5.4E, 5.3E). This product was not present in reactions with recombinant CPR only (Figure 5.4A). In a single trial, recombinant
CYP6DH1 did not accept Δ-3-carene, α-phellandrene, β-pinene, γ-terpinene, myrcene, terpinolene and R-(+)-limonene as substrates (not shown).
Recombinant CYP6DH2 hydroxylated (+)-α-pinene to yield a product that had a
similar retention time (27.8 min) (Figure 5.5B) and mass spectrum (Figure 5.5D)
identical to the myrtenol standard (Figure 5.5C, 5.5E). This assay confirmed the
preliminary result from Gorzalski (2010) that CYP6DH2 produced myrtenol when
incubated with (+)-α-pinene. The same enzyme preparation incubated with (-)-α-pinene
yielded a product with a similar retention time (24.59 min) (Figure 5.6B) and identical
mass spectrum (Figure 5.6D) to the standard of trans-verbenol (24.6 min) (Figure 5.6C;
5.3E). Recombinant CYP6DH2 converted Δ-3-carene and R-(+)-limonene to three unknown products that eluted at 20.11, 22.59, and 25.04 min (Figure 5.7B ), and 20.11,
22.57 and 25.01 min (Figure 5.8B), respectively. None of these products were detected in reactions run with recombinant CPR (negative control, Figure 5.5A; 5.6A; 5. 7A; 5.8A).
The retention time and mass spectra of the three products from limonene (Figure 5.8B) were similar or identical to the products from Δ-3-carene (Figure 5.7B). No products were observed in reactions of recombinant CYP6DH2 incubated with α-phellandrene, β- pinene, γ-terpinene, myrcene or terpinolene (not shown, n=2). The stock Δ-3-carene substrate contained about 1.9% limonene, but limonene did not show Δ-3-carene contamination (not shown). 147
Recombinant CYP9T2 converted (+)-α-pinene to a product with a retention time
similar (24.57 min) (Figure 5.9B) and mass spectrum (Figure 5.9D) identical to trans- verbenol at (24.6 min) (Figure 5.9C; 5.9E). Reactions with (-)-α-pinene yielded a product at 27.69 min (Figure 5.10B) with a similar retention time and mass spectrum (Figure
5.10C) identical to the myrtenol standard at 27.74 min (Figure 5.10C; 5.5E). A single product of CYP9T2-catalyzed reactions with Δ-3-carene eluted at 26.71 min (Figure
5.11B). Similarly, CYP9T2-catalyzed reactions with R-(+)-limonene yielded a single product that eluted at 26.76 min (Figure 5.12B). The mass spectra of products from Δ-3 carene and limonene were identical (Figure 5.11C; 5.12C). Negative controls for these substrates were incubated with recombinant CPR (Figures 5.9A; 5.11A and 5.12A) or
CYP6CR1-CPR (not shown) and did not yield any products, with the exception of a small
27.67 min peak from reactions of recombinant CPR incubated with (-)-α-pinene (Figure
5.10A).
To test CYP9T2 substrate preferences, enzyme assays were performed with the mixture of (+)-α-pinene and myrcene (Figure 5.13) or Δ-3-carene and myrcene (Figure
5.14) and the 2X NADPH regeneration system, and incubated for 3 hours or 6 hours.
Increasing the incubation time resulted in an increase of product. The percentage of substrate converted to product was calculated according to the areas of peaks of products and substrates. For 6-hour incubations, the reaction with the mixture of (+)-α- pinene and myrcene converted about 60.5% (+)-α-pinene to verbenol and hydroxylated about 78% myrcene to ipsdienol (Figure 5.13B), while with the mixture of Δ-3-carene and myrcene, about 21.7% 3-carene was converted to an unknown product and 55.4% myrcene was hydroxylated to ipsdienol (not shown). For 3-hour incubations, reactions containing the mixture of (+)-α-pinene and myrcene converted about 25% (+)-α-pinene to verbenol and about 37.3% myrcene to ipsdienol (not shown), while with the mixture of 148
Δ-3-carene and myrcene, about 4.2% 3-carene was converted to an unknown product and 28% myrcene was hydroxylated to ipsdienol (Figure 5.14B). Negative controls with recombinant CPR did not yield these products (Figure 5.13A; 5.14A). The concentration of myrcene in these two experiments was much less than (+)-α-pinene and Δ-3-carene.
Comparison of substrate profiles from CYP6DH2 and CYP9T2 showed that they both accept (+)-, (-)-α-pinene, Δ-3-carene and limonene as substrates, and CYP9T2 also accepted myrcene (Table 5.1). Interestingly, (+)-α-pinene was converted to myrtenol by
CYP6DH2, and to verbenol by CYP9T2, while (-)-α-pinene was converted to verbenol by
CYP6DH2, and to myrtenol by CYP9T2 (Table 5.1). The three products from Δ-3- carene catalyzed by CYP6DH2 are the same as those from R-(+)-limonene. CYP9T2 converted both Δ-3-carene and limonene to identical products (Table 5.1).
V. Discussion
Beetles survive in a toxic monoterpene-rich environment (Smith, 1964, 1966) in part through efficient detoxification and pheromone biosynthesis enzyme systems.
Pierce et al. (1987) interpreted in vivo monoterpene metabolism assays to suggest three monoterpene hydroxylating “systems,” two of which were concerned with α-pinene: the
“MP-1” system was characterized as hydroxylating of monoterpenes on an allylic methyl group E to a methylene or vinyl group, while “MP-2” produces trans-verbenol from (-)-α- pinene specifically and was hypothesized to be involved in pheromonal (-)-trans- verbenol production (Pierce et al., 1987). Microarray and qRT-PCR analyses suggested
CYP6DH2 as a candidate detoxification P450 (Aw et al., 2010; Gorzalski, 2010), while its paralog, CYP6DH1, was hypothesized to play a pheromone-biosynthetic role
(Gorzalski, 2010). Similarly, CYP9T2 is an identified pheromone-biosynthetic P450 in I. pini (Sandstrom, 2006), and is hypothesized to have evolved from a detoxification 149
ancestor (Blomquist et al., 2010; Sandstrom, 2007). Comparing substrate profiles of all three P450s may explain the properties of detoxification enzymes and the evolution of pheromone biosynthesis enzymes.
Functional recombinant CYP6DH1, CYP6DH2 and CYP9T2 were produced in
Sf9 cells (Figure 5. 2A, 5.2B, 5.2C). The peak at 450 nm indicates the presence of functional P450 for each sample (Omura & Sato, 1964). The significant peak at 420 nm suggests that the protein was not completely reduced, or that a fraction of recombinant proteins did not properly fold (Lambalot et al., 1995; Wen et al., 2003). Recombinant
CYP6DH1 yielded the weakest 450 nm peak, likely reflecting the quality of the
baculoviral stock (not shown). For this reason, sufficient enzyme was available for one
trial for each substrate. Thus, data from CYP6DH1 should be considered preliminary
until confirmed.
Enzyme assays with various monoterpenes showed that CYP6DH1 only
accepted (+)- and (-)-α-pinene as substrates and produced verbenol from both
enantiomers (Figure 5.3B, 5.4B). Other tested monoterpenes were not substrates (not
shown). These products were not present in negative control reactions with recombinant
CPR only (Figure 5.3A, 5.4A), suggesting that products were from CYP6DH1 activity,
not from endogenous activity in Sf9 cells. The enantiomers of verbenol from (+)- and (-)-
α-pinene are unknown. They can be identified by comparisons with (-)-, (+)-trans-
verbenol or cis- and trans-verbenol standards using an enantioselective GC column. It
is likely that both products from (+)- and (-)-α-pinene are trans-verbenol because female
D. ponderosae exposed to both substrates produced trans-verbenol in vivo (Greis et al.,
1990). It is also possible that products from (+)- and (-)-α-pinene are (+)-trans-verbenol
and (-)- trans-verbenol, respectively, because D. brevicomis beetles produced both (+)
and (-) enantiomers when exposed to (+)-α-pinene and (-)-α-pinene, respectively (Byers, 150
1983). Because MPBs also produce a small amount of cis-verbenol (Pureswaran et al.,
2000), it is necessary to confirm if cis-verbenol was produced by this P450.
