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Harnessing synthetic biology for drug discovery & waste valorization

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Harnessing synthetic biology for drug discovery & waste valorization

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Content Harnessing Synthetic Biology for Drug Discovery & Waste Valorization
by Chris Rabot
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
PHARMACEUTICAL SCIENCES
December 2022
Copyright 2022 Chris Rabot
ii

Epigraph

“The advance of genetic engineering makes
it quite conceivable that we will begin to
design our own evolutionary process”

Isaac Asimove

iii

Acknowledgements

I am incredibly fortunate to have had such a supportive network of people standing by me
over these past five years during my Ph.D. First, I would like to thank my family – George, Jackie,
Mom, & Dad for believing in my vision of becoming a scientist. I am lucky to have you as my
biggest supporters throughout this journey.
I owe a special thank you to Dr. Yi-Ming Chiang for truly being one of the most helpful
scientists that I have had the pleasure of working with. His breadth of knowledge, spanning from
fungal biology to molecular genetics, drug discovery, and analytical chemistry is genuinely stunning.
He has helped me troubleshoot problems more time than I could possibly count. I can say with
confidence that not only I, but our entire lab, would not be nearly as successful as it is if it were not
for Yi-Ming.
To my lab mates – both current and former – I look forward to my day ahead in lab thanks
to you. Eva – this journey has been difficult for us both. We’ve seen each other experience the highs
and lows of science, which I think has been a valuable learning experience for both of us. You are as
amazing of a scientist as you are a person, and I look forward to seeing what you do with the
knowledge that you’ve gained during your Ph.D. To Dean – I have been incredibly impressed seeing
how quickly you’ve built up your scientific repertoire. I’m looking forward to keeping track of your
progress as a professor in Taiwan.
To the newer Ph.D students Michael & Jennifer and to our resident Pharm.D Yijun – the
first few months (or years) in lab are particularly difficult, but it gets better. The number of
techniques that you must learn can be overwhelming, but I’ve seen both of you consistently
progressing – both technically and intellectually. I am fortunate to have spent my days in lab with
you.
To the undergraduates – Maria, Ben, Salma, and Celis – and the high school students –
Anoushka, Henry, and Tristen – that I’ve mentored in the lab environment, I hope I did my job of
showing you what it’s like to do scientific research. I really enjoyed seeing you develop your skills
and I look forward to seeing your next career steps.
To the lab alumni – Michelle, Ada, Jillian, Jan, Kevin, Simon, Swati, Sue, Jingyi, Frank, Ngan
– you have been immeasurably helpful to me. Thank you for always teaching me and helping me
troubleshoot problems whenever they would arise. Seeing what each of you are doing now inspires
me to continue to learn and grow.
iv

To my collaborators – Berl, Liz, Richard, Emily, Carly, Cameron, Jan, Moh, and Ivan – I am
fortunate to have had the opportunity to work with you. The depth of your helpful insights have
been incredibly helpful to me and my work. I will continue to track the research that you continue to
conduct in the future.
To Travis – I initially thought the limit of my interaction with you would be during my
qualifying exam defense. I am glad that this was not the case. I’ve been inspired by you and your
profound knowledge in chemistry. Your obvious enthusiasm regarding scientific inquiry and
communication is infectious. To be honest, I initially did not look forward to our Friday calls,
because consistently communicating negative results eventually became frustrating. I think, though,
that the success that we’ve created regarding the plastic projects exemplifies well what it means to be
scientist. We did not give up, and for that, we have landed upon some genuinely groundbreaking
results. Projects in this area seem to be multiplying, and I’m excited to see the future of the systems
that we have created. Thank you for being a valuable mentor to me.
To Yuhao – we were faced with a frankly ridiculous task regarding the plastic project. It felt
like years of repeatedly trying, and trying, and trying, only to have to repeat on our Friday calls that
this project still is not working. Now, though, we can see obviously what perseverance can yield.
Thank you for sticking by this project with me. You are an excellent chemist and an amazing friend.
Congratulations on the progress that you’ve made. I cannot wait to see what other incredible
research you will conduct.
Finally – to Clay: I am lucky to have such a knowledgeable, encouraging, and supportive
mentor for these past five years. It is fascinating seeing you come up with project ideas. Apparently,
crazy ideas sometimes work. They can also lead to some of the most innovative research projects
that have come out of this school, or even this field. I am fortunate to have had such a highly
respected and intellectually rigorous mentor to guide me through my research. You undoubtedly
have influenced my life trajectory in ways that I cannot begin to comprehend. Thank you for
believing in me.

v

Table of Contents
Epigraph .............................................................................................................................................................. ii
Acknowledgements ........................................................................................................................................... iii
List of Tables .................................................................................................................................................... vii
List of Figures .................................................................................................................................................. viii
Abbreviations ..................................................................................................................................................... xi
Abstract ............................................................................................................................................................. xiii
Chapter 1: Introduction ................................................................................................................................ 1
1.1 The ubiquity and diversity of fungi .......................................................................................... 1
1.2 Fungal secondary metabolism and implications of biosynthesis ......................................... 1
1.3 Applications of fungi in biotechnology, medicine, and agriculture ..................................... 3
1.4 Harnessing synthetic biology to rewire the fungal phenotype ............................................. 3
1.5 Leveraging fungal biology to utilize noncanonical carbon sources ..................................... 4
1.6 References .................................................................................................................................... 5
Chapter 2: Rapid and Efficient Conversion of Polyethylenes to Fungal Secondary Metabolites ..... 6
2.1 Abstract ......................................................................................................................................... 6
2.2 Introduction ................................................................................................................................. 7
2.3 Results & Discussion ................................................................................................................ 10
2.4 Conclusions ................................................................................................................................ 18
2.5 Materials & Methods ................................................................................................................ 19
2.6 References .................................................................................................................................. 26
2.7 Supplementary text .................................................................................................................... 32
2.8 Supplementary figures .............................................................................................................. 34
2.9 Supplementary tables ................................................................................................................ 55
Chapter 3: Polystyrene Upcycling into Fungal Natural Products and a Biocontrol Agent .............. 64
3.1 Abstract ....................................................................................................................................... 64
3.2 Introduction ............................................................................................................................... 64
3.3 Results & Discussion ................................................................................................................ 69
3.4 Materials & Methods ................................................................................................................ 78
3.5 References .................................................................................................................................. 84
3.6 Supplementary figures .............................................................................................................. 90
3.7 Supplementary tables ................................................................................................................ 99
Chapter 4: Transcription Factor Engineering Leads to Discovery of a Novel Orsellinaldehyde
Derivative in Aspergillus nidulans ............................................................................................................... 105
4.1 Abstract ..................................................................................................................................... 105
4.2 Introduction ............................................................................................................................. 105
4.3 Results & Discussion .............................................................................................................. 109
4.4 Materials & Methods .............................................................................................................. 117
4.5 References ................................................................................................................................ 121
4.6 Supplementary figures ............................................................................................................ 127
4.7 Supplementary tables .............................................................................................................. 144
vi

Chapter 5: Molecular Genetic Identification of the Terrecyclic Acid and Quadrone Biosynthetic
Gene Cluster in Aspergillus terreus ............................................................................................................. 147
5.1 Abstract ..................................................................................................................................... 147
5.2 Introduction ............................................................................................................................. 147
5.3 Results & Discussion .............................................................................................................. 150
5.4 Materials & Methods .............................................................................................................. 157
5.5 References ................................................................................................................................ 161
Chapter 6: Conclusions and Future Directions .................................................................................... 165

vii

List of Tables

2.1 Screening of different catalytic systems for LDPE degradation ..................................................... 55
2.2 GCMS integration factors of methanol-esterified products ............................................................ 56
2.3 Optimization of reaction conditions with reduced O 2 and catalysts loading ................................ 57
2.4 Yields of diacid products after model LDPE degradation under different catalytic
conditions ................................................................................................................................................. 58
2.5 The degradation of post-consumer PE waste using optimized conditions ................................... 59
2.6 The yield of diacid products after post-consumer PE waste degradation using optimized
conditions ................................................................................................................................................. 59
2.7 Names, genotypes, genetic engineering techniques, and SM products of strains used in
Chapter 2 .................................................................................................................................................. 60
2.8 Chemical shifts of
1
H NMR spectrum obtained for asperbenzaldehyde compared to
literature values ....................................................................................................................................... 61
2.9 Chemical shifts of
1
H NMR spectrum obtained for citreoviridin compared to literature
values ........................................................................................................................................................ 62
2.10 Chemical shifts of
1
H NMR spectrum obtained for mutilin compared to literature values ....... 63
3.1 Screening of different catalytic conditions for PS degradation ....................................................... 99
3.2 Benzoic acid production from the various post-consumer PS waste sources .............................. 100
3.3
1
H NMR chemical shifts of ergothioneine compared to literature values ..................................... 101

3.4
1
H NMR chemical shifts of pleuromutilin compared to literature values ..................................... 102
3.5
1
H NMR chemical shifts of mutilin compared to literature values ................................................. 104
4.1 Gene names, lengths, protein product lengths, and predicted functions of genes adjacent to
the AN6791 PKS .................................................................................................................................... 110
4.2 Names of strains and genotypes used in Chapter 4 .......................................................................... 114
4.3 Experimental
1
H and
13
C chemical shift data of orsellinaldehyde compared to literature
values ........................................................................................................................................................ 144
4.4 Experimental
1
H and
13
C chemical shift data for diorsellinaldehyde compared to
literature values ....................................................................................................................................... 145
4.5 Experimental
1
H and
13
C chemical shift data for triorsellinaldehyde ............................................. 146
5.1 Putative terpene BGCs predicted by antiSMASH ............................................................................. 151
5.2 Media & buffer recipes .......................................................................................................................... 157
viii

List of Figures

2.1 The upcycling of polyethylenes to SMs ............................................................................................... 10
2.2 Post-consumer plastics degraded in Chapter 2 .................................................................................. 12
2.3 Overview of the novel promoter system driving production of asperbenzaldehyde in strain
LO10050 .................................................................................................................................................. 15
2.4 Paired HPLC-DAD-MS profiles of engineered strains cultured in GMM and PMM ................ 17
2.5
1
H NMR showing evidence of symmetrical dicarboxylic acids ...................................................... 34
2.6 GC-MS spectrum showing methanol-esterified products of aerobic digestion .......................... 35
2.7 Calibration curves determined by the standard methanol-esterified products ............................ 36
2.8 The correlation between carbon count of the methanol-esterified product and
corresponding GC-MS integration factors ........................................................................................ 37
2.9 The distribution of diacid products after different catalytic conditions of LDPE ..................... 38
2.10 Toxicity of dicarboxylic acids and reaction catalysts to fungi ........................................................ 39
2.11 Fungal utilization of dicarboxylic acids as sole carbon sources .................................................... 40
2.12 Paired HPLC-DAD-MS traces of crude and extracted polyethylene digest ............................... 41
2.13 Mass spectra of crude and extracted polyethylene digest ............................................................... 42
2.14 Comparative asperbenzaldehyde mass recoveries of strains LO2955, LO8355, and
LO10050 ................................................................................................................................................ 43
2.15 Standard curve of asperbenzaldehyde generated via HPLC-DAD .............................................. 44
2.16 Standard curve of citreoviridin generated via HPLC-DAD .......................................................... 45
2.17 Standard curve of mutilin generated via HPLC-DAD-MS ............................................................ 46
2.18
1
H NMR spectrum of asperbenzaldehyde purified from LO10050 ............................................. 47
2.19
1
H NMR spectrum of citreoviridin purified from YM192 ............................................................ 48
2.20
1
H NMR spectrum of mutilin purified from YM283 ..................................................................... 49
2.21 Tandem mass spectra of asperbenzaldehyde produced in GMM and PMM .............................. 50
2.22 Tandem mass spectra of citreoviridin produced in GMM and PMM .......................................... 51
2.23 Tandem mass spectra of mutilin produced in GMM and PMM .................................................. 52
2.24 Asperbenzaldehyde mass recovery from strain LO10050 when cultured in liquid GMM
and PMM generated from post-consumer waste ............................................................................ 53
2.25 Phase contrast micrographs of LO10050 in GMM and PMM ...................................................... 54
3.1 Upcycling PS into structurally diverse SMs and spores of biocontrol agents .............................. 68
ix

3.2 Post-consumer PS sources used in Chapter 3 ..................................................................................... 70
3.3 Genetic engineering strategy to develop an A. nidulans strain capable of ergothioneine
production, and relative yields from strains YM267, YM820, and YM847 ................................... 72
3.4 Strategy to enable heterologous mutilin and pleuromutilin production in A. nidulans ................. 75
3.5 Comparative metabolomics of engineered strains of A. nidulans when cultured in GMM and
PSMM ........................................................................................................................................................ 77
3.6 The generation of spores of A. flavus Af36 from PS-derived BA ................................................... 78
3.7
1
H NMR spectrum of chloroform-extracted BA derived from PS ................................................ 90
3.8 Fungal metabolism and toxicity of BA and phthalic acid ................................................................ 91
3.9 Asperbenzaldehyde production in GMM compared to MM with different BA
concentrations ......................................................................................................................................... 92
3.10 Standard curve of ergothioneine generated via HPLC-DAD ......................................................... 93
3.11 Standard curve of pleuromutilin generated via HPLC-DAD-MS .................................................. 94
3.12 Standard curve of mutilin generated via HPLC-DAD-MS ............................................................. 95
3.13
1
H NMR spectrum of ergothioneine .................................................................................................. 96
3.14
1
H NMR spectrum of pleuromutilin ................................................................................................... 97
3.15
1
H NMR spectrum of mutilin .............................................................................................................. 98
4.1 Overall strategy and results of Chapter 4 ........................................................................................... 109
4.2 Strategy to generate a strain of A. nidulans that expresses a hybrid AN6788 TF to promote
SM biosynthesis ...................................................................................................................................... 113
4.2 HPLC-DAD-MS profiles showing induced compounds upon overexpression of TFs
adjacent to the AN6791 PKS ............................................................................................................... 116
4.3 Prediction of the AfoA DNA-binding motif using InterProScan .................................................. 127
4.4 Prediction of AfoA coiled-coil dimerization motifs using PCOILS .............................................. 128
4.5
1
H NMR spectrum of orsellinaldehyde ............................................................................................... 129
4.6
13
C NMR spectrum of orsellinaldehyde .............................................................................................. 130
4.7
1
H NMR spectrum of diorsellinaldehyde ............................................................................................ 131
4.8
13
C NMR spectrum of diorsellinaldehyde ........................................................................................... 132
4.9
1
H NMR spectrum of triorsellinaldehyde ........................................................................................... 133
4.10
13
C NMR spectrum of triorsellinaldehyde ......................................................................................... 134
4.11 gCOSY NMR spectrum of triorsellinaldehyde ................................................................................. 135
4.12 gHSQC NMR spectrum of triorsellinaldehyde ................................................................................. 136
x

4.13 gHMBC NMR spectrum of triorsellinaldehyde ................................................................................ 137
4.14 NOESY NMR spectrum of triorsellinaldehyde ................................................................................ 138
4.15 DEPT-90 NMR spectrum of triorsellinaldehyde ............................................................................. 139
4.16 DEPT-135 NMR spectrum of triorsellinaldehyde ........................................................................... 140
4.17 Mass spectrum for orsellinaldehyde .................................................................................................... 141
4.18 Mass spectrum for diorsellinaldehyde ................................................................................................ 142
4.19 Mass spectrum for triorsellinaldehyde ................................................................................................ 143
5.1 Chemical structures of terpenes and terpenoids from Aspergillus spp. .......................................... 150
5.2 Comparative metabolomics showing elimination of terrecyclic acid biosynthesis upon TS
deletion ..................................................................................................................................................... 155
5.3 The proposed biosynthetic pathway of quadrone and terrecyclic acid .......................................... 156

xi

Abbreviations

SM Secondary metabolite
BGC Biosynthetic gene cluster
PKS Polyketide synthase
NRPS Non-ribosomal peptide synthetase
TS Terpene synthase
TC Terpene cyclase
ATCC American Type Culture Collection
FGSC Fungal Genetics Stock Center
DNA Deoxyribonucleic acid
RNA Ribonucleic acid
mRNA Messenger RNA
ATP Adenosine triphosphate
GMM Glucose minimal media
LMM Lactose minimal media
PMM Polyethylene minimal media
PSMM Polystyrene minimal media
MM Minimal media
HPLC-DAD High-performance liquid chromatography-photodiode array
detection
HPLC-DAD-MS High-performance liquid chromatography-photodiode array
detection-mass spectrometry
TIC Total ion chromatogram
EIC Extracted ion chromatogram
HRESIMS High resolution electrospray ionization mass spectrometry
GC-MS Gas chromatography mass spectrometry
NMR Nuclear magnetic resonance
gCOSY Gradient correlation spectroscopy
gHSQC Gradient heteronuclear single quantum coherence
gHMBC Gradient heteronuclear multiple bond correlation
DEPT Distortionless enhancement by polarization transfer
xii

NOESY Nuclear Overhauser effect spectroscopy
MeOH Methanol
DCM Dichloromethane
EtOH Ethanol
EA Ethyl acetate
DMSO Dimethylsulfoxide
AcOH Acetic acid
IPP Isopentenyl pyrophosphate
DMAPP Dimethylallyl pyrophosphate
LDPE Low-density polyethylene
HDPE High-density polyethylene
PET Polyethylene terephthalate
FRP Fiber-reinforced plastic
PP Polypropylene
PS Polystyrene
PVC Polyvinyl chloride
BA Benzoic acid
CoA Coenzyme A
NHPI N-hydroxyphthalimide
NO Nitric oxide
alcA(p) alcA promoter
gpdA(p) gpdA promoter
CYP450 Cytochrome P450 monooxygenase
SDR Short-chain dehydrogenase/reductase
EU European Union

xiii

Abstract

Fungi represent one of the most versatile groups of organisms in existence. They and their
fermentation products have proven to be instrumental toward the advancement of medicine,
agriculture, and biotechnology. As evidence of their importance, even their transient absence in
ecological systems would lead to catastrophic environmental failure. What is more, most applications
in society that fungi confer simply result from wild-type strains. We are fortunate to exist in the
post-genomic era, which enables us to dramatically manipulate the behavior of fungal strains
through synthetic biology and genetic engineering approaches. The work detailed herein describes
approaches that I have taken to exploit the abilities of fungi to the fullest.
The principal goal of the research detailed herein is to exploit synthetic biological methods
toward two ends: (1) to discover new fungal SMs and/or to elucidate the mechanisms of their
biosynthesis, and (2) to utilize fungi as efficient cell factories the upcycle plastic waste into high-
value fungal products. I employ genetic engineering and analytical chemistry techniques to achieve
these goals.
In Chapter 2, I describe a system that I have developed in collaboration of the lab of Dr.
Travis Williams (USC, Department of Chemistry) to utilize fungi to upcycle polyethylenes into
structurally diverse and pharmacologically active SMs. Post-consumer and ocean-sourced
polyethylenes are first subjected to aerobic, catalytic degradation into populations of carboxylic
diacids. These diacids are then utilized as a sole carbon source by engineered strains of fungi to
generate SMs in high yield. The work in this chapter has been published in Angewandte Chemie
International Edition.
In Chapter 3, I describe the application of similar oxidative chemistry described in Chapter 2
to polystyrenes. We rapidly generate benzoic acid in very high yield from post-consumer polystyrene,
which is then upgraded by fungi to other interesting SMs. Further, we show that we can generate
xiv

spores of the agriculturally relevant biocontrol agent A. flavus Af36 directly from this polystyrene-
derived substrate. Apart from the relatively scare carbon atoms originating from starter fungal
cultures, essentially every carbon atom that incorporates into both SMs and entire organisms that we
produce originate entirely from post-consumer plastics. The work in this chapter has been submitted to a
peer-reviewed journal.
In Chapter 4, I focus on my efforts to activate a silent BGC encoding a putative highly-
reducing PKS (HR-PKS) in A. nidulans. To achieve this, we conduct a series of genetic
manipulations to activate this putative BGC through the modulation of transcription factor
expression. As a result of these efforts, we discovered an unexpected and novel product of a non-
reducing PKS that we term triorsellinaldehyde. In addition, this work revealed a complex network of
in trans biosynthetic regulation in A. nidulans. The work in this chapter is being prepared for
submission in a peer-reviewed journal.
In Chapter 5, I discuss genetic engineering techniques that I employed to generate a
genome-wide TS/TC knockout library in A. terreus. These efforts permitted the identification of the
BGC responsible for the biosynthesis of the sesquiterpenoids quadrone and terrecyclic acid A. We
also found that bioinformatic software packages used to identify SM BGCs vary widely in their
predictive abilities. The work in this chapter is being prepared for submission in a peer-reviewed
journal.
Finally, I discuss in Chapter 6 future directions for the work that I have conducted during
my dissertation research. I detail approaches to drug discovery that I find particularly exciting and
promising. I also explore subsequent projects that I would hope to see completed regarding fungal
upcycling of post-consumer plastics efforts that I have initiated.

