Application of biological and chemical approaches to generate new and diverse fungal natural products

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Content APPLICATION OF BIOLGICAL AND CHEMICAL APPROACHES TO GENERATE
NEW AND DIVERSE FUNGAL NATURAL PRODUCTS

by

Amber Dorothy Somoza

A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(CHEMISTRY)

August 2012

Copyright 2012 Amber Dorothy Somoza
ii

Dedication

I would like to dedicate this to my parents, Exiquio and Dorothy Valencia, whom
have inspired and supported me along my academic journey. Likewise, I also dedicate
this to my brother, Jonathan Valencia, for his encouragement and engaging scientific
conversations and most importantly to my husband, Christian Somoza, for his loyal
support and immense understanding.

iii
Acknowledgements
Completing the Ph.D. program at USC could not have been accomplished without
the help of key individuals and a vast network of support. The most important individual
in my academic journey has been my advisor Clay Wang. He welcomed me into his lab
in the summer of 2008 and has since taught me to think scientifically, manage a
laboratory, and given me a wealth of advice and knowledge in our field of study.
Moreover, he has provided me the tools I need to be successful after graduate school. I
would also like to thank Hannah Reisler for her pivotal role in my career. Her
encouragement and support kept me on my path to completing the program. It is with
immense gratitude to Dr. Wang and Dr. Reisler for all your help these past five years.
I also wish to thank everyone who served on my committee Michael Kahn, Richard
Roberts, Surya Prakash, Kyung Jung and Bogdan Olenyuk for their contributions. Also, I
would like to extend my gratitude to our collaborator, Berl Oakley, for his valuable
insight and help on experiments and preparation of manuscripts. Additionally, I would
like to recognize the National Institutes of Health for their financial support.
Additionally, I would like to gratefully acknowledge the visiting academics, graduate
students, and undergraduates in the Wang lab. I would like to specially thank Yi-Ming
Chiang for his kindness in teaching me a variety of techniques in the laboratory, and
furthermore for his insight and guidance on experiments. It was a pleasure and honor to
work with the following bright and talented individuals: Kuan-Han Lee, James Sanchez,
Shu-Lin Chang, Hsu-Hwa Yeh, Vessela Ensberg, Ting Liu, Ken Lo, Chunjun Guo, Mike
Praseuth and Junko Yaegashi. And, of course, I would like to mention undergraduates,
iv
Sarah Ifrah and Detrick Hudson for their help with experiments and improving my
teaching skills.
Chapter 2 is incorporated from Somoza, A. D.; Lee, K. H.; Chiang, Y. M.; Oakley,
B.; Wang, C. C. C. (2012). Reengineering an azaphilone biosynthesis pathway in
Aspergillus nidulans to create lipoxygenase inhibitors. Organic Letters, 14(4), 972-975.
Gene deletion and diagnostic PCR was carried out by the Oakley lab. Kuan-Han Lee
provided advice and helpful insight on the synthetic route to azaphilone analogs, and
assisted in the synthesis and chiral resolution of (±)-sclerotiorin into respective
diastereomers. Yi-Ming Chiang provided insight and guidance on the project.
Chapter 3 is incorporated from Liu T.; Chiang, Y. M.; Somoza, A. D.; Wang, C. C.
C.; (2011) Engineering of an “Unnatural” Natural Product by Swapping Polyketide
Synthase Domains in Aspergillus nidulans. Journal of the American Chemical Society.
133, 13314-13316. Ting Liu made the most contributions for this work, including gene
deletion and diagnostic PCR, feeding studies, natural product isolation and
characterization, and analyzed data on the HRMS. Yi-Ming Chiang assisted in the
characterization of compound 1. Amber Somoza carried out the syntheses of the
hexanoyl-SNAC and D-11 hexanoyl-SNAC compounds.

v
Table of Contents
Dedication ii
Acknowledgements iii
List of Tables vi
List of Figures vii
Abstract xi
Chapter I: Introduction
1. Significance of Aspergillus 1
2. The Status of Aspergillus Genomic Sequencing Efforts 3
3. Technological Progress In Genome Mining Endeavors 4
4. Genetic Characterization of Other Aspergillus Secondary Metabolites 16
5. Expanding Natural Product Diversity 17
Chapter II: Reengineering An Azaphilone Biosynthesis Pathway In Aspergillus
nidulans To Create Lipoxygenase Inhibitors
1. Abstract 54
2. Introduction 54
3. Results & Discussion 56
4. Supplemental Information 59
Chapter III: Engineering Of An "Unnatural" Natural Product By Swapping
Polyketide Synthase Domains In Aspergillus Nidulans
1. Abstract 107
2. Introduction 107
3. Results & Discussion 109
4. Supplemental Methods 113
Chapter IV: Connecting The Pieces - Common Motifs In Aspergillus nidulans
Secondary Metabolome Research 132

Bibliography 136

vi
List of Tables
Table 1-1 Features of Aspergillus Genomes 30
Table 1-2 Secondary metabolism gene clusters in A. nidulans 31
Table 1-3 Secondary metabolism gene clusters in A. fumigatus 32
Table 1-4 Secondary metabolism gene cluster in A. terreus 33
Table 1-5 Secondary metabolism gene clusters in A. niger 34
Table 2-1 Lipoxygenase-1 Inhibitory activity 77
Table 2-2 A. nidulans strain used in the study 78
Table 3-1 A. nidulans strains used in this study 118
Table 3-2 Primers used in this study 119
Table 3-3 NMR data for compound 1 (400 and 100 MHz) in CD
3
OD 129

vii
List of Figures
Figure 1-1 Structures of emericellamides A, C-F produced by A. nidulans 36
Figure 1-2 The biosynthetic pathway for austinol and dehydroaustinol 37
Figure 1-3 The biosynthetic pathway for acetylaszonalenin 38
Figure 1-4 A) Structure of pyripyropene A B) Structures of aspyridone A, B and
asperfuranone discovered by controlling expression of cluster-specific
transcriptional activators 39

Figure 1-5 The biosynthetic pathway for terrequinone A 40
Figure 1-6 Structures of secondary metabolites A) discovered from chromatin
restructuring B) discovered through co-incubation with microorganisms 41

Figure 1-7 Structures of aspoquinolines A-D and aspernidine A-B discovered
by OSMAC strategy 42

Figure 1-8 The biosynthetic pathway for prenyl xanthones in A. nidulans 43

Figure 1-9. a) A biosynthesis pathway in a WT organism b) In PDB the biosynthesis
pathway uses both the natural substrate and the mutasynthon to produce
the natural product and the unnatural product c) In MBS the biosynthesis
pathway is engineered to eliminate the availability of the natural substrate
and only uses the mutasynthon to produce only the unnatural product d)
In modified semi-synthesis the pathway is engineered to prematurely stop
at the intermediate stage and then structurally modified through chemical
synthesis to the natural product analog e) In CB (i) polyketide domain
swap takes a domain from the PKS of organism #2 to replace the same
PKS domain in organism and produces an unnatural polyketide different
from the polyketide of organism #1 and #2 ii) replacement of post-PKS
gene takes a tailoring gene from one pathway and adds the gene to a
different biosynthesis pathway to produce a modified natural product 44

Figure 1-10 Precursor-directed biosynthesis of A) rapamycin analogs with
modification at the cyclohexane moiety B) rapamycin analogs with
modification at the pipecolate ring 45

Figure 1-11 Precursor-directed biosynthesis of hormaomycin analogs 46

Figure 1-12 Mutasynthesis of pre-rapamycin analogs 47

viii

Figure 1-13 A) Structures of natural products FK523, FK520 and FK506 produced
by S. hygroscopicus NRRL5491 B) mutasynthesis of FK506 analogs
with selected modification at C-21 48

Figure 1-14 A) Structures of natural products ansamitocin P-2, P-3 and P-4 B)
mutasynthesis of ansamitocin P-3 analogs C) Synthetic transformation
under Stille conditions to install an alkene on an aromatic ring of
ansamitocin scaffold 49

Figure 1-15 A) Structure of avermectin analog, doramectin B) structure of spinosyn
A and selected modification on the western shore of the spinosyn
scaffold using combinatorial biosynthesis 50

Figure 1-16 A) Structure of tenellin and B) Combinatorial biosynthesis generated
bassianin using the hybrid PKS of tenellin 51

Figure 1-17 The engineered taxol pathway (dark arrows indicate genes overexpressed)
A general semisynthesis of the baccatin III intermediate is transformed in
two steps to taxol 52

Figure 1-18 A modified semisynthesis of artemisinin. The genes in the pathway are
heterologously expressed in S. cerevisiae to produce the advanced
intermediate artemisinic acid and further elaborated by semisynthesis
to the drug, artemisinin 53

Figure 2-1 Azaphilone natural products 73

Figure 2-2 Reengineered Biosynthetic Pathway for the Synthesis of (+)-Sclerotiorin
and 7-epi-Sclerotiorin (8) and Non-natural Azaphilone Polyketides 74

Figure 2-3 Concise Synthesis of (+)-Sclerotiorin, 7-epi-Sclerotiorin (8) and Analogs 75

Figure 2-4 A Short Route to Azaphilone Analogs 76

Figure 2-5 Optimization of Induction Time 79

Figure 2-6 Optimization of Culture Time (post-induction) 80

Figure 2-7 to Figure 2-17
1
H NMR and
13
C NMR spectra 81

Figure 2-18 Soybean lipoxygease-1 assay 92

ix
Figure 2-19 to Figure 2-32 LOX-1 Kinetics and Initial velocity graphs 93

Figure 3-1 HPLC profiles of extracts of A. nidulans stcAΔ, hybrid B2 double mutant
under non-inducing (a) and inducing (b) conditions as detected by UV
absorption at 254 nm. The y-axis of each profile was at the same order
of magnitude. *: metabolites that are non-specific to this study 120

Figure 3-2 Proposed biosynthetic pathway of asperfuranone and 1 from native
AfoE (upper) and hybrid AfoE (lower) 121

Figure 3-3 (A) HPLC profiles of extracts of A. nidulans stcAΔ, hybrid B2,
stcJΔstcKΔ triple mutant fed with DMSO (a) and hexanoyl-SNAC (b),
as detected by UV absorption at 254 nm. The y axis of each profile was
at the same order of magnitude. (B) Summary of metabolites produced
by hybrid AfoE based on feeding experiments. Each row lists the
predicted ion formula, retention time (t
R
), m/z of unlabeled form, m/z of
D11-incorporated form, and overall percentage of each metabolite in
negative ionization mode. *: assuming all molecules are equally
ionizable in negative mode 122

Figure 3-4 HPLC profiles of extracts of A. nidulans wild-type strain (a) and stcAΔ
mutant (b) under noninducing (A) and inducing (B) conditions as
detected by UV absorption at 254 nm. Wild type produces terrequinone
(TQ) as well as sterigmatocystin (ST). The y axis of each profile was at
the same order of magnitude. *: metabolites that are non-specific to this
study 123

Figure 3-5 Protein sequence homology alignment of PksA and StcA (only the first
400 amino acids are shown). *: PksA SAT domain cloning sites, which
correspond to K357, T363 and D379 on StcA 124

Figure 3-6 HPLC profiles of extracts of the stcAΔ, hybrid (A1-C3) strains 125

Figure 3-7 UV-Vis and HRESIMS spectra (negative mode) of compound 1 126

Figure 3-8 Key HMBC correlations of compound 1 127

Figure 3-9 High-resolution MS spectra (negative mode) of hybrid AfoE-derived
metabolites (a-i) detected by respectively feeding an equal volume of
(A) DMSO, (B) 0.2 mM hexanoyl-SNAC, (C) 0.2 mM D11-hexanoyl
-SNAC,(D) 0.2mM 1:1 hexanoyl-SNAC: D11-hexanoyl-SNAC to the
stcAΔ, hybrid B2, stcKΔstcJΔ strain. Deuterium-incorporated forms are
highlighted in blue and non-incorporated forms are highlighted in pink.
The y axis (relative abundance) of (a), which is compound 1, is
x
normalized to 1 based on abundance of highlighted peak in B (a). The
y axis of (b)- (e) and (f)- (i) is respectively at 10x and 20x magnitude
of that of (a) 128

Figure 3-10
1
H NMR of 1 130

Figure 3-11
13
C NMR of 1 131

xi
Abstract
Filamentous fungi are prolific producers of bioactive secondary metabolites.
Recent genome sequencing reveals fungi harbor more secondary metabolites than are
currently known. Exploration of fungal secondary metabolism is more attractive today
with recent advancements in genomics and molecular biology. Efficient gene-targeting
technology is a powerful tool used to “mine” the genome for novel secondary metabolites
and identify the genes in the biosynthetic pathway.

Furthermore, this technology can be
applied to engineering pathways to generate “unnatural” natural products. Integration of
biosynthetic engineering with chemical synthesis can introduce greater structural
diversity into natural products, a promising avenue for discovering therapeutic drugs. The
current work describes strategies that utilize the strengths of biosynthetic engineering and
chemical synthesis to generate novel fungal natural products.
Genome mining efforts in Aspergillus nidulans revealed the novel azaphilone
polyketide, asperfuranone; the first azaphilone with its biosynthetic pathway elucidated.
Azaphilone natural products are structurally diverse and exhibit a variety of biological
activities. We hypothesized the asperfuranone pathway can be reengineered to synthesize
a putative intermediate to the azaphilone natural product, sclerotiorin, and apply chemical
synthesis to access additional azaphilone compounds both natural and “unnatural”. Our
strategy uses gene-targeting technology to replace the transcription factor, afoA, promoter
with an inducible promoter and to knock out afoD, a gene that encodes for a hydroxylase,
with the aim of leveraging the biosynthetic machinery to overproduce the putative
intermediate, dimethyloctadiene benzaldehyde. Structural modifications utilizing
xii
synthetic chemistry transform the advanced intermediate into “unnatural” azaphilones in
2-3 steps. They were evaluated for biological activity against lipoxygenase-1 and
provided a structure-activity relationship.
To illustrate the potential of introducing diversity and generating novel
compounds using a single biosynthetic pathway, a domain swapping strategy was applied
to the pathway of asperfuranone. Our strategy to replace the loading domain of the native
polyketide synthase (PKS), AfoE, with the loading domain of sterigmatocystin PKS
would select for a different starter unit and alter the structure of asperfuranone.
Identification of the linking regions between domains and utilization of gene-targeting
technology allowed for the successfully assembly of the hybrid PKS. The engineered A.
nidulans strain produced an unnatural naphthoquinone compound, revealing the hybrid
PKS altered the polyketide backbone. Moreover, this work also suggested two domains
within the non-reduced PKS are chiefly responsible for controlling chain length of the
polyketide. Altogether, a single fungal biosynthetic pathway can be easily manipulated in
multiple ways to produce potentially new bioactive metabolites highlighting the utility
and value of combining chemical synthesis with biotechnology.