Dendroctonus brevicomis beetles produced cis-verbenol when exposed to both enantiomers of α-pinene (Byers, 1983).
CYP6DH2 also accepted these two substrates, and in addition accepted Δ3- carene and (R)-(+)-limonene (Figure 5.5-8). The narrower substrate range of CYP6DH1 compared to CYP6DH2 supports the suggestion that CYP6DH1 is important for pheromone biosynthesis. Gorzalski (2010) showed that CYP6DH1 was highly expressed in female hindguts and not induced by any monoterpenes, further supporting a pheromone biosynthetic role. In contrast, CYP6DH2 is strongly induced in beetles exposed to monoterpene atmospheres (Gorzalski, 2010) and had an apparent broad substrate range (Figure 5.5-8). CYP6DH1 may prefer (-)-α-pinene over (+)-α-pinene as its substrate. These results do not necessarily contradict a two α-pinene hydroxylating systems hypothesis (e.g. Pierce et al., 1987) if, for example, a third P450 with a narrow substrate range is active. The fact that CYP6DH2 did not hydroxylate myrcene further supports the suggestion that additional monoterpene-hydroxylating P450s remain to be discovered in D. ponderosae.
Recent replication of these experiments, preformed during preparation of this manuscript, showed that CYP6DH1 only accepted (-)-α-pinene as substrate, but not (+)-
α-pinene or other monoterpenes (not shown), and CYP6DH1 hydroxylated (-)-α-pinene to verbenol. This further confirms that CYP6DH1 may be a pheromonal trans-verbenol biosynthesis enzyme. More experiments with (+)-α-pinene need to be conducted to confirm the apparent very strict substrate selectivity of CYP6DH1. If these data are confirmed, they provide a remarkable tool to analyze structural constraints on substrate profiles because CYP6DH1 and CYP6DH2 differ by only 16 amino acids. 151
It is also interesting that CYP6DH2 specifically produced myrtenol or trans- verbenol, depending on the enantiomer of the substrate (see below), so the product profile can vary depending on the beetles’ environment. This may explain the two α- pinene hydroxylating systems hypothesis. CYP6DH2 mRNA levels are induced by (+)-α
-pinene, but not by (-)-α-pinene, and only in males, but not females (Gorzalski, 2010).
Thus, when females enter a host tree, CYP6DH1 transcription is induced, likely by
feeding, leading to the conversion of dietary α-pinene to trans-verbenol. CYP6DH2 is
not strongly transcribed in females. Males, on the other hand, would respond to only (+)-
α-pinene by inducing CYP6DH2, resulting in myrtenol production. CYP6DH1
transcription is relatively low in males (Gorzalski, 2010). This separation of metabolism,
which retains a pheromonal role for trans-verbenol production in females, while
permitting α-pinene detoxification in both sexes, is thus accomplished by a combination
of gene regulation patterns and product profiles. Competition experiments with mixtures
of these enantiomers for both enzymes may clarify this relationship.
CYP6DH2 and CYP9T2 appear to work as monoterpene metabolizing enzymes.
Both have similar broad range of substrate profiles, except that myrcene is a substrate
for CYP9T2 but not CYP6DH2. Both accepted (+)-, (-)-α-pinene, Δ-3-carene, and R-(+)- limonene as their substrates (Figure 5.5B-12B). The products were not found in negative control assays with CPR alone (Figure 5.5A-12A except 5.10A), confirming that products were not from endogenous P450 activity in Sf9 cells. A small peak of product was produced in the reaction containing (-)-α-pinene and CPR only (Figure 5.10A), suggesting an endogenous activity from Sf9 cells or contamination. Interestingly,
CYP6DH2 oxidized (-)-α-pinene to verbenol (Figure 5.5B) and (+)-α-pinene to myrtenol
(Figure 5.6B), a product spectrum opposite to that of CYP9T2 (Figure 5.9B; 5.10B). The apparent stereo-selectivity of substrates resulting in different hydroxylation products (e.g. 152
myrtenol vs. trans-verbenol, see Figures 5.5, 5.9) suggests a mechanism influencing how the substrate docks in the active site and may further explain the observed hydroxylation patterns. It is possible that the dimethyl group provides a hydrophobic signature that interacts with one side of the binding site and orients the substrates. If that is true, then the “hydrophobic side” of CYP9T2 substrate binding site may be oriented opposite to the “hydrophobic side” of CYP6DH2 (Figure 5.15). This hypothesis can be tested by protein modeling. If CYP6DH1 product profiles are confirmed (i.e. verbenol from both α-pinene enantiomers), it will be very interesting to see how known sequence differences between CYP6DH1 and CYP6DH2 make a huge difference in the product profiles. However, substrate-profiling experiments for CYP6DH1 must first be confirmed. CYP6DH1, with only 13 amino acids different from CYP6DH2, most likely evolved from the detoxification enzyme, CYP6DH2, by duplication followed by genetic drift. A single amino acid substitution can be sufficient to change substrate specificity and catalytic efficiency of an enzyme, including monoterpene-hydroxylating P450s
(Miyazawa et al., 2000). Therefore, CYP9T2 and CYP6DH1 may be good examples of how minor changes to existing metabolic pathway are sufficient to generate a hugely diverse suite of pheromone components (Sandstrom et al., 2006; Symonds & Elgar,
2008).
CYP6DH2 converted Δ3-carene to three possible products eluting at 20.11,
22.59 and 25.04, respectively (Figure 5.7B). Spectral searches of the MS data in the library provided with Xcalibur software suggested that the 20.11(not shown) and 22.59 peaks (Figure 5.7C) may be limonene-epoxide, and the 25.04 peak (Figure 5.7D) may be caren-10-ol. The products from R-(+)- limonene at 20.11 min and 22.57 min (Figure
5. 8B) were suggested to be limonene epoxide, and the 25.01 min product also was suggested to be caren-10-ol. The mass spectra of the three products from Δ3-carene 153
are identical to those from limonene respectively (not shown). Δ-3-carene and R-(+)- limonene were also converted to identical products by recombinant CYP9T2 (Figure
5.11B; 5.12B), but the pattern is different from the products by CYP6DH2 (Figure 5.7;
5.8). However, identities of these products remain unknown. It is unlikely that three products from Δ-3-carene and limonene catalyzed by CYP6DH2 were derived from contaminated substrates because the observed fraction of contaminants in the substrates was very low. The Δ-3-carene substrate contained about 1.9 % limonene
(not shown). It is possible that the cyclopropane ring in Δ-3-carene could be opened to form limonene and further epoxidized by these P450s. Three different products yielded from Δ-3-carene or limonene by CYP6DH2 suggests that different substrate binding domain regions may be involved in orienting the substrate. This phenomenon has been reported for a chimeric plant cytochrome P450 (Mau et al., 2010) and for CYP9T2
(Sandstrom et al., 2006). Both enzymes accepted the same substrates, but yielded different products. P450s from different species converting the same monoterpene into different products have been reported (Duisken et al., 2005; Lupien et al., 1999; Mau et al., 2010; Miyazawa & Kano, 2010). The same substrate profile and different product
profile by CYP6DH2 and CYP9T2 reflect similarities and differences of their structures.
Therefore, these P450s are useful subjects for structure-function studies to help
understand their properties.