1

Chapter 1: Introduction

1.1 The ubiquity and diversity of fungi

Fungi represent a kingdom of eukaryotes that are exceptionally diverse in their capabilities.
They have, throughout history, created incalculable opportunities both for humans and for other
groups of organisms. They have dispersed to essentially every corner of the Earth (and beyond):
they can withstand the harshest of environments, including high temperatures and extreme salinities;
they have been found on the International Space Station, inhabiting hydrothermal vents, in our gut
microbiome, throughout the arctic and Antarctic; they have even been found growing towards the
Chernobyl Nuclear Power Plant. Evidently, fungi represent a group of impressively resilient
organisms. The development of a thorough understanding of their abilities to adapt and thrive
represents an opportunity with incalculable potential.
1.2 Fungal secondary metabolism and implications of biosynthesis

One major factor that confers in fungi the ability for them to thrive in essentially any
environment is their potential to biosynthesize a vast repertoire of secondary metabolites (SMs).
SMs represent a diverse class of low-molecular weight organic compounds that are synthesized by
fungi, in addition to bacteria and plants. In contrast to primary metabolites, SMs are not strictly
required for the growth, survival, or reproduction of the organism that produces them
1
. However,
they frequently provide a selective advantage to the host by increasing its competitive fitness. Many
SMs (or derivatives thereof) exhibit antibacterial (i.e. penicillin) and antifungal (i.e. echinocandin B,
griseofulvin) properties to corroborate this claim. In fact, over two-thirds of SMs have been shown
to exhibit bioactivity
2
, which are not limited to antibacterial/antifungal properties. Indeed, many
SMs have been found to possess antineoplastic, anticholesterolemic, antiviral, and
2

immunosuppressive activities. As will be discussed, the research community has presently revealed
only a small fraction of this enormous catalog of natural products.
A driving force behind the potency and selectivity of SMs is made clear when one considers
evolutionary implications of the mechanism of their biosynthesis. SMs are enzymatically produced
3

and can be divided into classes based on the class of enzymes that direct their biosynthesis. The
most common types include the polyketides, non-ribosomal peptides, and terpenes. Each of these
classes of SMs are respectively produced by polyketide synthases (PKSs), non-ribosomal peptide
synthetases (NRPSs), and terpene synthases/cyclases (TS/TCs). In fungi, the genes that direct SM
biosynthesis are generally colocalized within biosynthetic gene clusters (BGCs). “Core” genes, which
typically are PKSs, NRPSs, or TS/TCs, synthesize the “backbone” of the SM, which is then further
functionalized by tailoring enzymes such as CYP450 monooxygenases, oxidases, reductases,
hydrolases, isomerases, prenyltransferases, etc. Collectively, the biosynthetic machinery of these
enzymes enables the production of remarkably diverse compounds that are subject to meticulous
regio- and stereocontrol. Further complexity arises from developmental regulation of SM
biosynthesis. When considering that these SMs originate from very simple building blocks, such as
acyl-CoA thioesters, peptides, and/or isoprenoid units, the structural diversity of SMs is staggering.
The efficacy of SMs is not an accident. The enzymatic nature of their biosynthesis drives
activity through chemical evolution
4,5
. Biosynthetic enzymes are some of the largest ever discovered
(often reaching into the MDa range) and they function by redirecting pools of primary metabolites
into secondary metabolic pathways. The production of the enzymes themselves, as well as the SMs
that they synthesize, is an energetically demanding process that must favor evolutionary selection -
the fungal host must be able to justify SM biosynthesis. If a host exists in an environment in which
the SM that it produces is not competitively advantageous, then populations of this host that express
fully intact and functional biosynthetic machinery would not be selected for. In contrast, mutations
3

in genes encoding biosynthetic enzymes that positively alter SM structure and/or activity would be
selected for in certain environmental circumstances. Thus, the enzymatic, and ultimately genetic
basis for SM biosynthesis has created a context conducive to the production of potent and selective
SMs.
1.3 Applications of fungi in biotechnology, medicine, and agriculture

In addition to SMs, many other products originating from fungi have been critical to society.
Ascomycetes, such as Aspergillus, Penicillium, and Trichoderma spp. have been fruitful sources of
products with diverse applications. Some species, such as A. niger, are used to produce a range of
industrially relevant chemicals, including citric acid, itaconic acid, and gluconic acid. Other strains of
A. niger, as well as A. oryzae and Trichoderma spp., are used to produce enzymes including proteases,
glucoamylases, and cellulases. Many fungal strains are used in the food and drink industry to
manufacture a wide variety of consumables including alcohols (beer, wine, sake, etc.), cheeses,
breads, vitamins, preservatives, and chemicals. Other fungal strains have found applications in
agriculture by increasing plant biomass and crop yields, deterring plant pathogens, and inducing
plant defenses. Clearly, the benefits that fungi offer are enormous, and we must leverage them to
fully realize their capabilities.
1.4 Harnessing synthetic biology to rewire the fungal phenotype

Critically, the vast applications of fungi detailed above are possible even with wild-type
fungal strains. The more recent development of molecular genetic toolboxes, largely attributable to
advances in genome sequencing, grants us the unprecedented opportunity to alter the fungal
genotype and resulting phenotype in ways that were not otherwise possible. Synthetic biology
approaches are now just beginning to be used to engineer fungal strains to increase fermentation
titers, rewire metabolic pathways, and create transgenic organisms. Indeed, the recent development
of efficient gene targeting techniques now permits specific genetic permutations, including promoter
4

replacement, targeted gene deletions, heterologous expression, knock-ins, knockdowns, and protein
engineering. Taken together, these techniques dramatically expand the possible applications of fungi.
1.5 Leveraging fungal biology to utilize noncanonical carbon sources

As is true for most organisms, sugars are optimal carbon sources suitable for growth. Fungi
grow readily using sugars, such as glucose, sucrose, lactose, and fructose. However, some research
has documented the fungal utilization of noncanonical substrates as alternative carbon sources.
These include acetic acid, caffeic acid, benzoic acid, and carboxylic diacids. As we have found, and
as will be detailed in two chapters in this thesis, these seemingly trivial observations have profound
consequences. Fungal metabolism of these substrates is a particularly useful ability if the substrates
are sourced from problematic sources, such as from plastics and plant biomass.

5

1.6 References

1. Keller, N. P. (2019). Fungal secondary metabolism: Regulation, function and drug discovery.
Nature Reviews Microbiology, 17(3), 167–180. https://doi.org/10.1038/s41579-018-0121-1

2. Pelaez, F. (2005). Handbook of Intustrial Mycology (Vol. 22). Marcel Dekker.

3. Oakley, B. R. (2017). Aspergillus nidulans. In Reference Module in Life Sciences (p.
B9780128096338061000). Elsevier. https://doi.org/10.1016/B978-0-12-809633-8.06093-3

4. Wink, M. (2003). Evolution of secondary metabolites from an ecological and molecular
phylogenetic perspective. Phytochemistry, 64(1), 3–19. https://doi.org/10.1016/S0031-
9422(03)00300-5

5. Firn, R. D., & Jones, C. G. (2000). The evolution of secondary metabolism—A unifying
model. Molecular Microbiology, 37(5), 989–994. https://doi.org/10.1046/j.1365-
2958.2000.02098.x

6

Chapter 2: Rapid and Efficient Conversion of Polyethylenes to
Fungal Secondary Metabolites

Authors: Chris Rabot, Yuhao Chen, Swati Bijlani, Yi-Ming Chiang, C. Elizabeth
Oakley, Berl R. Oakley, Travis J. Williams, Clay C. C. Wang

2.1 Abstract

Plastics have delivered countless improvements to society; their excellent physicochemical
properties enable them to be remarkably useful and versatile in essentially every industry today.
Indeed, plastics exhibit highly versatile characteristics that have allowed them to deliver
immeasurable benefits. These features have resulted in plastics being essentially indispensable to
society, and it is therefore likely that they will remain in use for the foreseeable future. It is thus
imperative that we can safely and responsibly mitigate the risks posed by plastics, which themselves
are expansive and pressing.
Waste plastics represent major environmental and economic burdens due to their ubiquity,
slow breakdown rates, and inadequacy of current recycling routes. Polyethylenes are particularly
problematic, because they lack robust recycling approaches despite being the most abundant plastics
in use today. We report a novel chemical and biological approach for the rapid conversion of
polyethylenes into structurally complex and pharmacologically active compounds. We present
conditions for aerobic, catalytic digestion of polyethylenes collected from post-consumer and
oceanic waste streams, creating carboxylic diacids that can then be used as a carbon source by the
fungus Aspergillus nidulans. As a proof of principle, we have engineered strains of A. nidulans to
synthesize the fungal secondary metabolites asperbenzaldehyde, citreoviridin, and mutilin when
grown on these digestion products. This hybrid approach considerably expands the catalog of
products to which polyethylenes can be upcycled.

7

2.2 Introduction

Plastic production is currently accelerating at a rate faster than any other material on the
planet
1,2
, and is estimated to reach a global production volume of 1.1 billion tons annually by 2040.
Only 9% of plastics were recycled as of 2015
3
. Millions of tons of plastics, in the form of trillions of
plastic particles
4
, leak from waste management systems into the environment, posing increasing
threats to our food supply and ecosystems
5
. Generally, polyesters are frequently recycled (ca. 30% of
polyethylene terephthalate (PET)), unlike polyolefins (ca. 6% of low-density polyethylene (LDPE))
2,3
.
Due to their robust microstructures and excellent physicochemical properties, polyethylenes have
been utilized to deliver countless improvements to quality of life and health. Polyethylenes are, thus,
likely to remain ubiquitous in society. To minimize the environmental hazards they present, we must
reclaim value embedded in these materials by developing viable upcycling approaches for them.
The same physicochemical properties that make polyethylenes useful also hinder their
degradation and recycling. Further exacerbating this problem are additives that necessarily
accompany any post-consumer waste stream, e.g. colorings and plasticizers. Unlike polyesters and
nylons, the chemical methods known to recycle or remanufacture polyethylenes are limited. Some
forcing methods (e.g. refluxing nitric acid, deep UV radiation) are known to cleave polyethylenes,
the former to give carboxylic acids
6-8
. This type of strategy was exemplified in an oxidative process in
which O 2 and nitric oxide (NO) were shown to cleave polyethylenes to carboxylic acids, nitrates, and
other oxygenates at 170 °C and 40 atm with 65% total yield
9
.
Separately, oxidant-free, catalytic approaches are emerging for polyethylene upcycling,
including alkane metathesis, hydrogenolysis, and related pathways to convert polyethylenes to light
alkanes
10,11
. While these have modest yields and require energy-intensive conditions, they avoid the
potential uncontrolled reactions that can result from heating organics with O 2. Still, oxidative
conditions have the important advantage of tolerance to impurities associated with post-consumer
8

polymer waste. These impurities, especially salts, are a particular concern in samples recovered from
the oceans or recycling centers.
Chemical approaches to polyethylene degradation generate a diverse distribution of products
because there are no functional handles in their pure hydrocarbon structures to direct a catalyst to
where the polymer should be cleaved. We’ve shown, by contrast, that thermoset epoxies and fiber-
reinforced polymers (FRPs) can be upcycled selectively, owing to the special chemistry of their
linking nitrogen atoms
12-14
. In polyethylenes, the diversity of products arising from cleavage either
limits the value of these products or creates a challenge of separating them. Thus, there is growing
interest in employing biological systems to break down plastics.
Over the past two decades, several groups have made tremendous progress toward the
biological upcycling of plastic degradation products. The discovery of enzymes capable of
depolymerizing PET has inspired extensive interest in both PET degradation and upcycling
15-18
.
Separately, whole-cell biocatalysts have also been employed to reclaim value embedded in PET.
Several groups have demonstrated the microbe-facilitated generation of polyhydroxyalkanoates
(PHAs) or related products from plastic-derived substrates
19-21
. Others have shown that PET-
derived PHAs can be converted to both alkenoic acids and hydrocarbon fuels
22
. One group utilized
E. coli to convert PET-derived terephthalic acid into diverse aromatics, including gallic acid and
catechol
23
. Others have engineered E. coli to upcycle terephthalic acid into vanillin
24
.
In contrast to PET, however, fewer biological upcycling approaches have been developed
for polyolefins such as LDPE and HDPE. While several groups have investigated the chemical
25-27

and biological degradation of these polymers
28-32
, approaches to biologically valorize these polymers
are limited.
We aimed to exploit fungi, which produce products worth billions of dollars each year
33
to
biologically upcycle polyethylenes. Their biosynthetic products include medically valuable secondary
9

metabolites (SMs) including antibiotics, the cholesterol-lowering statins, immunosuppressants, and
antifungals
34
. Because they have been reported to use diacids as carbon sources
35,36
, we sought to
generate structurally diverse and pharmacologically active SMs directly from polyethylene-derived
substrates.
We show here that post-consumer polyethylenes can be rapidly degraded to generate
substrates that are suitable for upgrading by fungal metabolism. As a proof of principle, we
demonstrate that these plastic-derived substrates can be used to produce the diverse SMs
asperbenzaldehyde, citreoviridin, and mutilin in useful yields (Scheme 1). We also demonstrate
robust genetic engineering strategies that permit the expression of biosynthetic gene clusters (BGCs)
from many different organisms. Thus, in principle, this method expands the catalog of products to
which polyethylenes can be upcycled to thousands of SMs.

10

Scheme 1. The upcycling of polyethylenes to SMs. Polyethylenes are chemically degraded using
metal catalysts and pressurized oxygen to generate a distribution of diacids, which are metabolized
by fungi to rapidly produce structurally diverse SMs.
2.3 Results & Discussion

2.3.1 Optimization of Polyethylene Digestion

By adapting conditions for the conversion of cyclohexane to adipic acid
37
, we were able to
optimize an initial system for polymer cleavage. Using O 2 consumption and
1
H NMR integration for
indicative signals as our characterization handles (Fig. S1), we eventually found conditions based on
cobalt and manganese salts and a phthalamide-based NO source
9
that give useful oxidative cleavage
results (Table S1). The distribution of α,ω-diacid products that are produced by the oxidative
chemistry was further be quantified by GCMS (see supplementary materials, Table S2 and Figs. S2-
4).
We observed that re-charging our reactor with additional O 2 did not restart the polymer
cleavage reaction and hypothesized that N-hydroxyphthalimide (NHPI) serves as a source of NO,
which is vented from the reactor headspace upon O 2 recharge. We see rapid hydrolysis of NHPI to
11

phthalic acid upon reaction initiation. We further observed that our metallic catalysts lost reactivity
in the recharge process (Table S3, entries 4-6), and that a better result was obtained when metal salts
were added portion-wise along with O 2 and NHPI at recharge (compare entries 6-7). Under
conditions optimized for full polymer conversion to relatively small diacid products (Fig. S5), we
observed 86 wt% mass recovery (entry 8) from a 5 g sample of clean polymer. Note that branched
products were not tabulated in this yield, because they could not be unambiguously identified by
identity to an authentic sample. Further, addition of oxygen to the polymer adds weight, so the
molar yield of carbon atoms was 52% to the named products.
Its highly tolerant O 2-based conditions give this method the critical and distinguishing
feature of tolerance of post-consumer wastes. We demonstrate that feature here with four examples
(Fig. 1 & Table S4). Note that plasticizers and branched fragments from this LDPE film were
omitted from the product accounting, although a large portion of these products are likely suitable
for fungal digestion. A plastic grocery bag was homogenized into dicarboxylic acids of length C4-
C12 with 36 wt% mass recovery (Table S4, entry 1). A plastic milk jug and laboratory squeeze bottle
(entries 2-3) were homogenized to heavier diacid products, respectively in 63 and 54 wt% mass
recovery. The higher and lower recoveries are explained by the difference of HDPE versus LDPE:
the HDPE milk jug does not have polymer branches that are omitted from the recovery calculation.

12

Figure 1. Post-consumer plastics degraded in this study. (A) From left to right: LDPE plastic
grocery bag, HDPE milk jug, LDPE laboratory squeeze bottle, Pacific gyre waste collected from
Santa Catalina Island, CA; (B) the distribution of diacid products after post-consumer PE waste
degradation using our optimized reaction.
The unique geography of Catalina Harbor at Santa Catalina Island, CA captures plastics that
wash out of the North Pacific gyre. With mixed plastics collected from Catalina Harbor (Fig. 1),
degradation afforded 31 wt% to C4-C6 diacids and 27 wt% heavier diacids. While we selected only
trash from this collection that visually appeared to be rigid like polyethylene, this experiment shows
that the conditions tolerate mixed plastics and impurities such as salts, tar, and biomass (barnacles).
13

2.3.2 Engineering Aspergillus nidulans to upgrade polyethylene digestion products
into secondary metabolites

Fungi represent attractive candidates for diacid upgrading due to their robust growth
capabilities, inexpensive cultivation requirements, engineerable metabolic pathways, and potential to
synthesize metabolites with potent and diverse bioactivities. Short chain diacids, however, have been
reported to inhibit fungal growth
38
. We confirmed that C4-C8 (studied individually, Fig. S6) were
toxic to the model filamentous fungus A. nidulans (strain FGSC A4) even when glucose was present
as a carbon source. We found, however, that A. nidulans utilizes C10 and C12 diacids as sole carbon
sources (Fig. S7) without signs of toxicity. We thus devised a system to separate polyethylene
digestion products of ≥10 carbons from those <10 carbons. A series of pH-controlled liquid-liquid
extractions permitted the rapid separation of C10+ diacids from light diacids and metal salts (Figs.
S8-9).
In a representative example (vide supra), 27 wt% of ocean-sourced polyethylenes were
converted to diacids that were discretely identifiable using authentic standards. It should be noted
that light diacids are not waste products. They may be used in large-market applications such as in
the synthesis of PBCx, a biodegradable plastic emerging in agricultural applications
39
. Our data also
suggest that these light diacids possess antifungal properties (Fig. S6) that may be exploited.
For attempts to produce SMs from polyethylene-derived diacids, the heavy diacid extract
was added to liquid minimal media at a concentration of 10 g L
-1
. Liquid cultures were inoculated
with fungal strains and incubated for several days (see ESI for a full extraction protocol, culture
conditions, and media recipes). SMs were analyzed and quantified from culture extracts via HPLC-
DAD and HPLC-DAD-MS.
Initial attempts to elicit SM production from various wild-type fungal strains resulted in only
small amounts of SMs as detected via HPLC-DAD-MS. We consequently genetically engineered A.
14

nidulans to overexpress SM biosynthetic genes or biosynthetic gene clusters (BGCs) and this proved
effective, allowing robust and efficient SM production.
In order to determine the versatility of this system, we attempted to engineer fungal strains
to produce various SMs using several BGC activation/expression approaches (Table S5). The SM
used as a readout for the first of these systems was asperbenzaldehyde, a major polyketide
intermediate in asperfuranone biosynthesis
40
. Asperbenzaldehyde and its derivatives disassemble tau
filaments, inhibit lipoxygenases, and inhibit the interactions of the oncogenic RNA-binding proteins
HuR and Musashi-1 with their target mRNAs
41-43
. We chose to target a biosynthetic intermediate
because it can serve as a discovery platform that can easily be synthetically modified.
The Oakley lab developed three strains with different systems for driving asperbenzaldehyde
production: LO2955, LO8355, and LO10050. All molecular genetic modifications were executed
using previously described fusion PCR-based construct generation and transformation protocols
44
.
In strain LO2955, the afoD gene was deleted, blocking asperfuranone biosynthesis such that
asperbenzaldehyde, its biosynthetic precursor, accumulates. Further, the promoter of the afoA gene
that encodes the transcription factor (AfoA) that drives expression of the asperfuranone BGC was
replaced with the alcA promoter (alcA(p)), which is highly inducible with a variety of alcohols and
ketones, including methyl ethyl ketone
45
. To increase expression of AfoA, we next replaced the
promoter of the alcR gene with the strong, constitutive gpdA promoter
46
in LO2955, creating strain
LO8355. The alcR sequence encodes a transcription factor that drives expression of alcA
47
.
In addition, we developed a new, strong constitutive promoter system that employs a
positive feedback loop (Fig. 2) and incorporated it in strain LO10050. This system requires no
induction and should drive strong expression on any carbon source, whereas the AlcA system is
repressed by a number of sugars including glucose. The positive feedback system is designed to
drive very high levels of transcription. In addition to the new promoter system and the deletion of
15

afoD, LO10050 also carries deletions of the entire sterigmatocystin BGC (genes AN7804-AN7825)
and the emericellamide BGC (genes AN2545-AN2549). Deletion of these highly expressed BGCs
increases the pool of SM precursors, which are then free to feed into asperbenzaldehyde
biosynthesis.