1
Chapter I. Introduction

1. Significance of Aspergillus

Aspergillus, a genus of filamentous fungi, is renowned for its medical and
commercial importance. Species in Aspergillus have been sources of lifesaving drugs,
devastating toxins, or mass-produced industrial enzymes. Some species are pathogenic
and pose a danger to immunocompromised patients. Laboratory research on Aspergillus
has also contributed much knowledge about fundamental cell biology and biochemistry.
The significance of Aspergillus was cause for the sequencing of the genomes of some of
the most well known members of this genus. A. fumigatus is a common airborne
pathogen, threatening susceptible patients with infection and life-threatening illness.
Neosartorya fischeri (anamorph A. fischerianus) is genetically closely related to A.
fumigatus, but it is rarely pathogenic. A chief motivation for its sequencing was thus to
learn more about A. fumigatus pathogenicity. In parallel, A. oryzae is a close genetic
cousin to A. flavus. Remarkably, whereas A. flavus is a contaminant of food stocks and a
generator of the potent toxic and carcinogenic aflatoxins, A. oryzae has been safely used
in East Asian cuisine for centuries.
A. terreus, like A. fumigatus, is a significant cause of aspergillosis, but it is also
the main source of the anticholesterol drug lovastatin, with worldwide sales topping $10
billion annually. A. nidulans is a model organism that has been used for the past 60 years
to study genetics and cell biology. Unlike many other Aspergilli, it has a well-
characterized sexual cycle. A. niger has also served as an important model organism; it is

2
also a major producer of enzymes and metabolites, including citric acid. The genomes of
these species, and in addition A. clavatus and A. carbonarius, are now publicly available.
One of the predictions coming from the genome sequencing projects was that our
understanding of the Aspergillus secondary metabolism would profit from the provided
data. Sequence information greatly facilitates the identification of natural product genes,
the function of which can be demonstrated by molecular biological and biochemical
approaches. When a set of genes involved in the formation of the same secondary
metabolite are recognized, a biosynthesis can be proposed. Down the road, such advances
should be useful for enhanced production of secondary metabolites of interest and the
development of second-generation compounds with improved pharmacodynamics and
pharmacokinetic characteristics.
This review examines the benefits genomic sequencing has brought to Aspergillus
secondary metabolite research. As will be detailed below, one of the major findings from
the data is that, given the number of putative secondary metabolite genes that have been
found, many corresponding natural products have yet to be discovered. This untapped
potential of Aspergillus has inspired researches to undertake various strategies to induce
the generation of previously unknown natural products. And for compounds that had
already been reported, the genomic data has been instrumental in identifying genes and
biosynthetic pathways at a rapid pace.
Unquestionably, much important work on Aspergillus secondary metabolism was
completed or in progress before genome sequencing information became available. The
study of the aforementioned aflatoxins (Payne, 1993) and lovastatin (Kennedy, 1999)

3
were among the many works that not only taught us much about secondary metabolite
regulation and organization but were part of the inspiration to sequence genomes in order
to learn more.

2. The Status of Aspergillus Genomic Sequencing Efforts

The task of fully sequencing the first Aspergillus genomes was a community
effort, involving private companies, academic laboratories and research institutes and
funded by private and public entities. For example, A. nidulans FGSC A4 was initially
sequenced by Cereon Genomics (Monsanto) in 1998, the threefold coverage becoming
publicly available in 2003. Shortly thereafter the Whitehead Institute/MIT Center for
Genome Research rereleased the sequence with additional coverage sequence to afford
13-fold coverage. A seminal article on the genome sequence was published in 2005, with
authors representing The Broad Institute of MIT and Harvard, The Institute for Genomic
Research (TIGR), and over 20 universities around the world (Galagan, 2005).
The work and the teamwork, do not stop with the sequence data. Many times the
initial gene annotations are incomplete or contain inaccuracies, such as merged genes and
missed exon calls. On the subject of A. nidulans, a goal spearheaded by the Eurofungbase
in collaboration with TIGR/J. Craig Venter Institute and university laboratories is to raise
the number of functionally assigned proteins. Since then, over 2500 genes have been
edited, and the percent of gene products with an informative name has increased from
about 3% to 19% (Wortman, 2009). Table 1-1 lists several features of sequenced

4
Aspergillus genomes.
The future will likely bring additional Aspergillus sequencing and annotation data
that is the result of collaborations among institutions around the world. But there may
also be a trend in which individual laboratories alone may sequence an Aspergillus
genome (over 180 species left). Because fungal genomes tend to be compact, with few
repetitive sequences, they may be amenable to assembly from the short sequence reads
that come from low-cost, next-generation sequencing techniques, as demonstrated by the
de novo assembly of raw sequence data from Sordaria macrospora (Nowrousian, 2010).
The bioactive secondary metabolite profile of Penicillium aethiopicum motivated
researchers to undertake 454 shotgun sequencing of a P. aethiopicum strain, covering
approximately 90% of the genome (Chooi, 2010).

3. Technological Progress In Genome Mining Endeavors

Methods To Study Existing Secondary Metabolites
Published sequenced genomes allow investigators to pinpoint putative secondary
metabolite genes based on sequence similarity to established genes from other species.
BLAST analysis allows assignment of putative polyketide synthase (PKS), non-
ribosomal peptide synthetase (NRPS), and hybrid PKS-NRPS genes, coding for the core,
backbone structures of many fungal natural products. What is more, these genes together
with tailoring and regulatory genes tend to exist in clusters, which is not the case for
fungal primary metabolic genes or for secondary metabolism genes from other kingdoms,

5
such as plants. Alternative explanations for this phenomenon have been proposed, (Bok,
2004; Rosewich, 2000; Walton, 2000) and although no one explanation has been
universally accepted, it is unmistakable that this feature is fortuitous for researchers, as
the identification of one secondary metabolite gene automatically implicates neighboring
genes as suspects in the formation of the same metabolite. Web-based tools assist in
systematically locating secondary metabolite clusters. One of them, Secondary
Metabolite Unknown Regions Finger (SMURF; www.jcvi.org/smurf), is exclusively
tailored to fungi (Khaldi, 2010). Another, antibiotics and Secondary Metabolite Analysis
Shell (antiSMASH), locates biosynthetic loci involved with the entire range of secondary
metabolite classes (Medema, 2011). Guided by SMURF, Tables 1-2 to 1-5 lists backbone
genes in A. nidulans, A. fumigatus, A. terreus and A. niger, and actual or predicted
products. When candidate genes are identified, several approaches can be taken to study
their function in relation to a natural compound.
The Development of Efficient Genetic Deletion Systems
Gene targeting is a useful technique to study fungal secondary metabolism.
Deletion of the gene, or the replacement of its promoter with a regulatable promoter,
should indicate its significance (if any) in the formation of a particular product. Knocking
out genes has the additional advantage in that intermediates in a biosynthesis may
accumulate in a deletant strain and offer clues to how a natural product is pieced together.
However, the susceptibility of some intermediates to spontaneous rearrangements,
degradation or catalysis by endogenous enzymes can complicate analysis.

6
High rates of correct gene targeting will hasten progress in studying gene
function. However, in Aspergillus species, rates hover as low as 0-20% (Yu, 2004).
Human Ku protein is a heterodimer of two polypeptides, Ku70 and Ku80. Its plays a
chief role in nonhomologous endjoining repair. In Neurospora crassa, correct
homologous integration occurs only 3-5% of the time, but when the homologs to Ku70
and Ku80 were separately disrupted, each strain was able to integrate exogenous DNA
into homologous sequences at a frequency of 100% (Ninomiya, 2004). Based on this
finding, Ku homologs in Aspergillus have been deleted with the effect of substantially
improving homologous integration efficiency. This approach has been demonstrated in A.
nidulans (nkuAΔ) (Nayak, 2006), A. fumigatus (akuAΔ and akuB
KU80
Δ) (Ferreira, 2006;
Krappmann, 2006), A. niger (kusAΔ) (Meyer, 2007), A. terreus (akuBΔ) (Gressler, 2011),
and A. oryzae (ku70Δ and ku80Δ) (Takahashi, 2006a, 2006b). Because correct gene
targeting frequencies are somewhat lower in A. oryzae compared with other Aspergilli, an
alternative method was developed in which a homolog of human DNA ligase IV (ligD)
was disrupted (Maruyama, 2008; Mizutani, 2008). As a further aid to experiments, fusion
PCR allows transforming sequences to be developed without ligation, and, if the genomic
sequence is known, the targeted gene does not have to be cloned (Kuwayama, 2002;
Yang, 2004; Yu, 2004).
Efficient gene knockout was used in the study of the emericellamides (1-5) in A.
nidulans (Chiang, 2008). The emericellamides are molecules of mixed polyketide/peptide
origin that were shown to have antibiotic activity, first found from a marine Emericella
species (Oh, 2007). Emericellamide A and four other related molecules (thus labeled C-

7
F) were soon after isolated from A. nidulans. As expected, both a polyketide and a
nonribosomal peptide synthetase gene are responsible for the formation of the
emericellamides, as deletion of these genes eliminated the compounds’ production. Two
additional crucial genes (EasC and EasD) were identified within the cluster.
Efficient genetic targeting also allowed investigators to delete 32 known and
putative PKS genes and search for any link to observed metabolites (Nielsen, 2011). The
study revealed that inactivating the PKS gene AN8383 eliminated production of two
compounds of mixed polyketide-terpene origin, austinol (6) and dehydroaustinol (7).
AN8383 under the control of an inducible promoter generated 3,5-dimethylorsellinic acid
(8), a likely precursor in the synthesis of the meroterpenoids (Figure 1-2).
Gene Amplification And Overexpression
A common method to study a secondary metabolite gene is to amplify the gene of
interest, overexpress it in a heterologous host, purify the enzyme, and then examine the
enzyme’s function, often by incubating it with a supposed secondary metabolite
intermediate and any cofactors that are predicted to be necessary. Examples provided
below illustrate that the approach has filled in many gaps in our knowledge of various
biosynthetic pathways. Given the substrate promiscuity of many secondary metabolite
enzymes, however, it may be difficult to assign specific function when the encoding gene
is not clustered with other secondary metabolite genes that give context to the overall
biosynthesis.

8
As one example, a consideration of the structure of the mycotoxin
acetylaszonalenin (9), a dipeptide derivative of tryptophan and anthranilic acid that has
been prenylated and acetylated, prompted experimenters to mine the N. fischeri genome
for a cluster containing NRPS, prenylation and acetylation genes (Yin, 2009). The
prenyltransferase gene AnaPT in such a cluster was cloned and overexpressed. To serve
as the starting molecule, the predicted first intermediate, R-benzodiazepinedione (10) was
acquired through synthesis. Recombinant AnaPT catalyzed reverse prenylation at the C-3
position, and at this stage ring formation between C-2 of the indole ring and N-12 of the
diketopiperazine seems to occur as well (Figure 1-3). Isolated product aszonalenin (11)
and acetyl coenzyme A were then incubated with the acetylating protein in the cluster,
AnaAT. The enzymatic product was confirmed to be acetylaszonalenin.
Heterologous Production of Natural Products
A whole set of genes from one organism may be transferred to a host organism
that does not naturally contain these genes. This is a well-established method to induce
the expression of genes from a strain that is less-than-optimal in generating the associated
natural product (Zhang, 2011). Ideally, heterologous expression can be used to assess the
function of these transferred genes. However, the approach has some limitations,
including the difficulty of handling large genes and gene clusters, as well as finding a
suitable host.

9
Despite these challenges, researches succeeded in reconstituting up to five steps
of the biosynthesis of meroterpenoid pyripyropene A (12) from A. fumigatus, a potent
inhibitor of acyl-coenzyme A: cholesterol acyltransferase, using A. oryzae M-2-3 as a
host (Itoh, 2010). In the search for a terpene cyclase they discovered a protein with
unusual sequence and primary structure. The protein bears homology to an efflux pump,
raising the possibility that it in fact serves a dual purpose: catalyzing a late-stage
cyclization step and, perhaps as a self-resistance move, exporting the metabolite from the
cell. Further, the experimenters fed benzoic acid, instead of the natural precursor
nicotinic acid, to a transformant and ultimately obtained a close analog to a compound
that is an inhibitor of inducible nitric-oxide synthase in human cells. Their approach
points to the flexibility of secondary metabolite biosynthetic enzymes, allowing the
formation of novel metabolites with distinct biological properties.
Uncovering Hidden Biosynthetic Pathways
It came as a surprise to many researchers that Aspergillus genomes were
indicating that the number of putative biosynthetic genes far exceed the number of
secondary metabolites that had been acquired from those species. Because of the
extensive investigation of some of these fungi, it is unlikely that many secondary
metabolites have simply escaped notice. Rather, it is plausible to conclude that most of
the genes that have not yet been associated with a secondary metabolite are not expressed
or expressed in very low amounts in a laboratory culture setting. The discovery of the
untapped metabolic potential of Aspergillus both inspires excitement and poses a

10
challenge: How can researchers unearth the hidden secondary metabolites of Aspergillus,
especially with an incomplete knowledge of their regulation?
Remarkably, various strategies have been successful to this end, many of which
take advantage of some interesting characteristics of fungal secondary metabolism.
Besides the aforementioned clustering phenomenon, many (but not all) secondary
metabolite gene clusters contain a class of zinc binuclear (Zn(II)
2
Cys
6
) transcription
factor genes, which are unique to fungi (Yin, 2011).
Controlling Expression of Cluster-Specific Transcriptional Activators
In A. nidulans a plasmid carrying the transcriptional gene apdR under the control
of the inducible alcA promoter was ransformed, leading to the upregulation of the
putative entire cluster to which the gene belongs (Bergmann, 2007). As a result, two new
related polyketide-nonribosomal peptide hybrids, aspyridones A and B (13-14), were
acquired.
In another approach that also took note of the fact that many secondary metabolite
gene clusters contain their own regulatory gene, the native promoter of one of these
genes, afoA, was replaced by an inducible promoter, and as a result the novel polyketide
asperfuranone (15), with structural similarities to the azaphilone class of natural products
was generated (Chiang, 2009). Under alcA-inducing conditions, the nearby genes could
be deleted and analyzed for their effect on asperfuranone formation. Asperfuranone was
found to come from the product of a highly reduced PKS, which was then loaded onto a
non-reduced PKS and then further tailored to lead to the final product.

11
LaeA, A Global Regulator of Secondary Metabolism
LaeA (Loss of aflR expression A), a nuclear protein, was first found in A.
nidulans (Bok, 2004; Butchko, 1999), and orthologs have since been identified in A.
fumigatus, A. terreus, A. flavus (Kale, 2008), as well as in fungi beyond Aspergillus
(Kosalkova, 2009; Wiemann, 2010; Xing, 2010; Zhang, 2009). Its deletion was found to
cause the decrease of sterigmatocystin and penicillin production in A. nidulans and of
gliotoxin production in A. fumigatus (Bok, 2005) and lovastatin in A. terreus (Bok, 2004).
Deletion also leads to reduced virulence in A. fumigatus (Bok, 2005; Dagenais, 2009) and
A. flavus (Amaike, 2009). Overexpression of LaeA, on the other hand, boosted the
generation of the above products. The data suggest that the protein has a broad, global
role in secondary metabolite regulation. Indeed, microarray data from A. fumigatus
revealed that up to 13 of 22 studied gene clusters were affected by LaeA (Perrin, 2007).
Comparing the deletion and overexpression mutants of LaeA in A. nidulans was
instrumental in defining the terrequinone A (16) gene cluster (Bok, 2006; Bouhired,
2007).
Each of the five terrequinone A genes was characterized by another group through
heterologous expression (Balibar, 2007). TdiD was established as a pyridoxal-5’-
phosphate-dependent L-tryptophan aminotransferase that converted tryptophan to indole
pyruvic acid (17). Next, the tridomain NRPS TdiA couples two tethered units together to
yield dimethylasterriquinone D (18). The combination of TdiB, TdiC, and TdiE was
reported to be necessary to proceed substantially to terrequinone A, and that TdiC is an
NADH-dependent quinone reductase, generating a hydroquinone that is primed for two

12
prenylations catalyzed by TdiB (Balibar, 2007). TdiE is necessary to direct the pathway
away from a shunt metabolite (Figure 1-5). Concurrent independent research was in
agreement about the roles of TdiA and TdiD (Schneider, 2007) and characterizes TdiB as
a catalyst for reverse prenylation of the indole moiety (Schneider, 2008).
Recent methodology, including the employment of suppressor screens that can
remediate loss of laeA, may uncover new global regulators of secondary metabolism. For
examples, such a screen identified a bZIP protein called RsmA (Restorer of secondary
metabolism A), which was independently able to enhance secondary metabolite
production in A. nidulans (Shaaban, 2010).
Based on its homology to histone and arginine methyltransferases, LaeA is
believed to function by influencing chromatin. As chromatin can exist in an “open”
(euchromatin) or “closed” (heterochromatin) state, experimenters considered the
possibility that the manipulation of the interplay between these two states may open up
previously closed sections of the genome to the transcriptional machinery. LaeA, in fact,
was shown to reverse the establishment of heterochromatic marks (Reyes-Dominguez,
2010). Subsequent research has confirmed that control of chromatin remodeling in
Aspergillus and elsewhere may affect its metabolic profile.
Chromatin Restructuring Through Enzyme Inhibitors or by Deletion of A
Chromatin Remodeling Protein