The CYP6DH2 substrate profile did not completely match its gene expression:
qRT-PCR experiments showed CYP6DH2 is induced by non-substrates including
myrcene and terpinolene as well as putative substrates, Δ-3-carene, (+)- and (-)-α-
pinene (Gorzalski, 2010). This suggests that gene induction by xenobiotics may be not
a reliable indicator of substrate range. Alternatively, it is possible that CYP6DH2 could
not metabolize some substrates in the reaction conditions used here. Nevertheless, the 154
broad range of substrates and induction by monoterpenes confirm that CYP6DH2 works
as a detoxification enzyme in the MPB.
Competition experiments revealed that CYP9T2 preferred myrcene as substrate
rather than (+)-α-pinene or Δ-3-carene (Figure 5.13; 5.14) because it converted 78%
myrcene to ipsdienol compared with 60.5% for (+)-α-pinene, and converted 21.7% 3-
carene to an unknown product compared with 55.4% for myrcene. These results
support that CYP9T2 or its ancestor originally worked as a detoxification enzyme
because it can use non-pheromonal precursors as substrates, but it is now dedicated to
pheromone production (Blomquist et al., 2010; Sandstrom, 2007; Sandstrom et al.,
2006). The concentration of myrcene in these experiments was much less than (+)-α-
pinene and Δ-3-carene. This may be due to incorrect concentration of the stock of myrcene purified by our lab.
CYP6DH2 and CYP9T2 did not accept α-phellandrene, β-pinene, γ-terpinene, terpinolene as substrates, implying that other detoxification P450s are present to metabolize other monoterpenes that CYP6DH2 and CYP9T2 do not accept. Orthologs or paralogs with high sequence similarity to CYP9T2 and CYP6DH2 would be good candidates to screen for this role.
VI. Acknowledgments
I thank C Keeling (UBC, BC, Canada) for CYP6DH1 bacterial clone; D. Quilici and R. Woolsey at the Nevada Proteomics for GC/MS analysis; B. Perrino
(Department of Physiology, University of Nevada, Reno) for Sf9 cells; M. Schuler
(U. Illinois at Urbana-Champaign) for kindly providing the housefly reductase baculoviral clone; D. Schooley (Department of Biochemistry, University of
Nevada, Reno) for generously offering Maxsoft Diode-Array Spectrophotometer 155
for using; members of GJB and CT’s laboratories, especially A. Gorzalski for
previous work, S. Buzby for making pENTR4(-NOCI)-6DH1 clone, S. Young and
L. Plaugher for generating CYP6DH1 P3 viral stock and R. Figueroa Teran for
helpful advice.
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VIII. Figure legends:
Figure 5.1. Proposed routes for trans-verbenol biosynthesis in the MPB. (-)-α-Pinene, likely from host tree, is converted to (-)-trans-verbenol by a cytochrome P450,
CYP6DH1. CYP6DH2 likely accepts (-)-α-pinene or other monoterpenes as substrates.
Figure 5.2. CO-difference absorption spectra of Sf9 microsomes infected with
CYP6DH1 (A), CYP6DH2 (B) and CYP9T2(C) recombinant baculovirus, respectively.
The characteristic absorbance peak at 450 nm indicated functional P450 produced. The
concentration of functional P450 was approximately 6.4 X10-4mM for CYP6DH1, 3.173
X10-4 mM for CYP6DH2 and 4.73 X10-4 mM for CYP9T2.
Figure 5.3. GC-MS analysis of verbenol formed from (-)-α-pinene by recombinant
CYP6DH1 (n=2). Reactions contained (A) microsomes from Sf9 cells with recombinant
CPR and (-)-α-pinene, (B) microsomes from Sf9 cells with recombinant CYP6DH1 and
(-)-α-pinene. (C) Standard of trans-verbenol. (D) Mass spectrum with mass to charge
(m/z = 79) of the product eluting at 24.60 min in B, is identical to (E) mass spectral from
24.60 min in C. The substrate is indicated by an arrow, and product by an arrowhead.
Figure 5.4 GC-MS analysis of verbenol formed from (+)-α-pinene by recombinant
CYP6DH1 (n=2). Reactions contained (A) microsomes from Sf9 cells with recombinant
CPR and (+)-α-pinene, (B) microsomes from Sf9 cells with recombinant CYP6DH1 and 164
(+)-α-pinene. (C) Standard of cis-verbanol. (D) Mass spectrum with mass to charge
(m/z =79) from the product eluting at 24.61 min in B is identical to (E) the standard of cis-
verbanol at 24.60 min in C. The substrate is indicated by an arrow, and product by an
arrowhead.
Figure 5.5. GC-MS analysis (m/z =79) of myrtenol formed from (+)-α-pinene by
recombinant CYP6DH2 (n=2). Reactions contained (+)-α-pinene and (A) microsomes
from Sf9 cells with recombinant CPR and (B) microsomes from Sf9 cells with
recombinant CYP6DH2. (C) The standard of myrtenol. (D) Mass spectral analysis from
the retention time of 27.8 min in B, is identical to (E) mass spectral from the standard of
myrtenol at 27.73 min in C. n-Octonal: 22.01min in A and 22.03 min in B. The substrate
is indicated by an arrow and product by an arrowhead.
Figure 5.6. GC-MS analysis (m/z =79) of verbenol formed from (-)-α-pinene by
recombinant CYP6DH2 (n=2). Reactions contained (+)-α-pinene and (A) microsomes from Sf9 cells with recombinant CPR or (B) microsomes from Sf9 cells with recombinant
CYP6DH2. (C) The standard of trans-verbenol. (D) Mass spectral analysis from the retention time of 24.59 min in B. The arrow indicates (-)-α-pinene, and the arrowheads indicate product and trans-verbenol.
Figure 5.7. GC-MS analysis (m/z =79) of unknown products formed from Δ-3-carene by recombinant CYP6DH2 (n=2). Reaction contained Δ-3-carene and (A) microsomes from
Sf9 cells with recombinant CPR, or (B) microsomes from Sf9 cells with recombinant
CYP6DH2. Mass spectral from unknown products in B: (C) 22.59 min and (D) 25.04 min. Δ-3-carene is indicated by an arrow and products by arrowheads. 165
Figure 5.8. GC-MS analysis (m/z =77) of unknown products formed from limonene by
recombinant CYP6DH2 (n=2). Reactions contained limonene with (A) microsomes from
Sf9 cells with recombinant CPR, or (B) microsomes from Sf9 cells with recombinant
CYP6DH2. (C) Mass spectral from products at 20.11 min in B. Limonene is indicated
by an arrow and products by arrowheads.
Figure 5.9. GC-MS analysis (m/z =79) of verbenol formed from (+)-α-pinene by
recombinant CYP9T2 (n=2). Reactions contained (+)-α-pinene with (A) recombinant
CPR, or (B) recombinant CYP9T2. (C) The standard of trans-verbenol at 24.60 min. (D)
mass spectral from 24.57 min in B and 24.60 min in C. (+)-α-pinene is indicated by an
arrow and product and trans-verbenol by arrowheads.
Figure 5.10. GC-MS analysis (m/z =93) of myrtenol produced from (-)-α-pinene by recombinant CYP9T2 (n=2). Reactions were conducted with (-)-α-pinene and (A) recombinant CPR or (B) recombinant CYP9T2. (C) The standard of myrtenol at 27.74
min. (D) Mass spectral from 27.69 peak in B. (-)-α-pinene is indicated by an arrow and product and trans-verbenol by arrowheads.
Figure 5.11. GC-MS analysis (m/z =93) of unknown products formed from Δ-3-carene by recombinant CYP9T2 (n=2). Reactions were conducted with Δ-3-carene and (A) microsomes from Sf9 cells with recombinant CPR, or (B) microsomes from Sf9 cells with recombinant CYP9T2. (C) Mass spectra from the peak eluting at 26.71 min in B. Δ-3- carene is indicated by an arrow and product by an arrowhead.