Figure 2. Overview of the novel promoter system driving production of asperbenzaldehyde in
strain LO10050. The constitutive promoter gpdA(p) drives expression of afoA, encoding the AfoA
transcription factor. AfoA binds to the promoter regions of genes afoG, afoE, and afoC within the
asperbenzaldehyde BGC, leading to their expression and subsequent asperbenzaldehyde production.
AfoA also binds to the afoE promoter (afoE(p)) controlling a second copy of afoA inserted
elsewhere in the genome, driving additional AfoA production. This results in a positive feedback
loop that generates high levels of both AfoA and asperbenzaldehyde. Note that afoD is deleted,
halting conversion of asperbenzaldehyde to downstream metabolites. Other genes responsible for
conversion of further downstream products to asperfuranone, the final product of the pathway, are
not shown.

Yields of each strain grown in liquid lactose minimal media (LMM) were quantified via
HPLC-DAD (Fig. S10). Each strain gave substantial yields but yields from LO10050 were the
highest (4.3 g L
-1
from 15 g L
-1
lactose, or 29% mass conversion of lactose to asperbenzaldehyde).
We therefore selected this strain to assay for asperbenzaldehyde production on polyethylene digest.
To determine the general utility of the system, we also attempted to express the diterpene
antibiotic platform mutilin from the basidiomycete Clitopilus passeckerianus and the F1-ATPase β-
16

subunit inhibitor citreoviridin from A. terreus var. aureus
48
. Mutilin is an intermediate in the
biosynthetic pathway for pleuromutilin, which binds to the peptidyl transferase center of the
bacterial ribosome, thus halting protein synthesis
49
. Mutilin is therefore an attractive platform for
medicinal discovery efforts toward overcoming bacterial antibiotic resistance. Further,
basidiomycetes are phylogenetically distant from ascomycetes such as A. nidulans and the ability to
produce mutilin would indicate that this system works for BGCs from very diverse fungi.
Citreoviridin is a potent mycotoxin that uncompetitively and noncompetitively inhibits ATP
hydrolysis and ATP synthesis, respectively, by binding to the β-subunit of F1-ATPase
50
. Compounds
in this class of mycotoxins have been investigated for the treatment of cancer
51
. In total, four genes
from A. terreus var. aureus and five genes from C. passeckerianus were transferred into an A. nidulans
recipient strain and placed under control of alcA(p) to generate robust producers of citreoviridin and
mutilin, respectively.
2.3.3 Fungal Metabolism of Polyethylene Digest

Engineered fungal strains were incubated in liquid minimal media supplemented with 10 g L
-
1
polyethylene digest extracts (PMM, polyethylene minimal media). Culture media and/or mycelia
were extracted with appropriate organic solvents (see Materials and Methods), which were then
analyzed via HPLC-DAD or HPLC-DAD-MS (Fig. 3). Standard curves were generated (Figs. S11-
13) using purified standards to quantify SM yields in PMM relative to glucose minimal media (GMM)
or minimal media controls. SMs were purified from polyethylene digest cultures and confirmed via
1
H NMR (Figs. S15-17) and tandem MS (Figs. S18-20).

17

Figure 3. (A) Paired extracted ion chromatograms generated via HPLC-DAD-MS.
Asperbenzaldehyde production in (I) GMM and (II) PMM; citreoviridin production in (III) GMM
and (IV) PMM; mutilin production in (V) GMM and (VI) PMM. Intensities are normalized for
metabolites in each condition (B) SM yields produced by engineered fungal strains when grown in
PMM and GMM liquid media. Bars represent means ± SD (n = 3). *P ≤ 0.05; **P ≤ 0.01; ***P ≤
0.005.
Our results indicate that engineered fungal strains can efficiently produce useful quantities of
each target SM in under one week. Interestingly, microscopic examination of LO10050 when
cultured in liquid PMM revealed initial stunted growth relative to GMM controls (Fig. S21).
However, we observed ample hyphal growth after 48 hours and abundant asperbenzaldehyde
18

crystals after 72 hours of incubation in PMM, which is consistent with our findings regarding
asperbenzaldehyde titers.
These yields are in contrast to other metabolic engineering efforts; while high-yielding strains
have been reported following extensive engineering
52
, ample SM production typically requires much
larger quantities of carbon source(s) to achieve comparable yields
53-55
. It is also noteworthy that our
yields were obtained from shake flasks with minimal optimization. Alteration of other culture
parameters known to influence fermentation titers (e.g. culture length, media components, etc.)
should permit significantly higher yields. Use of the strong constitutive promoter system may
increase production of citreoviridin and mutilin and codon optimization may further increase
mutilin production.
We further note that it was not necessary to employ metabolic engineering strategies to
confer the ability to metabolize polymer-derived diacids to the fungi; rather, simple extraction
protocols selectively isolated diacids suitable for fungal metabolism. Finally, it is quite likely that
polyethylene degradation products can be used as a carbon source in the production of other SMs.
The BGCs that we have expressed are from diverse fungi and the approaches we have developed
should permit the expression of BGCs from many sources. The combination of the catalytic
degradation of polyethylenes with genetic engineering of filamentous fungi represents a promising
strategy to plastic upcycling.
2.4 Conclusions

We present a method to rapidly upcycle post-consumer polyethylenes into structurally
diverse and medically useful SMs. We degrade these polyethylenes using oxidative catalysis to
generate a distribution of diacids. These diacids are rapidly isolated and upgraded by engineered
strains of A. nidulans to synthesize bioactive SMs. Taken together, this two-step process dramatically
expands the catalog of products to which polyethylenes can be upcycled to thousands of SMs.
19

2.5 Materials & Methods

2.5.1 General methods

All commercially available chemicals were purchased from TCI America and used as
received, except for cobalt nitrate and manganese nitrate, each obtained from Alfa Aesar. Acetic acid
and hexanes were purchased from EMD Millipore. The NMR solvents chloroform-d, methanol-d 4,
and acetone-d 6 were purchased from Cambridge Isotopes Laboratories. Solvents and metal salts
were used as received.
Model low-density polyethylene was purchased from LyondellBasell (model LDPE, brand
NA 270001) and used as received. The plastic grocery bag (LDPE), milk jug (HDPE) and acetone
squeeze bottle (LDPE) were collected from the waste stream. The mixed plastic waste samples were
collected from Catalina Harbor at Santa Catalina Island, CA. All the plastic waste samples were
roughly washed and then shredded using a benchtop coffee grinder before use.
1
H NMR spectra were collected on a Varian VNMRS 600 or 400MR spectrometer and
processed via MestreLab Mnova. All the chemical shifts are shown by the units of ppm and
referenced to the residual
1
H solvent peak; and line-listed according to (s) singlet, (d) double, (t)
triplet, etc.
2.5.2 General procedure for catalyst screening

In a 300 mL Parr reactor, ground model LDPE powder (5 g) was mixed with N-oxide
catalyst (NHPI: N-hydroxyphthalamide) and metal catalysts (0.5 g of each). Acetic acid (75 mL) was
added to the mixture, and the Parr reactor was pressurized to 16 bars with oxygen and stirred at
150 °C. The reaction was manually terminated when no more oxygen consumption was observed as
determined by change of reactor head pressure. Upon cooling to room temperature, the internal
pressure of the reactor was released. Volatiles including acetic acid were removed via distillation; the
reaction mixture was dissolved in 1 M NaOH solution (ca. 50 – 100 mL), and then filtered by
20

vacuum filtration. The resulting translucent solution was acidified using concentrated HCl droplets
to pH 1 and extracted with ethyl acetate (3 x 100 mL). The combined organic layers were dried over
Na 2SO 4, and the solvent was removed by rotary evaporation to afford a yellowish, oily product
mixture.
A known complication of our oxidative strategy is uncontrolled solvent oxidation
56
: this is a
tradeoff between safety/scalability versus tolerance of impurities in the polymer waste stream. To
scale these conditions, the process must operate at low enough O 2 pressure such that the vapor
pressure of acetic acid is above its upper explosivity limit so that uncontrolled reactions cannot
occur. Lowering O 2 pressure affords proportional reaction conversion (compare Table S3, entries 1-
3), but not a decrease in reaction rate (entries 2-3). The system can be restarted by recharging the
reactor with O 2 and catalysts (entry 4). These are consistent with a view that while [O 2] is not
kinetically relevant, positive pressure is necessary to maintain catalyst life.
Safety notes: all pressurized reactions should be isolated behind appropriately rated blast shields and conducted at the
minimal requisite scale. All reactions involving pressurized oxygen gas must have appropriately specified burst disks.
All operations involving heating acetic acid should be conducted in a chemical fume hood. An analysis of upper and
lower flammability limits should be conducted before heating and combination of oxygen gas and organic solvent. No
scaling of such procedures should be attempted without consulting appropriate experts in safety engineering.
2.5.3 Analytical qualification and purification

In a representative analytical protocol, the obtained degradation product mixture was
suspended in 100 mL methanol in a 250 mL round bottom flask, followed by adding a catalytic
amount (ca. 1 mL) of concentrated H 2SO 4. A reflux condenser was placed on the flask and the
mixture was stirred at 50 °C for 20 hours. Excess methanol was removed by distillation and the
system was neutralized by adding NaHCO 3 powder until no gas evolution was seen. The residue was
poured into diluted NaHCO 3 solution and extracted with ethyl acetate. The combined organic layers
21

were dried over Na 2SO 4, and the solvent was removed under vacuum to afford a dark-yellow oil.
The resulting mixture was dissolved in chloroform for mass-selective gas chromatography analysis
(GC-MS, Agilent HP 6890 GC network with an Agilent 5973 mass selective detector).
The calibration curve (R
2
> 0.990 and standard error < 0.05) of each methanol-esterified
product was generated by the corresponding standard dimethyl ester (C5 to C10 and C12;
synthesized from the corresponding diacids, respectively) at different concentrations (500, 833, 1000,
1250 and 1428 ppm, respectively; also see supplementary text section below).
2.5.4 General procedure for condition optimization

To a 300 mL Parr reactor was added ground model LDPE powder (5g), NHPI (0.5g), cobalt
nitrate (0.1 – 0.5 g), manganese nitrate (0.1 – 0.5 g) and acetic acid (75 mL). The reactor was
pressurized with O 2 (2 - 18 bars, see safety notes above) and stirred at 150 °C. Once the pressure gauge
reached zero bars, the reactor was removed from the heat and cooled to room temperature (50 – 60
min). The cooled reactor was then re-charged with O 2 and placed back to the heat to continue the
reaction. After various cycles (see supporting tables), the reactor was cooled to room temperature
and the remaining oxygen was released to the air once no further oxygen consumption was observed.
The reactor was then opened, recharged with catalysts, sealed, pressurized back with O 2 and heated
at 150 °C again to restart the reaction. After the reaction was completed, the resulting crude mixture
was treated by the same work-up procedure as the initial studies to obtain a yellow-tinted, oily
product mixture.
2.5.5 Digest purification/analyticals

Following ethyl acetate extraction, polyethylene digest was dried under vacuum, resuspended
in 100 mL deionized H 2O, and pH-adjusted to 13.0 using 5.5 M KOH to maximize solubility. The
solution was pH-adjusted to 8.0 using concentrated HCl (ca. 37% aq) and extracted with hexanes (3
x 1000 mL, technical grade), then further acidified to 1.0 and again extracted with hexanes (3 x 1000
22

mL). The combined hexane fractions were dried under vacuum and the obtained products were used
to generate liquid media for fungal cultures.
2.5.6 Media recipes

All recipes are based on that of minimal medium (MM): 12.0 g L
−1
NaNO 3, 3.04 g L
−1

KH 2PO 4, 1.04 g L
−1
KCl, 1.04 g L
−1
MgSO 4·7H 2O, and 1 mL L
−1
Hutner’s trace element solution
57
.
MM is supplemented with 10 g L
-1
d-glucose, lactose monohydrate, or polyethylene digest extract to
create glucose minimal media (GMM), lactose minimal media (LMM), or polyethylene minimal
media (PMM), respectively. Media was supplemented with riboflavin (2.5 mg L
-1
), pyridoxine (0.5
mg L
-1
), uracil (1.0 g L
-1
), and/or uridine (10 mM) when necessary. For solid cultures, agar was added
at a concentration of 1.5% w/v. For comparative metabolic experiments using liquid cultures, all
media was pH-adjusted to 8.0 using 5.5 M KOH.
2.5.7 Diacid toxicity assay (GMM + diacids)

To determine the extent of toxicity of diacids of various chain lengths, 5.0 x 10
5
spores were
inoculated into 24-well plates containing 2 mL of GMM plus 10.0, 1.0, and 0.1 g L
-1
of diacid
standards of length C4, 5, 6, 7, 8, 9, 10, and 12. Phthalic acid and NHPI were also included in the
analysis. Diacids standards and catalysts were initially dissolved in 60 µL (3% v/v) DMSO. GMM,
GMM + 3% v/v DMSO, MM, and MM + 3% v/v DMSO were used as controls. 24-well plates
were incubated for seven days at 37 °C (Fig. S6).
2.5.8 Diacid uptake assay (MM + diacids)

To determine if A. nidulans can utilize diacids of various lengths as sole carbon sources, 1.0 x
10
6
spores were inoculated into 12-well plates containing 10 mL MM plus 10 g L
-1
of diacid
standards of length C4, 5, 6, 7, 8, 9, 10, and 12. Diacids standards were initially dissolved in 300 µL
(3% v/v) DMSO. GMM, GMM + 3% v/v DMSO, MM, and MM + 3% v/v DMSO were used as
controls. 12-well plates were incubated for seven days at 37°C (Fig. S7).
23

2.5.9 Culture conditions

Spores (ca. 1.0 x 10
7
) of each strain were inoculated into 25 mL Erlenmeyer flasks containing
10 mL of media. Cultures were incubated at 37°C with shaking at 180 rpm. In the case of YM192
and YM283, 50 mM methyl ethyl ketone (MEK) was added 42 hours after inoculation for alcA(p)
induction. YM192 and YM283 were then incubated for an additional 72 hours. LO10050, which
does not require induction, was incubated for a total of 144 hours. For comparative metabolomics
experiment to determine relative production of asperbenzaldehyde, citreoviridin, and mutilin in
PMM vs. GMM, all strains were cultured in triplicate.
2.5.10 Comparative metabolomics of asperbenzaldehyde-producing strains

Spores (ca. 3.0 x 10
7
) of strains LO2955, LO8355, and LO10050 were inoculated in triplicate
into 125 mL Erlenmeyer flasks containing 30 mL of GMM. Cultures were incubated at 37 °C for
144 hours with shaking at 180 rpm. For strains LO2955 and 8355, 50 mM methyl ethyl ketone
(MEK) was added 42 hours after inoculation for alcA(p) induction.
2.5.11 Secondary metabolite analysis
For LO2955/LO8355/LO10050, 10 mL of methanol was added to culture flasks, which
were then sonicated for one hour. Culture media with methanol was filtered and dried (TurboVap
LV). Dried extracts were resuspended in 25 mL ddH 2O, which was extracted three times with 25
mL ethyl acetate. The organic phase was then dried, resuspended in 10 mL methanol, and diluted
1:99 in methanol. For YM192 and YM283, mycelia were filtered from the culture medium. The
media was then extracted three times with 25 mL of dichloromethane (for YM192) or ethyl acetate
(for YM283). Extracts were dried and resuspended in 10 mL methanol. Extracts from YM192 were
diluted 1:9 in methanol. Extracts from YM283 were analyzed without further dilution.
Extracts of LO2955/LO8355/LO10050 and YM192 (10 µL each) were analyzed via HPLC-
DAD (Agilent 1200 Series). Analysis was performed with an RP-18 column (Kinetex® 5 µm EVO
24

C 18 100 Å LC Column, 150 x 4.6 mm) at a flow rate of 1.0 mL min
-1
with detection using a DAD
detector. The solvents for both asperbenzaldehyde and citreoviridin analysis were 100% AcN
(solvent B) in 5% AcN−H 2O (solvent A), both with 0.05% TFA. For asperbenzaldehyde, the
solvent gradient used was: 0 to 60% solvent B from 0 to 12 min, 60 to 100% solvent B from 12 to
15 min, 100% solvent B from 15 to 20 min, 100 to 0% solvent B from 20 to 21 min, and
reequilibration with 0% solvent B from 21 to 26 min. For citreoviridin, the solvent gradient used
was: 0 to 100% solvent B from 0 to 17 min, 100% solvent B from 17 to 22 min, 100 to 0% solvent
B from 22 to 23 min, and reequilibration with 0% solvent B from 23 to 28 min.
As mutilin is not UV-active, 10 µL of YM283 culture extracts were analyzed via HPLC-
DAD-MS. Spectra were acquired with a ThermoFinnigan LCQ Advantage ion trap mass
spectrometer equipped with a reverse phase C 18 column (Alltech Prevail C 18; particle size, 3 μm;
column, 2.1 x 100 mm) with a flow rate of 125 μL min
−1
. The solvents used were 95% AcN-H 2O
(solvent B) in 5% AcN-H 2O (solvent A) plus 0.05% formic acid. The solvent gradient used was 0%
solvent B from 0 to 5 min, 0 to 100% solvent B from 5 to 35 min, 100% solvent B from 35 to 40
min, 100 to 0% solvent B from 40 to 45 min, and reequilibration with 0% solvent B from 45 to 50
min. MS conditions were as follows: 5.0 kV capillary voltage, sheath gas flow rate of 60 arbitrary
units (AUs), auxiliary gas flow rate of 10 AUs, and ion transfer capillary temperature at 350 °C.
2.5.12 Standard curve generation

Purified standards of SMs were analyzed via HPLC-DAD (for asperbenzaldehyde and
citreoviridin) or HPLC-DAD-MS (for mutilin). Each SM was injected at concentrations of 1.0, 0.3,
0.1, 0.03, and 0.01 g L
-1
to generate standard curves (Figs. S11-13). Concentrations were plotted
against AUCs for each metabolite. For mutilin, the measured AUC corresponded to a positive-mode
extracted ion chromatogram (EIC) at m/z = 303, corresponding to the [M+H-H 2O]
+
ion. Microsoft
Excel was used to generate standard curves, R
2
values, and regression formulae.
25

2.5.13 Compound purification

For asperbenzaldehyde purification, ethyl acetate extracts from LO10050 grown in 30 mL
PMM were purified via HPLC-DAD equipped with a reverse phase C 18 column (Luna C 18(2); 100 Å;
250 x 10 mm). The solvents used were 100% ddH 2O (solvent A) and 100% AcN (solvent B), both
containing 0.05% trifluoroacetic acid with a flow rate of 4 mL min
-1
. The solvent gradient was: 30%
to 100% solvent B from 0 to 25 minutes, 100% solvent B from 25 to 30 minutes, 100% to 30%
solvent B from 30 to 31 minutes, and reequilibration at 30% solvent B from 31 to 36 minutes. 13.3
mg of asperbenzaldehyde was isolated.
For citreoviridin purification, YM192 was cultured in 30 mL PMM. Extracts were purified
via normal-phase preparative thin layer chromatography (Merck) using 3:1 ethyl acetate:hexanes as a
mobile phase to yield 8.6 mg of citreoviridin.
For mutilin purification, YM283 was cultured in 200 mL PMM. Culture medium was
extracted (vide supra), and extracts were purified via normal-phase preparative thin layer
chromatography using 3:7 ethyl acetate:hexanes as a mobile phase to yield 7.1 mg of mutilin.