Various enzymes catalyze the addition or removal of small functional groups,
such as acetyl, methyl, and phosphoryl, onto the tails of histone proteins. These groups
appear to influence the architecture of the surrounding chromatin; for instance, acetylated

13
histone tails are generally associated with the open state, whereas deacetylated histones
are related with closed chromatin. It was postulated that blocking the enzymes that
catalyze deacetylation could potentially open up previously inaccessible secondary
metabolite genetic loci and in turn bring about its associated natural product. Indeed,
treatment of Alternaria alternate and Penicillium expansum with the histone deacetylase
(HDAC) inhibitor trichostatin A (TSA) induced the production of a number of undefined
metabolites (Shwab, 2007). In A. niger, the addition of the HDAC inhibitor
suberoylanilidehydroxamic acid (SAHA) generated the production of nygerone A (19),
containing a unique 1-phenyl-pyridin-4(1H)-one core (Henrikson, 2009). Investigators
also considered the effects of removing critical members of chromatin regulatory proteins
through targeted gene deletion. A. nidulans contains CclA, a putative ortholog of the
Saccharomyces cerivisiae Bre2 protein, belonging to a complex (COMPASS), which
both activates and represses chromatin-related processes through lysine 4 histone H3
methylation. Two classes of secondary metabolites emerged when the cclA gene was
knocked out: emodin (20), monodictyphenone (21), and related compounds, which all
depend on the PKS gene AN0150; and F9775A/B (22-23), which stems from the gene
AN7909 (Bok, 2009).
The research so far on the epigenetic link to secondary metabolites suggests that
alternations in the chromatin landscape can have a dramatic effect on a fungus’s
metabolic profile, although no work to date has reported that one single approach yields
the entire array of hidden natural products from one fungus. Nevertheless, epigenetic
strategies, if they cannot access an entire genome, are still useful in opening up at least

14
part of the secondary metabolome that had previously been closed off to researchers. In
addition to histone proteins, other targets may touch upon secondary metabolite
regulation. The single sumoylation gene of A. nidulans, sumO, was shown to affect the
production of several metabolites (Szewczyk, 2008). The gene’s removal led to decreased
amounts of austinol (6) and dehydroaustinol (7) production but also a significant increase
in the generation of the polyketide asperthecin (24). The substantial amount of
asperthecin permitted researchers to identify a cluster of three genes containing
responsible polyketide synthase gene, a gene thought to be needed for the release of the
polyketide from its enzyme, and a monooxygenase gene. It is not yet clear how the
deletion of sumO affects A. nidulans secondary metabolism, but as the small product
SUMO is added post-translationally to a variety of proteins, it may play a role in the
regulation of at least some secondary metabolites.
Mimicking Ecological Systems Through Co-Incubation With Microorganisms
Another successful strategy was based on the realization that, in nature, fungi
often co-inhabit ecosystems with other species, and indeed the purpose of many
secondary metabolites apparently to is to assist the producing organism to secure its
niche, whether the interaction is antagonistic or symbiotic. This has indeed translated to
the induction of secondary metabolites when a fungus is co-incubated with another
organism (Cueto, 2001; Oh, 2005, 2007; Zuck, 2011). Again in A. nidulans, it was
demonstrated that cultivation with a particular actinomycete, Streptomyces
rapamycinicus, prompted the upregulation of 248 genes including a PKS and NRPS gene

15
(Schroeckh, 2009). It was demonstrated that, rather than arising from a chemical signal,
the induction was triggered by the physical interaction of the two organisms. Metabolic
profiling revealed the emergence of the polyketide orsellinic acid (25) as well as
lecanoric acid (26) and F-9775 A and B (22-23), which may be derived form the simple
tetraketide, found to be encoded by orsA (AN7909) (Sanchez, 2010; Schroeckh, 2009).
One Strain Many Compounds (OSMAC)
Despite the unmistakable value of sophisticated molecular biological research in
uncovering previously undetected secondary metabolites, a potentially equally powerful
method involves little more than a change in recipe. Articulated as OSMAC (One Strain-
Many Compounds), the strategy endeavors to expose the metabolic diversity of fungi
(among other organisms) by subjecting the same strain to a number of different culture
conditions (Bode, 2002). A frequent approach is to alter the broth composition in which
the organism is cultivated. Other parameters may be adjusted, such as temperature,
shaking, speed, and even flask shape and size.
Our understanding of the mechanisms that make OSMAC effective can only be as
complete as our knowledge of the complex regulatory networks that influence secondary
metabolism. It is realized that heightened glucose, ammonium, or phosphate
concentrations often (but not necessarily) curb secondary metabolism formation
(Masuma, 1986; Omura, 1982). However, in many cases it is not clear why a certain
environment results in the desired effect of a new metabolite. Considering the facts that
altering culture conditions is usually not labor intensive, but that many attempts will not

16
lead to a difference metabolic profile, a sensible course of action is to test out many
different conditions at once.
OSMAC inspired the discovery that simple culture conditions, namely A. nidulans
in a sucrose-based media without shaking, led to the formation of orsellinic acid (25) and
F-9775 A and B (22-23) (Sanchez, 2010). Interestingly, F-9775A and B has now been
demonstrated to be a result of three separate approaches: chromatin remodeling (Bok,
2009), co-incubation with an actinomycete (Schroeckh, 2009), and alternation of culture
conditions. The straightforward nature of changing culture conditions facilitated the
discovery of a three-gene cluster through genetic deletion analysis and the acquisition of
two bioactive compounds, gerfelin and diorcinol, from two of the knockout strains. As
other examples, one group prepared 40 different conditions and found that one (and not
the 39 others) yielded four related quinoline-2-one alkaloids named aspoquinolines (27-
30) (Scherlach, 2006). An additional round of investigation was rewarded with the
discovery of the prenylated isoindolinone alkaloids, aspernidine A and B (31-32)
(Scherlach, 2010).

4. Genetic Characterization of Prenylated Xanthones From A. nidulans

Investigation into the biosynthesis of the naturally occurring prenyl xanthones of
A. nidulans (33-36) revealed that the identified genes are separated into three distinct
genomic loci (Sanchez, 2011). The two necessary prenyltransferase genes are distant
from a main cluster, and another gene is proximal to one of the prenyltransferase genes
(Figure 1-8). This and other examples of “non-cluster” in the literature (Inderbitzin, 2010;

17
Nicholson, 2009; O'Donnell, 2000) suggest that fungal secondary metabolite genetic
lustering should be thought of as a guiding principle rather than an unbreakable rule. The
study also detailed that the main cluster of genes was the same as the one responsible for
emodin (17), monodictyphenone (18), and related compounds, which were only detected
in a chromatin remodeling mutant strain. The genes outside the cluster complete a
biosynthesis in which emodin and monodictyphenone may be intermediates.
These findings suggest that applications of strategies that aim to upregulate a
genetic cluster may not equally influence genes that pertain to the biosynthesis but reside
outside the cluster. Consequently, the “real” natural product may not be formed, but
rather one or more molecules representing a portion of the biosynthesis. However, since
many intermediates on their own possess potent biological activity, an original aims of
these aforementioned approaches, the generation of bioactive secondary metabolites, may
be satisfied after all.

5. Expanding Natural Product Diversity

Nature has, over millions of years, methodically constructed vast numbers of
complex secondary metabolites that are optimized for interaction with biological systems
to secure the niche of the organism. Natural products, however, are not optimized for
drug-like properties, yet still they remain an important source for therapeutic agents with
notable examples such as penicillin, lovastatin, and rapamycin. Despite past successes,
natural product-based drug discovery still has challenges in isolation and optimization of
leads by standard chemical synthesis or high-throughput approaches largely due to the

18
structural complexity of the natural product (Li, 2009). This has prompted the utilization
of combinatorial biosynthesis methods (Figure 1-9). These approaches integrate
biotechnology with chemical synthesis for development of analog libraries of natural
product leads. And over the years they have proven to be valuable tools for drug
discovery.
Combinatorial biosynthesis, in the larger sense, is applying genetic engineering to
modify biosynthetic pathways to leverage the potential of nature’s biosynthetic
machinery for producing novel bioactive compounds. Methods for generating “unnatural”
products involve a combination of chemical synthesis and some form of genetic
manipulation that consist of either disruption of a gene, overexpression of a gene, or
expressing a gene or a combination of different genes in the producing organism or
heterologous host. These engineering strategies are remarkable feats that arise from the
advancements in genome sequencing, bioinformatics, and molecular biology. Here
described are the in vivo strategies for engineering pathways in microbes to expand the
structural diversity of natural products with examples in literature that highlight both the
opportunities and limitations. For additional coverage on biosynthetic engineering the
reader is referred to excellent reviews on polyketides (Floss, 2006) and non-ribosomal
peptides (Sussmuth, 2011).

19
Precursor-Directed Biosynthesis
Precursor-directed biosynthesis (PDB) constructs secondary metabolite analogs
by synthesizing simple precursors and then outsourcing the chemical demanding step(s)
to the wild-type microbe to produce the modified metabolite. Typically, an experiment
supplements the growth medium with the surrogate precursor (also called mutasynthon),
then after cellular-uptake it has to compete or out-compete the natural substrate for
selection by the enzyme(s). Of course, success of this strategy and related strategies
depends on toxicity of the unnatural intermediates, and the degree of flexibility of the
downstream enzyme(s) toward the mutasynthon. This strategy has been used to modify
many different natural products (Thiericke, 1993). Although it does not involve
biosynthetic engineering, it still is an important method and may be the best method in
cases where the gene of interest is essential for survival of the organism.
A common approach to PDB and others is using alternative PKS starter units
(Moore, 2002) to interrogate the flexibility of enzymes. The polyketide natural product,
rapamycin (37) is produced by Streptomyces hygroscopicus and has potent
immunosuppressant activity. In 1994, a group of researchers sequenced the entire
biosynthetic gene cluster of 37 (Aparicio, 1996; Schwecke, 1995) and later identified the
starter unit of rapamycin as 4,5-dihydroxycyclohex-1-enecarboxylic acid (38) (Lowden,
2001). Moreover, significant incorporation of 38 suggested a relaxed substrate specificity
of the loading module in the PKS. The previous work paved the way for researchers to
explore modifying the structure of the cyclohexane ring on rapamycin by feeding 21
different carboxylic acids separately to S. hygroscopicus (Lowden, 2004). The alternative

20
substrates tolerated by the rap PKS were limited to simple precursors but generated new
rapamycin analogs (42-44), albeit in lower titer, and in addition to rapamycin (Figure 1-
10A). Later, a mutasynthesis example expands the scope of surrogate precursors
incorporated in the scaffold of 37.
Although PDB is amenable to structural alternations at early stages of the
biosynthesis, it is not limited to only starter unit modifications. Researchers discovered
rapamycin production is reduced in the presence of proline molecules because they
inhibit the production of the natural building block, L-pipecolate (Graziani, 2003). A
strategy was devised to modify the pipecolate ring on 37, where a culture of S.
hygroscopicus was supplemented with the enzyme inhibitor (±)-nipecotic acid (45) and
two unusual pipecolate analogs (46-47) that produced new rapamycin derivatives
(Figure1-10B). These new metabolites, however, showed weaker binding to FKBP12
(Ritacco, 2005).
Even though the structural modification did not improve upon the biological
activity of rapamycin in some cases it may lead to the development of a completely new
type of bioactivity. The peptide lactone, hormaomycin (50) produced by Streptomyces
griseoflavus strain W384 displays antibacterial activity. Interestingly, 50 contains a
unique cyclopropyl moiety found to originate from lysine. Preliminary feeding
experiments suggested that the unnatural amino acid, 3-nitrocyclopropyl alanine, is a
suitable surrogate precursor for the bacterial NRPS (Figure 1-11) (Kozhushkov, 2005).
Feeding the unnatural amino acid to the producing organism generated new
hormaomycin analogs (Zlatopolskiy, 2006). Interestingly, when substrate (54) was

21
enriched in the medium the microbe generated multiple unexpected analogs resulting
from an unseen metabolic fate. Secondly, these metabolites were difficult to separate
from the competitively produced hormaomycin, an inherent limitation of PDB. Despite
the complications, hormaomycin analogs were tested for antibacterial activity and in an
unlikely turn of events, 57a showed potent activity against the fungal pathogen C.
albicans, with comparable activity to the antimycotic agent nystatin.
Mutational Biosynthesis (Mutasynthesis, MBS)
A more attractive approach to augment natural product diversity and the most
successful of combinatorial biosynthesis methods is mutational biosynthesis or
mutasynthesis (MBS) as coined by Rinehart in the late 1970’s (Rinehart, 1977). In
mutasynthesis alternative biosynthetic precursors or early stage intermediates are fed to a
mutant that is blocked in the ability to biosynthesize the natural precursor or intermediate
to exclusively produce the modified natural product which is then evaluated for
biological activity. It was a novel concept over 40 years ago and first demonstrated with
the aminoglycoside antibiotic, neomycin (Shier, 1969). Genetically engineered microbes
can be created either by mutagenesis, strain evolution, or if there is sequence information
available by targeted gene deletion or inactivation. MBS improves upon precursor-
directed biosynthesis by removing internal competition, which facilitates the uncontested
incorporation of mutasynthons thereby expanding the number of possible metabolites
generated, and simplifying the isolation of the analog.