166
Figure 5.12. GC-MS analysis (m/z =79) of unknown products yielded from limonene by recombinant CYP9T2 (n=2). Reactions were conducted with limonene and microsomes containing (A) recombinant CPR or (B) recombinant CYP9T2 and CPR. (C) Mass spectral from 26.76 peak in B. Limonene is indicated by an arrow and product by an arrowhead.
Figure 5.13. GC-MS analysis (m/z=79) of products produced from the mixture of (+)-α-
pinene and myrcene by recombinant CYP9T2 with 6-hour incubation. Reactions were
conducted with the mixture of (+)-α-pinene and myrcene and (A) recombinant CPR or
(B) recombinant CYP9T2 and CPR. (+)-α-pinene in thick arrow and myrcene in thin arrow; product from (+)-α-pinene is indicated by the larger arrowhead (24.57) and product from myrcene by the smaller arrowhead (24.85).
Figure 5.14. GC-MS analysis (m/z=79) of products formed from the mixture of Δ-3- carene and myrcene by recombinant CYP9T2 following 3-hour incubation. Reactions were run with a mixture of Δ-3-carene and myrcene and (A) recombinant CPR or (B) recombinant CYP9T2 and CPR. Δ-3-carene is shown by the thick arrow and myrcene by the thin arrow; product from Δ-3-carene by the larger arrowhead (26.62) and product from myrcene by the smaller arrowhead (24.75).
Figure 5.15. Cartoon representation to show possible hydrophobic interactions between the substrate (especially the dimethyl group) and one side of the substrate-binding site of CYP6DH2. The binding site of CYP6DH2 orients different enantiomers in the active site, and the enzyme hydroxylates different carbons of the substrate and results in different products: verbenol and myrtenol. The red hexagon is the heme in P450. 167
Host Tree
O O P O O- OH pyruvate glyceraldehyde-3-phosphate
DOX-P pathway
P P GPP
monoterpenes
CYP6DH2 (-)-α-pinene
CYP6DH1 monoterpenoid alcohols
HO verbenol
Figure 5.1 168
(A) CYP6DH1 0.06
0.04
0.02
0
-0.02
-0.04
-0.06 400 410 420 430 440 450 460 470 480 490 500
-0.005 (B) CYP6DH2
-0.015
-0.025
-0.035 ΔAbsorbance
-0.045 400 410 420 430 440 450 460 470 480 490 500
0.015 (C) CYP9T2
-0.005
-0.025
-0.045
-0.065 400 410 420 430 440 450 460 470 480 490 500 Wavelength (nm)
Figure 5.2 169
(A) CPR & (-)-α-pinene 8.14 21.98
(B) CYP6DH1 & (-)-α-pinene
8.14 21.96
24.60 Abundance Relative (C) Std of trans-verbenol 24.60
8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 Time (min)
(D) 24.6 in B 67.1 79.1 91.1 100 109.1
40.9 119.1
55.1 137.2 0 (E) 24.6 in C
79.2 100 67.2 91.2 41 109.1 119.1 Abundance Relative 55.2 137.2 0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 m/z
Figure 5.3 Figure 5.4 Figure Relative Abundance Relative Abundance 100 100 0 0
40 7 (C) (B) CYP6DH1 & (+) & CYP6DH1 (B) &(+) CPR (A) (E) 24.55 in C 24.55 in (E) (D) 24.61 in B 40.9
40.9 8.14 8
8.15 50 Std 9 55.1 55.1
10 of of 60 11 67.1 cis 67.1 70 12 - verbenol
- α 13 79.1 79.1 - 80 pinene 14 - 15 91.1 90 α 91.1 - 16 pinene
100 17 109.1 109.1 18 Time (min) Time 110 m/z 19 119.1 119.1
120 20 21 130 22.02 21.99
22 137.2 137.2 23 140 24 24.61 24.55 150 25 26 160 27 28 170 29 180 30 170 171
(A) CPR & (+)-α-pinene
8.17
22.01
(B) CYP6DH2 & (+)-α-pinene
8.16
27.8 22.03
Abundance Relative (C) Std. of myrtenol
27.73
8 10 12 14 16 18 20 22 24 26 28 20 Time (min)
(D) 27.8 in B 100 79.1 91.1
108.1 41 67.1 119.1 135.2 152.1 0 (E) 27.73 in C 100 79.3 91.3 108.2
Abundance Relative 135.1 119.1 41.1 67.3 152 0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 m/z
Figure 5.5 172
(A) CPR & (-)-α-pinene
22.16 24.67 7.84
(B) CYP6DH2 & (-)-α-pinene
22.18 24.59
7.83
(C) Std. of trans-verbenol 24.60 Abundance Relative 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 Time (min)
(D) 24.59 in B 79.1 100 91.1 67.1 109.1
40.9 119.1
55.1 137.1 0 152.1 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 m/z
Figure 5.6 173
(A) CPR & Δ3-carene 11.20
21.98
12.53 25.85
(B) CYP6DH2 & Δ3-carene 11.20
21.97
Abundance Relative 22.59 25.04 25.84 20.11
12.52
10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 Time (min)
(C) 22.59 in B 100 67.4 109.2 43.1
79.2 91.2
55.2 119.1 137.1
0 (D) 25.04 in B
100 79.2 91.1 41.0 67.2 109.1 Relative Abundance Relative
55.2 119.1 137.2
0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 m/z
Figure 5.7 174
(A) CPR & R-(+)-limonene
12.53 21.95
(B) CYP6DH2 & R-(+)-limonene 21.95 22.57 25.01 12.52 25.84
20.11
10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 Time (min) (D) 20.11 in (B) Abundance Relative
100 79.1 91.1 107.2 43.1 117.2 135.1 55.1 65.2 149.2 0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 m/z
Figure 5.8 175
(A) CPR & (+)-α-pinene
8.14 21.98
(B) CYP9T2 & (+)-α-pinene 14.23 8.14 21.95
24.57 25.01
(C) Std. of trans-verbenol 24.60 Abundance Relative
7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 Time (min)
(D) 24.57 in (B) 100 40.9
91.2 55.1 67.1 109.2 79.1 119.2
0 (E) 24.60 in (C) 79.1 91.1 100 67.1 109.1
40.9 119.1 Abundance Relative 55.1
0 40 50 60 70 80 90 100 110m/z 120 130 140 150 160 170 180 m/z
Figure 5.9 176
(A) CPR & (-)-α-pinene
8.14 21.93
24.99 27.67
(B) CYP9T2 & (-)-α-pinene 21.96 8.14 27.69
(C) Std. of myrtenol 27.74 Abundance Relative
8 10 12 14 16 18 20 22 24 26 28 30 Time (min)
(C) 27.69 in (B)
100 79.1 40.9 91.1
55.1 67.1 108.1 119.0 133.1 152.0 0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 m/z
Figure 5.10 177
(A) CPR & Δ3-carene
11.19 21.94
25.75
(B) CYP9T2 & Δ3-carene 25.77 11.19 21.95
26.71
9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 Abundance Relative Time (min) (C) 26.71 in (B)
67.2 91.1 100 109.1 40.9 119.2 81.1 137.1
152.1
0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 m/z
Figure 5.11 178
(A) CPR & R-(+)-limonene
12.53 21.95
(B) CYP9T2 & R-(+)-limonene
12.52 21.95
26.76
10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 Time (min) Abundance Relative (C) 26.76 in (B) 83.1 100
40.9 56.1 45.1 109.2 91.1 119.2 137.1 67.2 152.1 0 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 m/z
Figure 5.12 179
(A) CPR & (+)-α-pinene & myrcene 21.95 8.13
11.51 25.84 24.99
(B) CYP9T2 & (+)-α-pinene & myrcene
21.96 24.85
8.14 24.57 Abundance Relative 11.52 25.02
25.86
7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 Time (min)
Figure 5.13 180
(A) CPR & Δ3-carene & myrcene
11.17
25.70 11.48 24.93 21.86
(B) CYP9T2 & Δ3-carene & myrcene
11.17
Abundance Relative 25.71
11.48 24.75 26.62 21.88
10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 Time (min)
Figure 5.14 181
Table 5.1: Substrates and products from recombinant CYP6DH2 and CYP9T2 Products Substrates CYP6DH2 CYP9T2 Identification GC Elution Identification GC Elution Time (min) Time (min) OH
27.8 24.57 OH (+)-a-pinene myrtenol verbenol OH
24.59 27.69 HO (-)-a-pinene verbenol myrtenol Unknown* 20.11 Δ3-carene O 22.59 limonene epxide* HO 25.04 unknown 26.71 caren-10-ol* Unknown* 20.11 R-(+)-limonene O 22.57 limonene epxide* HO 25.01 unknown 26.76 caren-10-ol* OH No product --- 24.85 myrcene ipsdienol * Tentative identification based on EXCALIBUR library comparison. 182
OH OH
(-)-a-pinene
(+)-a-pinene
verbenol
myrtenol
Figure 5.15 183
Chapter 6
Discussion and future directions
I. Discussion II. Future directions III. References
I. Discussion
A bark beetle’s successful attack on a host tree is due in part to the ability to metabolize host monterpenes efficiently and to produce aggregation pheromones, which induce mass attack. Investigating enzymes involved in pheromone biosynthesis and monoterpene metabolism in the MPB may identify unique molecular targets and provide information for developing new means to deal with their infestations.