26

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32

2.7 Supplementary text

2.7.1 Product quantification

We thought it necessary to quantify accurately the distribution of α,ω-diacid products that
are produced by the aerobic digest chemistry before we could develop the reaction more fully. We
addressed this analytical problem by converting mixed diacid products resulting from aerobic
polymer digestion to esters by acidic methanolysis to enable GC-MS analysis (Fig. S2). Products
identified for linear diacids are readily assigned by mass and confirmed by identity to standard diester
samples of the C5 (glutaric), 6 (adipic), 7, 8, 9, 10 and 12 diacids. Further, integration response
factors can be calibrated for these seven known compounds, and uncertainties can be measured for
each. Observing that these response factors are a linear function of carbon count with high co-
variance (Pearson’s R
2
> 0.99), we can predict with quantifiable certainty those response factors (eg.
C11) for which we have no authentic sample (see Fig S3-4 & Table S2). Response factors for the
inaccessible diesters are each within 2% standard error. Integration data from
1
H NMR, particularly
for dimethyl phthalate, dimethyl succinate, and methylene groups adjacent to product esters groups
enabled us to corroborate our GC assignments within the confidence of the NMR integration (d1 >
5 T1). This analysis neglects branched products resulting from pieces of LDPE that themselves
contain branches.
2.7.2 Purification of C10+ diacids

Based on the observation that fungi utilize C10+ diacids as sole carbon sources, we chose to
go forward with the less reactive oxidative system (Table S1, entry 6). In order to favor the
generation of heavy diacids, we reduced the initial oxygen loading to 4 bars without any further
recharging. In this case, 1.8 g of the initial 5.0 g model LDPE reacted after three hours. Following
workup and pH-controlled liquid-liquid extraction, roughly 150-180 mg of heavy diacids were
collected (confirmed by HPLC-DAD-MS, Figs. S8-9). Replacing model LDPE with oceanic waste
33

resulted in the recovery of 200-220 mg of heavy diacids post-extraction. This extraction permitted
the rapid and facile separation of C10+ diacids from diacids known to be toxic to the fungus. We
observed that, following a final hexanes extraction of aqueous medium at pH 1, insoluble material
that represented the majority of the ca. 1.8 g reacted starting material was also able to be utilized by
fungi as carbon sources. We nevertheless chose to utilize the hexanes extracts for metabolomics
experiments due to their absence of diacids C9 and below.

34

2.8 Supplementary figures

Figure S1.
1
H NMR showing evidence of symmetrical dicarboxylic acids. Dicarboxylic acids were
dissolved in methanol-d 4.
35

Figure S2. GC-MS spectrum showing methanol-esterified products of aerobic digestion (in
chloroform). The tall peak represents dimethyl phthalate.
36

Figure S3. Calibration curves determined by the standard methanol-esterified products.
37

Figure S4. The correlation between carbon count of the methanol-esterified product and
corresponding GC-MS integration factor.
38

Figure S5. The distribution of diacid products after model LDPE degradation under different
catalytic conditions. (Referring to entries in Table S3, see below).

39

Figure S6. 24-well plates showing toxicity of dicarboxylic acids and reaction catalysts to fungi. 1.0 x
10
6
spores of A. nidulans FGSC A4 were inoculated into each well containing 2 mL (0) GMM; (1)
MM; (2) GMM + 3% v/v DMSO; (3) MM + 3% v/v DMSO; (4-6) GMM + 10/5/1 g L
-1
NHPI;
(7-9) GMM + 10/5/1 g L
-1
phthalic acid; (10-12) GMM + 10/5/1 g L
-1
succinic acid; (13-15) GMM
+ 10/5/1 g L
-1
glutaric acid; (16-18) GMM + 10/5/1 g L
-1
adipic acid; (19-21) GMM + 10/5/1 g L
-1

pimelic acid; (22-24) GMM + 10/5/1 g L
-1
suberic acid; (25-27) GMM + 10/5/1 g L
-1
azelaic acid;
(28-30) GMM + 10/5/1 g L
-1
sebacic acid; (31-33) GMM + 10/5/1 g L
-1
dodecanedioic acid. All
diacid standards were initially dissolved in 60 μL (3% v/v) DMSO. Cultures were incubated for
seven days at 37 °C.

40

Figure S7. Fungal utilization of dicarboxylic acids as sole carbon sources. 1.0 x 10
6
spores of A.
nidulans FGSC A4 were inoculated into each well containing 10 mL (A) GMM; (B) MM; (C) GMM
+ 3% v/v DMSO; (D) MM + 3% v/v DMSO; (E) MM + 10 g L
-1
succinic acid; (F) MM + 10 g L
-1

glutaric acid; (G) MM + 10 g L
-1
adipic acid; (H) MM + 10 g L
-1
pimelic acid; (I) MM + 10 g L
-1

suberic acid; (J) MM + 10 g L
-1
azelaic acid; (K) MM + 10 g L
-1
sebacic acid; (L) MM + 10 g L
-1

dodecanedioic acid. All diacid standards were initially dissolved in 300 μL (3% v/v) DMSO.
Cultures were incubated for seven days at 37 °C.

41

Figure S8. Paired HPLC-DAD-MS traces of (A) crude polyethylene digest and (B) polyethylene
digest following pH-controlled liquid-liquid extraction. Diacid and phthalic acid peaks are annotated.

42

Figure S9. Mass spectra of (A) crude polyethylene digest and (B) polyethylene digest following pH-
controlled liquid-liquid extraction. Differences of m/z = 14 indicate consecutive additions of
methylene groups. The peak representing phthalic acid is highlighted in blue.
43

Figure S10. Comparative asperbenzaldehyde mass recoveries of strains LO2955, LO8355, and
LO10050. Yields were 1.27, 1.85, and 4.30 g L
-1
from 15 g L
-1
of carbon source for LO2955,
LO8355, and LO10050, respectively.

44

Figure S11. Standard curve of asperbenzaldehyde generated via HPLC-DAD.

45

Figure S12. Standard curve of citreoviridin generated via HPLC-DAD.
46

Figure S13. Standard curve of mutilin generated via HPLC-DAD-MS.
47

Figure S14.
1
H NMR spectrum of asperbenzaldehyde purified from LO10050 when cultured in
liquid PMM medium. 13.3 mg of asperbenzaldehyde was dissolved in acetone-d 6.
48

Figure S15.
1
H NMR spectrum of citreoviridin purified from YM192 when cultured in liquid PMM
medium. 8.6 mg of citreoviridin was dissolved in methanol-d 4.
49

Figure S16.
1
H NMR spectrum of mutilin purified from YM283 when cultured in liquid PMM
medium. 7.1 mg of mutilin was dissolved in methanol-d 4.
50

Figure S17. Tandem mass spectra of asperbenzaldehyde produced in (A) GMM and (B) PMM
liquid medium.
51

Figure S18. Tandem mass spectra of citreoviridin produced in (A) GMM and (B) PMM liquid
medium.
52

Figure S19. Tandem mass spectra of mutilin produced in (A) GMM and (B) PMM liquid medium.
The m/z ≈ 303 represents the [M+H-H 2O]
+
ion.
53

Figure S20. Asperbenzaldehyde mass recovery from strain LO10050 when cultured in liquid GMM
vs. PMM. PMM was generated using post-consumer polyethylene digest. Yields were 1.88 and 4.23 g
L
-1
asperbenzaldehyde from 10 g L
-1
of GMM and PMM, respectively.

54

Figure S21. Phase contrast micrographs of asperbenzaldehyde-producing strain LO10050 in GMM
and PMM. Cultures were grown in 10 mL liquid cultures in 30 mL Erlenmeyer flasks. Incubation
time is displayed on the left. In GMM (left column), hyphal growth is extensive by 24 h. Hyphae
remain healthy at 48 h but show some darkening at 72 h indicating a loss of viability. In PMM,
germination of the same strain is delayed; at 24 h only short germlings are present. By 48 hours
hyphal growth is robust. By 72 hours most hyphae are dark indicating that they are no longer viable.
However, crystals are abundant (one of which is indicated with an arrow). These crystals match the
previously reported description of asperbenzaldehyde crystals
40
. From the large amounts of
asperbenzaldehyde produced under these conditions as detected via HPLC-DAD, we are confident
that the crystals are asperbenzaldehyde. Asperbenzaldehyde crystals are seen occasionally in GMM,
but they are much less abundant than in PMM. Scale in each panel = 25 μm.

55

2.9 Supplementary tables

Table S1. Screening of different catalytic systems for LDPE degradation. (a) In a pressurized 300-
mL Parr reactor, model LDPE (5g) was reacted with molecular oxygen (16 bars) with the presence
of N-oxide catalyst (0.5g) and metal catalyst (0.5g of each) in acetic acid (75mL) at 150
o
C. The
reaction was manually terminated when no more oxygen consumption happened (b) NHS = N-
Hydroxysuccinimide; NHPI = N-Hydroxyphthalimide (c) Based on the relative NMR peak intensity
of alpha, beta and gamma protons: short (C4-C6), mid-short (C7-C8), medium (C9-C10), mid-long
(C11-C12) and long (C12+) (d) No reaction.

56

Table S2. GCMS integration factors of corresponding methanol-esterified products: (a) Integration
factors of C11, 13-18 diesters are calculated based on the correlation curve (also see Fig. S4).

57

Table S3a. Optimization of reaction condition with reduced O2 and catalysts loading: (a) Reaction
conditions unless specified mentioned otherwise: A mixture of model LDPE (5g), catalysts, and
acetic acid (75mL) was stirred in a 300-mL, oxygen-pressurized Parr reactor (with active refilling) at
150
o
C (b) Mass Recovery Yield (wt%) = [(Mass of diacid products)/(Mass of starting LDPE)] x 100%
(c) Molar Yield (%) = [(Mass of carbon in diacid products)/(Mass of carbon in starting LDPE)] x
100% (d) Extra 2 wt% of each metal catalyst and 10 wt% of NHPI was added after 6th oxygen
delivery cycle (e) Additional 4 wt% of each metal catalyst and 10 wt% of NHPI were added after 6th
oxygen delivery cycle (f) Extra 4 wt% of each metal catalyst and 20 wt% of NHPI were added after
6th oxygen delivery cycle.

58

Table S3b. The yield of diacid products after model LDPE degradation under different catalytic
conditions: (a) All entries are referred to entries in Table S3a (b) “Heavy” portion = a mixture of
diacids comprises linear products of mass > C10.

59

Table S4a. The degradation of post-consumer PE waste using our optimized condition (entry 8 in
table S3): (a) Various of post-consumer PE waste was investigated using our optimized protocol
(entry 8 in table S3) (b) LDPE Plastic grocery tag was used (c) HDPE Milk jug was used (d) LDPE
Laboratory squeeze bottle was used (e) Plastic waste collected from Santa Catalina Island was used.

Table S4b. The yield of diacid products after post-consumer PE waste degradation using our
optimized condition: (a) All entries are referred to entries in Table S4a.
60

Table S5. Names, genotypes, genetic engineering techniques, and SM products of strains used in
this study.

61

Table S6. Chemical shifts of 1H NMR spectrum obtained for asperbenzaldehyde compared to
literature values.

62

Table S7. Chemical shifts of
1
H NMR spectrum obtained for citreoviridin compared to literature
values.

63

Table S8. Chemical shifts of
1
H NMR spectrum obtained for mutilin compared to literature values.

64

Chapter 3: Polystyrene Upcycling into Fungal Natural Products and
a Biocontrol Agent

Authors: Chris Rabot, Yuhao Chen, Shu-Yi Lin, Ben Miller, Yi-Ming Chiang, C.
Elizabeth Oakley, Berl R. Oakley, Clay C. C. Wang, Travis J. Williams

3.1 Abstract

Polystyrene (PS) is one of the most used, yet infrequently recycled plastics. Although
manufactured on the scale of 300 million of tons per year globally, current approaches toward PS
degradation are energy- and carbon-inefficient, slow, and/or limited in the value that they reclaim.
We recently reported a scalable process to degrade post-consumer polyethylene-containing waste
streams into carboxylic diacids. Engineered fungal strains then upgrade these diacids biosynthetically
to synthesize pharmacologically active secondary metabolites. Herein, we apply a similar reaction to
rapidly convert PS to benzoic acid in high yield. Engineered strains of the filamentous fungus
Aspergillus nidulans then biosynthetically upgrade PS-derived crude benzoic acid to the structurally
diverse secondary metabolites ergothioneine, pleuromutilin, and mutilin. Further, we expand the
catalog of plastic-derived products to include spores of the industrially relevant biocontrol agent
Aspergillus flavus Af36 from crude PS-derived benzoic acid.
3.2 Introduction

Plastic polymers have delivered immeasurable benefits to society. Attributable to their
variable densities, plasticities, and other physicochemical properties, they have been extensively
applied to virtually every industry today. The profound benefits conferred by plastics necessitate that
they will remain an integral part of our economy. It is therefore imperative that we continue to
develop the expanding portfolio of diverse and effective methods to maximize the value that we
reclaim from them at the end of their first use.
65

Polystyrene (PS) is one of the most common, but least frequently recycled plastics.
Current recycling rates of PS are extremely low; only 0.9% of PS was recycled in 2018 in the United
States.
1
PS is manufactured in several forms: about 10% is expanded polystyrene foam (EPS), 50% is
pure PS, and the remaining 40% is blended with other plastics to form copolymers.
2
These different
forms of PS have widely varied densities, from 0.016 to 1.04 g/cm3,
3
and as a result, they create
complications in waste sorting when they are managed in recycling centers. Attributable to its highly
stable and hydrophobic chemical structure, natural PS degradation is extremely slow.
4,5
Taken together, the volume of PS manufacturing, slow environmental degradation rate, and
complex recycling challenges lead to its abundance both in landfills and the environment, the latter
ultimately arriving in the oceans. In fact, PS occupies approximately one-third of landfills
worldwide,
6
and leaked PS waste in the environment
7,8
cause adverse health effects both to humans
9-
12
and wildlife.
13-15
Several groups have recently reported methods to convert PS to fine chemicals, such as
graphene
16
and styrene
17
. In the past year, multiple groups have independently reported the
conversion of PS to benzoic acid (BA).
18-21
In addition to chemical upcycling approaches, biological
solutions to plastic degradation have also been of increasing interest. Several fungal
22,23
and
bacterial
24-26
species have been shown to degrade PS or related polymers. Mealworms can also
consume PS.
27,28
Two groups have shown that Pseudomonas putida can convert styrene to
biodegradable polyhydroxyalkanoates (PHAs).
29,30
Recently, one group showed that mixed plastics,
including PS, can be oxidatively degraded and upgraded by an engineered strain of P. putida to β-
ketoadipate or PHAs.
31

We recently reported an oxidative catalytic method rapidly to degrade post-consumer
polyethylenes into distributions of carboxylic diacids.
32
Diacids suitable for fungal metabolism are
then upgraded by engineered strains of fungi to high quantities of pharmacologically active
66

secondary metabolites (SMs), all in under one week. Inspired by reports that fungi can utilize BA as
a carbon source,
33-35
we hypothesized that the same oxidative chemistry used to degrade
polyethylenes can likewise be applied to PS to generate BA or oligomers suitable for digestion. If
confirmed, this would represent a major step toward our ability to upcycle mixed plastics.
As such, we collected post-consumer PS, including ocean waste sourced from Catalina
Harbor on Santa Catalina Island, CA and subjected it to degradation. By employing similar oxidative
conditions, we demonstrate that our catalytic system can be used to convert PS sourced from post-
consumer and oceanic waste streams to BA (illustrated by
1
H NMR, Fig. S1) with up to 71% molar
yield in 12 hours. While pristine BA can be isolated from this method, we collected crude product
containing oligomeric material, thus enabling collection of > 71% of this PS material. This
combined fraction was used for fungal upgrading.
Having demonstrated that we can generate BA from PS, we then implemented genetic
engineering techniques to promote SM biosynthesis. Fungi are ideal candidates for plastic upgrading
due to their rapid growth, robust biosynthetic capabilities, genetic tractability, inexpensive cultivation
requirements, and high fermentation titers. Collectively, these characteristics confer the ability to
produce diverse products worth billions of dollars annually.
36
Attributable to advances in genome
sequencing, our understanding of fungal biosynthesis has rapidly progressed and facilitated the
development of efficient and customizable fungal fermentation platforms. We leveraged these
advances to enable production of the structurally diverse and pharmacologically active fungal SMs
ergothioneine, pleuromutilin, and mutilin from engineered strains of the filamentous fungus
Aspergillus nidulans using PS-derived BA. By introducing exogenous genes encoding biosynthetic
enzymes into an A. nidulans host and placing them under the inducible alcA promoter (alcA(p)), we
generated strains capable of upgrading PS digestion products to useful quantities of these SMs in
under one week.
67

We further use PS-derived BA to generate spores of the agriculturally relevant fungal strain
A. flavus Af36. Most strains of A. flavus are opportunistic pathogens that frequently infect maize,
peanuts, cottonseed, and tree nuts.
37
These strains produce a family of highly carcinogenic and
mutagenic polyketide-derived SMs called aflatoxins. Aflatoxins inflict massive crop losses and
account for an estimated 28% of global cases of liver cancers. Roughly 4.5 billion people have been
exposed to unsafe levels of aflatoxins.
38
Accordingly, many academic, industrial, and government
groups have focused on mitigating risks of aflatoxins over the preceding decades. One such method
is the use of A. flavus strain Af36. This natural strain, first isolated from Arizona, lacks the ability to
produce aflatoxin due to a single nucleotide polymorphism in the polyketide synthase aflC within the
aflatoxin biosynthetic gene cluster (BGC).
39-41
A. flavus Af36 can competitively displace aflatoxin-
producing strains of A. flavus, thereby limiting aflatoxin exposure. This strain is currently approved
for use in the United States by the Environmental Protection Agency as a biocontrol agent for
aflatoxin accumulation in peanuts, maize, and cottonseed.
42
The generation of spores of this strain
directly from PS-derived BA represents a unique and sustainable application of PS upcycling that has
potential mass demand that could address a meaningful portion of PS waste.
Our overall route to upgrade PS exploits crude PA digest, which is then converted to either
medically relevant SMs or spores of the biocontrol agent A. flavus Af36 (Scheme 1). Taken together,
this approach enables upcycling of post-consumer PS into multiple high-value products.
68

Scheme 1. Upcycling PS into structurally diverse SMs and spores of biocontrol agents. Post-
consumer PS was collected and subjected to catalytic, oxidative cleavage to generate BA in high yield.
This BA is then utilized as a sole carbon source by engineered strains of A. nidulans to generate the
SMs ergothioneine, mutilin, and pleuromutilin. Furthermore, this BA is also used to generate high
quantities of spores of the atoxigenic biocontrol agent A. flavus Af36.

69

3.3 Results & Discussion

3.3.1 Catalytic PS degradation conditions

We initiated our investigation of conditions for PS degradation using styrofoam insulated
boxes (Table S1) that we shredded by hand. We monitored BA generation via
1
H NMR integration
as a readout for the optimization of our catalytic conditions (Fig. S1). Low conversion efficiencies
were observed with manganese(II) acetoacetonate, while the corresponding nitrate salt afforded a
more reactive cleavage system (entries 1-3). Interestingly, we found that the introduction of cobalt
together with manganese synergistically promoted BA generation. For example, 5 wt% each of two
nitrate salts afforded a 26% yield of BA after four hours, but 10 wt% of either catalyst alone could
not permit a comparable conversion (entries 3-5).
Based on these results, we hypothesize that manganese is likely to act as a Lewis acid in
catalyzing electron-transfer oxidation from tertiary carbons in the polymer backbone, and cobalt
tends to catalyze the β-scission for C-C bond cleavage.
43
Furthermore, under optimal conditions
with portion-wise recharging of O 2, we observed the complete degradation of PS with up to 71% of
starting polymer recovered as BA and 84% total mass recovery (entries 5-7). The added balance
comprises incompletely digested oligomers. We also note that a reduction in reaction time did not
afford increased product yield, even under optimized conditions (entries 8-10).
We tested our conditions on four additional post-consumer PS sources: a styrofoam plate,
waste collected from Santa Catalina Island, CA, a disposable coffee cup lid, and a red drink cup (Fig.
1). The styrofoam plate and Catalina Island waste (Table S2, entries 2-3) were efficiently
homogenized into BA, with 51 and 39% molar yield, respectively. The coffee lid and red drink cup
(entries 4-5) were also degraded into BA, with 15 and 21% molar yield, respectively. The differences
in recoveries can be explained by the presence of additives, such as free radical scavengers and
composite polymers, that may inhibit catalytic oxidation. Trash was collected specifically from
70

Catalina Harbor, because the harbor’s unique geography accumulates pieces of the Great Pacific
Garbage Patch that wash up from the North Pacific Gyre. This enables us to demonstrate our
method as an approach to recycle the garbage patch itself.
Following catalysis optimization, we developed a simple procedure to isolate polymer digest
for downstream fungal upgrading. A series of liquid-liquid extractions followed by recrystallization
afforded BA in high purity as indicated by
1
H NMR. This isolated BA was then used for
downstream metabolomics experiments. For a detailed extraction protocol, see Materials & Methods.

Figure 1. (A) Post-consumer PS sources used in this study; (B) Mass recoveries corresponding to
each PS source. (I) Styrofoam cold box; (II) Styrofoam plate; (III) Catalina Island waste; (IV) Coffee
lid; (V) Red drink cup.