22
In MBS, a common strategy to introduce diversity is the inactivation or deletion
of a gene responsible for the biosynthesis or regulation of the starter or primer unit
(Eichner, 2009). Rapamycin has been the subject of many studies because it represents a
lead structure for the development of immunosuppressive and anti-cancer agents. And all
rapamycin analogs in clinical trials are structurally modified in the starter unit, as such
Biotica explored application of mutasynthesis to generate a broader range of analogs
(Gregory, 2005). To apply mutasynthesis a mutant blocked in ability to form the
polyketide primer unit needs to be created and serendipitously, the desired mutant was
made when the post-PKS genes (rapKIJMNOQL) were excised from the genome of the
wild-type and the mutant, MG2-10, could not produce rapamycin. However, pre-
rapamycin (58) production was restored when fed 59, subsequently a variety of
carboxylic acids (Figure 1-12) were then supplemented to the mutant and exclusively
incorporated into the rapamycin scaffold. Moreover, mutasynthesis made possible the
incorporation of a norborane carboxylic acid, whereas only simple acids incorporated in
the PDB approach. In the same study, production of pre-rapamycin was also restored
with expression of the rapK gene. Sequence analysis revealed that rapK regulates
production of the starter unit.
Extender units of PKSs are also amenable to manipulation in mutasynthesis.
FK506 (64) is similar to rapamycin except it contains an allyl moiety at C-21 found to be
critical for biological activity. A recent study investigated the allylmalonyl-CoA
(coenzyme A) pathway in the producing organism of FK506 and found a dedicated PKS
(TcsB) biosynthesizes the unique extender unit (Mo, 2011). The deletion of tcsB gene

23
abolished production of FK506 but complementation of allylmalonyl-SNAC restored
FK506. In addition, the PKS of 64 suggests broad substrate tolerance based on the trace
amounts of 65 and 66 in the wild-type organism. In an effort to create FK506 analogs,
mutasynthesis was employed to derivatize the scaffold at C-21 using a ΔtcsB strain.
When culture was supplemented with carboxylic acids (67-69) the mutant successfully
produced 70, 71, and 72 which are not readily accessible by semisynthesis (Figure 1-13).
The immunosuppressive activity was not improved; however, 71 did exhibit improved
neurite outgrowth activity over FK506.
Improvement of the natural product’s pharmacological properties is the primary
aim to generate compound libraries from natural products, an example are the
ansamitocins (73-75). These highly potent antitumor compounds were targeted for
modification at the starter unit, 3-amino-5-hydroxybenozic acid (AHBA). Analogs of
ansamitocins were created by supplementing a mutant blocked in forming AHBA with
halogenated benzoic acid derivatives produced the corresponding halogenated
ansamitocin (76-78) (Taft, 2008). The researchers extended their work by combining
mutasynthesis with semisynthesis to demonstrate this approach as a powerful way to gain
access to potentially more bioactive analogs not available by mutasynthesis alone. To
illustrate the potential of the combined approach, researchers performed a Pd-catalyzed
Stille reaction with the brominated metabolite to install a vinyl moiety at C-19 (Figure 1-
14), consequently 79 was one of most active compounds tested amongst the analogs
mutasynthesized. Further investigations proved this approach can access folic
acid/ansamitocin conjugates that target cancer cells (Taft, 2012). Likewise, the Hertweck

24
group integrated mutasynthesis, biotransformation and combinatorial biosynthesis to
achieve (theoretically) up to 77 different aureothin analogs (Werneburg, 2010).
Interestingly, the producing organism limited the formation of aureothin derivatives due
their toxicity on the microbe. To get around this hurdle, deoxyaureothin intermediates
were mutasynthesized and subsequently fed to a heterologous host expressing the last
enzyme in the pathway, AurH to transform the intermediate to the final product.
One of the most successful examples to come from mutasynthesis is the discovery
of the commercial antiparasitic drug, doramectin (80). Streptomyces avermitilis produces
a group of macrolides called avermectins. With the biosynthesis of avermectins
elucidated, a mutant strain ATCC 53568 lacking the ability to create starter units
isobutyric and S-2-methylbutyric acids, presented the opportunity to supplement the
organism with other acids. The incorporation of alternative precursors produced a series
of novel avermectins, which included doramectin (Dutton, 1990). In another study, a
cluster of genes was discovered to encode the biosynthesis of cyclohexanoic (CHC) acid
(Cropp, 2000). To bypass issues of insufficient amounts of precursor and transport, the
set of five genes for CHC acid biosynthesis were heterologously expressed in the S.
avermitilis mutant and successfully produced doramectin without exogenous addition of
acid (Cropp, 2000). This is one example of combinatorial biosynthesis (Figure 1-9e),
which will be discussed in detail later.
Literature is scattered with examples of employing mutasynthesis as a tool to
produce structural analogs of bioactive metabolites, other prominent examples are
salinosporamide A (Eustaquio, 2008), geldanamycin (Eichner, 2009), and borrelidin

25
(Moss, 2006). The rationale for using mutasynthesis or any other combinatorial
biosynthetic tool is to generate analogs difficult to access by conventional synthetic
methods. In some cases, like borrelidin these tools eliminated labor-intensive steps and in
other cases chemical synthesis is a more appropriate method for analog construction.
Combinatorial Biosynthesis
Combinatorial biosynthesis is another drug discovery tool that rationally alters the
structure of a polyketide by swapping biosynthesis genes in a pathway, to either modify
functional groups on the polyketide or reconfigure the polyketide backbone. From an
enzymatic perspective, either a specific enzyme can be structurally manipulated (e.g.
modules within a polyketide synthase) or a tailoring enzyme is replaced with a homolog
(Olano, 2010). The latter technique is most commonly used to generate analogs but both
have been applied to generating unnatural polyketides (Hertweck, 2009) and macrolide
antibiotics.
The former approach constructs hybrid PKSs and several ways have been
successful including loading module swaps, module insertions (Rowe, 2001) and acyl
transferase (AT) domain swaps (Oliynyk, 1996). The loading module swap is a common
strategy used for generating unnatural polyketides. In one example, production of
doramectin was devised by replacing the native loading module of the Ave PKS with the
loading module of the phoslactomycin B PKS, which selects for the CHC-CoA starter
unit. The experiment revealed a 6-fold increase in production of 80 (Wang, 2011).
Likewise, spinosyn A (81) is a macrolide with potent insecticidal activity where

26
semisynthetic methods could not access analogs with modifications at the “western
shore” of the 81. Previous work identified that the erythromycin (ery) PKS loading
module and the avermectin (avr) PKS loading module they accept a range of substrates.
Investigators individually swapped loading modules with the spinosyn’s PKS loading
module creating ery/spn and avr/spn hybrid enzymes which generated a library of novel
compounds modified at C-21 (Sheehan, 2006). A limitation of this method was a
reduction in titer of the new metabolites using the engineered strains attributed to non-
optimal protein-protein interaction or the substrate tolerance of downstream domains.
Despite this, selected analogs display improved insecticidal activity towards other species
illustrating the value of biosynthetic engineering.
In addition to generating potentially bioactive compounds, combinatorial
biosynthesis can be a useful tool in answering fundamental mechanistic questions about
enzymes. Fungal polyketides are assembled by iterative PKS’s that can be further
classified by non-reducing, partially-reducing or highly-reducing. The Cox group
investigated TenS, a highly-reducing PKS (HR-PKS) that biosynthesizes tenellin (82), in
a domain swap approach to uncover the origins of control on polyketide chain length and
methylation patterns (Fisch, 2011). During the study the KR domain of TenS, which
catalyzes the reduction of a keto group, was replaced by dmbS-KR and coexpressed with
other enzymes in the tenellin pathway (Figure 1-16). The result was the in vivo
production of a so-called extinct metabolite, bassianin (83), which was discovered in a
Beauveria strain in 1977 (Wat, 1977) but since has never been found in any Beauveria
species. Secondly, results also revealed CMET and KR domains play a dominant role in

27
determining methylation and chain length.
Modified Semi-Synthesis
There are circumstances where natural products cannot proceed toward clinical
development due to an unsustainable supply from the producing organism. In such cases,
traditional semisynthesis has been able to address this issue. A historical example is the
semisynthetic route developed for the production of Taxol (84), one of the most potent
anticancer drugs used commercially. Taxol was first isolated from the bark of a yew tree;
however, the problem of obtaining enough of the compound was soon realized. It was
then reported that a biosynthetic intermediate baccatin III (85) could be isolated from the
needles of the tree and then chemical synthesis transforms the complex terpene in several
steps to the drug (Figure 1-17) (Holton, 1995). An effort to find an improved way of
generating 84 was the recent investigation in the overproduction of a Taxol precursor
(86) in a microbial host via metabolic engineering (Ajikumar, 2010). Previous studies at
reconstitution of the isoprenoid pathway and overexpression of the pathway enzymes
provide less than 10 mg/liter of 84. Researchers changed their approach and applied a
multivariate modular engineering strategy, which partition the terpenoid pathway and
varied expression of these modules simultaneously (Figure 1-17). The study identified
parameters that influenced pathway flux and boosted production of taxadiene (86) by
15,000-fold over the control. Moreover, the engineered E. coli strain also improved the
production of taxdien-5α-ol (87) by 2400-fold over yeast.
Semisynthesis over recent years has incorporated metabolic engineering to

28
address the issue of economic production of a drug or drug candidates. For example,
artemisinin (88) is a sesquiterpene anti-malarial drug extracted from the plant Artemisia
annua. Although a chemical synthetic route exists, it cannot meet the demand for 88 at a
low cost for those in the developing world. To resolve this issue, researchers engineered a
yeast strain to overproduce the building block, farnesyl diphosphate, and in addition, co-
express amorphadiene synthase and cytochrome P450 monooxygenase to produce
artemisinic acid (89) (Figure 1-18) (Ro, 2006). This biosynthetic intermediate was
converted to 88 by chemical synthesis in two steps (Roth, 1989). This example
showcases an advanced intermediate of a drug can be cheaply produced by leveraging the
biosynthetic machinery in microbes and minimal synthesis is required to obtain the
bioactive compound.
The interplay between biosynthetic engineering and chemical synthesis has
illuminated new tools to structurally modify complex natural products that were
inaccessible by total synthesis. Judicious application of these methods has successfully
led to the advancement of natural product clinical candidates (Koehn, 2012); however,
there remain hurdles to overcome. A common drawback in most studies is the insufficient
knowledge of substrate specificity, the inefficient processing of unnatural precursors, as
well as other contributing factors, often lead to low titers (less than 2 mg/l), sometimes
amounts only detectable by high-resolution mass spectrometry (HRMS). However, low
titer can be alleviated by utilizing different metabolic engineering strategies (Pickens,
2011). Despite the limitation, useful insights can still be gleaned from the structure-
activity relationships (SAR) of natural product libraries and the flexibility of the

29
biosynthetic machinery in a pathway. Other hurdles are toxicity from unnatural
intermediates, which may be resolved with the utilization of a heterologous host. Two
other bottlenecks are the limited availability of precursors and the regulatory controls
operating within the organism. Moreover, the strategies mentioned herein do not fully
encompass all current methods, such as in vitro pathway engineering (Kwon, 2012), and
protein engineering (Zabala, 2012). Additionally, biosynthetic engineering of pathways in
fungi and plants (Runguphan, 2009) is less common, however, progress is being made
(Ajikumar, 2010). Overall combinatorial biosynthetic methods will continue to advance
in ways to generate novel compounds from natural product leads, but also gain a deeper
understanding of how pathways can be manipulated with aid from ongoing advances in
genome sequencing, informatics, and molecular biology of microorganisms.

30

31
33

32

33
33

34

35
33

36

Figure 1-1. Structures of emericellamides A, and C-F produced by A. nidulans

37

Figure 1-2. The biosynthetic pathway for austinol and dehydroaustinol

38

Figure 1-3. The biosynthetic pathway for acetylaszonalenin

39

Figure 1-4. A) Structure of pyripyropene A B) Structures of aspyridone A, B and
asperfuranone discovered by controlling expression of cluster-specific transcriptional
activators

40

Figure 1-5. The biosynthetic pathway for terrequinone A

41

Figure 1-6. Structures of secondary metabolites A) discovered from chromatin
restructuring B) discovered through co-incubation with microorganisms

42

Figure 1-7. Structures of aspoquinolines A-D and aspernidines A-B discovered by
OSMAC strategy

43

Figure 1-8. The biosynthetic pathway for prenyl xanthones in A. nidulans

44

Figure 1-9. a) A biosynthesis pathway in a WT organism b) In PDB the biosynthesis
pathway uses both the natural substrate and the mutasynthon to produce the natural
product and the unnatural product c) In MBS the biosynthesis pathway is engineered to
eliminate the availability of the natural substrate and only uses the mutasynthon to
produce only the unnatural product d) In modified semi-synthesis the pathway is
engineered to prematurely stop at the intermediate stage and then structurally modified
through chemical synthesis to the natural product analog e) In CB (i) polyketide domain
swap takes a domain from the PKS of organism #2 to replace the same PKS domain in
organism and produces an unnatural polyketide different from the polyketide of organism
#1 and #2 ii) replacement of post-PKS gene takes a tailoring gene from one pathway and
adds the gene to a different biosynthesis pathway to produce a modified natural product

45

Figure 1-10. Precursor-directed biosynthesis of A) rapamycin analogs with modification
at the cyclohexane moiety B) rapamycin analogs with modification at the pipecolate ring

46

Figure 1-11. Precursor-directed biosynthesis of hormaomycin analogs

47

Figure 1-12. Mutasynthesis of pre-rapamycin analogs

48

Figure 1-13. A) Structures of natural products FK523, FK520 and FK506 produced by S.
hygroscopicus NRRL5491 B) mutasynthesis of FK506 analogs with selected
modification at C-21

49

Figure 1-14. A) Structures of natural products ansamitocin P-2, P-3 and P-4 B)
Mutasynthesis of ansamitocin P-3 analogs C) Synthetic transformation under Stille
conditions to install an alkene on an aromatic ring of ansamitocin scaffold

50

Figure 1-15. A) Structure of avermectin analog, doramectin B) Structure of
spinosyn A and selected modification on the western shore of the spinosyn scaffold
using combinatorial biosynthesis

51

Figure 1-16. A) Structure of tenellin and B) Combinatorial biosynthesis generated
bassianin using the hybrid PKS of tenellin

52

Figure 1-17. The engineered taxol pathway (dark arrows indicate genes overexpressed)
A general semisynthesis of the baccatin III intermediate is transformed in two steps to
taxol

53

Figure 1-18. A modified semisynthesis of artemisinin, where select genes in the
pathway are heterologously expressed in S. cerevisiae to produce the advanced
intermediate artemisinic acid and further elaborated by semisynthesis to the drug,
artemisinin

54
Chapter II: Reengineering An Azaphilone Biosynthesis Pathway In Aspergillus
nidulans To Create Lipoxygenase Inhibitors

1. Abstract

Sclerotiorin, an azaphilone polyketide, is a bioactive natural product known
toinhibit 15-lipoxygenase and many other biological targets. To readily access
sclerotiorin and analogs, we developed a 2-3 step semisynthetic route to produce a variety
of azaphilone starting from an advanced, putative azaphilone intermediate (5)
overproduced by an engineered strain of Aspergillus nidulans. The inhibitory activities of
the semisynthetic azaphilones against 15-lipoxygenase were evaluated with several
compounds displaying low micromolar potency.
2. Introduction
Lipoxygenases (EC 1.13.11.12) are ubiquitous enzymes widely distributed within
plants, fungi, and mammals (Brash, 1999). They are nonheme iron dioxygenases that
catalyze the addition of molecular oxygen to polyunsaturated fatty acids with a cis,cis-1,4
pentadiene to generate a hydroperoxydiene formed through a radical, regio- and
stereoselective mechanism (Schneider, 2007). Reaction products of lipoxygenases are
involved in several common human such as allergies, inflammation, and asthma.
Lipoxygenases are responsible for the oxidation of lipids in foods subsequently reducing
the foods’ nutritional value (Gordon, 2001).

55
Several natural products from microbial sources inhibit 15-lipoxygenase (15-
LOX) (Komoda, 2003; Rao, 2002). More recently the fungal pigment, (+)-sclerotiorin
(1), was found to inhibit lipoxygenase-1, also known as 15-LOX (Chidananda, 2007).
Sclerotiorin was first isolated from P. sclerotiorum in 1940 (Curtin, 1940). Since then, 1
has been found to inhibit multiple therapeutic targets (Osmanova, 2010).
Sclerotiorin belongs to an important class of natural products called azaphilones.
Azaphilones are structurally diverse polyketides that share a highly oxygenated bicyclic
core and chiral quaternary center. These polyketides are known for their 4H-pyran motif,
which reacts with amines to produce the corresponding vinylogous γ-pyridones (Wei,
2005). Early synthetic studies by Whalley and co-workers reported the total synthesis of
several azaphilones, which included compound 8 prepared in 14 steps (Chong, 1971).
Recent synthetic efforts by Porco and co-workers have shown assembly of the azaphilone
core through a copper-mediated enantioselective dearomatization route. The application
of their asymmetric methodology was demonstrated on (-)-S-15183a, (-)-mitrorubin, and
more recently with 1 (Figure 2.1) (Germain, 2011; Zhu, 2006, 2005).
Although many azaphilones have been isolated and identified, their biosynthetic
pathways remained unknown until our recent identification of the asperfuranone (4)
biosynthetic pathway in Aspergillus nidulans (Chiang, 2009). A mutant strain from the
previous study provided aldehyde 5 as a stable intermediate, which has been isolated
from other azaphilone-producing organisms. Our work aims to enhance the production of
the putative azaphilone intermediate (5) and use synthetic chemistry to structurally
diversify 5 into natural and non-natural azaphilones.