There are three major MPB pheromone components: frontalin, exo-brevicomin, and trans-verbenol. Frontalin is a male-specific anti-aggregation pheromone (Borden,
1985; Pureswaran et al., 2000; Ryker & Libbey, 1982). It is likely derived from its precursor, 6-methylhept-6-en-2-one (Brand et al., 1979; Perez et al., 1996; Vanderwel & and Oehlschlager, 1987) via the mevalonate pathway in the anterior midgut (Barkawi et al., 2003; Hall et al., 2002b). exo-Brevicomin, a male-specific aggregation pheromone component (Conn et al., 1983; Libbey et al., 1985; Pureswaran et al., 2000) is likely synthesized de novo by epoxidation followed by cyclization from its precursor (6Z)-non-
6-en-2-one (Figure 2.1) (Francke et al., 1996; Vanderwel et al., 1992). (6Z)-Non-6-en-2- one is likely fatty acid derived (Figure 2.1) (Rafaeli & Jurenka, 2003; Vanderwel et al.,
1992). trans-Verbenol is a female-specific aggregation pheromone component (Billings et al., 1976; Libbey et al., 1985; Pitman et al., 1968; Pitman et al.,1969). trans-Verbenol production increases with feeding and ceases upon mating (Pureswaran et al., 2000). It 184
is most likely produced via cytochrome P450-mediated hydroxylation of α-pinene from the host tree (Figure 5.1) (Greis et al., 1990; Hunt & Smirle, 1988; Pierce et al., 1987) although de novo biosynthesis may be possible (Blomquist et al., 2010; Renwick et al.,
1973).
Understanding pheromone biosynthesis is important for the development of new management techniques, both by providing targets for strategies and by informing how the beetles interact with each other and their environment. Although major pheromone components in the MPB were identified several decades ago, none of their biosynthetic pathways were studied in detail and none of the involved enzymes were identified or characterized beyond the most general terms. For example, a P450 with epoxidase activity was proposed for exo-brevicomin biosynthesis by Vanderwel et al. (1992), and tentatively identified as CYP6CR1 by Aw et al. (2010), but not confirmed. Similarly, α- pinene hydroxylation to trans-verbenol was predicted to be catalyzed by at least two different P450 “systems” (Pierce et al., 1987), but the responsible enzymes have not been identified. Enzymes near the ends of pheromone biosynthetic pathways are most likely to be species-specific and therefore more valuable as potential targets for new pest management strategies than those involved in more general metabolism. As their orthologs and paralogs are identified, comparisons can help explain how these systems evolve, and thus predict impacts of future management efforts. Their identification and characterization are a necessary step to help manage these pests.
Biochemical studies of pheromone biosynthesis can be considered with the evolutionary relationship to resin detoxification in mind. MPBs survival in a toxic monoterpene environment (Byers, 1995; Raffa et al., 1985; Smith, 1965, 1966) is mostly
due to efficient enzyme systems which metabolize a wide range of toxic host substances
rapidly to compounds that are less toxic and or more easily excretable (Brattsten et al., 185
1977). Because some detoxification products are identical to pheromone components, enzymes involved in both processes should be related. However, little is known about monoterpene metabolism in bark beetles. Monoterpene metabolism by cytochromes
P450 has been studied extensively in plants (Karp et al., 1987; Karp et al., 1990;
Kjonaas et al., 1985; Lupien et al., 1999) and, to a lesser extent, in other insects. Δ3-
Carene metabolism by larvae of the common cutworm (Spodoptera litura) implicated a cytochrome P450 (Miyazawa & Kano, 2010). Cytochromes P450 have been proposed for monoterpene metabolism in various bark beetles (White et al., 1980; White et al.,
1979). Cytochromes P450 involved in α-pinene and myrcene metabolism by MPBs have been reported (Hunt & Smirle, 1988). Blocking P450 using piperonyl buxtoxide decreased the level of certain terpene alcohol phermonone components and increased the levels of their monoterpene precursors (Hunt & Smirle, 1988). The only bark beetle
P450s with known functions are myrcene hydroxylases (CYP9T1 and CYP9T2 from I. confuses and I. pini, respectively) (Sandstrom, 2007; Sandstrom et al., 2008; Sandstrom et al., 2006). CYP9T2 is also likely involved with monoterpene metabolism because it hydroxylates α-pinene to verbenol (Sandstrom, 2007)(Chapter 5).
This dissertation explored, for the first time, biochemical details of two important
MPB pheromone components: exo-brevicomin and trans-verbenol. I presented new in vivo information regarding exo-brevicomin biosynthesis, characterized four new enzymes – three P450s and an oxidoreductase either clearly involved or else implicated in exo-brevicomin and trans-verbenol biosynthesis, and explored bark beetle monoterpene metabolism in the context of specific enzymes. Therefore, this dissertation is the first effort to reveal exo-brevicomin biosynthesis pathway at the molecular level. It also offers new insights on the evolution of pheromone production and detoxification in the MPB and I. pini. 186
Chapter 2 explored exo-brevicomin biosynthesis in vivo. Although this chapter did not identify new enzymes, it provided information about terminal steps that helped clarify enzyme candidates and assisted in their eventual identification. My work revealed that exo-brevicomin was specifically produced in male fat bodies, but not in females, male carcasses or alimentary canals (Figure 2.2). This is the first report of a bark beetle pheromone produced in fat body and is in contrast to the precedent that bark beetle pheromones are produced in the midgut (Barkawi et al., 2003; Hall et al., 2002a; Hall et al., 2002b). Thus, pheromone component biosynthesis is more complex than originally thought. These data suggest a new paradigm in which lipid-based pheromone components are synthesized in the fat body, while de novo-synthesized monoterpenoid components are synthesized in the midgut. This chapter also revealed new details about terminal steps in the exo-brevicomin biosynthetic pathway. There is likely no desaturase activity for short chain fatty acids in the male fat body because decanoic acid was converted to nonan-2-one, not (6Z)-non-6-en-2-one (Figure 2.3). Thus, the direct precursor of (6Z)-non-6-en-2-one is 10:1 fatty acid, which is likely derived from a long chain fatty acid (Figure 2.1). Furthermore, the male fat body probably contains a decarboxylase that may take both saturated and unsaturated precursors (Figure 2.1). A
P450 inhibitor, piperonyl butoxide, greatly reduced epoxide produced by male fat bodies
(Figure 2.4) and thus confirmed the hypothesized role of a P450 (Vanderwel et al., 1992) in the conversion of (6Z)-non-6-en-2-one to 6,7-epoxynonan-2-one, the direct epoxide precursor of exo-brevicomin. This keto-epoxide was very stable in physiological conditions and required a pH below 3.0 to be cyclized into exo-brevicomin (Figure 2.5), implying a cyclase is required for the final step of exo-brevicomin production. The conversion of epoxide to exo-brevicomin does not require hydride transfer.