71

3.3.2 Fungal metabolism of benzoic acid

We first confirmed that fungi can utilize BA as a sole carbon source using the model
filamentous fungus A. nidulans FGSC A4 (Fig. S2). We observed a slight discoloration in the
presence of BA relative to glucose minimal media (GMM) positive controls, indicating some degree
of BA-induced toxicity. We separately determined that BA was not significantly toxic to the fungus.
We repeated the above experiments with phthalic acid, the hydrolysis product of the NHPI catalyst,
to confirm that it is both unable to be metabolized and is nontoxic to the fungus (Fig. S2).
For initial metabolomics experiments, we utilized the strain A. nidulans LO10050 to
determine if SMs can be generated from a BA standard as a carbon source. Reported previously,
32

LO10050 has been engineered to express certain genes from the asperfuranone biosynthetic gene
cluster (BGC). The incorporation of a positive feedback promoter system into this strain drives
production of the biosynthetic intermediate asperbenzaldehyde to very high levels. The high
production yield of asperbenzaldehyde allowed us to easily compare initial culture conditions
permissive of SM production from BA.
We took advantage of this robust asperbenzaldehyde production system to determine
preliminary culture conditions for fungal metabolism of PS-derived BA. We noticed that incubation
of fungal strains in the presence of a BA standard affected the morphology of the strain; spherical
mycelia characteristic of filamentous fungi cultured in shake flasks were not observed in these
culture conditions. Nevertheless, microscopic examination of culture medium revealed the presence
of extensive hyphae, revealing that growth attributable to BA was still occurring. We therefore chose
to use asperbenzaldehyde production as a surrogate endpoint for fungal growth during these initial
experiments.
Due to its reported toxicity to fungi,
34
we next determined the concentration of a BA
standard that permitted the highest yield of asperbenzaldehyde in liquid cultures. We found that
72

LO10050 could dose-dependently utilize BA as a sole carbon source to generate asperbenzaldehyde
(Fig. S3): production plateaued when LO10050 was cultured in MM supplemented with 12.5 g L
-1

BA, with areas under the curve (AUCs) reaching ca. 270% that of GMM controls. We eventually
observed a decrease in asperbenzaldehyde yield when cultures were supplemented with 15.0 g L
-1

BA, presumably due to BA toxicity.

Figure 2. Strategy to enable heterologous ergothioneine production in A. nidulans. (A) Top: genetic
architecture of the na-tive afo regulon in A. nidulans. AN1029 (afoA) encodes a TF that regulates
expression of each gene in the BGC, leading to production of asperfuranone, the final product of
the pathway. Bottom: replacement of the coding regions of various genes in the afo regulon with
endogenous (AN7620 and AN6227 from A. nidulans) and exogenous (Afu2g15650 and Afu2g13295
from A. fumigatus, NCU04343 and NCU11365 from N. crassa) ergothioneine biosynthetic genes.
Expression of AfoA is driven to high levels by alcA(p), which then binds to the native promoter
regions of genes within the afo regulon, leading to their expression. The egt1 and egt2 genes from A.
nidulans, A. fumigatus, and N. crassa are shown in gold, green, and pur-ple, respectively; (B) the
biosynthetic pathway of ergothioneine in N. crassa; (C) relative ergothioneine yields from strains
YM267, YM812, YM820, and YM847. Bars represent means and error bars represent SDs. **** p ≤
0.0001; *** p ≤ 0.0005.

73

3.3.3 Strain engineering – ergothioneine

To promote the efficient bioconversion of waste-derived BA in high yield, several genetic
engineering strategies were employed to generate three medically- and industrially-relevant SMs:
ergothioneine, pleuromutilin, and mutilin. The first SM that we aimed to generate from BA was
ergothioneine, an unusual thio-histidine betaine amino acid.
44
Ergothioneine is a natural antioxidant
that can be microbially synthesized by certain species of fungi and actinobacteria. It has been
reported to exhibit anti-inflammatory and cytoprotective properties, leading to its growing
application in the pharmaceutical and cosmetic industries.
45
Although discovered more than a century ago, there has recently been exponential growth in
publications related to ergothioneine.
46
The genetic basis of its biosynthesis has been elucidated both
in fungi and actinobacteria. In the latter, five genes (egtABCDE) direct its biosynthesis from histidine
and cysteine.
47
However, fungi can synthesize ergothioneine using only two genes: egt1 and egt2.
48,49

To engineer a strain to produce ergothioneine in useful quantities, we took advantage of the robust
A. nidulans afo regulon
50
to express ergothioneine biosynthetic genes. This regulon natively governs
production of the polyketide SM asperfuranone; overexpression of afoA, encoding a cluster-specific
transcription factor, has been shown strongly to activate all genes within the BGC, leading to
production of very high levels of products of the cluster. To exploit the robust expression profile of
this regulon, we replaced the coding regions of genes within the BGC with genes involved in
ergothioneine biosynthesis.
First, BLASTp was used to identify putative A. nidulans homologs using the Neurospora crassa
egt1 and egt2 as queries. This recovered two sequences bearing moderate homology to egt1 and egt2:
AN7620 (63% similarity/50% identity) and AN6227 (58% similarity/43% identity). AN7620 and
AN6227 were then amplified using A. nidulans genomic DNA and inserted into the coding regions
of afoG and afoF, respectively. Maintenance of the native promoters of each of these genes allows for
74

afoA to bind to them and drive their expression. We additionally replaced the native promoter of
afoA with alcA(p) to create strain YM267.
To further increase yields, we replaced the coding regions of afoE and afoD with the A.
fumigatus egt1 (Afu2g15650) and egt2 (Afu2g13295) homologs, respectively, to yield strain YM812. We
inserted a third pair of ergothioneine biosynthetic genes into the regulon by replacing the coding
regions of afoC and afoB with the N. crassa egt1 (NCU04343) and egt2 (NCU11365) to yield strain
YM820. Finally, we deleted the agsB gene encoding an α-1,3-glucan synthase
51
to create strain
YM847. Deletion of genes encoding α-1,3-glucan synthases have been shown to improve
fermentation titers by reducing hyphal clumping when grown in liquid cultures.
52
Collectively, these
genetic engineering strategies enabled the generation of a strain expressing an inducible system
controlling three pairs of ergothioneine biosynthetic genes (Fig. 2A). The full biosynthetic pathway
for ergothioneine in N. crassa is shown in Fig. 2B. All strains described above were cultured in
triplicate to determine their relative ergothioneine titers (Fig. 2C). Gratifyingly, YM847 was able to
produce over 170 mg L
-1
ergothioneine and was therefore selected for subsequent upgrading of PS-
derived BA.

75

Figure 3. Strategy to enable heterologous mutilin and pleuromutilin production in A. nidulans. (A)
Top: genetic architecture of the native afo regulon in A. nidulans. Middle: replacement of the coding
regions of various genes in the afo regulon with exogenous mutilin biosynthetic genes. Expression of
AfoA is driven to high levels by alcA(p), which then binds to the native promoter regions of genes
within the afo regulon, leading to their expression. In total, five genes (orange) from C. passeckerianus
were incorporated into the afo regulon to enable mutilin biosynthesis. Bottom: heterologous
expression of two additional genes (gold) from C. passeckerianus into the afo regulon enables total
reconstitution of the pleuromutilin biosynthetic pathway; (B) the biosynthetic pathway of mutilin
and pleuromutilin. Dimethylallyl pyrophosphate (DMAPP) and isopentenyl pyrophosphate (IPP)
first condense head-to-tail to form geranyl pyrophosphate (GPP). An additional IPP subunit
condenses with GPP to form farnesyl pyrophosphate (FPP). Geranylgeranyl pyrophosphate
synthase (Pl-ggs) condenses a final IPP subunit to form geranylgeranyl pyrophosphate (GGPP).
GGPP undergoes intramolecular cyclization catalyzed by the gene product of Pl-cyc encoding a
terpene cyclase. Two cytochrome p450s encoded by Pl-p450-1 and Pl-p450-2 catalyze the installation
of two hydroxyl groups. The hydroxyl group bound to the cyclopentane ring is then oxidized to a
ketone by the gene product of Pl-sdr, encoding a short-chain dehydrogenase/reductase, to form
mutilin. An additional hydroxyl group bound to the cyclooctane ring is acetylated by the
acetyltransferase encoded by Pl-atf. Finally, a cytochrome P450 encoded by Pl-p450-3 hydroxylates
76

the primary carbon within the acetyl group to yield pleuromutilin, the final product of the
pathway.
57,59

3.3.4 Strain engineering – pleuromutilin & mutilin

We next utilized strains of A. nidulans engineered to synthesize the SMs pleuromutilin and
mutilin from BA. Pleuromutilin, a diterpene natural product produced by the basidiomycete
Clitopilus passeckerianus, was first discovered in 1950
53
. It and its derivatives function by selectively
inhibiting bacterial translation by binding to the peptidyl transferase center of the bacterial
ribosome.
54,55
Recently, the semisynthetic pleuromutilin derivative lefamulin was approved by the
FDA for the treatment of community-acquired bacterial pneumonia.
56
Its biosynthetic pathway,
involving seven genes in total, was elucidated in 2017.
57
We utilized strain YM343, reported recently
by our group,
58
that was engineered to reconstitute the entire pleuromutilin biosynthetic pathway to
produce pleuromutilin from PS- derived BA. Each gene of interest was placed under control of the
afo regulon to drive expression to very high levels. We also utilized strain YM283, which expresses
only five of the seven genes within the pleuromutilin BGC, to produce its precursor, mutilin.
Production of a biosynthetic precursor of the final product in the pathway should enable late-stage
synthetic derivatization. Details regarding the biosynthetic pathway of mutilin and pleuromutilin are
shown in Fig. 3.
3.3.5 Comparative metabolomics

We next determined if engineered fungal strains can biosynthesize SMs from PS-derived BA.
Metabolic profiling of strains YM847, YM283, and YM343 revealed that all three metabolites can be
produced in useful quantities from engineered strains of A. nidulans when grown in minimal media
with PS-derived BA (PSMM) as a carbon source (Fig. 4). To quantify SM yields, standard curves for
each SM were generated via HPLC-DAD or HPLC-DAD-MS (Figs. S4-6). Further, we isolated each
of these SMs from large-scale cultures and confirmed their structures via
1
H NMR (Figs. S7-9 and
Tables S3-5). We note that, although SM yields in GMM are higher than in PSMM, the generation of
77

non-trivial quantities of these valuable SMs from post-consumer PS still represents a transformative
approach to PS upcycling. We further note that these strains were able to produce these SMs with
minimal optimization in shake flasks. Optimization studies along with alteration of culture
parameters should easily enable higher SM yields.

Figure 4. Comparative metabolomics of engineered strains of A. nidulans when cultured in GMM vs.
PSMM. 1.0 x 10
7
spores of YM847, YM343, or YM283 were cultured in liquid GMM or PSMM.
Relative SM levels were quantified using HPLC-DAD (for ergothioneine) or HPLC-DAD-MS (for
pleuromutilin and mutilin). Bars represent means and error bars represent SDs. ns, not significant;
** p ≤ 0.01.

3.3.6 Biocontrol agent spore generation

Finally, we sought to determine if spores of an agriculturally-relevant biocontrol agent can be
generated from PS-derived BA. This is of particular importance to the field of plastics upcycling,
because the biocatalytic products produced to date, while more valuable than the parent polymer,
are not frequently used on scales that approach the quantity of plastic that will ultimately need to be
78

reclaimed. Thus, it is valuable to add widely-used agrichemicals to our product portfolio. To
determine feasibility of this application, we cultured A. flavus Af36 on solid GMM and PSMM agar
plates and quantified spore generation after a seven-day incubation period. Our results indicate that
spores of A. flavus Af36 can readily be generated using PS-derived BA (Fig. 5), with yields being 5.2-
fold higher in PSMM relative to GMM.

Figure 5. The generation of spores of A. flavus Af36 from PS-derived BA. A. flavus Af36 is an
atoxigenic strain that lacks the ability to produce aflatoxins. It is currently used agriculturally by
inoculation onto crops at various stages of their development. Following inoculation, it outcompetes
toxigenic strains of A. flavus, thereby mitigating aflatoxin levels. Bars represent means and error bars
represent SD. **** p < 0.0001.

3.4 Materials & Methods

3.4.1 General methods

All commercially available chemicals were obtained from TCI America except for cobalt
nitrate and manganese nitrate, which were purchased from Alfa Aesar, and BA, which was
purchased from Sigma Aldrich. Acetic acid, hexane, and chloroform were obtained from EMD
Millipore. The NMR solvents chloroform-d and methanol-d 4 were purchased from Cambridge
79

Isotopes Laboratories. All solvents and metal salts were used as received, without any further
purification.
All polystyrene (PS) waste including the styrofoam cold box, styrofoam plate, coffee lid, and
red solo cup were collected from the waste stream. The mixed plastic waste samples were collected
from Catalina Harbor at Santa Catalina Island, CA. All plastic waste samples were roughly cleaned
with acetone, dried and then shredded using a benchtop coffee grinder before use.

1
H NMR spectra were obtained by Varian VNMRS 600 or 400MR spectrometers and
processed via MestreLab Mnova. All the chemical shifts are shown by the units of ppm and
referenced to the residual
1
H solvent peak; and line-listed according to (s) singlet, (d) double, (t)
triplet, etc.
3.4.2 General procedure for catalyst screening

In a 300 mL Parr reactor, ground styrofoam PS powder (5 g) was mixed with NHPI (0.5 g),
metal catalysts (0.5 g total) and acetic acid (75 mL). The reactor was then sealed and pressurized to
four bars with molecular O 2 and stirred at 150
o
C for four hours. The reactor was then cooled to
room temperature and unsealed to release remaining O 2. Any volatiles, including acetic acid, were
removed by rotary evaporation and the resulting product mixture was resuspended in 1 M NaOH
(ca. 80-100 mL). Insoluble particles were removed by vacuum filtration. The resulting solution was
acidified to pH 1 with concentrated HCl and extracted with ethyl acetate (EA) (3 x 150 mL). The
combined organic fractions were dried over Na 2SO 4 and the solvent was removed by rotary
evaporation to afford a dark yellow solid product mixture. The yield of BA was determined by
1
H
NMR with 1,3,5-trimethoxybenzene as an internal standard.
3.4.3 General procedure for condition optimization

In a 300 mL Parr reactor, ground styrofoam PS powder (5 g) was mixed with cobalt nitrate
(0.25 g), manganese nitrate (0.25 g), and acetic acid (75 mL). The reactor was pressurized with four
80

bars O 2 and stirred at 150
o
C for the specified period (see Table 1, entries 5-10). The reactor was
then removed from the heat and cooled to room temperature (30 - 60 min). After recharging with
O 2, the reactor was again heated to continue the reaction. After time periods specified in Table 1, the
Parr reactor was cooled to room temperature and the internal pressure was released. The obtained
crude mixture was processed using the same procedure described above to yield a dark yellowish
solid mixture.
3.4.4 Digest purification

The obtained PS degradation mixture was resuspended in 50 mL chloroform in a 100 mL
round-bottom flask, stirred (50 - 60 min) and then filtered. The chloroform was removed in vacuo to
generate a light brown powder, which was washed with hot hexanes (ca. 50 °C, 3 x 50 mL) over a
filter. The hexane extract was concentrated in vacuo to obtain a chalky powder.
To recrystallize BA, the hexane extract was dissolved in ca. 100 mL boiling water and passed
through a fritted filter funnel. The flow-through was collected into a 250 mL Erlenmeyer flask
placed in an ice bath and allowed to cool (60 min). The contents of the flask were passed through a
clean filter funnel, and the obtained residue was flushed with ca. 100 mL cold water. The obtained
residue was dried over a filter paper to yield a crystalline white powder.
3.4.5 Media & buffer recipes

All recipes are based on MM: 12.0 g L
−1
NaNO 3, 3.04 g L
−1
KH 2PO 4, 1.04 g L
−1
KCl, 1.04 g
L
−1
MgSO 4·7H 2O, and 1 mL L
−1
Hutner’s trace element solution (60). GMM is MM supplemented
with 10 g L
-1
d-glucose. PSMM is MM supplemented with 10 g L
-1
purified PS digest. Solid plates
follow the same recipes as above with the addition of 15 g L
-1
agar. All media were adjusted to pH
8.0 using 5.5 M KOH. ST buffer: 8.5 g L
-1
NaCl, 1 mL L
-1
Tween 80.
3.4.6 Asperbenzaldehyde production from BA

81

In order to confirm if fungal strains can utilize BA as a sole carbon source to generate SMs,
ca. 3.0 x 10
7
spores of LO10050 were inoculated, in triplicate, into 125 mL Erlenmeyer flasks
containing 30 mL MM supplemented with increasing concentrations (2.5, 5.0, 7.5, 10.0, 12.5, and
15.0 g L
-1
) of a BA standard. Cultures were incubated for six days at 37 °C with shaking at 180 rpm.
To lyse mycelia to release intracellular asperbenzaldehyde, 30 mL MeOH was added to each culture
flask, which were sonicated for one hour. 10 µL aliquots of LO10050 extracts were then analyzed via
HPLC-DAD-MS. Extracted ion chromatograms corresponding to asperbenzaldehyde were
measured for each condition.
3.4.7 Fungal metabolism of PS digestion products

To determine if other products of the PS digestion reaction apart from BA are suitable for
fungal metabolism, 3.0 x 10
7
spores of LO10050 were inoculated into a 125 mL Erlenmeyer flask
containing 30 mL PSMM. Cultures were incubated for six days at 37 °C with shaking at 180 rpm.
200 µL aliquots of the culture medium were collected daily throughout a six-day incubation period.
These aliquots were extracted with ca. 3 mL EA, dried, and resuspended in 4:1 MeOH:DMSO.
Extracted ion chromatograms corresponding to each compound of interest were measured for each
condition (Fig. S5B).
3.4.8 SM extraction and quantification

For ergothioneine quantification, cultures of YM847 were heated in a 100 °C water bath for 20
minutes to lyse mycelia containing intracellular ergothioneine. 10 µL of this lysate was analyzed with
HPLC-DAD without dilution. Extracts were analyzed with a Venusil HILIC column (4.6 x 250 mm,
5 µm). The solvent used was 4:1 AcN:20 mmol L
-1
ammonium acetate (pH 6) at a flow rate of 1 mL
min
-1
.
For pleuromutilin and mutilin quantification, mycelia were filtered from the culture media
and sonicated in 1:1 DCM:MeOH for one hour. The organic extracts were then filtered and dried
82

(TurboVap LV). Once dried, extracts were resuspended in 20 mL ddH 2O and extracted three times
with 20 mL EA. Organic extracts were then dried as above. Culture media (ca. 25 mL) were
extracted three times with 25 mL EA. Organic extracts from the culture media were combined with
those from the mycelia and dried. Extracts were resuspended in 10 mL 4:1 MeOH:DMSO and 10
µL of this extract was injected for HPLC-DAD-MS analysis.
3.4.9 Compound purification & characterization

For ergothioneine purification, ca. 1.0 x 10
7
spores of YM847 were inoculated into three 25
mL Erlenmeyer flasks, each containing 10 mL PSMM. Culture and induction parameters were the
same as described above. To lyse mycelia, 10 mL MeOH was added to culture flasks, which were
then sonicated for one hour. Culture media were then combined and filtered to remove fungal
biomass. Culture media were concentrated in vacuo and subjected to reverse-phase column
chromatography with gradient elution. The mobile phases used were 100% AcN followed by 9:1,
17:3, 4:1, and 7:3 AcN:ddH 2O. Fractions containing ergothioneine were combined, dried in vacuo,
and subjected to final purification with reverse-phase preparative thin layer chromatography using
17:3 AcN:ddH 2O as a mobile phase. A band corresponding to ergothioneine was excised and
flushed with ddH 2O over a fritted filter funnel. The aqueous extract was concentrated in vacuo to
yield ergothioneine.
For pleuromutilin and mutilin purification, ca. 3.0 x 10
7
spores of YM343 and YM283,
respectively, were inoculated into five 125 mL Erlenmeyer flasks, each containing 30 mL PSMM.
Culture and induction parameters were the same as described above. Following incubation, mycelia
were filtered from the culture media. Culture media were extracted three times with 150 mL DCM
and organic extracts were dried in vacuo. Extracts were then subject to purification by normal-phase
column chromatography using the following mobile phase gradient: 9:1, 7:3, 1:1, 3:7, 1:9. Fractions
containing pleuromutilin or mutilin were combined and subjected to final purification using normal-
83

phase preparative thin layer chromatography (TLC). The mobile phases used were 1:1 for
pleuromutilin and 3:7 EA:hexanes for mutilin. Bands corresponding to pleuromutilin or mutilin
were excised and flushed with DCM over a fritted filter funnel. Organic extracts were concentrated
in vacuo to yield pleuromutilin and mutilin.
84

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Natural Products. ACS Synthetic Biology, 10(1), 173–182.
https://doi.org/10.1021/acssynbio.0c00536

59. Bailey, A. M., Alberti, F., Kilaru, S., Collins, C. M., de Mattos-Shipley, K., Hartley, A. J.,
Hayes, P., Griffin, A., Lazarus, C. M., Cox, R. J., Willis, C. L., O’Dwyer, K., Spence, D. W.,
& Foster, G. D. (2016). Identification and manipulation of the pleuromutilin gene cluster
from Clitopilus passeckerianus for increased rapid antibiotic production. Scientific Reports, 6(1),
25202. https://doi.org/10.1038/srep25202

60. Hutner, S. H., Provosoli, L., Schatz, A., & Haskins, C. P. (1950). Some approaches to the
study of the role of metals in the metabolism of microorganisms. Proceedings American Phil
Society, 94(2), 152–170.