56
3. Results & Discussion

In this study, the fungal strain used to overproduce compound 5 contains two
genetic alterations (Figure 2-2). The native promoter of afoA, the gene that encodes the
pathway-specific transcription activator of the asperfuranone pathway, was replaced with
the inducible alcohol dehydrogenase promoter, alcA. The afoD gene, which codes for the
hydroxylase in the asperfuranone pathway, was deleted to enable the accumulation of
intermediate, compound 5 (Chiang, 2009). It should be noted that the afo cluster is silent
under normal laboratory growing conditions. The wild type strain, thus, did not produce
detectable quantities of compound 5 and asperfuranone (4) by LC/MS analysis. The
mutant strain was initially cultured in a liquid lactose minimal medium under inducing
conditions (refer to Supporting Information) at 37°C for 3 days to produce nearly 200
mg/L of 5 without need for further purification since 5 is poorly dissolved in aqueous
media.
We altered culture conditions in several ways to optimize the titer of compound 5.
First, culture time prior to the induction of alcA was investigated. Cultures were
incubated from 12 to 36 h before the chemical inducer, cyclopentanone, necessary to
induce the alcA promoter, was introduced. Thereafter the culture remained under
inducing conditions for an additional 72 h. The experiment revealed the production of 5
was enhanced by growth 30-36 h before induction (Figure 2-5). We next examined a
second parameter, the culture time post induction. The A. nidulans strain was cultured for
7 days after induction, and samples were collected at 1-day intervals from day three

57
through seven (Figure 2-6). An increase and then decline was observed over the period,
with the accumulation of 5 peaking on day five. Under optimized expression conditions
our engineered strain produced the polyketide (5) abundantly, providing a titer of 900
mg/L. The elevated production of this advanced metabolite allowed us to employ it fro
the preparation of a small library of azaphilones.
We focused on applying our semisynthetic route to prepare (+)-sclerotiorin by
treating 5 with p-toluenesulfonic acid (Figure 2-3) to form the 2-benzopyrilium salt (6),
which is then oxidized by lead tetraacetate to generate the non-halogenated azaphilone
(7) (Chong, 1971). Although the acetoxylation at C-7 would be nonstereospecific, the
diastereomers were indistinguishable (t.l.c.,
1
H and
13
C NMR) and could not be separated
by HPLC.
Electrophilic chlorination of azaphilone 7 introduces a chlorine atom at C-5 by
using a slight excess of N-chlorosuccinimide to provide the natural product (+)-
sclerotiorin and 7-epi-sclerotiorin (8) in 61% yield. Despite the recalcitrant purification of
8, the diastereomers were separated by analytical chiral HPLC to reveal close to a 1:1
ratio of (+)-sclerotiorin and 7-epi-sclerotiorin.
Additionally, several azaphilone analogs were also prepared from 5 (Figure 2-4).
To create a more efficient synthetic route, we were interested in hypervalent-iodine-
mediated phenol oxidative dearomatization with o-iodoxybenzoic acid (IBX), a method
developed by Pettus and co-workers (Marsini, 2006). The reaction proceeds with the
formation of 6, which subsequently is treated with IBX and catalyst Bu
4
NI at room
temperature to form 11. We observed 14 as a major side product of the reaction. It is

58
plausible that the generation of 14 could arise from tetrabutylammonium triiodide or IOH
formed in the presence of the residual acetic acid with adventitious water (Moorthy,
2009; Perez-Benito, 1990). To assist in the regioselective halogenation of 11, the
corresponding N-halosuccinimides were employed to produce compounds 12, 13, and 14.
Then to further functionalize the scaffold of the azaphilone core, a wittig olefination was
performed with carbethoxymethylenetriphenylphosphorane. It was observed that the
ylide selectively coupled with the less hindered ketone and produced a mixture of E/Z
isomers (1:1.05) as determined by
1
H NMR data. Due to the difficulties in separation, the
isomeric mixtures of 15-17 were tested toward the inhibition of lipoxygenase-1.
Based on a report that (+)-sclerotiorin has potent LOX-1 inhibition (Chidananda,
2007), the biological activities of all semisynthetic azaphilones were evaluated for
soybean LOX-1 inhibition to provide a preliminary structure-activity relationship (SAR).
In screening for inhibition, azaphilones 7 and 13 displayed the highest lipoxygenase-1
inhibition (Table 2-1). Azaphilones 9, 11, and 12 showed similar inhibition activities.
Iodinated azaphilones displayed less inhibition toward LOX-1, which may be due, in
part, to putative chemical instability. The other compounds (15-17) showed no
appreciable LOX-1 inhibition.
The LOX-1 inhibition screening suggested that the C-8 ketone might be an
important structural feature for activity against lipoxygenase targets. The azaphilone
analogs also indicated that halogenation at C-5 is not essential to maintain low
micromolar activity, except that when C-5 was iodinated a loss of inhibition was
observed. Since (+)-sclerotiorin has similar IC
50
with 8 containing both (+)-sclerotiorin

59
and 7-epi-sclerotiorin, the chiral center C-7 is not critical for the LOX-1 inhibition.
It has been suggested that 1 inhibits LOX-1 by trapping lipid radicals formed at
the active site of the enzyme-substrate complex. Although, we have not measures the
reductive properties of our semisynthetic azaphilones, it is reasonable that they have
similar antioxidant properties to (+)-sclerotiorin.
The metabolic engineering of a biosynthetic pathway in the filamentous fungus,
Aspergillus nidulans, demonstrates the feasibility of producing copious amounts of the
advanced polyketide (5). Coupled with existing synthetic methodology, this provides
facile synthetic access to derivatives of the natural product sclerotiorin. Azaphilone
analogs 7 and 13 were the most effective to inhibit the therapeutic target, LOX-1.
Preliminary SAR indicates the importance of the C-8 ketone for inhibition of
lipoxygenase. This may also provide insight into the further development of more potent
LOX-1 inhibitors.

4. Supplemental Information

General Information
1
H and
13
C NMR spectra were recorded on a Varian Mercury Plus 400 spectrometer.
Chemical shifts are reported in ppm relative to CDCl
3
(
1
H, δ 7.26;
13
C, δ 77.0) or
acetone-d
6
(
1
H, δ 2.05;
13
C, δ 30.0, 206.0).
1
H NMR data is reported as: chemical shift,
multiplicity (s = singlet, d = doublet, t = triplet, q = quartet, m = multiplet), coupling
constant, and integration. Infrared spectra were recorded on a JASCO FTIR-4100.

60
Infrared frequencies are reported in reciprocal centimeters. High resolution electrospray
ionization mass was obtained on an Agilent 6210 time of flight LC-MS. Reactions were
monitored by analytical thin-layer chromatography on EMD silica gel-60
F254
plates. Flash
chromatography was performed on EMD silica gel 60, 70-230 mesh. All reagents were
used without further purification unless otherwise noted. Reagents were purchased from
Sigma-Aldrich, and Alfa Aesar. Diethyl ether (anhydrous) and 1,2-dichloroethane
(anhydrous) were used directly from the bottle. Water sensitive reactions were performed
under a nitrogen atmosphere with oven-dried glassware. Kinetic experiments for
lipoxygenase inhibition were recorded on a Shimadzu UV-2401PC at 26˚C. Materials for
the lipoxygenase assay such as linoleic acid, 15-Lipoxygenase (soybean P
1
), and (+)-
sclerotiorin were purchased from Cayman Chemicals.
Optimization Of Cyclopentanone Inducing Time
Five flasks inoculated with alcA(p)-afoA, afoDΔ A. nidulans containing 1 x 10
6
spores
per mL in a liquid lactose minimal medium (lactose 15g, NaNO
3
6g, KCl 0.52g,
MgSO
4
7H2O 0.52g, KH
2
PO
4
1.52g, H
2
O 1 L supplemented with 1 mL of trace elements
and adjusted to a pH of 6.5) were incubated at 37°C in a Barnstead/Lab-Line MaxQ 4000
rotary shaker at a speed of 180 rpm. After incubation for the times shown, the chemical
inducer, cyclopentanone was added in a single dose to a final concentration of 30 mM.
After induction, the strain was cultured for three days under the same conditions.

61
Extraction And Isolation
The mycelium was collected by filtration and then immersed in 50 mL of acetone The
mycelium-acetone mixture was subjected to stirring for 0.5 h and filtered to isolated the
organic solution. The solvent was removed in vacuo to leave a light yellow solid residue
analyzed by LC-MS and
1
H NMR to be compound 5.
Optimization Of Culture Time
The A. nidulans strain, alcA(p)-afoA, afoDΔ , at a concentration of 1 x 10
6
spores per mL
was incubated 37°C in 25 mL of liquid lactose minimal medium (lactose 15g, NaNO3
6g, KCl 0.52g, MgSO47H2O 0.52g, KH2PO4 1.52g, H
2
O 1 L supplemented with 1 mL
of trace elements and adjusted to a pH of 6.5) at a speed of 180 rpm. For induction,
cyclopentanone at a final concentration of 30 mM was introduced to the medium at 30 h
of incubation. The strain was cultured for several days, and the mycelia were collected
from day 3 to day 7 and analyzed for production of 5.
Construction of Mutant Strain alca(p)-afoA, afoDΔ
In a previous study (Chiang, 2009) the mutant strain was prepared using an nkuΔ strain
and fusion PCR (Szewczyk, 2006).

62
Experimental Procedures
Compound 7
To 2,4-dihydroxy-6-(5,7dimethyl-2-oxo-trans-3-trans-5-nonadienyl)-3-
methylbenzaldehyde 5 (151 mg, 0.48 mmol) in 60 mL of acetic acid at room temperature
was added in one portion p-TsOH·H
2
O (940 mg, 4.94 mmol). The reaction was warmed
in an oil bath to 100˚C and stirred for 2 hours. Then the reaction was cooled to room
temperature and purged with nitrogen for 1.5 hours. It was further cooled to 18˚C and
then treated with lead tetraacetate (276 mg, 0.621 mmol) in three portions over a 15-
minute period. The solution stirred for 1 hour, and was quenched with 200 mL of ice
water. The reaction mixture was extracted with dichloromethane 3x, and the combined
organic layers were dried over magnesium sulfate, filtered and concentrated under
reduced pressure. The crude product was purified by flash chromatography on silica gel
(15% EtOAc/DCM) to afford 50.3 mg of compound 7. Isolated yield: 30%
1
H NMR (CDCl
3
, 400 MHz) δ 7.91 (s, 1H), 6.98 (d, J = 16 Hz, 1H), 6.17 (s, 1H), 5.96 (d,
J = 15.6 Hz, 1H), 5.65 (d, J = 9.6 Hz, 1H), 5.57 (d, J = 1.2 Hz, 1H), 2.47 (m, 1H), 2.18 (s,
3H), 1.82 (d, J = 1.2 Hz, 3H), 1.55 (s, 3H), 1.43 (m, 1H), 1.30 (m, 1H), 1.01 (d, J = 5.6
Hz, 3H), 0.86 (t, J = 7.2 Hz, 3H);
13
C NMR (CDCl
3
, 400 MHz) δ 193.3, 192.5, 170.1,
156.5, 153.5, 147.9, 142.8, 141.6, 131.8, 115.6, 114.8, 109.2, 107.3, 84.3, 35.0, 30.1,
22.4, 20.25 20.18, 12.3, 11.9; IR [acetone solution, ν
max
cm
-1
] 3079, 2965, 2933, 2877,
1746, 1717, 1639, 1247; HRMS (EIS) m/z calculated for C
21
H
25
O
5
357.1697,
experimental 357.1696 [M+H]
+

63
Compound 11
To 2,4-dihydroxy-6-(5,7-dimethyl-2-oxo-trans-3-trans-5-nonadienyl)-3-
methylbenzaldehyde 5 (200 mg, 0.635 mmol) in 2.5 mL of acetic acid at room
temperature was added p-TsOH (425 mg, 2.47 mmol). The suspension stirred for 1.5
hours under nitrogen leaving an orange precipitate, 2-benzopyrilium salt (6). The acetic
acid was removed and the precipitate redissolved in 1,2-dichloroethane (5mL) and then
treated with freshly prepared 2-iodoxybenzoic acid (IBX) (Frigerio, 1999)(182 mg, 0.65
mmol), which was followed by the addition of tert-n-butyl ammonium iodide (23 mg,
0.062 mmol). The solution stirred for six hour and quenched with saturated NaS
2
O
3
and
ethyl acetate. The mixture was extracted with ethyl acetate 4x, and the combined organic
layers were washed with saturated NaHCO
3
, water and brine. It was subsequently dried
with anhydrous sodium sulfate, filtered and concentrated under reduced pressure. The
crude product was purified by flash chromatography on silica gel (60% EtOAc/n-
hexanes) to afford 79.4 mg of compound 11. Isolated yield: 40%
1
H NMR (CDCl
3
, 400 MHz) δ 7.91 (s, 1H), 7.01 (d, J = 15.6 Hz, 1H), 6.19 (s, 1H), 5.97
(d, J = 16 Hz, 1H), 5.67 (d, J = 10 Hz, 1H), 5.57 (d, J = 0.8 Hz, 1H), 3.94 (br 1, 1H), 2.46
(m, 1H), 1.82 (d, J = 1.2 Hz, 3H), 1.56 (s, 3H), 1.43 (m, 1H), 1.30 (m, 1H), 1.01 (d, J =
6.8 Hz, 3H), 0.85 (t, J = 7.6 Hz, 3H);
13
C NMR (CDCl
3
, 400 MHz) δ 196.0, 195.7, 156.9,
152.5, 148.3, 144.2, 142.1, 131.8, 117.9, 115.5, 108.8, 105.5, 83.3, 35.1, 30.1, 28.7, 20.2,
12.3, 11.9; IR [acetone solution, ν
max
cm
-1
] 3431, 2965, 2930, 2874, 1715, 1624, 1236;
HRMS (ESI) m/z calculated for C
19
H
23
O
4
315.1591, experimental 315.1595 [M+H]
+

64
Compound 8
To a stirring solution of compound 7 (28.4 mg, 0.078 mmol) in acetic acid (0.93 mL) was
added N-chlorosuccinimide (14 mg, 0.105 mmol) at room temperature. The solution
stirred for 3 hours and then quenched with saturated Na
2
S
2
O
3
aqueous solution and
diluted with ethyl acetate. The resulting mixture was extracted with ethyl acetate and the
combined organic layers washed with NaHCO
3
, water, and brine. It was dried over
anhydrous sodium sulfate, filtered and concentrated under reduced pressure. The crude
product was purified by flash chromatography on silica gel (25% EtOAc/n-hexanes) to
afford 16 mg compound 8 (
1
H ,
13
C NMR, IR and HRMMS below). Isolated yield: 52%.
The diastereomers of 8 were separated by analytical chiral HPLC (Diacel CHIRALCEL
® OD column 0.46 x 25 cm, 10% EtOH-hexanes, 1.0 mL/min, 360 nm, t
R =
7.5 min 7-
epi-sclerotiorin, 9.5 min (+)-sclerotiorin compared with commercial available (+)-
sclerotiorin.