Homogenized fat bodies did not convert epoxide to exo-brevicomin without any 187
cofactors. Very preliminary data suggested that the conversion of epoxide to exo-
brevicomin in the fat body may require NADH or NADPH to activate the cyclase: an un-
replicated assay showed fat bodies likely converted exo-brevicomin to epoxide in the
presence of NAD+ or NADP+ (Figure 2.6).
Chapter 3 presented the molecular and biochemical characterization of a new
P450, CYP6CR1, and demonstrated its role in exo-brevicomin biosynthesis. CYP6CR1
was highly expressed in unfed males, but not in females (Figure 3.3). Feeding
decreased the level of its expression, and basal expression levels were highest in the
male fat body (Figure 3.3). Every aspect of this expression profile is consistent with exo-
brevicomin production (Figure 2.2). Enzyme assays demonstrated that microsomes
expressing recombinant CYP6CR1 and housefly NADPH-cytochrome P450 reductase
(CPR) converted (6Z)-non-6-en-2-one into 6,7-epoxynonanone, the direct epoxide precursor of exo-brevicomin (Figure 3.5). CYP6CR1 did not accept (6E)-non-6-en-2-one or various monoterpenes as substrates (not shown) confirming that CYP6CR1 is specific in the penultimate step of exo-brevicomin biosynthesis. CYP6CR1 is the first characterized MPB pheromone biosynthetic enzyme.
Male MPBs also produce endo-brevicomin, which is not an aggregation pheromone (Pureswaran et al., 2000; Rudinsky et al., 1974), from its precursor, (6E)- non-6-en-2-one (Vanderwel et al., 1992) with the same pattern as exo-brevicomin production (Pureswaran et al., 2000). This suggests that endo- and exo-isomers may
depend on related desaturases that produce trans- or cis- unsaturated fatty acid
precursors and these precursors may not be discriminated by the downstream enzyme,
CYP6CR1. However, CYP6CR1 did not accept (6E)-non-6-en-2-one as its substrate,
implying that a different P450 facilitates the conversion of (6E)-non-6-en-2-one to endo-
brevicomin. (+)-exo-Brevicomin is the natural chirality produced by male MPB, with the 188
ratio of 98% of the blend of (+)- and (-)-exo-brevicomin (Pureswaran et al., 2000). It is
likely derived from the cis-epoxide (Silverstein et al., 1968). MPBs exposed to (6Z)-non-
6-en-2-one produced (+)-exo-brevicomin, not (-)-exo-brevicomin (Vanderwel et al.,
1992). Non-enzymatic, acid hydrolysis of the cis-and trans- epoxides yields exo- and
endo-bicylic products, respectively (Silverstein et al., 1968). The chirality of the epoxide
produced by CYP6CR1 is not clear and may include different enantiomers, such as (-)-
and (+)-cis-epoxide.
Chapter 4 investigated the role of a novel oxidoreductase in exo-brevicomin
biosynthesis. First named by its EST identifier, DPG022F04 (Aw et al., 2010), this
enzyme was renamed “ZnoDH’ (for (6Z)-non-6-en-2-ol dehydrogenase) following
characterization. Similar to CYP6CR1, the expression pattern of ZnoDH is consistent
with exo-brevicomin production (Figure 4.3). Recombinant ZnoDH converted (6Z)-non-
6-en-2-ol to (6Z)-non-6-en-2-one in the presence of NAD+ or NADP+ (Figure 4.5, 4.6), with a preference for NAD+. ZnoDH did not convert n-octanol, epoxide or C10:0 fatty
acid derived precursors to any products (not shown). These results suggest ZnoDH is
likely involved in exo-brevicomin biosynthesis by specifically activating (6Z)-non-6-en-2-
one production. However, these data also present an intriguing mystery regarding the
source of nonen-2-ol. Standard decarboxylation of a C10 precursor should yield the methylketone (Skiba & Jackson, 1994), so the requirement for ZnoDH retains unclear. It is possible that C10: 1 fatty acid is decarbonylated to 3-nonene via fatty acyl-CoA reductase and oxidative decarbonylase – the standard metabolic pathway from fatty acid
to hydrocarbon in insects (Reed et al., 1994). 3-Nonene could then be hydroxylated to
(6Z)-non-6-en-2-ol by a different P450 (G. Blomquist, personal communication).
MPBs exposed to (6Z)-non-6-en-2-one produced (6Z)-non-6-en-2-ol (Vanderwel et al., 1992), implying that a dehydrogenase is required for this reduction process. 189
ZnoDH is most likely the enzyme for this reduction. The possibility that ZnoDH acts as a decarboxylase should be also examined because decarboxylations are commonly catalyzed by dehydrogenases (Rule & Roelofs, 1989; Skiba & Jackson, 1994; Vaz et al.,
1988). However, a preliminary assay of ZnoDH-recombinant Sf9 cells cultured with
C10:0 fatty acid did not produce a 9-carbon product. One possible reason is that Sf9 cells might not produce the substrate of ZnoDH from C10 fatty acid. The other reason is
ZnoDH may not be the enzyme for the decarboxylation.
It is worth noting that Chapters 3 and 4 explored pheromone-biosynthetic enzyme candidates suggested from microarray experiments. Aw et al. (2010) noted a cluster of three genes, including CYP6CR1 and ZnoDH (identified by its EST,
DPG022F04), with apparent coordinate regulation consistent with exo-brevicomin biosynthesis. Here, I demonstrated that CYP6CR1 and ZnoDH are both strongly implicated in exo-brevicomin biosynthesis, both confirming the hypothesis by Aw et al.
(2010) and validating the use of clustering expression profiles to identify useful genes.
This “functional genomics followed by enzyme characterization” approach parallels studies in I. pini that led to the identification of important pheromone-biosynthetic enzymes in that species (Figueroa-Teran et al., 2011; Sandstrom et al., 2006).
Chapter 5 presented new data regarding monoterpene metabolism by cytochromes P450 in D. ponderosae and I. pini which, when combined with prior expression and biochemical analyses (Gorzalski, 2010; Sandstrom, 2007; Sandstrom et al., 2006), reveals new insights on the evolution of pheromone production and resin detoxification. This chapter described substrate and product profiles of CYP6DH1 and
CYP6DH2 from D. ponderosae and CYP9T2 from I. pini. CYP6DH1 was highly expressed in female hindguts and not induced by monoterpenes (Gorzalski, 2010).
Enzyme assays revealed that recombinant CYP6DH1 hydroxylated (+)- and (-)-α-pinene 190
into verbenol, but did not accept other monoterpenes as substrates. Earlier studies
showed two distinct α-pinene hydroxylating pathways in female D. ponderosae (Greis et
al., 1990; Pierce et al., 1987): the first pathway does not discriminate between the two
enantiomers and likely plays a detoxification role, while second pathway appears
specific for (-)-α-pinene and therefore produces the bulk of the female specific
aggregation pheromone component, (-)-trans-verbenol (Pierce et al., 1987). Thus,
CYP6DH1 is likely a pheromone biosynthetic enzyme. However, chapter 5 showed that
both CYP6DH2 and CYP6DH1 accepted both enantiomers of α-pinene. This result does
not necessarily contradict a two α-pinene hydroxylating systems hypothesis if, for
example, a third P450 with a narrow substrate range is active. Also, CYP6DH2
specifically produced myrtenol or trans-verbenol, depending on the substrate (see
below), so the product profile can vary depending on the beetles’ environment.