61. Szewczyk, E., Nayak, T., Oakley, C. E., Edgerton, H., Xiong, Y., Taheri-Talesh, N., Osmani,
S. A., & Oakley, B. R. (2006). Fusion PCR and gene targeting in Aspergillus nidulans. Nature
Protocols, 1(6), 3111–3120. https://doi.org/10.1038/nprot.2006.405
90

3.6 Supplementary figures

Figure S1.
1
H NMR spectrum of chloroform-extracted BA derived from PS. The sample was
dissolved in CDCl 3.
91

Figure S2. Fungal metabolism and toxicity of BA and phthalic acid. 1.0 x 10
6
spores of A. nidulans
FGSC A4 were inoculated into 6-well plates containing 10 mL of (A) GMM, (B) GMM + 10 g L
-1

BA, (C) GMM + 10 g L
-1
phthalic acid, (D) MM, (E) MM + 10 g L
-1
BA, (F) MM + 10 g L
-1
phthalic
acid. Cultures were incubated at 37 °C for seven days.

92

Figure S3. Asperbenzaldehyde production in GMM compared to MM supplemented with
increasing concentrations of BA. 3.0 x 10
7
spores of LO10050 were inoculated into 125 mL
Erlenmeyer flasks containing 30 mL of media. Each condition was cultured in triplicate. Cultures
were incubated for six days at 37 °C with shaking at 180 rpm. Following incubation, 30 mL MeOH
was added to each culture flask, which were then sonicated for one hour. 10 µL aliquots of extracts
were then analyzed via HPLC-DAD-MS. Extracted ion chromatograms corresponding to
asperbenzaldehyde were measured for each condition.

93

Figure S4. Standard curve of ergothioneine generated via HPLC-DAD.

94

Figure S5. Standard curve of pleuromutilin generated via HPLC-DAD-MS.

95

Figure S6. Standard curve of mutilin generated via HPLC-DAD-MS.

96

Figure S7.
1
H NMR spectrum of ergothioneine. Solvent = D 2O.

97

Figure S8.
1
H NMR spectrum of pleuromutilin. Solvent = chloroform-d.

98

Figure S9.
1
H NMR spectrum of mutilin. Solvent = methanol-d 4.

99

3.7 Supplementary tables

Table S1. Screening of different catalytic conditions for PS degradation (PS source: waste styrofoam
cold box). (a) In a 300-mL Parr reactor, a mixture of PS (5 g), metal catalyst (10 wt%), N-
Hydroxyphthalimide (NHPI, 10 wt%) and acetic acid (75 mL) was stirred with molecular oxygen (4
bars, with active refilling) at 150
o
C (b) Equivalents of O 2 per monomer unit in PS (c) Either 0.5 g of
one catalyst or 0.25 g of two catalysts (d) Mass recovery yield (wt%) = [(mass of benzoic acid
products)/(mass of starting PS)] x 100% (e) molar yield (%) = [(mass of carbon in BA
products)/(mass of carbon in starting PS)] x 100%.

Entry
a

O2
Aliquots
O2
Consumed
Equiv
b

Time
(hrs)
Metal
Catalyst
c

Mass
Recovery
(wt%)
d

Molar
Yield
(%)
e

1 1 0.75 4 Mn(acac)3 3.4 2.8
2 1 0.67 4 Mn(acac)2 4.5 3.8
3 1 0.67 4 Mn(NO3)2 14.1 12.0
4 1 0.75 4 Co(NO3)2 6.0 5.1
5 1 0.75 4 Co(NO3)2 + Mn(NO3)2 30.7 26.2
6 2 1.50 8 Co(NO3)2 + Mn(NO3)2 54.6 46.5
7 3 2.06 12 Co(NO3)2 + Mn(NO3)2 84.1 71.4
8 1 0.75 3 Co(NO3)2 + Mn(NO3)2 19.2 16.3
9 2 1.50 6 Co(NO3)2 + Mn(NO3)2 47.2 40.1
10 3 1.98 9 Co(NO3)2 + Mn(NO3)2 72.5 61.5

100

Table S2. The production of benzoic acid from the degradation of various post-consumer PS waste
sources under optimized conditions (entry 7 in table S1). (a) Styrofoam cold box; (b) Styrofoam
plate; (c) Catalina Island waste; (d) Coffee lid; (e) Red drink cup.
Entry
O2
Aliquots
O2
Consumed
Equiv.
Time
(hrs)
Mass
Recovery (wt%)
Molar
Yield (%)
1
a
3 2.06 12 84.1 71.4
2
b
3 1.80 12 60.2 51.1
3
c
3 1.89 12 45.7 38.8
4
d
2 1.23 8 17.6 15.0
5
e
2 1.32 8 25.2 21.4

101

Table S3.
1
H NMR chemical shifts of ergothioneine compared to literature values
59
. Solvent = D 2O.
δH
(literature)
(ppm)
Splitting
pattern
J (Hz)
δH (experimental)
(ppm)
Splitting
pattern
J (Hz)
ΔδH
(ppm)
3.10 m - 3.10 m - 0.00
3.19 s - 3.19 s - 0.00
3.83 dd
4.6,
11.0
3.81 dd
3.6,
11.8
0.02
6.70 s - 6.72 s - 0.02

102

Table S4.
1
H NMR chemical shifts of pleuromutilin compared to literature values
57
. Solvent =
chloroform-d.
δH
(literature)
(ppm)
Splitting
pattern
J (Hz)
δH (experimental)
(ppm)
Splitting
pattern
J (Hz)
ΔδH
(ppm)
1.41 - 1.52 m - 1.42 - 1.52 m - 0.00
1.61 - 1.73 m - 1.61 - 1.72 m -
0.00 -
0.01
2.16 - 2.30 m - 2.16 - 2.30 m - 0.00
2.16 - 2.30 m - 2.16 - 2.30 m - 0.00
2.11 s - 2.11 s - 0.00
1.61 - 1.73 m - 1.63 - 1.70 m -
0.02 -
0.03
1.55 dd 13.8, 2.7 1.57 m - 0.02
1.40 ddd
13.8, 6.0,
2.7
1.39 m - 0.01
1.79 dq 14.5, 3.1 1.78 dq 14.4, 3.2 0.01
1.15 td 14.3, 4.4 1.14 td 14.2, 4.5 0.01
2.29 - 2.40 m - 2.31 - 2.37 m -
0.02 -
0.03
3.34 dd 10.8, 6.6 3.37 d 6.6 0.03
2.10 dd 16.0, 8.7 2.10 dd 16.0, 8.7 0.00
1.33 d 16.0 1.33 d 16.1 0.00
5.85 d 8.6 5.84 d 8.6 0.01
1.44 s - 1.44 s - 0.00
0.71 d 7.1 0.71 d 7.1 0.00
0.90 d 7.1 0.90 d 7.1 0.00
1.19 s - 1.18 s - 0.01
6.50 dd 17.4, 11.0 6.50 dd
17.4,
11.0
0.00
5.37 dd 11.0, 1.3 5.37 dd 11.0, 1.5 0.00
5.22 dd 17.4, 1.4 5.22 dd 17.4, 1.5 0.00
103

4.05 qd 17.1, 5.4 4.04 q 17.0, 5.5 0.01

104

Table S5.
1
H NMR chemical shifts of mutilin compared to literature values
57
. Solvent = methanol-
d 4.
δH
(literature)
(ppm)
Splitting
pattern
J (Hz)
δH (experimental)
(ppm)
Splitting
pattern
J (Hz)
ΔδH
(ppm)
1.27 - 1.70 m - 1.29 - 1.68 m - 0.02
1.07 - 1.16 m - 1.07 - 1.15 m -
0.00 -
0.01
2.06 - 2.27 m - 2.08 - 2.26 m -
0.02 -
0.01
2.20 s - 2.21 s - 0.01
1.72 - 1.78 m - 1.73 - 1.79 dq 14.6, 3.1 0.01
1.27 - 1.70 m - 1.29 - 1.68 m - 0.02
1.07 - 1.16 m - 1.07 - 1.15 m -
0.00 -
0.01
1.27 - 1.70 m - 1.29 - 1.68 m - 0.02
1.07 - 1.16 m - 1.07 - 1.15 m -
0.00 -
0.01
2.14 m - 2.17 m - 0.03
3.44 d 6.1 3.44 d 6.2 0.00
1.95 d 15.8, 7.6 1.95 dd 15.8, 7.7 0.00
1.64 d 15.8 1.64 d 15.7 0.00
4.27 d 7.6 4.27 d 7.7 0.00
1.33 s - 1.33 s - 0.00
0.90 d 7.3 0.90 d 7.2 0.00
0.91 d 7.0 0.92 d 7.0 0.01
1.10 s - 1.10 s - 0.00
6.15 dd
17.8,
11.2
6.15 dd
18.0,
11.3
0.00
5.34 dd 17.8, 1.4 5.32 dd 18.0, 1.5 0.02
5.20 dd 11.2, 1.4 5.20 dd 11.2, 1.5 0.00

105

Chapter 4: Transcription Factor Engineering Leads to Discovery of a
Novel Orsellinaldehyde Derivative in Aspergillus nidulans

Authors: Chris Rabot
,
Michelle F. Grau, Ruth Entwistle, Yi-Ming Chiang, Yamilex
Zamora Roberts, Manmeet Ahuja, C. Elizabeth Oakley, Clay C. C. Wang, Richard B.
Todd, & Berl R. Oakley

4.1 Abstract

Fungal secondary metabolites are valuable and abundant sources of medically and
industrially relevant compounds. Genome sequencing efforts have revealed that transcription factors
are frequently colocalized within biosynthetic gene clusters. Overexpression or deletion of these
transcription factors represents a facile approach to simultaneously activate all genes within a gene
cluster, leading to production of its downstream metabolite(s). Unfortunately, the modulation of
cluster-specific transcription factor expression is not always a successful activation approach. Herein,
we attempted to activate a cryptic gene cluster in Aspergillus nidulans containing a highly-reducing
polyketide synthase using a combination of transcription factor engineering and overexpression.
This approach resulted in unexpected, in trans polyketide biosynthesis and discovery of a novel
polyketide we term triorsellinaldehyde. Targeted deletion of a non-reducing polyketide synthase
elsewhere in the genome confirmed its role in its biosynthesis.
4.2 Introduction

Secondary metabolites (SMs) are structurally diverse, low-molecular weight organic
compounds produced by bacteria, plants, and fungi. In contrast to primary metabolites, SMs are not
directly required for the growth, survival, or reproduction of the organisms that produce them
1
.
Rather, they typically provide selective advantages to the organism that produces them. Their
structural and stereochemical complexities frequently confer diverse and potent bioactivities, making
them remarkably useful in medicine and agriculture
2,3
. Notable examples of SMs include the
106

chemotherapeutic doxorubicin, the anticholesterolemic lovastatin, and the immunosuppressant
cyclosporine.
A hallmark of fungal genetics is that the genes that encode production of proteins involved
in SM biosynthesis are usually clustered within the genome
4
. Recent genome sequencing studies have
revealed that the number of these biosynthetic gene clusters (BGCs) vastly exceed the number of
known SMs
5,6,7
, suggesting that many more SMs remain to be discovered. Unfortunately, most
putative BGCs are silent under standard laboratory conditions
8
. A number of strategies have been
developed in an effort to activate these BGCs, including, but not limited to: (1) the “one strain many
compounds” (OSMAC) approach
9
,
10
, (2) epigenetic modulation
11,12
, (3) co-culturing methods
13,14
, (4)
heterologous expression
15,16
, and (5) transcriptional regulator overexpression
17
or deletion
18
. While
each of these approaches have shown moderate success in natural product discovery efforts, no one
approach is universally successful. New approaches toward the activation of cryptic BGCs are still
needed.
In order to expand the catalog of techniques that can be used to activate silent BGCs, we
sought to exploit the occasional presence of multiple transcription factors (TFs) located within
BGCs
4,19,20
. Modulation of cluster-specific TF expression has been shown in many cases to activate
expression of cryptic BGCs
21,22,23
. However, this approach is not universally successful. Previously,
we attempted to activate 18 silent BGCs in A. nidulans by placing cluster-specific TFs or related
biosynthetic genes under control of the inducible alcA promoter (alcA(p))
24
. Surprisingly, only three
of these cases resulted in the production of adequate levels of SMs to permit isolation and
characterization. Other cases resulted either in the production of SMs in levels too low to permit
characterization, or no product at all.
This approach may have only been partly successful for several reasons. First, TFs often
must homo- or heterodimerize in order to function fully
25
. Furthermore, it is possible that some TFs
107

were incorrectly annotated, such that the promoter would not have properly driven expression, or
would have driven expression of only a portion of the TF
26
. Even if the annotation were correct, the
TF activation domain (AD), which is responsible for recruitment of transcriptional machinery to
target loci
27
, may have been insufficiently active to drive expression of the BGC to high levels. The
TF may also have lacked a natural inducer that would have driven strong activation. Finally, some
TFs require extensive post-translational modifications (PTMs), which, if not applied to the TFs of
interest, would not permit proper activation and subsequent expression of the BGC
28
.
We aimed to address several of the potential limitations described above by employing
simultaneous TF overexpression and engineering strategies in an attempt to activate a putative HR-
PKS BGC in A. nidulans. This putative BGC contains two putative Zn(II) 2Cys 6 TFs (AN6788 and
AN6790) colocalized with an HR-PKS (AN6791). We exploited the modularity of TFs
29
by creating
a hybrid AN6788 TF, in which its native AD has been replaced by that of AfoA, the TF that
governs expression of the asperfuranone BGC
30
. We recently utilized this approach to activate the
silent BGC responsible for (+)-asperlin production in A. nidulans
26
. We reasoned that the native AD
of AN6788 may be insufficient in its ability to recruit transcriptional machinery to the genes that it
regulates. Replacement of this AD with that of the highly efficient AfoA should result in a TF that is
highly active. As TF dimerization often occurs through the DNA-binding domain (DNA-BD)
31
, this
approach should not interrupt heterodimerization between AN6788 and AN6790, a second TF
colocalized within the BGC.
Considering that some TFs must dimerize to activate target loci
27
, we reasoned that the
simultaneous activation of both AN6788 and AN6791 may be required to activate the AN6791
BGC. To explore this approach, we also replaced the native promoters of AN6788 and AN6790
either separately or together with alcA(p), a promoter that is strongly activated in the presence of
ethanol or related substrates
32
. To increase the expression of TFs under control of alcA(p), we also
108

replaced the promoter of alcR with the constitutive gpdA promoter (gpdA(p)). The alcR gene encodes
a TF that positively regulates alcA(p) expression
33,34
. Taken together, we generated a hybrid AN6788
TF and overexpressed it together with AN6790 to determine if this approach can activate the
putative AN6791 BGC.
Metabolic profiling of strains that overexpress a hybrid AN6788 alone revealed production
of orsellinaldehyde (1) and 3-(2,4-dihydroxy-6-methylbenzyl)-orsellinaldehyde (2), hereafter termed
diorsellinaldehyde. Interestingly, 1 and 2 were previously found to be produced upon replacement of
the promoter of pkfA, a NR-PKS located on chromosome VI
24
. In contrast, sole overexpression of
the native AN6790 did not result in production of any metabolites detectable via HPLC-DAD-MS.
However, simultaneous activation of both a hybrid AN6788 and a native AN6790 resulted in
production of the novel polyketide that we term triorsellinaldehyde (3) in addition to 1 and 2
(Scheme 1). As 3 appears to be structurally related to 1 and 2, we hypothesized that its biosynthesis
is attributable to the pkf BGC rather than a gene or genes within the putative AN6791 BGC.
Targeted deletion of AN3230 in the hybrid AN6788-native AN6790 overexpression
background eliminated production of 1, 2, and 3, confirming its role in biosynthesis of these SMs. In
contrast, deletion of the AN6791 HR-PKS in the same background had no effect on the production
of these SMs. Taken together, these results suggest that AN6788 can, alone or as a heterodimer with
AN6790, activate in trans the NR-PKS AN3230. This approach may be successful in the activation
of other cryptic BGCs which harbor multiple TFs.
109

Scheme 1: Overall strategy and results of this study. The putative AN6791 BGC, located on
chromosome I, harbors two putative Zn(II) 2Cys 6 TFs (AN6788 and AN6790). We fused the native
AD of AN6788 with that of AfoA to create a hybrid TF. We also replaced the native promoters of
both TFs with the inducible alcA(p). Metabolic profiling of this strain revealed the production of 1, 2,
and 3. Targeted deletions revealed these compounds do not originate from AN6791. Rather, they
are synthesized by one or more genes within the pkf BGC located on chromosome VI.

4.3 Results & Discussion

4.3.1 Hybrid TF rationale

Previously, our group attempted to activate the A. nidulans AN6791 BGC, encoding a
putative highly reducing PKS, by placing the putative TF AN6788 under control of alcA(p)
24
. Details
of the genes in the putative AN6791 BGC are listed in Table 1. This approach was not sufficient to
promote enough SM production to permit purification and characterization. We reasoned that
AN6788, even if expressed to high levels through alcA(p) induction, may not harbor an AD
sufficient to strongly drive expression of its regulon. Thus, maintenance of the native AN6788
DNA-BD to retain target specificity, but replacement of its AD with one that is highly active, should
generate a hybrid TF that can drive expression of its target loci to appreciable levels.
110

Table 1. Gene names, lengths, protein product lengths, and predicted functions of genes
adjacent to the AN6791 PKS. Gene names are based on designations from FungiDB.