1
H NMR (CDCl
3
, 400 MHz) δ 7.93 (s, 1H), 7.06 (d, J = 15.6 Hz, 1H), 6.64 (s, 1H), 6.08
(d, J = 15.6 Hz, 1H), 5.70 (d, J = 9.6 Hz, 1H), 2.47 (m, 1H), 2.17 (s, 3H), 1.85 (d, J = 0.8
Hz, 3H), 1.57 (s, 3H), 1.42 (m, 1H), 1.32 (m, 1H), 1.01 (d, J = 6.8 Hz, 3H), 0.86 (t, J =
0.8 Hz, 3H);
13
C NMR (CDCl
3
, 400 MHz) δ 191.8 185.9, 170.1, 158.1, 152.6, 148.8,
142.8, 138.6, 131.9, 115.6, 114.5, 110.8, 106.4, 84.5, 35.1, 30.0, 22.5, 20.2, 20.1, 12.3,
11.9; IR [acetone solution, ν
max
cm
-1
] 2961, 2927, 2872, 1740, 1719, 1634, 1527, 1246,
1131, 1089; HRMS (ESI) m/z calculated for C
21
H
24
ClO
5
391.1307, experimental 391.131
[M+H]
+

65

Compound 9
To a stirring solution of compound 7 (13.9 mg, 0.39 mmol) in acetonitrile (0.650 mL)
was added N-bromosuccinimide (10 mg, 0.056 mmol) at room temperature. The solution
stirred for 4 hours after which it was quenched with water and extracted with ethyl
acetate. The organic layers were combined, washed with brine, dried over anhydrous
sodium sulfate, filtered and concentrated under reduced pressure. The crude product was
purified by flash chromatography on silica gel (20% EtOAc/n-hexanes) to afford 68.9 mg
of compound 9. Isolated yield: 41%
1
H NMR (CDCl
3
, 400 MHz) δ 7.91 (s, 1H), 7.07 (d, J = 15.6 Hz, 1H), 6.68 (s, 1H), 6.08
(d, J = 15.6 Hz), 5.71 (d, J = 10 Hz, 1H), 2.48 (m, 1H), 2.17 (s, 3H), 1.85 (d, J = 0.8 Hz,
3H), 1.57 (s, 3H), 1.42 (m, 1H), 1.32 (m, 1H), 1.03 (d, J = 6.8 Hz, 3H), 0.86 (t, J = 7.6
Hz, 3H);
13
C NMR (CDCl
3
, 400 MHz) δ 191.9, 186.0 170.0, 158.4, 152.7, 148.9, 142.9,
140.9, 132.0, 115.6, 115.2, 109.1, 103.1, 84.4, 35.1, 30.0, 22.6, 20.2, 20.1, 12.3, 11.9; IR
[acetone solution, ν
max
cm
-1
] 2962, 2928, 1735, 1718, 1633, 1520, 1245, 1126; HRMS
(ESI) m/z calculated for C
21
H
24
BrO
5
435.0802, experimental 435.0807 [M+H]
+

Compound 10
To a stirring solution of compound 7 (19.1 mg, 0.054 mmol) in acetonitrile (1.3 mL) was
added N-iodosuccinimide (14 mg, 0.059 mmol) at room temperature. The solution stirred
for 30 minutes and quenched with water. The mixture was extracted with ethyl acetate;

66
combined organic layers were washed with brine, dried over anhydrous sodium sulfate,
and concentrated under reduced pressure. Crude extract was purified by flash
chromatography on silica gel (20% EtOAc/n-hexanes) to afford 7.2 mg of compound 10.
Isolated yield: 28%

1
H NMR (CDCl
3
, 400 MHz) δ 7.81 (s, 1H), 7.07 (d, J = 15.6 Hz, 1H), 6.67 (s, 1H), 6.10
(d, J = 15.6 Hz, 1H), 5.71 (d, J = 10 Hz, 1H), 2.49 (m, 1H), 2.17 (s, 3H), 1.86 (d, J = 1.2
Hz, 3H), 1.56 (s, 3H), 1.42 (m, 1H), 1.31 (m, 1H), 1.01 (d, J = 6.8 Hz, 3H), 0.86 (t, J =
7.6 Hz, 3H);
13
C NMR (CDCl
3
, 400 MHz) δ 192.4, 187.5, 170.0, 158.8, 152.7, 149.9,
144.9, 142.9, 132.0, 115.9, 115.6, 114.2, 83.5, 83.4, 35.1, 30.0, 22.7, 20.2, 20.1, 12.4,
11.9; IR [acetone solution, ν
max
cm
-1
] 2961, 2927, 2872, 1739, 1718, 1630, 1509, 1245;
HRMS (ESI) m/z calculated for C
21
H
24
IO
5
483.0663, experimental 483.0667 [M+H]
+

Compound 12
To a stirring solution of compound 11 (59.3 mg, 0.188 mmol) in 3.0 mL of acetonitrile at
room temperature was added N-chlorosuccinimide (27 mg, 0.202 mmol). The solution
stirred for 48 hours and quenched with water then extracted with ethyl acetate. The
combined organic layers were washed with brine, dried over anhydrous sodium sulfate,
filtered and concentrated under reduced pressure. Crude extract was purified by flash
chromatography on silica gel (45% EtOAc/n-hexanes) to afford 29.3 mg of compound
12. Isolated yield: 45%

67
1
H NMR (CDCl
3
, 400 MHz) δ 7.94 (s, 1H), 7.09 (d, J = 15.6 Hz, 1H), 6.62 (s, 1H), 6.08
(d, J = 15.6 Hz, 1H), 5.72 (d, J = 9.6 Hz, 1H), 3.90 (br s, 1H), 2.48 (m, 1H), 1.85 (d, J =
1.2 Hz, 3H), 1.59 (s, 3H), 1.42 (m, 1H), 1.32 (m, 1H), 1.01 (d, J = 6.4 Hz, 3H), 0.86 (t, J
= 7.6 Hz, 3H);
13
C NMR (CDCl
3
, 400 MHz) δ 194.0, 189.6, 158.6, 151.7, 149.4, 143.4,
140.0, 132.0, 115.6, 115.1, 108.9, 105.9, 83.9, 35.2, 30.1, 28.8, 20.2, 12.4, 12.0; IR
[acetone solution, ν
max
cm
-1
] 3444, 3085, 3065, 3039, 2871, 1724, 1635, 1527, 1178, 670;
HRMS (ESI) m/z calculated for C
19
H
22
ClO
4
349.1201, experimental 349.1205 [M+H]
+

Compound 13
To a stirring solution of compound 11 (122.09 mg, 0.39 mmol) in acetonitrile (6.5 mL)
was added N-bromosuccinimide (77.0 mg, 0.43 mmol) at room temperature. The solution
stirred for 45 minutes and was then quenched with water. The mixture was extracted with
ethyl acetate and the combined organic layers were dried over anhydrous sodium sulfate,
filtered and concentrated under reduced pressure. The crude product was purified by flash
chromatography on silica gel (20% EtOAc/n-hexanes) to afford 71 mg of compound 13.
Isolated yield: 46%
1
H NMR (CDCl
3
, 400 MHz) δ 7.92 (s, 1H), 7.11 (d, J = 15.6 Hz, 1H), 6.66 (s, 1H), 6.10
(d, J = 15.6 Hz, 1H), 5.74 (d, J = 9.6 Hz, 1H), 3.92 (s, 1H), 2.49 (m, 1H), 1.86 (d, J = 1.2
Hz, 3H), 1.56 (s, 3H), 1.43 (m, 1H), 1.32 (m, 1H), 1.02 (d, J = 6.8 Hz, 3H), 0.87 (t, J =
7.6 Hz, 3H);
13
C NMR (CDCl
3
, 400 MHz) δ 194.1, 190.0, 158.9, 151.7, 149.4, 143.5,
142.2, 132.0, 115.9, 115.6, 108.6, 100.5, 83.9, 35.2, 30.0, 28.8, 20.2, 12.4, 12.0; IR

68
[acetone solution, ν
max
cm
-1
] 3451, 3081, 2964, 2931, 2875, 1719, 1633, 1518, 1172,
1144; HRMS (ESI) m/z calculated for C
19
H
22
BrO
4
393.0696, experimental 393.0700 [M
+ H]
+

Compound 14
To a stirring solution of compound 11 (33.2 mg, 0.106 mmol) in acetonitrile (2.1 mL)
was added N-iodosuccinimide (28 mg, 0.124 mmol) at room temperature. The solution
stirred for 1 hour and quenched with water. The mixture was extracted with ethyl acetate;
organic layers were combined and washed with brine, dried over anhydrous sodium
sulfate and concentrated under reduced pressure. The crude product was purified by flash
chromatography on silica gel (35% EtOAc/n-hexanes) to afford 23.7 mg of compound
14. Isolated yield: 51%
1
H NMR (CDCl
3
, 400 MHz) δ 7.81 (s, 1H), 7.10 (d, J = 16 Hz, 1H), 6.64 (s, 1H), 6.10 (d,
J = 15.6 Hz, 1H), 5.74 (d, J = 9.6 Hz, 1H), 3.94 (s, 1H), 2.49 (m, 1H), 1.86 (d, J = 1.2 Hz,
3H), 1.57 (s, 3H), 1.42 (m, 1H), 1.32 (m, 1H), 1.02 (d, J = 6.8 Hz, 3H), 0.86 (t, J = 7.6
Hz, 3H);
13
C NMR (CDCl
3
, 400 MHz) δ 194.4, 191.6, 159.3, 151. 6, 149.4, 146.2, 143.5,
132.0, 116.7, 115.4, 113.5, 83.2, 79.3, 35.2, 30.0, 28.9, 20.2, 12.4, 11.9; IR [acetone
solution, ν
max
cm
-1
] 3458, 3070, 2963, 2928, 2873, 1721, 1633, 1504, 1166; HRMS (ESI)
m/z calculated for C
19
H
22
IO
4
441.0557, experimental 441.056 [M+H]
+

69
Compound 15
To a stirring solution of compound 11 (55 mg, 0.175 mmol) in anhydrous diethyl ether (6
mL) was added Wittig reagent, Ph
3
=CHCO
2
Et, (68 mg, 0.193 mmol). The solution stirred
for 7 hours at room temperature. The solvent was removed under reduced pressure and
crude extract purified by flash chromatography on silica gel (15% acetone/n-hexanes) to
afford 11 mg of compound 15. Isolated yield: 17%
1
H NMR (acetone-d
6
, 400 MHz) δ 8.13 (s, 1H), 7.07 (d, J = 16 Hz, 1H), 6.47 (s, 1H),
6.27 (d, J = 15.6 Hz, 1H), 5.71 (d, J = 9.6 Hz, 1H), 5.38 (s, 1H), 4.6 (s, 1H), 4.18 (q, J =
7.2 Hz, 2H), 2.53 (m, 1H), 1.87 (d, J = 1.2 Hz, 3H), 1.49 (m, 1H), 1.37 (s, 3H), 1.33 (m,
1H), 1.25 (t, J = 7.2 Hz, 3H), 1.01 (d, J = 6.4 Hz, 3H), 0.86 (t, J = 7.6 Hz, 3H);
13
C NMR
(acetone-d
6
, 400 MHz) δ 197.1, 166.5, 157.0, 150.8, 147.2, 147.1, 141.4, 133.4, 117.9,
115.1, 110.0, 109.3, 104.6, 77.7, 61.0, 35.7, 31.0, 29.2, 20.8, 14.6, 12.7, 12.4; IR [acetone
solution, ν
max
cm
-1
] 3441, 2967, 2955, 2877, 1716, 1623, 1543, 1179; HRMS (ESI) m/z
calculated for C
23
H
29
O
5
385.201, experimental 385.2014 [M+H]
+

Compound 16
To a stirring solution of compound 13 (20.5 mg, 0.052 mmol) in anhydrous diethyl ether
(2.6 mL) was added Ph
3
=CHCO
2
Et, (23 mg, 0.066 mmol). The solution stirred for 6
hours at room temperature. The solvent was removed under reduced pressure and crude
extract purified by flash chromatography on silica gel (15% acetone/n-hexanes) to afford
16.3 mg of compound 16. Isolated yield: 68%

70
1
H NMR (acetone-d
6
, 400 MHz) δ 8.12 (s, 1H), 7.18 (d, J = 16 Hz, 1H), 6.78 (s, 1H),
6.51 (d, J = 16 Hz, 1H), 6.50 (s, 1H), 5.78 (d, J = 9.2 Hz, 1H), 4.86 (s, 1H), 4.18 (q, J =
7.2 Hz, 2H), 2.55 (m, 1H), 1.91 (d, J = 1.2 Hz, 3H), 1.45 (m, 1H), 1.40 (s, 3H), 1.33 (m,
1H), 1.24 (t, J = 7.2 Hz, 3H), 1.02 (d, J = 6.8 Hz, 3H), 0.86 (t, J = 7.2 Hz, 3H);
13
C NMR
(acetone-d
6
, 400 MHz) δ 190.9, 166.3, 159.4, 150.0, 148.3, 145.7, 144.9, 142.9, 133.6,
118.0, 116.1, 109.4, 108.6, 99.2, 78.6, 61.2, 35.9, 30.9, 28.8, 20.7, 14.6, 12.7, 12.4; IR
[acetone solution, ν
max
cm
-1
] 3464, 2961, 2933, 2874, 1717, 1627, 1554, 1515, 1174, 669;
HRMS (ESI) m/z calculated for C
23
H
28
BrO
5
463.1115, experimental 463.1118 [M+H]
+

Compound 17
To a stirring solution of compound 14 (48 mg, 0.109 mmol) in diethyl ether (5.0 mL) was
added Ph
3
=CHCO
2
Et, (46 mg, 0.132 mmol). The solution stirred for 6 hours at room
temperature. The solvent was removed under reduced pressure and crude extract purified
by flash chromatography on silica gel (15% acetone/n-hexanes) to afford 31.8 mg of
compound 17. Isolated yield: 57%
1
H NMR (acetone-d
6
, 400 MHz) δ 8.01 (s, 1H), 7.18 (d, J = 16 HZ, 1H), 6.76 (s, 1H),
6.53 (d, J = 16 Hz, 1H), 6.50 (s, 1H), 5.78 (d, J = 9.6 Hz, 1H), 4.82 (s, 1H), 4.18 (q, J =
7.2 Hz, 2H), 2.55 (m, 1H), 1.97 (d, J = 1.2 Hz, 3H), 1.44 (m, 1H), 1.40 (s, 3H), 1.32 (m,
1H), 1.24 (t, J = 7.2 Hz, 3H), 1.02 (d, J = 4.8 Hz, 3H), 0.87 (t, J = 7.6 Hz, 3H);
13
C NMR
(acetone-d
6
, 400 MHz) δ 192.8, 166.3, 159.7, 149.9, 149.0, 148.3, 142.9, 133.6, 118.0,
115.9, 114.3, 113.6, 105.8, 78.0, 61.2, 35.9, 30.9, 28.9, 20.7, 14.6, 12.7, 12.4; IR [acetone

71
solution, ν
max
cm
-1
] 3456, 2965, 2930, 2873, 1717, 1624, 1555, 1502, 1176, 530; HRMS
(ESI) m/z calculated for C
23
H
28
IO
5
511.0976, experimental 511.0977 [M+H]
+

Soybean Lipoxygenase-1 Assay
The enzyme assay was performed according to Axelrod et. al. (Axelrod, 1981). Each
solution was prepared and measured at room temperature. Enzyme kinetics was measured
on a Shimadzu UV-2401PC at a wavelength of 234 nm in a quartz cuvette. For the assay,
the linoleic acid substrate was prepared by mixing 20 µL linoleic acid (Cayman Chemical
#760716) with 20 µL KOH (Cayman Chemical #760713) and 40 µL double distilled
water. Lipoxygenase-1 (Cayman Chemical #60712) was diluted from original stock
concentration to a concentration of 100 U/mL in the cuvette. The semisynthetic
azaphilones were prepared as stock concentrations in dimethylsulfoxide (DMSO) from 20
µM to 10 mM; 5 µL was added to the assay mixture to give the desired final
concentration of the inhibitor with 1% of DMSO in the assay.
Lipoxygenase-1 Inhibition Assay
The spectrophotometric assay is based on monitoring the reaction, catalyzed by LOX-1 to
convert linoleic acid (L) to the hydroperoxide (LOOH), at 234 nm. The substrate L does
not absorb at the indicated wavelength, so that any absorption is due to the generation of
LOOH. The reaction is initiated when the non- heme iron of LOX-1 in the ferric state is
reduced by the substrate L. While in the enzyme-substrate complex, L is oxidized and
converted to the peroxide free radical (LOO), and subsequently released from the

72
enzyme as LOOH. Inhibitors can interfere in the process in several different ways to
prevent the generation of LOOH by the chelation of iron, removal of oxygen or
prevention of free radical formation.
Procedure
Prior to measurements, a sample with buffer was used as a blank. In a quartz cuvette,
each sample was prepared with 482.5 µL 0.2 M borate buffer (at a pH of 9.0) mixed with
5 µL inhibitor and 5 µL substrate by pipetting the solution. The cuvette was placed in the
spectrophotometer and measured for 120 seconds and then rapidly 7.5 µL lipoxygenase-1
was added and mixed to initiate the reaction. Measurements were recorded every 30
seconds for 7.5 minutes. All measurements were carried out in triplicate.
IC
50
Calculation
Inhibition of an enzyme at 50 % can be experimentally measured by monitoring enzyme
kinetics. A kinetic study generates a curve where the slope provides the initial velocity
(V
o
) of the enzyme. For each sample with different inhibitor concentrations the initial
velocity was derived from a linear segment between 210-300 nm. The absorbance was
used directly and not converted into concentration units. These data points (V
o
,
[Inhibitor]) were fitted to a curve using the software Curve Expert Professional 1.5.0 to
provide an equation for the curve; the best mathematical model for the data was the MMF
model. The concentration of inhibitor at which 50% inhibition occurred (as measured by
change in V
o
) was reported as the IC
50
.