Competition experiments with mixtures of these enantiomers may answer these
questions.
CYP6DH2 and CYP9T2 appear to work as monoterpene metabolism enzymes.
Both have similar substrate profiles, except that myrcene is a substrate for CYP9T2, but not for CYP6DH2. Both accepted (+)-, (-)-α-pinene, Δ3-carene, and R-(+)-limonene as their substrates. Interestingly, CYP6DH2 and CYP9T2 converted the same substrate into different products. This may be contributed by the difference of their structures.
The apparent stereo-selectivity of substrates resulting in different hydroxylation products
(e.g. myrtenol vs. trans-verbenol, see Figures 5.5, 5.9) suggested a mechanism influencing how the substrate docks in the active site and may further explain the observed hydroxylation patterns. P450s from different species converting the same monoterpenes into different products have been reported (Duisken et al., 2005; Lupien
et al., 1999; Mau et al., 2010; Miyazawa & Kano, 2010). 191
Competition experiments revealed that CYP9T2 preferred myrcene as its substrate over α-pinene and Δ3-carene (Figure 5.13, 5.14). These results support that
CYP9T2 or its ancestor originally worked as a detoxification enzyme, but it is now dedicated to pheromone production (Sandstrom, 2007; Sandstrom et al., 2006). These data also implicate that other detoxification P450s are present to metabolize other monoterpenes that CYP6DH2 or CYP9T2 do not accept. Orthologs or paralogs with high sequence similarity to CYP9T2 and CYP6DH2 would be good candidates to screen for this role.
A detoxification enzyme may have a broad range of substrates and be induced by those substrates, whereas a pheromone-biosynthetic enzyme may show a strong preference for the pheromone precursor (Blomquist et al., 2010). The difference between the two may conceivably occur following simple point mutations in coding or regulatory regions or duplication. CYP6DH1, with only 16 amino acids different from
CYP6DH2, most likely evolved from detoxification enzyme, CYP6DH2, by duplication followed by genetic drift. A single amino acid substitution can be sufficient to change substrate specificity and catalytic efficiency of an enzyme, including monoterpene- hydroxylating P450s (Miyazawa & Kano, 2010; Schalk & Croteau, 2000). Therefore,
CYP9T2 and CYP6DH1 may be good examples of how minor changes to existing metabolic pathway are sufficient to generate a hugely diverse suite of pheromone components (Sandstrom et al., 2006; Symonds & Elgar, 2008).
It is interesting that CYP6DH2 converted Δ3-carene and limonene into three different products respectively (Figures 5.7, 5.8). Three different products from Δ3- carene are the same as products from limonene. Similarly, Δ3-carene and limonene were converted into the same product by CYP9T2 (Figure 5.11, 5.12). Three different products from Δ 3-carene or limonene by CYP6DH2 suggests that different substrate 192 binding regions may be involved in orienting the substrates. This may explain multiple products formed from the single substrate by a P450. This phenomena has been reported when using a chimeric plant cytochrome P450 (Mau et al., 2010) and CYP9T2
(Sandstrom et al., 2006). Furthermore, the cyclopropane ring in Δ3-carene could be opened to form limonene, thereby broadening the product profile from 3-carene.
Therefore, these P450s are useful subjects for structure-function studies to help understand their properties.
Using molecular and biochemical tools, this dissertation has advanced our understanding of exo-brevicomin biosynthesis in the MPB, and the expected evolutionarily related pathways for pheromone biosynthesis and monoterpene detoxification in D. ponderosae and I. pini. CYP6CR1 and ZnoDH, mostly likely involved in the final step of exo-brevicomin biosyntheisis, may be targeted for controlling beetles by RNAi. Knocking down these enzymes should abolish exo-brevicomin production to terminate pheromonal attraction. Because of substrate specificity of CYP6CR1, it is possible that recombinant CYP6CR1 can be an ideal enzyme to produce epoxide, in turn cyclized into exo-brevicomin on acid hydrolysis (Silverstein et al., 1968), which can be used as bait since chemical synthesis seems labor intensive and complicated (Mori,
1997; Silverstein et al., 1968). ZnoDH and CYP6DH1 may also be potential enzymes for commercial purposes.
II. Future directions
exo-Brevicomin is most likely derived from long-chain fatty acids via a shortening process in the fat body. To confirm this, a long-chain fatty acid, such as C16 or C18, cultured with male fat bodies can be conducted. Furthermore, the enantiomer of the epoxide produced by CYP6CR1 has not been examined, but could readily be 193
determined by separating products on an enantioselective column coupled with GC.
This information will be useful for commercial application because chemically
synthesized exo-brevicomin contains equal proportions of (+)- and (-)-isomers (C.
Oehlschlager, personal communication). Furthermore, kinetic analyses should help
understand the enzyme’s preferred substrate(s) and help determine conditions for
potential commercial application. Although ZnoDH converted (6Z)-non-6-en-2-ol into
(6Z)-non-6-en-2-one efficiently, more alcohol substrates need to be tested to confirm its
specificity. The reverse reaction from (6Z)-non-6-en-2-one to (6Z)-non-6-en-2-ol has not been observed though experiments are in progress. More experiments with different conditions for this reverse step need to be conducted. cis-3-Nonene will be used to test the hypothesis that (6Z)-non-6-en-2-ol is derived from 3-nonene in the male fat body.
Other possible ten-carbon C10 fatty acid derivatives also should be used to further examine if ZnoDH plays a role in decarboxylation. The final step of exo-brevicomin biosynthesis requires a cyclase. Searching microarray data may find the candidate of cyclase.
CYP6DH1 is likely a trans-verbenol biosynthetic enzyme. In this study, biochemical assays should be replicated to confirm the substrate and product profiles.
Enantiomers of the products from (+) and (-)-α-pinene need to be further identified although female MPBs convert both (+) and (-)-α-pinene to trans-verbenol (Greis et al.,
1990). Time-course and kinetic studies also need to be conducted to further understand this enzyme activity.
Frontalin is derived from mevalonate pathway. Frontalin and exo-brevicomin are often grouped together because of the similarity of their and their precursors’ structures
(Symonds & Elgar, 2004; Symonds & Wertheim, 2005), even though their precursors are derived from different pathways and in different tissues (Barkawi et al., 2003; Hall et al., 194
2002b; Perez et al., 1996; Vanderwel et al., 1992). The two chemicals may be metabolically linked and may be epoxidized by the same P450. However, CYP6CR1 did not convert 6-methylhept-6-en- 2-one to frontalin or its intermediates. This single enzyme assay was conducted with a low concentration of functional CYP6CR1 (not shown). More replicates with high concentration of functional CYP6CR1 should be examined. It is also possible that a different, unknown P450 besides CYP6CR1 is responsible for frontalin production.
The similarity and difference of substrate and product profiles from CYP6DH1,
CYP6DH2, and CYP9T2 indicate the similarity and difference of their structures that influence substrate binding and product forming. Protein modeling may explore their properties. The preference of substrates for these P450s also needs to be conducted to better understand their roles.