Gene name Gene length (bp) Protein length (residues) Predicted function
AN6785 1906 476 Hypothetical protein
AN11527 339 88 Hypothetical protein
AN6786 1062 244 Beta-1,4-endoglucanase
AN6787 2003 504 Cytochrome P450
AN6788 2221 612 Zn(II) 2Cys 6 transcription factor
AN6789 1670 351 Hypothetical protein
AN6790 2222 609 Zn(II) 2Cys 6 transcription factor
AN11907 943 153 Hypothetical protein
AN6791 7977 2568 Polyketide synthase
AN6792 1438 407 Glycerol-3-phosphate dehydrogenase
AN6793 1283 370 Hydrolase
AN6794 1626 446 DMATS-type prenyltransferase

Selection of a candidate AD to fuse to a native DNA-BD of a TF requires consideration of
several factors. First, the hybrid TF must not be susceptible to post-translational inactivation to any
significant extent. Second, it should not require an unknown inducer in order to be activated. Lastly,
it should be able to function during late stationary phase, when SM, and particularly polyketide,
biosynthesis can be driven to high levels
35,36
.
AfoA is a transcription factor that regulates the asperfuranone gene cluster
30
. Previously, we
showed that upregulation of afoA permits the isolation of very high levels (>2 g L
-1
) of purified
asperbenzaldehyde, an intermediate in asperfuranone biosynthesis
37
. To our knowledge, it is not
111

inhibited by PTMs to any significant extent, and it is able to drive expression to high levels in late
stationary phase growth. We recently utilized the AfoA AD to generate a hybrid TF to successfully
activate the (+)-asperlin BGC
26
. For these reasons, the AfoA AD was selected as a candidate to fuse
to the native AN6788 DNA-BD in an attempt to activate the BGC and characterize its product.
AfoA contains a Zn(II) 2Cys 6 DNA-binding motif. The DNA-BD of this type of TFs is
comprised of six cysteine residues that coordinate two Zn(II) ions to form a cluster, which is
followed by amino acid residues that provide specificity for the DNA-binding site and usually a
coiled-coil domain that mediates dimerization
38,39
. These TFs bind symmetrical DNA-binding sites
that are comprised of inverted, directly repeated, or everted triplet amino acid residues that are
separated by several nucleotides. The specificity region and dimerization motif generally determine
the spacing between the triplet residues.
4.3.2 Generation of a strain expressing a hybrid AN6788 TF

To generate a hybrid AN6788 TF, we first used InterProScan to predict that the DNA-
binding motif of AfoA is located at residues 16-43
40
(Fig. S1). Next, we used PCOILS to locate two
predicted coiled-coil dimerization motifs at residues 55-80 and 98-129
41
(Fig. S2). Finally, PSORT II
was used to locate a putative nuclear localization signal (NLS) of the SV40 Large T-antigen-type at
residues 236-242
42,43
. Previous work verified the boundaries of the AfoA AD by fusing residues 130-
666 of AfoA to the DNA-BD of the FacB TF (26). This hybrid TF was expressed in strains lacking
native facB; the gene product of which permits utilization of acetate as a sole carbon source. The
hybrid AfoA-FacB TF activated acetate utilization genes, confirming that the AfoA AD can
functionally be utilized as a candidate for a hybrid TF.
4.3.3 Strain construction

In order to activate the silent BGC containing AN6791, we fused the predicted DNA-BD of
AN6788 (residues 1-200, as annotated by FungiDB) to the AfoA AD (residues 130-666). This
112

hybrid TF was placed under control of alcA(p) and inserted into the yA locus using the A. terreus
pyrG selectable marker. The recipient strain, LO9577, is derived from the parent strain LO8030,
which contains deletions of eight of the most highly-expressed SM BGCs
44
. This serves two
purposes; first, it minimizes the SM background, facilitating detection of new SMs. Second, it
theoretically should free up SM precursor molecules that can then feed into other biosynthetic
pathways. LO9577 also carries a replacement of the yA gene with a fragment containing the A.
fumigatus pyroA gene (AfpyroA) and the aldA promoter (aldA(p)), which drives expression of afoA.
The resulting strain LO10141 thus contained a hybrid TF, comprised of the AfoA AD and
the DNA-BD of AN6788, that was placed under control of alcA(p). Finally, we placed the alcR gene
under the control of the strong constitutive gpdA promoter (gpdA(p)). The alcR gene encodes the
transcriptional activator AlcR, which, in the presence of an inducer, binds to alcA(p) to strongly
drive its expression
45
. To do this, a transforming fragment was generated which contained the A.
fumigatus riboB selectable marker (AfriboB) fused to alcR, which was placed under control of gpdA(p).
This construct was transformed into recipient strain LO10141 to generate LO10240. Taken together,
these genetic manipulations permitted the construction of a strain that robustly expresses a hybrid
AN6788 TF (Fig. 1).
113

Figure 1. Strategy to generate a strain of A. nidulans that expresses a hybrid AN6788 TF to
promote SM biosynthesis. (A) The organization of the AN6791 (left) and the afo (right) BGCs in
A. nidulans. Relevant TFs are labeled. (B) The DNA-binding domain of the AN6788 transcription
factor (translated to residues 1-200, solid line) and the afoA AD (translated to residues 130-666,
dotted line). (C) The AN6788 DNA-BD, the afoA AD, and the inducible alcA(p) were fused via
fusion PCR and transformed into a parent strain of A. nidulans. The fusion PCR construct also
contained the A. terreus AfpyrG selectable marker (not shown). Upon induction, the hybrid TF binds
to the promoters of target loci and recruits transcriptional machinery to initiate transcription and
subsequent SM biosynthesis.

4.3.4 Subsequent strain engineering

We also generated a set of strains by the same methodology that expresses either native or
hybrid forms of the AN6790 TF by the same methodology described above. Finally, we constructed
a strain that places both a hybrid AN6788 and native AN6790 under control of alcA(p). To do this,
a transforming fragment was generated which contained the A. fumigatus pyroA selectable marker as
well as AN6790 places under control of alcA(p). This construct was transformed into recipient strain
LO10240 to generate LO11940.
Finally, we generated two sets of strains in the LO11940 background that contained
deletions of the AN6791 HR-PKS and the AN3230 NR-PKS to determine if they are responsible
114

for production of the detected SMs detailed above. To do this, a transforming fragment was
generated which contained the pyrithiamine (ptrA
+
)
46,47
resistance gene targeting either AN6791 or
AN3230. The resulting strains were named LO12110 (AN6791Δ) and LO12139 (AN3230Δ). A full
list of strains and corresponding genotypes used in this study is shown in Table 2.
Table 2. Names of strains and genotypes used in this study.

Strain name Genotype
LO8030
44

pyroA4, riboB2, pyrG89, nkuA::argB, stc(AN7804-AN7825)Δ, eas(AN2545-
AN2549)Δ, afo(AN1039-AN1029)Δ, mdp(AN10023-AN10021)Δ, tdi(AN8512-
AN8520)Δ, aus(AN8379-AN8384, AN9246-AN9259)Δ, ors(AN7906-AN7915)Δ,
apt(AN6000-AN6002)Δ
LO9577 yA::AfpyroA-aldA(p)afoA in LO8030
LO10141 AtpyrG-alcA(p)-AN6788(1−200)-afoA(130−666) in LO9577
LO10193 AtpyrG-alcA(p)-AN6790(1-200)-afoA(130−666) in LO9577
LO10240 AfriboB-gpdA(p)-alcR in LO10141
LO11940 Afpyro-alcA(p)-AN6790 in LO10240
LO11974 AfpyroA-alcA(p)-AN6790 in LO8030
LO12110 ptrA
+
-AN6791Δ in LO11940
LO12139 ptrA
+
-AN3230Δ in LO11940
LO8923 Afpyro-alcA(p)-AN6793; AfpyrG-alcA(p)-AN6791 in LO8030
LO11902 AtriboB-gpdA(p)-alcR in LO8923

4.3.5 Single TF activation

We first attempted to activate the AN6791 BGC by placing individual TFs within the cluster
under control of alcA(p). Upon induction of strain LO10240, which expresses a hybrid AN6788
under control of alcA(p) along with alcR under control of gpdA(p), we detected two major induced
compounds via HPLC-DAD-MS. Following purification and characterization, we identified these
compounds as 1 and the related 2. 1 is an important aromatic polyketide precursor that functions as
115

an anti-inflammatory agent that also induces apoptosis
48,49
. 2 resembles a dimer of 1 that was
previously isolated by placing the A. nidulans NR-PKS AN3230 under control of alcA(p)
24
. In
contrast, strain LO11974, which expresses a native AN6790 under control of alcA(p), did not
produce any compounds detectable via HPLC-DAD-MS upon alcA(p) induction. We also note that
we could not detect 1 or 2 upon induction of LO11974, which expressed a hybrid AN6790 (data not
shown).
4.3.6 Dual TF activation

We next overexpressed both a hybrid AN6788 and a native AN6790 simultaneously by
placing them both under control of alcA(p). Upon alcA(p) induction, we again detected both 1 and 2
as major products. Interestingly, the yields of both compounds were higher upon dual TF activation;
negative-mode extracted ion chromatogram AUCs revealed that the titers of 1 and 2 were increased
ca. 5.4- and 12.0-fold, respectively, in strain LO11940 relative to LO10236. We also detected a third
induced product 3 with m/z 424. Importantly, we observed that this compound was exclusively
produced when both TFs were overexpressed (Fig. 2). Detailed 1D and 2D NMR analysis revealed
that 3 resembles a trimer of 1. For NMR spectra (Figs. S3-S14), chemical shift data (Table S1), and
mass data (Figs. S15-17) see Supporting Information.

116

Figure 2. Negative-mode extracted ion chromatograms (m/z = 150-152, 286-288, 422-424)
showing induced compounds upon overexpression of TFs adjacent to the AN6791 PKS. The
promoters of the hybrid AN6788 or native AN6790 TFs, separately or together, were replaced with
the alcA(p) promoter. Strains were screened in 125 mL Erlenmeyer flasks containing 30 mL of
lactose minimal media (LMM; see below for recipe). Cultures were incubated at 37 °C with shaking
at 180 rpm. For alcA(p) induction, 50 mM methyl ethyl ketone (MEK) was added after 42 hours.
Strains were then incubated for another 72 hours, at which point culture media were extracted with
ethyl acetate (EA), dried, and analyzed via HPLC-DAD-MS. All y-axes are normalized to the same
intensities.

As mentioned above, 1 and 2 are known products of the pkf BGC. In order to determine if
these induced SMs, along with 3, are the products of the pkf BGC or the AN6791 PKS colocalized
with the TFs that were overexpressed, we generated deletions of each PKS in the LO11940
background. As expected, deletion of AN3230 abolished the production of 1, 2, and 3. In contrast,
the deletion of AN6791 had not effect on the production of these SMs. Taken together, these
results indicate that the combination of TF engineering and overexpression can lead to the
production of novel, albeit unexpected SMs. Our understanding of TFs that function in trans is still
weak; a more thorough understanding of these TFs should permit the discovery of new SMs, even
from BGCs for which the final products are thought to be known.
117

4.3.7 Direct activation of AN6791

Our results regarding overexpression of the TFs adjacent to the AN6791 PKS indicate that
they likely do not regulate its expression. It is possible that AN6791, together with another BGC
such as the pkf BGC, coordinates the production of products yet to be discovered. In order to
determine, at a minimum, the direct product of AN6791, we replaced its native promoter with
alcA(p). Metabolic profiling of the AN6791 overexpression strain revealed induced metabolites with
m/z = 284 and 258 (Figs. S16-17). Purification and characterization of these metabolites are
ongoing.
4.3.8 Future prospects regarding activation of cryptic A. nidulans BGCs

This study has revealed several useful findings regarding the regulation of SM biosynthesis in
filamentous fungi. First, TF engineering may be a successful approach toward BGC activation,
especially in cases in which overexpression of native TFs permitted insufficient SM production.
Further, the combination of TF engineering with simultaneous overexpression of colocalized TFs
may promote the production of novel, although potentially unexpected SMs.
We also note that the overexpression of alcR can markedly increase SM titers, provided
biosynthetic genes are placed under control of alcA(p). In five of the 18 attempts we made to
activate BGCs, purification and characterization of resulting SMs were limited by low fermentation
titers. The approaches described above may prove useful in future attempts to identify these elusive
SMs as well as those from other species.
4.4 Materials & Methods

4.4.1 Molecular Genetic Manipulations

Transforming constructs were generated by fusion PCR as previously described
50,51
. All
constructs were verified by diagnostic PCR. The selectable markers used were the A. fumigatus riboB
gene (AfriboB), the A. fumigatus pyroA gene (AfpyroA), the A. terreus pyrG gene (AtpyrG), and the A.
118

oryzae pyrithiamine resistance gene (ptrAI
+
). Transformation procedures were conducted as
previously described
50
.
4.4.2 Fermentation and HPLC-DAD-MS Analysis

For initial screening of strains, 3.0 x 10
7
spores of each strain were inoculated into 125 mL
Erlenmeyer flasks containing 30 mL lactose minimal media (15.0 g L
-1
lactose, 0.31 g L
-1
KOH, 6.0 g
L
-1
NaNO 3, 0.52 g L
-1
KCl, 0.52 g L
-1
MgSO 4*7 H 2O, 1.52 g L
-1
KH 2PO 4, and 1.0 mL L
-1
Hutner’s
trace element solution
52
). When needed, media was supplemented with 0.5 mg L
−1
pyridoxine, 2.5
mg L
−1
riboflavin, uracil (1 g L
-1
), and/or uridine (10 mM). Cultures were incubated at 37 °C with
shaking at 180 rpm. For alcA(p) induction, 50 mM MEK was added after 42 hours. Following
induction, cultures were incubated for another 72 hours.
Following incubation, mycelia were filtered from the culture medium by vacuum filtration.
The culture medium was extracted with 20 mL EA. Organic extracts were isolated, dried (TurboVap
LV) and resuspended in 1 mL 4:1 MeOH:DMSO. 10 µL of the extract was analyzed via HPLC-
DAD-MS.
HPLC-DAD-MS spectra were obtained using a ThermoFinnigan LCQ Advantage ion trap
mass spectrometer with a reverse phase C 18 column (Alltech Prevail C 18; particle size, 3 μm; column,
2.1 by 100 mm) at a flow rate of 125 μL min
−1
. The solvent gradient was 95% MeCN–H 2O (solvent
B) in 5% MeCN–H 2O (solvent A), both of which contained 0.05% formic acid, as follows: 0%
solvent B from 0 to 5 min, 0 to 100% solvent B from 5 min to 35 min, 100% solvent B from 35 to
40 min, 100 to 0% solvent B from 40 to 45 min, and reequilibration with 0% solvent B from 45 to
50 min. Conditions for MS included a capillary voltage of 5.0 kV, a sheath gas flow rate at 60
arbitrary units, an auxiliary gas flow rate at 10 arbitrary units, and the ion transfer capillary
temperature at 350 °C.

119

4.4.3 Compound Purification

For isolation of 2, 5.0 x 10
8
spores of LO10236 were inoculated into seven 2 L Erlenmeyer
flasks containing 500 mL LMM supplemented with 0.5 mg L
-1
pyridoxine. Note that the titer of 1
was too low to permit purification from this strain; its later identification was facilitated by the
higher titers of LO11940. Cultures were incubated at 37 °C with shaking at 180 rpm. For alcA(p)
induction, 50 mM methyl ethyl ketone (MEK) was added after 42 hours. Following induction,
cultures were incubated for another 72 hours.
Following incubation, mycelia were filtered from the culture medium by vacuum filtration.
The culture medium was extracted three times with a volume of EA equal to the volume of the
medium. The EA extract was evaporated in vacuo (660 mg) and fractionated via isocratic normal-
phase column chromatography using 49:1 DCM:MeOH as a mobile phase. Fractions containing the
compound of interest as indicated by thin-layer chromatography (TLC) were combined and dried in
vacuo (35 mg). This subfraction was fully purified via HPLC-DAD (Agilent 1200) equipped with an
RP-18 column (Phenomenex® Luna 5 μm C 18, 250 x 10 mm) at a flow rate of 4.0 mL min
-1
. The
solvents used were 100% AcN (solvent B) and 100% H 2O (solvent A), both containing 0.05% TFA.
The solvent gradient used was: 0 to 100% solvent B from 0 to 19 min, 100% solvent B from 19 to
24 min, 100 to 0% solvent B from 24 to 25 min, and reequilibration with 0% solvent B from 25 to
30 min. 4.0 mg of 2 was isolated and analyzed by
1
H and
13
C NMR (Oxford NMR AS400).
For isolation of 1 and 3, 5.0 x 10
8
spores of LO11940 were inoculated into eight 2 L
Erlenmeyer flasks containing 500 mL LMM. Culture conditions and induction parameters were the
same as above. The ethyl acetate extract (395 mg) was fractionated via normal-phase column
chromatography using EA:hexanes as a mobile phase in the following ratios: 1:9, 3:7, 1:1, 7:3, 1:0.
Fractions containing both 1 and 3 as indicated by TLC were combined and dried in vacuo (155.5 mg).
This subfraction was further purified via normal phase preparative TLC using 1:19 MeOH:DCM.
120

Bands were excised from the preparative TLC plate with organic solvent and dried in vacuo. 1 was
analyzed with no further purification. 3 was further purified using HPLC-DAD using isocratic
elution. The solvents used for purification of 3 were 100% AcN (solvent B) and 100% H 2O (solvent
A), both containing 0.05% TFA. The ratio of solvent A to solvent B was 1:1. 25.7 mg of 1 and 10.5
mg of 3 were isolated.
4.4.4 Compound Spectral Data

NMR spectra were acquired on a Varian Mercury Plus 400 spectrophotometer. NMR spectra
for 1, 2, and 3 are available in Figs. S3-14. Mass spectra are available in Figs. 15-17. The high-
resolution electrospray ionization mass spectrum (HRESIMS) of 3 was obtained on a ThermoFisher
Q-Exactive Orbitrap mass spectrometer.
Triorsellinaldehyde (3): Red-orange amorphous powder; For 1D and 2D NMR (MeOH-d 4),
see Figures S7-14; for mass spectra, see Figure S17; HRESIMS, [M + H]
+
m/z found 425.16092,
calc. for C 24H 24O 7: 424.15221.

121

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study of the role of metals in the metabolism of microorganisms. Proceedings American Phil
Society 94(2), 152–170.

126

53. Solladié, G., Rubio, A., Carreño, M. C., & García Ruano, José L. (1990). Asymmetric
synthesis of orsellinic acid type macrolides: The example of lasiodiplodin. Tetrahedron:
Asymmetry, 1(3), 187–198. https://doi.org/10.1016/0957-4166(90)90013-Z
127

4.6 Supplementary figures

Figure S1. Prediction of the AfoA DNA-binding motif using InterProScan.

128

Figure S2. Prediction of AfoA coiled-coil dimerization motifs using PCOILS.

129

Figure S3.
1
H NMR spectrum of orsellinaldehyde (1). The solvent used was acetone-d 6.
130

Figure S4.
13
C NMR spectrum of orsellinaldehyde (1). The solvent used was acetone-d 6.
131

Figure S5.
1
H NMR spectrum of diorsellinaldehyde (2). The solvent used was acetone-d 6.
132

Figure S6.
13
C NMR spectrum of diorsellinaldehyde (2). The solvent used was acetone-d 6.
133

Figure S7.
1
H NMR spectrum of triorsellinaldehyde (3). The solvent used was methanol-d 4.
134

Figure S8.
13
C NMR spectrum of triorsellinaldehyde (3). The solvent used was methanol-d 4.
135

Figure S9. gCOSY NMR spectrum of triorsellinaldehyde (3). The solvent used was methanol-d 4.
136

Figure S10. gHSQC NMR spectrum of triorsellinaldehyde (3). The solvent used was methanol-
d 4.
137

Figure S11. gHMBC NMR spectrum of triorsellinaldehyde (3). The solvent used was
methanol-d 4.
138

Figure S12. NOESY NMR spectrum of triorsellinaldehyde (3). The solvent used was methanol-
d 4.
139

Figure S13. DEPT-90 NMR spectrum of triorsellinaldehyde (3). The solvent used was
methanol-d 4.
13
C NMR is shown in red for reference.
140

Figure S14. DEPT-135 NMR spectrum of triorsellinaldehyde (3). The solvent used was
methanol-d 4.
13
C NMR is shown in red for reference.

141

Figure S15. Mass spectrum for orsellinaldehyde (1). (A) Positive-mode ESI; (B) Negative-mode
ESI.
142

Figure S16. Mass spectrum for diorsellinaldehyde (2). (A) Positive-mode ESI; (B) Negative-
mode ESI.

143

Figure S17. Mass spectrum for triorsellinaldehyde (3). (A) Positive-mode ESI; (B) Negative-
mode ESI.

144

4.7 Supplementary tables

Table S1. Experimental
1
H and
13
C chemical shift data of orsellinaldehyde (1) compared to
literature values
53
. Splitting patterns and coupling constants (in Hz) are indicated. The solvent used
was acetone-d 6.

Position ðH (exp) ðH (lit) ΔðH ðC (exp) ðC (lit) ΔðC
1 10.10 (s) 9.55 (s) 0.55 194.3 194.1 0.2
2 - - - 113.8 113.7 0.1
3 - - - 166.4 167.1 0.7
4 6.19 (d, 2.3) 6.16 (d, 2.0) 0.03 111.6 111.5 0.1
5 - - - 164.2 166.1 1.9
6 6.31 (d, 2.3) 6.29 (d, 2.0) 0.02 101.4 101.3 0.1
7 - - - 146.0 145.9 0.1
8 2.54 (s) 2.53 (s) 0.01 18.1 18.2 0.1
3-OH 12.49 12.49 0.00 - - -
5-OH 9.78 10.09 0.31 - - -

145

Table S2. Experimental
1
H and
13
C chemical shift data for diorsellinaldehyde (2) compared
to literature values
24
. Splitting patterns and coupling constants (in Hz) are indicated. The solvent
used was acetone-d 6.

Position ðH (exp) ðH (lit) ΔðH ðC (exp) ðC (lit) ΔðC
1 10.08 (s) 10.08 (s) 0.00 194.6 194.5 0.1
2 - - - 113.5 113.5 0.0
3 - - - 164.6 164.6 0.0
4 - - - 112.6 112.6 0.0
5 - - - 164.6 164.6 0.0
6 6.29 (s) 6.30 (s) 0.01 112.1 112.0 0.1
7 - - - 143.4 143.3 0.1
8 2.49 (s) 2.50 (s) 0.01 18.0 17.9 0.1
1’ 3.86 (s) 3.86 (s) 0.00 18.6 18.5 0.1
2’ - - - 116.3 116.2 0.1
3’ - - - 155.7 155.6 0.1
4’ 6.33 (d, 2.0) 6.35 (d, 2.3) 0.02 101.2 101.1 0.1
5’ - - - 157.3 157.2 0.1
6’ 6.24 (d, 2.0) 6.24 (d, 2.4) 0.00 111.0 110.9 0.1
7’ - - - 140.8 140.6 0.2
8’ 2.29 (s) 2.29 (s) 0.00 20.7 20.6 0.1
3-OH 13.34 (s) 13.34 (s) 0.00 - - -

146

Table S3. Experimental
1
H and
13
C chemical shift data for triorsellinaldehyde (3). Splitting
patterns and coupling constants (in Hz) are indicated. The solvent used was methanol-d 4.