73

Figure 2-1. Azaphilone natural products

74

Figure 2-2. Reengineered Biosynthetic Pathway for the Synthesis of (+)-Sclerotiorin and
7-epi-Sclerotiorin (8) and Non-natural Azaphilone Polyketides

75

Figure 2-3. Concise Synthesis of (+)-Sclerotiorin, 7-epi-Sclerotiorin (8) and Analogs

76

Figure 2-4. A Short Route to Azaphilone Analogs

77

78

79

Figure 2-5. Optimization of Induction Time

80

Figure 2-6. Optimization of Culture Time (post-induction)

81

Figure 2-7.
1
H and
13
C NMR of compound 7

82

Figure 2-8.
1
H and
13
C NMR of compound 8

83

Figure 2-9.
1
H and
13
C NMR of compound 9

84

Figure 2-10.
1
H and
13
C NMR of compound 10

85

Figure 2-11.
1
H and
13
C NMR of compound 11

86

Figure 2-12.
1
H and
13
C NMR of compound 12

87

Figure 2-13.
1
H and
13
C NMR of compound 13

88

Figure 2-14.
1
H and
13
C NMR of compound 14

89

Figure 2-15.
1
H and
13
C NMR of compound 15

90

Figure 2-16.
1
H and
13
C NMR of compound 16

91

Figure 2-17.
1
H and
13
C NMR of compound 17

92

________________________________________________________________________

Figure 2-18. Soybean lipoxygease-1 assay

93

Figure 2-19a. LOX-1 enzyme kinetics
with varying concentrations of (+)-
sclerotiorin measured at λ 234 nm.

Figure 2-19b. Initial Velocity (Vo) of
LOX-1 with concentration, where the
initial velocity of enzyme incubated in the
absence of inhibitor designated a value of
1.

94

Figure 2-20a. LOX-1 enzyme kinetics
with varying concentrations of compound
4 measured at λ 234 nm.

Figure 2-20b. Initial Velocity (Vo) of
LOX-1 with concentration, where the initial
velocity of enzyme incubated in the absence
of inhibitor designated a value of 1.

95

Figure 2-21a. LOX-1 enzyme kinetics with
varying concentrations of compound 5
measured at λ 234 nm.

Figure 2-21b. Initial Velocity (Vo) of
LOX-1 with concentration, where the
initial velocity of enzyme incubated in
the absence of inhibitor designated a
value of 1.

96

Figure 2-22a. LOX-1 enzyme kinetics with
varying concentrations of compound 7
measured at λ 234 nm.
Figure 2-22b. Initial Velocity (Vo) of
LOX-1 with concentration, where the
initial velocity of enzyme incubated in the
absence of inhibitor designated a value of
1.

97

Figure 2-23a. LOX-1 enzyme kinetics with
varying concentrations of compound 8
measured at λ 234 nm.

Figure 2-23b. Initial Velocity (Vo) of
LOX-1 with concentration, where the
initial velocity of enzyme incubated in the
absence of inhibitor designated a value of
1.

98

Figure 2-24a. LOX-1 enzyme kinetics with
varying concentrations of compound 9
measured at λ 234 nm.

Figure 2-24b. Initial Velocity (Vo) of
LOX-1 with concentration, where the
initial velocity of enzyme incubated in the
absence of inhibitor designated a value of
1.

99

Figure 2-25a. LOX-1 enzyme kinetics with
varying concentrations of compound 10
measured at λ 234 nm.

Figure 2-25b. Initial Velocity (Vo) of
LOX-1 with concentration, where the
initial velocity of enzyme incubated in the
absence of inhibitor designated a value of
1.

100

Figure 2-26a. LOX-1 enzyme kinetics with
varying concentrations of compound 11
measured at λ 234 nm.

Figure 2-26b. Initial Velocity (Vo) of
LOX-1 with concentration, where the
initial velocity of enzyme incubated in the
absence of inhibitor designated a value of
1.

101

Figure 2-27a. LOX-1 enzyme kinetics with
varying concentrations of compound 12
measured at λ 234 nm
Figure 2-27b. Initial Velocity (Vo) of
LOX-1 with concentration, where the
initial velocity of enzyme incubated in the
absence of inhibitor designated a value of
1.

102

Figure 2-28a. LOX-1 enzyme kinetics with
varying concentrations of compound 13
measured at λ 234 nm.

Figure 2-28b. Initial Velocity (Vo) of
LOX-1 with concentration, where the
initial velocity of enzyme incubated in the
absence of inhibitor designated a value of
1.

103

Figure 2-29a. LOX-1 enzyme kinetics with
varying concentrations of compound 14
measured at λ 234 nm.

Figure 2-29b. Initial Velocity (Vo) of
LOX-1 with concentration, where the
initial velocity of enzyme incubated in the
absence of inhibitor designated a value of
1.

104

Figure 2-30a. LOX-1 enzyme kinetics with
varying concentrations of compound 15
measured at λ 234 nm
Figure 2-30b. Initial Velocity (Vo) of
LOX-1 with concentration, where the
initial velocity of enzyme incubated in the
absence of inhibitor designated a value of
1.

105

Figure 2-31a. LOX-1 enzyme kinetics with
varying concentrations of compound 16
measured at λ 234 nm.

Figure 2-31b. Initial Velocity (Vo) of
LOX-1 with concentration, where the
initial velocity of enzyme incubated in the
absence of inhibitor designated a value of
1.

106

Figure 2-32a. LOX-1 enzyme kinetics with
varying concentrations of compound 17
measuredat λ 234 nm.

Figure 2-32b. Initial Velocity (Vo) of
LOX-1 with concentration, where the
initial velocity of enzyme incubated in the
absence of inhibitor designated a value of
1.

107
Chapter III. Engineering Of An “Unnatural” Natural Product By Swapping
Polyketide Synthase Domains In Aspergillus nidulans

1. Abstract

An StcA-AfoE hybrid polyketide synthase (PKS), generated by swapping the
AfoE (asperfuranone biosynthesis) SAT domain with the StcA (sterigmatocystin
biosynthesis) SAT domain, produced a major new metabolite with the same chain length
as the native AfoE product. Structure elucidation allowed us to propose a likely pathway,
and feeding studies supported the hypothesis that the chain length of PKS metabolites
may be under precise control of KS and PT domains.

2. Introduction

Fungal polyketides are produced by multidomain iterative type I polyketide
synthases (PKSs). Fungal PKSs can be further grouped into non-reduced (NR), partially
reduced (PR), and highly reduced (HR) PKS by examining the domains encoded in the
gene (Cox, 2007). NR-PKSs contain starter unit ACP transacylase (SAT), β-ketoacyl
synthase (KS), acyl transferase (AT), product template (PT), and acyl carrier protein
(ACP) domains at a minimum and may also contain methyltransferase (CMeT) and
product-releasing domains such as thioesterase/Claisen-cyclase (TE/CLC), thioesterase
(TE), and reductase (R). The SAT domain has been determined to possess remarkable
selectivity toward starter units and to initiate polyketide synthesis by transferring the
starter unit onto the ACP domain (Crawford, 2006). The PT domain was found to

108
cooperate with the KS domain to exert precise control over production formation
(Crawford, 2008). The KS domain extends the starter unit to a fixed length, and the PT
domain transforms a reversible precursor to an irreversible form by specific aldol
cyclization and aromatization, thereby enhancing the flux to products (Crawford, 2008).
Recent crystallographic studies of the PksA PT monomer revealed that the PT catalytic
pocket is comprised of three regions: (1) a phosphopantetheine (PPT)-binding region at
the entry, (2) a hydrophobic hexyl-binding region at the bottom, perfectly adapting the
hexanoyl head, and (3) a cyclization chamber between regions 1 and 2. Conformational
changes of the PT domain lead to folding a fully extended C
20
-precursor anchored by
regions 1 and 2 to a “hairpin” conformation that fits and subsequently gets cyclized in the
chamber (Crawford, 2009).
Through a genome mining effort in Aspergillus nidulans, our group was able to
uncover the genes involved in asperfuranone biosynthesis, an azaphilone class of fungal
natural product (Chiang, 2009). Azaphilones are structurally diverse polyketides that
share a highly oxygenated bicyclic core and a chiral quaternary center. These polyketides
are known for the reaction of their 4H-pyran motif with amines to produce the
corresponding vinylogous γ-pyridones (Wei, 2005). Mostly of fungal origin, these
pigments display a broad range of biological activities, including inhibition of
monoamine oxidase, acyl-CoA:cholesterol acyltransferase, and cholesteryl ester transfer
protein (Osmanova, 2010). The asperfuranone biosynthesis pathway, similar to all
azaphilones, involves two PKSs: the HR-PKS AfoG and the NR-PKS AfoE (Chiang,
2009). AfoG is responsible for the biosynthesis of the dimethyloctadienone moiety,

109
which is loaded on the SAT domain of AfoE, extended with four malonyl-CoAs, and
modified with one S- adenosyl methionine (SAM) (Figure 3-2, upper) (Chiang, 2009).
We reasoned that if we could engineer the NR-PKS AfoE to accept different starter units,
we would have the opportunity to create new “unnatural” polyketide products in vivo. To
test this hypothesis, we replaced the SAT domain from StcA, the NR-PKS in the
sterigmatocystin (ST) biosynthesis pathway, which accepts the hexanoyl starter unit
produced by the fatty acid synthase (FAS) encoded by stcJ and stcK (Brown, 1996; Yu,
1995).

3. Results & Discussion

Our analysis of the metabolism profile of wild-type A. nidulans shows that the ST
biosynthesis pathway is highly expressed under normal laboratory culture conditions.
This means that it should not be necessary to artificially turn on the expression of the two
FAS genes stcJ and stcK for the production of the hexanoyl starter unit. However, it
would be necessary to delete stcA so the hexanoyl starter unit is not consumed by the ST
pathway. Metabolic profiles of an stcAΔ mutant showed that the production of ST was
abolished in either inducing or non-inducing conditions (Figure 3-4), and we used the
stcAΔ strain for our domain swap experiments. We also observed the production of ST
and terrequinone (TQ) in controls was decreased under the inducing conditions (Figure 3-
4B), probably due to growth inhibition of cyclopentanone that was used to induce the
alcA promoter.

110
Next, we replaced the SAT domain of AfoE with the SAT domain from StcA.
Since we did not know the optimal junction position of AfoE, we used the Udwary-
Merski algorithm (UMA) (Udwary, 2002) to predict the linker region between the SAT
and KS domains of AfoE and selected three sites, E386, Q410, and R424 (numbered as
sites 1-3), out of the linker as the SAT junction sites. Since the SAT domain of PksA, the
homologue of StcA in A. parasiticus, has been previously cloned from three sites as
individually expressed (Crawford, 2006), we selected three corresponding sites, K357,
T363, and D379 on StcA (labeled as sites A-C) as the SAT junction sites, based on
protein sequence alignment between StcA and PksA (Figure 3-5). Additionally, an
inducible alcA promoter was used to conditionally turn on the hybrid AfoE since the
asperfuranone gene cluster is normally silent. Together, nine hybrid constructs (A1, A2,
A3, B1, B2, B3, C1, C2, C3) were generated for metabolic analysis.
LC-DAD-MS analysis of the metabolites of the stcAΔ, hybrid (A1-C3) double
mutants revealed that all nine constructs were capable of producing 1 under induction,
which was a major metabolite for most of the constructs (Figure 3-1 and Figure 3-6). To
elucidate the structure of 1, the compound was isolated and purified froma large-scale
culture of the stcAΔ, hybrid B2 strain. HRESIMS (m/z 303.1245, [M-H]
-
) and
13
C NMR
spectroscopic data provided the molecular formula C
17
H
20
O
5
for 1, representing eight
indices of hydrogen deficiency (IHD).
1
H,
13
C, and HMQC NMR spectra showed signals
for an aliphatic side chain [δ
H
7.36 and 8.06 (each 1H, s)] (Table 3-1). Besides the
aliphatic side chain, the
13
C NMR spectrum also revealed that 1 contains four olefins (δ
C

111.4, 119.6, 125.0, 125.4, 130.7, 138.5, 155.2, and 158.7) and two carbonyl groups (δ
C

111
181.0, 185.6). This, together with the fact that 1 has eight IHD, suggested that 1 contains
a naphthaquinone chromophore. Tetrahydrosclerotoquinone, an alkali-degraded product
of tetrahydrosclerotiorin (Graham, 1957), has a UV-vis absorption maximum similar to
that of 1 (Figure 3-7), indicating that 1 has a naphthaquinone chromophore similar to that
of tetrahydrosclerotoquinone. HMBC correlations of 1 allowed us to fully construct its
structure as a 1,4-naphthaquinone derivative (Figure 3-8).
Structure elucidation allowed us to propose a putative biosynthetic pathway of 1
(Figure 3-2, lower). The hexanoyl starter unit loaded from StcJ/StcK is elongated to a
hexaketide intermediate. After C2-C7 cyclization, benzaldehyde intermediate 2 is
reductively released from the R domain of the hybrid AfoE. Intramolecular Knoevenagel
condensation followed by tautomerization produces naphthol 3, which is then oxidized to
become 4. Birchall et al. have observed a rapid oxidation of a similar naphthol with 3 to
1,4-naphthaquinone (Birchall, 1969). After reduction of 4, possibly by an endogenous
reductase, 1 is generated. Interestingly, 1 and the native AfoE product, asperfuranone,
both have C
16
chain length, despite having been initiated with starter units of different
lengths. Thus, it appears that the final chain length, not the number of extension rounds,
was fixed in the precursor elongation, seemingly supporting the idea of Crawford et al.
(Crawford, 2008) that the KS domain elongates the precursor to a fixed length, followed
by cyclization by the PT domain, which stabilizes the precursor and drives the production
flux.

112
To verify this hypothesis, we investigated other metabolites generated from the
hybrid AfoE also featured C
16
chain length by feeding deuterium-labeled starter units.
We deleted stcK and stcJ, which are required for hexanoyl starter production, to generate
stcAΔ, hybrid B2, stcKΔstcJΔ triple mutant and conducted feeding studies with hexanoyl-
N-acetylcysteamine (hexanoyl-SNAC). These results convincingly confirm that 1
originated from the hybrid PKS (Figure 3-3A). Besides 1, eight other metabolites were
also detected by comparing the extract profiles of SNAC-fed and D
11
-SNAC-fed cultures,
and they were shown to carry 16-19 carbons, based on high-resolution mass spectrometry
results (Figure 3-3B and Figure 3-9). The presence of C
19
metabolites could be explained
by a sixth extension round with malonyl-CoA. The C
16
and C
18
metabolites were possibly
derived from C
17
and C
19
due to poor functioning of the CMeT domain or post-
decarboxylation. However, the fact that up to 90% of metabolites were C
17
, i.e., a C
16
chain length, provided strong evidence that the length of metabolites is most likely under
strict control of the AfoE KS and PT domains. Tang’s group observed a similar length
control based on data obtained when they fed octanoyl-CoA to PKS4 in vitro (Ma, 2007).
For years, major obstacles in domain engineering have rested in the uncertainty of
domain boundaries and the time-consuming traditional cloning workflow. Recent
technological advances, such as UMA algorithm (Udwary, 2002) and the development of
genetic engineering protocols, allowed a precise and fast strategy for domain swap. We
presented here our work on NR-PKS SAT domain swap experiments in A. nidulans. The
successful swap between StcA and AfoE led to the production of compound 1, which had
the same length as native AfoE product. Feeding studies with deuterium-labeled

113
hexanoyl-CoA analogues in a hexanoyl starter-deficient mutant restored the production of
1 and verified that chain length of the metabolites may be strictly determined by the KS
and PT domains.