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Appendix A:
Figure A.1. Sequence of DPG017M13
Figure A.2. Sequence of DPG022F04
Figure A.3. Amino acid alignment of CYP6DH1 and CYP6DH2
204
DPG017M13
1 2 ACTGATAGTGACCTGTTCGTTGCACAAATTGATGAGCAATGCTTTTTTATAATGCCAACT 21 62 TTGTACAAAAAAGTTGGAGTTACTTCACACTAAGATTGAAGACCACACAATATTGAAGTA 41 122 GTAGGTGCCAAAAACATCCGAAACAAGTAAACCAGAAATTGTTGCACCAGAGAACACAAA 61 M L P V P A A L M I 182 ATTGATGAGCAACACATTGAAGAAGAAAAGATGTTACCAGTTCCAGCTGCTCTCATGATT 81 I I I I L L L I I F S I V I T F S T Y T 242 ATCATCATAATTTTACTCTTAATAATTTTCTCTATTGTTATAACCTTCTCTACGTACACT 101 F S Y W K K R K F N F L E P T I P F G N 302 TTTTCTTATTGGAAGAAACGAAAATTTAATTTCCTGGAGCCAACAATCCCATTTGGAAAT 121 A Q S F F L G K K G L G E L Y S D W Y L 362 GCTCAGAGTTTCTTTCTTGGAAAAAAGGGTTTAGGAGAACTTTATAGCGATTGGTATCTG 141 E M K A K G W D M G G A Y F G S K P V F 422 GAAATGAAAGCCAAGGGTTGGGATATGGGAGGTGCCTATTTCGGCAGCAAACCCGTTTTC 161 I P I D N K L I K T I L V K D F S N F Q 482 ATACCAATCGATAATAAGCTGATTAAAACGATATTGGTTAAAGATTTTTCGAATTTTCAA 181 N H G F Y I N E K I D P L S G H I Y N L 542 AATCACGGCTTTTACATCAACGAAAAAATCGATCCACTGTCTGGGCATATCTACAATTTG 201 E S S K W K N L R S K I L P A F S 602 GAAAGCAGCAAGTGGAAGAACCTGCGCTCCAAAATCCTCCCAGCTTTCTC
Figure A.1
205
DPG022F04
1 3 AGTGACCTGGTTCGTTGCAACAAATTGATGAGCAATGCTTTTTTATAATGCCAACTTTGT 21 63 ACAAAAAAGTTGGAGGTTAAACGCCTACTAATTAAGACCTTCCTTATAATTGCTCAACAG 41 M H F I L W F 123 TCTTCTTTCTAACATCATCTTTTAATTTTTTCTCTTAATATGCATTTTATTTTGTGGTTT 61 V S C L V L L K V V N A L T C F W C K S 183 GTTTCCTGCTTGGTTTTGTTGAAAGTGGTAAATGCATTGACGTGTTTCTGGTGCAAAAGC 81 K V C L V G K T A I I T G G A A G I G F 243 AAAGTGTGTTTAGTGGGCAAAACGGCAATCATTACAGGAGGGGCCGCAGGTATTGGTTTC 101 Q T A L A L A S K G C R V I I A D I S N 303 CAGACCGCACTAGCTCTAGCTTCCAAAGGATGTCGGGTGATTATAGCTGATATTTCCAAC 121 L T K A V D E L K N V S K N Q N I I G V 363 TTAACTAAAGCAGTGGATGAATTGAAAAATGTTTCGAAAAATCAAAATATCATTGGTGTA 141 E V D L A S F R S V R E F A K K I L D T 423 GAAGTTGATCTGGCATCTTTTAGATCCGTTAGGGAATTTGCGAAAAAAATTCTCGATACA 161 E P R L D I L V C N A G I G E H K M R I 483 GAGCCCAGGCTCGATATATTGGTTTGTAATGCAGGTATCGGTGAACACAAAATGCGGATA 181 M T G D G V E K T M Q I N Y Y S N F L M 543 ATGACTGGTGATGGGGTAGAAAAAACTATGCAAATTAATTACTACAGTAATTTCTTAATG 201 I H L L L D L L K 603 ATTCATTTGCTGTTGGATTTGTTGAAAAA
Figure A.2
206
CYP6DH1 & CYP6DH2
1 60 Cyp6DH1 ORF (1) MLVYILISLITLLYFFLKYKHSYWSRRKITQGNPRFLFGSVDQNIFGSGNPTEYVRKAYW Cyp6DH2 ORF (1) MLVYILIPVITLLYYFLKYKHSYWSRRKITQGNPRFLFGSVDQSIFGSGNPTEYVRKAYW Consensus (1) MLVYILI LITLLYFFLKYKHSYWSRRKITQGNPRFLFGSVDQ IFGSGNPTEYVRKAYW
61 120 Cyp6DH1 ORF (61) DLKKKGAKHGGIYLFYNPVWIPIDLKLIKKVLVTDYDHFSSHGFFHHKKDSLSENLFQKE Cyp6DH2 ORF (61) DLKKKGAKHGGIYIFYNPVWIPIDLKLIKKVLVTDYDHFSSHGFFHHKKDSMSDNLFHKE Consensus (61) DLKKKGAKHGGIYIFYNPVWIPIDLKLIKKVLVTDYDHFSSHGFFHHKKDSLSDNLF KE
121 180 Cyp6DH1 ORF (121) GDEWKVLRSHLSPTFTPSKLKNMYATLYKFGHRMEERISDSCKKGQPLNIRDTTTSYLIT Cyp6DH2 ORF (121) GDEWKVLRSHLSPTFTPSKLKNMYDTLYKFGHRMEERISDSCKKGQPLNIRDTTTSYLIN Consensus (121) GDEWKVLRSHLSPTFTPSKLKNMY TLYKFGHRMEERISDSCKKGQPLNIRDTTTSYLI
181 240 Cyp6DH1 ORF (181) VIASCFFGIESKSLDDPNSDFKHYGKLIAQSRPLRFLVESLVNWDLLAHLGYSFFPWAVR Cyp6DH2 ORF (181) VTASCFFGIESKSLDDPNSDFKHYGKLIAQSRPLRFLVESLVNWDLLAHLGYSFFPWAVR Consensus (181) V ASCFFGIESKSLDDPNSDFKHYGKLIAQSRPLRFLVESLVNWDLLAHLGYSFFPWAVR
241 300 Cyp6DH1 ORF (241) PFFTSLIKDVVDEREKNSIIRKDCLDTLYQMSKNGGPLTFQDVVATSTFLYAAGYETSSS Cyp6DH2 ORF (241) PFFTSLIKDVVDEREKNSIIRKDCLDTVFQMSKNGGPLTFQDVVATSTFLYAAGYETSSS Consensus (241) PFFTSLIKDVVDEREKNSIIRKDCLDTLFQMSKNGGPLTFQDVVATSTFLYAAGYETSSS
301 360 Cyp6DH1 ORF (301) TLSYLMYELAKNQDVQDKLRSEILSICKDNAELTYEDLSKMKYADLCLAEILRCYPALAQ Cyp6DH2 ORF (301) TLSYLMYELAKNQDVQDKLRSEILSICKDNAELTYEDLSKMRYADLCLAEILRCYPALAQ Consensus (301) TLSYLMYELAKNQDVQDKLRSEILSICKDNAELTYEDLSKMKYADLCLAEILRCYPALAQ
361 420 Cyp6DH1 ORF (361) LPRACTKEYRIPGTDQVIEKGTTILIPVWAIQNDPEYFRNPTMFDPENMSPENQNSNVED Cyp6DH2 ORF (361) LPRACTKEYRIPGTDQVIEKGTTILIPVWAIQNDPEYFRNPTMFDPENMSPENQNSNVED Consensus (361) LPRACTKEYRIPGTDQVIEKGTTILIPVWAIQNDPEYFRNPTMFDPENMSPENQNSNVED
421 480 Cyp6DH1 ORF (421) AWFAFGYGPRLCLGYKFAQMEIKVALVKYLKNHRYKLNIATPEELTFNYDTIVLYPAEDI Cyp6DH2 ORF (421) AWFAFGYGPRLCLGYKFAQMQIKVALVKYLKNHRYKLNIATPEELTFNYDTVVLYPAEDI Consensus (421) AWFAFGYGPRLCLGYKFAQM IKVALVKYLKNHRYKLNIATPEELTFNYDTIVLYPAEDI
481 Cyp6DH1 ORF (481) ILDIEAIN Cyp6DH2 ORF (481) ILDIEAIN Consensus (481) ILDIEAIN
Figure A.3 Amino acid differences are highlighted