4. Supplemental Methods

Fungal Strains And Molecular Genetic Manipulations

A. nidulans strains used in this study are listed in Table 3-1. All primers used in this study
are listed in Table 3-2. StcA was deleted by replacing it with an A. fumigatus pyroA gene
cassette (AfpyroA) in TN02A7 (Nayak, 2006). Deletion of a chromosomal region
including two adjacent genes, stcJ and stcK, was generated by replacing this region with
an A. fumigatus riboB gene cassette (AfriboB) in CW1030. Double mutant strains [stcA_,
hybrid (A1-C3)] were generated from CW1001. The AfoE SAT plus a 120bp promoter
region immediately upstream of the afoE start codon was replaced with a cassette
containing 1) the A. fumigatus pyrG gene (AfpyrG) followed by 2) a 401bp fragment
containing the A. nidulans alcA promoter [alcA(p)] followed by 3) an stcA SAT fragment
that started from the start codon and ended at the selected swapping sites A/B/C, such
that the hybrid coding sequence was placed under the control of the alcA promoter. Two
~1kb fragments upstream and downstream of each targeted DNA region were amplified
from A. nidulans genomic DNA by PCR and fused together with the replacement cassette
by fusion PCR (Szewczyk, 2006). Protoplast production and transformation were carried
out as described (Szewczyk, 2006). Three to five transformants for each genotype were

114
analyzed by diagnostic PCR with three primer sets. In the case that the external primers
in the first round of PCR were used, the difference in the sizes of targeted DNA region
before and after replacement allowed the determination of correct gene replacement. In
the cases that one of the external primers and the primer located inside the cassette were
used, the correct mutant gave the PCR product of the expected size, otherwise no product
was present.

Synthesis of Hexanoyl-SNAC
The synthesis was carried out using 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide
(EDC) coupling as described by Boddy et al (Sharma, 2007). To a solution of hexanoic
acid in anhydrous dichloromethane was added EDC (2 eq.) at 0°C under nitrogen. After
stirring for 20 min, N-acetylcysteamine (1.2 eq.) a few crystals of DMAP was added to
the solution, warmed to room temperature and stirred for additional 3 h. The reaction
mixture was quenched with water, and then extracted with ethyl acetate. Organic layers
were washed with brine, dried over anhydrous Na
2
SO
4
, concentrated, and purified by
flash chromatography (40% acetone/n-hexanes). Hexanoic acid D-11 (98%, purchased
from Cambridge Isotope Laboratories) was used as the precursor of D11-hexanoyl-
SNAC. The respective yield of hexanoyl-SNAC and D11-hexanoyl-SNAC was 98% and
77%.

115
Fermentation And LC-MS Analysis
2.5x10
7
spores of each A. nidulans strain were grown at 37°C, shaking at 200 rpm in 25
mL liquid LMM medium (Bok, 2009) supplemented when necessary with uracil (1 g/L)
and uridine (10 mM) and/or riboflavin (2.5 mg/L). For alcA(p)-inducing conditions,
cyclopentanone at a final concentration of 30 mM was added to the medium after 18 h of
incubation. For feeding studies, 4 aliquots of inoculated and induced CW1061 were
respectively fed with 7 µL DMSO, 7 µL 0.2 mM hexanoyl-SNAC, 7 µL 0.2 mM D11-
hexanoyl-SNAC, 7 µL 0.2 mM 1:1 hexanoyl-SNAC: D11-hexanoyl-SNAC successively
at 20 h, 38 h and 56 h. Culture medium was collected at 66 h by filtration and extracted
once with 25 mL of EtOAc. The EtOAc layer was evaporated in vacuo, redissolved in
0.35 mL of 20% DMSO/MeOH, and10 µL was injected for HPLC-DAD-MS analysis.
HPLC-MS analysis was carried out in the negative mode using an Agilent Technologies
1200 series high-resolution mass spectrometer with an RP C18 column (Alltech Prevail
C18 3 µm 2.1 Å~ 100 mm) at a flow rate of 125 µL/min. The solvent gradient for HPLC
was 95% MeCN/H2O (solvent B) in 5% MeCN/H2O (solvent A), both containing 0.05%
formic acid: 0% B from 0 to 5 min, 0 to 100% B from 5 to 35 min, maintained at 100% B
from 35 to 40 min, 100 to 0% B from 40 to 45 min, and re- equilibration with 0% B from
45 to 50 min. Conditions for MS included a capillary voltage at 4000 V, a nebulizer
pressure at 20 psig, a drying gas flow rate at 10 L/min, and the drying gas temperature at
350 °C.

116
Isolation And Identification of Secondary Metabolites
For structure elucidation, strain CW1030 was cultivated in 90 aliquots of 30 mL liquid
LMM medium at 37°C with shaking at 200 rpm for 66 h and induced with
cyclopentanone at 18 h. The culture medium was collected through filtration, extracted
twice with an equal volume of EtOAc, and evaporated as described above. The crude
extract (449.5 mg) was applied to a Merck Si gel column (230 to 400 mesh; ASTM) and
eluted with CHCl
3
/MeOH mixtures of increasing polarity (fraction A, 1:0; fraction B,
49:1; fraction C, 19:1; fraction D, 9:1; and fraction E, 7:3). Fraction C (42.1 mg) was
further purified by reverse-phase HPLC with a Phenomenex Luna C18 column (5-µm
particle size; 250 by 21.2 mm) at a flow rate of 10.0 mL/min and measured by a UV
detector at 254 nm. The gradient system was MeCN (solvent B) in 5% MeCN/H2O
(solvent A), both containing 0.05% trifluoroacetic acid, as follows: equilibration with
20% solvent B from 0 to 5 min, 20 to 80% solvent B from 5 to 30 min, 80 to 100%
solvent B from 30 to 33 min, 100% solvent B from 33 to 38 min, 100 to 20% solvent B
from 38 to 41 min, and reequilibration with 20% solvent B from 41 to 45 min. Compound
1 (2.2 mg) was eluted at 22.7 min.
Compound Identification
Optical rotation was measured on a JASCO P-2000 digital polarimeter. Infrared (IR)
spectra were recorded on a Nicolet MAGNA-IR 560 spectrometer.
1
H and
13
C nuclear
magnetic resonance (NMR) spectra were collected on a Varian Mercury Plus 400

117
spectrometer.
Compound 1: yellow solid; [α]
D
23
-50.7° (MeOH, c 0.2); IR (ZnSe) cm-1 3283,
1653,1570,1380, 1340, 1209, 1138, 1074, 977; For UV and ESI-MS data, see Figure 3-7;
For
1
H and
13
C NMR data, see Table 3-3.

118

119

120

Figure 3-1. HPLC profiles of extracts of A. nidulans stcAΔ, hybrid B2 double mutant
under non-inducing (a) and inducing (b) conditions as detected by UV absorption at 254
nm. The y-axis of each profile was at the same order of magnitude. *: metabolites that are
non-specific to this study

121

Figure 3-2. Proposed biosynthetic pathway of asperfuranone and 1 from native AfoE
(upper) and hybrid AfoE (lower)

122

Figure 3-3. (A) HPLC profiles of extracts of A. nidulans stcAΔ, hybrid B2, stcJΔstcKΔ
triple mutant fed with DMSO (a) and hexanoyl-SNAC (b) detected by UV absorption at
254 nm. The y axis of each profile was at the same order of magnitude. (B) Summary of
metabolites produced by hybrid AfoE based on feeding experiments. Each row lists the
predicted ion formula, retention time (t
R
), m/z of unlabeled form, m/z of D11-
incorporated form, and overall percentage of each metabolite in negative ionization
mode. *: assuming all molecules are equally ionizable in negative mode

123

Figure 3-4. HPLC profiles of extracts of A. nidulans wild-type strain (a) and stcAΔ
mutant (b) under noninducing (A) and inducing (B) conditions as detected by UV
absorption at 254 nm. Wild type produces terrequinone (TQ) as well as sterigmatocystin
(ST). The y axis of each profile was at the same order of magnitude. *: metabolites that
are non-specific to this study

124

Figure 3-5. Protein sequence homology alignment of PksA and StcA (only the first 400
amino acids are shown). *: PksA SAT domain cloning sites, which correspond to K357,
T363 and D379 on StcA

125

Figure 3-6. HPLC profiles of extracts of the stcAΔ, hybrid (A1-C3) strains

126

127

Figure 3-8. Key HMBC correlations of compound 1

128

Figure 3-9. High-resolution MS spectra (negative mode) of hybrid AfoE-derived
metabolites (a-i) detected by respectively feeding an equal volume of (A) DMSO, (B) 0.2
mM hexanoyl-SNAC, (C) 0.2 mM D11-hexanoyl-SNAC, (D) 0.2mM 1:1 hexanoyl-
SNAC: D11-hexanoyl-SNAC to the stcAΔ, hybrid B2, stcKΔstcJΔ strain. Deuterium-
incorporated forms are highlighted in blue and non-incorporated forms are highlighted in
pink. The y axis (relative abundance) of (a), which is compound 1, is normalized to 1
based on abundance of highlighted peak in B (a). The y axis of (b)- (e) and (f)- (i) is
respectively at 10x and 20x magnitude of that of (a)

129

130

131

132
Chapter IV: Connecting The Pieces – Common Motifs In Aspergillus nidulans
Secondary Metabolome Research

The previous chapters described different methods for generating new fungal
metabolites, unveiling a common theme that provides insight into the manipulation of
biosynthetic pathways in Aspergillus nidulans. Rationally manipulating a biosynthesis
pathway with the aim of diversifying the structure of a natural product requires
knowledge and tools. First, there needs to be increasing knowledge on the biosynthesis of
the natural product and the genes in the pathway. Acquiring this knowledge will be aided
by a sequenced genome of the organism and annotation of genes. Secondly, the
development of genetic tools (e.g. an efficient gene-targeting system) will facilitate in
manipulation of a secondary metabolite pathway. These essential components will then
expedite the process of accomplishing the intended goal.
In this case, both the biosynthesis knowledge and the genetic tools were in place
for studying the asperfuranone pathway in Aspergillus nidulans. Previous research
revealed the pathway consists of a cluster specific transcriptional activator, two
polyketide synthases, one hydroxylase, one oxidase and one oxidoreductase. Most
importantly, a gene-targeting system was used both to selectively knock out genes and to
artificially turn on the silent pathway. This provided groundwork for applying approaches
that use biosynthetic engineering with chemical synthesis.
Two different approaches were utilized on the asperfuranone pathway and each
had their own advantages and limitations. The first approach applied a modified
semisynthesis. This strategy benefited by utilizing an engineered strain to genetically

133
leverage the biosynthetic machinery to synthesize an advanced, chiral polyketide
intermediate cheaply and in large quantities. A second advantage was using synthetic
chemistry to achieve uniquely short syntheses of a natural product and “unnatural”
analogs, and it also provided enough product for evaluating biological activity. Although
the fungus produces an advanced intermediate for synthesis, the strategy falls short
because the advanced intermediate is limited in the synthetic transformations it can
undergo. In the second strategy a loading-domain swap was applied to the same pathway.
The unique advantage of this approach was the polyketide backbone could be structurally
reconfigured, whereas semi-synthesis is usually suited for late-stage modifications. The
limitation, and a common obstacle for most approaches, is the low yield of new
compound produced by the organism.
Aside from introducing greater diversity into natural products of A. nidulans, a
couple of intriguing discoveries were made. First, the fact that an engineered fungal strain
can produce nearly 1 g/L of a polyketide intermediate suggests that there is less
regulation of intermediates in a secondary metabolite pathway. Secondly, although the
biological activity of the new azaphilone analogs were not significantly improved from
the natural product (+)-sclerotiorin, the structure-activity relationships revealed new
insights on the structural motifs crucial for maintaining activity. In the domain swapping
strategy, precursor incorporation studies with a triple mutant revealed fundamental
insight into iterative NR-PKS programming. Typically, mutasynthesis studies are used
for generating chemical diversity, and incorporation studies with deuterium-labeled
precursors are a common strategy for elucidating the biosynthesis of a natural product.

134
However, the combination illuminated how a polyketide is processed by hybrid AfoE and
also suggested that the KS and PT domains are controlling the length of the polyketide in
a fungal NR-PKS.
Biotechnology plays an instrumental and prevalent role in this work. Each chapter
illustrates different biosynthetic engineering strategies applied to the asperfuranone
pathway using gene-targeting technology. One strategy showcases the titer of a selected
metabolite can be enhanced by two genetic mutations, 1) the promoter exchange from
native to inducible and 2) knocking out a gene downstream of the non-reduced PKS to
stop the biosynthesis of asperfuranone prematurely. The second strategy engineers the
pathway to produce an “unnatural” compound in vivo, which required two different
genetic mutations, 1) the knocking out of the stcA gene to eliminate this precursor sink,
and 2) the swapping of the SAT domain AfoE with the SAT domain of StcA to accept the
unnatural starter unit hexanoyl-CoA. Additionally, the domain swap was also facilitated
by another advancement, the UMA algorithm.
In summary, Aspergillus nidulans secondary metabolism is complex, yet
advancements in biotechnology have made it readily amenable toward biosynthetic
manipulation to generate new fungal compounds. To continue expanding natural product
diversity in vivo may likely require integrating multiple approaches (e.g. mutasynthesis
and combinatorial biosynthesis) to access a larger pool of potentially bioactive
compounds. And ultimately the design of the experiment will depend on each approach’s
capabilities and limitations. In many studies the main limitation is low titer, a significant
obstacle that may be alleviated by metabolic engineering such as introducing an inducible

135
or constitutive promoter or considering a semi-synthesis strategy. Of course, these
solutions can not be successfully applied to every case; however, it is hoped that a deeper
understanding of substrate flexibility of enzymes, secondary metabolism regulation, and
toxicity of “unnatural” products on microbes will be gained and culminate in creating
efficient methods to access diverse “unnatural” natural products.

136
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Asset Metadata

Creator Somoza, Amber Dorothy (author)

Core Title Application of biological and chemical approaches to generate new and diverse fungal natural products

Contributor Electronically uploaded by the author (provenance)

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Chemistry

Publication Date 08/02/2012

Defense Date 05/21/2012

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag Aspergillus,biosynthesis,fungi,genetic engineering,oai:digitallibrary.usc.edu:usctheses,OAI-PMH Harvest,Secondary metabolites

Language English

Advisor Wang, Clay C.C. (committee chair), Olenyuk, Bogdan (committee member), Roberts, Richard W. (committee member)

Creator Email amber.somoza@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c3-85291

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Legacy Identifier etd-SomozaAmbe-1125.pdf

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Document Type Dissertation

Rights Somoza, Amber Dorothy

Type texts

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Access Conditions The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...

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Abstract (if available)

Abstract Filamentous fungi are prolific producers of bioactive secondary metabolites. Recent genome sequencing reveals fungi harbor more secondary metabolites than are currently known. Exploration of fungal secondary metabolism is more attractive today with recent advancements in genomics and molecular biology. Efficient gene-targeting technology is a powerful tool used to “mine” the genome for novel secondary metabolites and identify the genes in the biosynthetic pathway. Furthermore, this technology can be applied to engineering pathways to generate “unnatural” natural products. Integration of biosynthetic engineering with chemical synthesis can introduce greater structural diversity into natural products, a promising avenue for discovering therapeutic drugs. The current work describes strategies that utilize the strengths of biosynthetic engineering and chemical synthesis to generate novel fungal natural products. ❧ Genome mining efforts in Aspergillus nidulans revealed the novel azaphilone polyketide, asperfuranone