Position ðH ðC Position ðH ðC
1 10.00 (s) 194.7 5’ - 155.2
2 - 113.9 6’ 6.27 (s) 101.0
3 - 164.8 7’ - 140.6
4 - 113.4 8’ 2.07 (s) 16.4
5 - 165.3 1’’ 3.84 (s) 23.3
6 6.17 (s) 112.3 2’’ - 118.9
7 - 143.7 3’’ - 156.5
8 2.43 (s) 18.1 4’’ 6.11 (d, 2.2) 101.3
1’ 3.82 (s) 19.4 5’’ - 140.6
2’ - 117.9 6’’ 6.04 (d, 2.1) 110.5
3’ - 153.7 7’’ - 156.9
4’ - 120.3 8’’ 1.96 (s) 20.4

147

Chapter 5: Molecular Genetic Identification of the Terrecyclic Acid
and Quadrone Biosynthetic Gene Cluster in Aspergillus terreus

Chris Rabot, Tzu-Shyang Lin, & Clay C. C. Wang

5.1 Abstract

Terpenes represent one of the most diverse and abundant classes of natural products known.
Genome sequencing efforts have revealed a plethora of biosynthetic gene clusters predicted to
encode enzymes involved in terpene biosynthesis. However, most products of these clusters remain
elusive. In order to link putative biosynthetic gene clusters to downstream terpene & terpenoid
products, we generated a genome-wide terpene synthase/cyclase knockout library in Aspergillus terreus.
Metabolic profiling of mutants permitted the identification of the biosynthetic gene cluster
responsible for production of the sesquiterpenoids terrecyclic acid and quadrone.
5.2 Introduction

Secondary metabolites (SMs) are structurally heterogeneous, low-molecular weight natural
products that often exhibit a broad range of bioactivities. In contrast to primary metabolites, SMs
are not directly required for the growth and survival of the producing organism. Rather, they usually
confer a selective advantage to the organism that produces them. SMs can be divided into classes
depending on the type of enzymes that direct their biosynthesis. The most common types of SM
classes include polyketides, nonribosomal peptides, and terpenes/terpenoids, which are synthesized
by polyketide synthases (PKSs), nonribosomal peptide synthetases (NRPSs), and terpene
synthases/cyclases (TS/TCs), respectively. Collectively, these enzymes are able to generate a vastly
diverse array of compounds from a relatively small subset of starting materials.
Filamentous fungi are widely regarded as prolific producers of SMs with potent and diverse
bioactivities. Notable examples of clinically relevant fungal SMs include the antineoplastic polyketide
doxorubicin, the anticholesterolemic HMG-CoA reductase inhibitor lovastatin, and the
148

immunosuppressant peptide cyclosporine. Genome sequencing efforts have revealed that genes that
orchestrate fungal SM biosynthesis are generally organized into biosynthetic gene clusters (BGCs)
1
.
The clustered nature of these biosynthetic genes facilitates the subsequent in silico identification of
putative BGCs. Using these computational approaches, subsequent targeted gene deletions and/or
heterologous expression approaches can then be employed to validate the role that a putative BGC
has in the biosynthesis of a given SM. Following this confirmation, modulation of biosynthetic gene
expression using an array of gene targeting techniques (promoter replacement, gene
deletions/duplications, domain swapping, etc.) can enable yield optimization, combinatorial
biosynthesis, and BGC activation efforts.
Terpenes/terpenoids are the most diverse and abundant class of SMs produced in nature
2
.
Produced by almost all forms of life, they exhibit remarkable functional diversity despite originating
from relatively simple building blocks: the C 5 isoprene units isopentenyl pyrophosphate (IPP) and
dimethylallyl pyrophosphate (DMAPP). In terpene biosynthesis, IPP is first isomerized by IPP
isomerase to DMAPP
3
. IPP and DMAPP typically then condense “head-to-tail” or “tail-to-tail” to
form larger carbon backbones. The “head-to-tail” coupling of isoprenoid precursors first yields the
linear C 10 substrate geranyl pyrophosphate (GPP; GPP synthase). In turn, additional isoprenoid
subunits can sequentially condense with GPP to yield farnesyl pyrophosphate (FPP; FPP synthase),
geranylgeranyl pyrophosphate (GGPP; GGPP synthase), geranylfarnesyl pyrophosphate (GFPP;
geranylfarnesyl pyrophosphate synthase), and so on. TSs then utilize these linear substrates to form
remarkably diverse structures through highly controlled carbocation rearrangements and cyclizations.
Tailoring enzymes can further expand the chemodiversity of terpenes through the installation of
various functional groups
4
.
Terpenes and terpenoids have found widespread utility in medicine, physiology, agriculture,
and industry. Many members of this class of compounds with interesting bioactivities have been
149

isolated from Aspergillus spp. alone, such as the herbicidal sesquiterpenoid aspterric acid
5
and the
antibacterial asperterpenoid A
6
(Fig. 1). Artesunate, a semisynthetic derivative of the sesquiterpene
lactone artemisinin, is FDA-approved and recommended by the World Health organization as a
first-line treatment for severe malaria
7,8
. Steroids and related compounds, such as cholesterol and
ergosterol, are also derived from terpenes
9,10
. Other terpenes have been shown to exhibit plant
protective properties through oviposition deterrence, pollinator attraction, or related mechanisms
11-14
.
Some terpenes, such as the sesquiterpene bisabolane, have been explored for biofuel applications
15,16
.
Of the 80,000+ members comprising over one-third of all characterized compounds in the
Dictionary of Natural Products
4
, the global terpenome represents an indispensable, and growing,
chemical arsenal. Thus, furthering our understanding of approaches to facilitate terpene discovery
and biosynthesis are of critical importance.
We sought to expand our understanding of terpene biosynthesis by generating a genome-
wide TS knockout library in the filamentous fungus Aspergillus terreus ATCC 20516. Fungi represent
rich sources of terpenes and terpenoids, and the genetic tractability of certain genera, especially the
Aspergilli, represents an opportunity both to reveal aspects of terpene biosynthesis and to facilitate
future discovery efforts. This approach enabled the identification of the SM BGC responsible for
the biosynthesis of the sesquiterpenoids quadrone and terrecyclic acid. Through these efforts, we
also found that bioinformatic software packages that detect BGCs, even amongst different versions
of the same software, can vary widely in their predictive capacities toward terpene biosynthesis.
150

Figure 1. Chemical structures of terpenes and terpenoids from Aspergillus spp. Boxed
structures indicate compounds included in this study.

5.3 Results & Discussion

5.3.1 Bioinformatic BGC prediction

Following genome sequencing, assembly, and annotation of the A. terreus ATCC 20516
genome (see Materials & Methods), we next used antiSMASH (v4.2.0
17
and v6.1.1
18
) to predict
BGCs. Interestingly, a comparison of predicted terpene BGCs between these two versions revealed
vastly different outcomes; antiSMASH v4.2.0 predicted 13 terpene BGCs, while v6.1.1 predicted
nine. Careful analysis of these putative BGCs revealed that only seven were predicted by both
versions. Six were exclusively predicted by v4.2.0, and two were exclusively predicted by v6.1.1.
151

In our analysis, we included TS/TCs colocalized with other core biosynthetic genes (PKSs,
NRPSs, etc.), predicted to comprise hybrid BGCs. We reasoned that TS/TCs located within a
certain proximity of these core biosynthetic genes may not guarantee their concerted biosynthetic
routes. A list of putative BGCs identified by both versions of antiSMASH along with relevant details
are listed in Table 1. Interestingly, we also used PRISM 4
19
to predict terpene BGCs, which failed to
recover any putative sequences.
Table 1. Putative terpene BGCs predicted by antiSMASH. Versions 4.2.0 and 6.1.1 were used
to predict putative BGCs. Other parameters relevant to the putative BGCs are included. Letters
indicate BGCs that match between versions.

BGC
antiSMASH
version
BGC class
TS
gene size
(bp)
TS
protein size
(residues)
TS
start site
TS
end site
9
a
4.2.0 Meroterpenoid 1593 471 701074 798378
16 4.2.0 Terpene 3170 574 430871 454041
17 4.2.0 Terpene 1540 364 535119 556659
26 4.2.0 Meroterpenoid 1389 368 73863 150623
27
b
4.2.0 Terpene 1178 319 391781 412959
35
c
4.2.0 Meroterpenoid 1091 320 83908 148222
40
d
4.2.0 Terpene 1128 355 229084 250212
41 4.2.0 Terpene 2568 677 295885 318453
46 4.2.0 Terpene 1153 327 257544 278697
47
e
4.2.0 Terpene 2394 714 188987 211381
60
f
4.2.0 Terpene 1573 375 45488 67061
66 4.2.0 Terpene 986 328 1 12329
67
g
4.2.0 Terpene 2090 696 1 13029
5.2
a
6.1.1 NRPS-like/terpene 3112 683 701074 746217
6.1 6.1.1 Terpene 1235 322 131790 153025
16.2
b
6.1.1 Terpene 1504 316 391781 413285
24.1
c
6.1.1 Meroterpenoid 1078 279 93908 148222
152

27.1
d
6.1.1 Terpene 1128 355 229084 250212
33.1
e
6.1.1 Terpene 4011 1109 189395 213406
45.2 6.1.1 Terpene 996 185 215175 230022
90.1
f
6.1.1 Terpene 930 229 1 12303
120.1
g
6.1.1 Terpene 2090 696 1 13029

5.3.2 Generation of a genome-wide TS knockout library

Having identified putative terpene BGCs in the A. terreus ATCC 20516 genome, we then set
out to establish a genetic system to permit the generation of a genome-wide TS knockout library. In
order to facilitate selection of correct A. terreus ATCC 20516 transformants with knocked out TSs,
we first knocked out the homolog of pyrG using a fusion PCR-based approach
20
. The gene product
of pyrG, orotidine-5'-phosphate decarboxylase, catalyzes the decarboxylation of orotidine
monophosphate (OMP) to uridine monophosphate (UMP)
21
. Further, an intact copy of pyrG renders
strains sensitive to 5-fluoroorotic acid (5-FOA)
22
. Thus, pyrG represents an attractive genetic target
for subsequent gene knockout efforts; the addition of uracil/uridine and 5-FOA to transformation
plates facilitates positive selection of correct transformants and negative selection of non-
transformants, respectively.
We used BLASTp to identify the homolog of pyrG in A. terreus ATCC 20516 using the A.
nidulans pyrG (AN6157) as a query. BLASTp recovered one sequence (gene 009089: 85%
similarity/76% identity) as a candidate pyrG homolog that was subsequently knocked out using
fusion PCR. Briefly, flanking regions approximately 1500 bp up- and downstream from the coding
sequence of the pyrG homolog were amplified and fused together. This construct was transformed
into A. terreus protoplasts (see Materials & Methods for transformation protocol). Transformants
were plated onto selective solid plates containing 5-FOA to facilitate negative selection of non-
transformants. Transformants were chosen and verified with diagnostic PCR.
153

Having established a correct pyrG mutant, we then used a similar fusion PCR-based
approach to knock out all nine candidate TSs based on antiSMASH v6.1.1. To do this, we amplified
approximately 1.0-1.5 kbp flanking regions up- and downstream of candidate TSs and fused it to the
A. fumigatus pyrG (AfpyrG) selectable marker. We also targeted all TSs localized within putative
hybrid BGCs for reasons stated above. Transformation of the A. terreus ATCC 20516 pyrGΔ mutant
with these constructs restores uracil & uridine prototrophy, facilitating subsequent selection of
correct transformants. All transformants were verified by diagnostic PCR.
5.3.3 Metabolic profiling of terpene synthase mutants

Metabolic profiling of the wild-type A. terreus ATCC 20516 along with the TS knockout
strains did not indicate that the production of putative terpenes or terpenoids was eliminated. We
did note, however, that we observed the putative production of terrecyclic acid when cultured in
MEB medium. Indeed, we confirmed this using an authentic terrecyclic acid standard (Fig. 2). As
quadrone is a known precursor of terrecyclic acid
23
, it is likely that quadrone also was produced in
these conditions. The similar polarities and identical masses likely made it difficult to distinguish
between these two compounds. As the production of terrecyclic acid and/or quadrone was not
eliminated upon deletion of all predicted TSs in the genome, we therefore reasoned
that antiSMASH v6.1.1 may not have accurately predicted all terpene BGCs.
Curious as to whether earlier versions of antiSMASH had different predictive capabilities
regarding terpene biosynthesis, we analyzed the same A. terreus genome with antiSMASH v4.2.0. As
discussed above, the two versions yielded markedly different predictions: v6.1.1 predicted 13 terpene
BGCs, while v4.2.0 predicted nine. Critically, six BGCs were exclusively predicted by version 4.2.0.
We hypothesized that one of these six BGCs was responsible for the biosynthesis of terrecyclic acid
and quadrone. We therefore employed the same knockout strategy as previously described to knock
out the remaining six TS/TCs predicted by antiSMASH v4.2.0.
154

Gratuitously, metabolic profiling revealed that terrecyclic acid production was eliminated
upon deletion of one of the remaining six putative TS/TCs (Fig. 2). Analysis of the composition of
this BGC revealed a TS bearing homology to several sesquiterpene TSs, including the penifulvin A
synthase (66% similarity, 47% identity), the presilphiperfolan-8-beta-ol synthase (63% similarity, 46%
identity), and the pentalenene synthase (64% similarity, 46% identity). We term this BGC qdr,
containing the TS qdrA that initiates the cyclization of FPP.
Quadrone, first discovered in 1978 in A. terreus
24
, was found to exhibit significant inhibitory
activity in vitro towards KB cells, derived from HeLA cells, with an EC 50 of 1.3 µg mL
-1
. It also
exhibits in vivo activity towards murine P388 lymphocytic leukemia cells
25
. Similarly, terrecyclic acid
has been shown to exhibit broad antimicrobial activity against gram-positive bacteria, yeasts, and
fungi. It also exhibits moderate activity in vivo against murine P388 lymphocytic leukemia cells
26
.
Furthermore, terrecyclic acid has been shown to modulate oxidative and inflammatory cellular stress
response pathways. It also acts as a small-molecule inducer of the transcriptional heat shock protein
response in 3LL Lewis lung carcinoma cells
27
. Due to their bioactivity and interesting structural
features, several total syntheses of quadrone and terrecyclic acid have been reported
28-32
.

155

Figure 2. Comparative metabolomics showing elimination of terrecyclic acid biosynthesis
upon deletion of a TS in A. terreus ATCC 20516. Negative-mode extracted ion chromatograms
(m/z = 246-248) of (i) a terrecyclic acid standard, (ii) the culture medium of A. terreus ATCC 20516
cultured in MEB liquid medium, and (iii) culture medium of A. terreus ATCC 20516 pyrGΔ with a
deleted TS (gene 009303). Y-axes are standardized to the same maximum intensities.

Bioinformatic analysis reveals that a putative cytochrome P450 monooxygenase (gene
009304) and a short-chain dehydrogenase/reductase (gene 009305) are colocalized with this TS
(gene 009303). Based on these observations, we propose the putative biosynthetic pathway for
quadrone and terrecyclic acid (Fig. 3). We hypothesize that FPP undergoes a 1,11-cyclization to
yield a humulyl cation, which then undergoes a series of hydride shifts and intramolecular bond
formations. This cyclization cascade may ultimately be quenched by a water molecule, as has been
observed in terpene biosynthesis
4
, to yield a hydroxylated tricycle 1. Oxidation of this hydroxyl
group, methyl migration & oxidation, and installation of a carboxylic acid to form terrecyclic acid
may be aided, at least in part, by the cytochrome P450 monooxygenase (gene 009304) encoded
within the BGC. Finally, an intramolecular proton transfer and bond formation between the
methylene and carboxylic acid functionalities may be catalyzed by the short-chain
156

dehydrogenase/reductase (gene 009305) to form quadrone. Although members of this class of
enzymes typically catalyze simple redox reactions, they have also been shown to initiate rather
unusual cyclizations
33-35
.

Figure 3. The proposed biosynthetic pathway of quadrone and terrecyclic acid. Brackets
indicate hypothetical intermediates.

Our current studies focus on the elucidation of the full biosynthetic pathway of quadrone
and terrecyclic acid by generating mutants of individual genes in the qdr BGC detailed above. Upon
elucidation of the biosynthetic pathway of terrecyclic acid and quadrone, targeted overexpression or
deletion of genes within the qdr BGC can enable yield optimization or combinatorial biosynthesis,
respectively. Taken together, we identified the BGC responsible for terrecyclic acid and quadrone
biosynthesis through the generation of a genome-wide TS knockout library in A. terreus ATCC
20516.

157

5.4 Materials & Methods

5.4.1 Media & buffer recipes

Medium/reagent Ingredients
Glucose minimal
medium (GMM)
10 g L
-1
d-glucose, 12 g L
-1
NaNO 3, 1.04 g L
-1
KCl, 1.04 g L
-1

MgSO 4*7H 2O, 3.04 g L
-1
KH 2PO 4, 1 mL Hutner’s trace element
solution
36
, 15 g L
-1
agar
Potato dextrose broth
(PDB)
4 g L
-1
potato extract, 20 g L
-1
d-glucose
Malt extract broth
(MEB)
20 g L
-1
malt extract, 1 g L
-1
peptone, 20 g L
-1
d-glucose
0.1% Tween 80 8.5 g L
-1
NaCl, 1 mL L
-1
Tween 80
YEPD medium 10 g L
-1
yeast extract, 20 g L
-1
Bacto-Peptone, 20 10 g L
-1
d-glucose
Digestion buffer
1.1 M KCl, 0.1 M citric acid monohydrate. Adjust final pH to 5.8 with
KOH. Sterilize via autoclave.
1.0 M ST 1.0 M Sorbitol, 50 mM Tris (pH 8.0). Sterilize via autoclave.
1.0 M STC
1.0 M Sorbitol, 50 mM Tris (pH 8.0), 50 mM CaCl 2. Sterilize via
autoclave.
40% PEG in STC 40% w/v polyethylene glycol 4000 in STC buffer. Sterilize via autoclave.

5.4.2 Strain information & genomic DNA extraction

A. terreus Thom ATCC 20516 (NRRL 11156) was purchased from ATCC
(https://www.atcc.org). To obtain genomic DNA suitable for genome sequencing, the strain was
cultivated on solid GMM agar plates and incubated for three days at 37 °C. Spores were harvested in
0.1% Tween 80 solution, and the resulting spore suspension was used for genomic DNA extraction
(QIAGEN DNeasy PowerMax Soil DNA Isolation Kit). Roughly 3 μg of purified gDNA was used
for microbial whole genome resequencing (Illumina Platform PE150, Novogene) using the A. terreus
NIH 2624 genome as a reference genome.
158

5.4.3 Genome sequencing, assembly, & annotation

Subsequent genome analysis using the raw reads was performed at the High-Performance
Computing Center (HPCC) at UC Riverside (https://hpcc.ucr.edu). AAFTF (v0.2.0) was used to
assemble the genome, which uses Trimmomatic (v0.36) to trim reads and Bowtie (v2.3.4.1) to filter
against a database of contaminants. Reads were assembled using SPAdes (v3.12.0). Vector sequences
were removed with BLASTN against a vector sequence database. Bacterial contamination was
filtered with sourmash (v3.5.0) and the assembly was polished with the short reads using Pilon
37-39
.
The assembly was masked using RepeatMasker (v. open-1.0.11) and annotated with the Funannotate
pipeline (v1.5.0)
40,41
. This approach uses HISAT2 (v2.2.1), Trinity (v2.11.0), and PASA (v2.4.1) to
predict genes and identify homology for annotation
42-44
.
5.4.4 Protoplasting protocol

1. Inoculate a 250 mL Erlenmeyer flask containing 50 ml YEPD with 5.0 x 10
7
spores of A.
terreus ATCC 20516 (10
6
spores mL
-1
). Incubate for 18-24 hours at 37 °C with shaking at 100
rpm.
2. Harvest the mycelia by filtering the culture through sterile Miracloth.
3. To a 50 mL Falcon tube, add 1.2 g VTP and dissolve in 20 mL Digestion Buffer. Vortex for
30 minutes to fully dissolve. Filter-sterilize and discard the first 1-2 mL of buffer that passes
through the filter to eliminate any residual detergent. Add the filter-sterilized digestion buffer
to a sterilized 50 mL Erlenmeyer flask.
4. Add 10 mL sterile YEPD (plus any required supplements) to the 50 mL Erlenmeyer flask.
5. Add the mycelia to the digestion buffer/YEPD solution and digest the mycelia at 30 °C, 80-
100 rpm with shaking for 3-4 hours.
6. Monitor the mycelia for protoplast formation microscopically. Protoplasts are large, circular
structures that are much larger than fungal spores. When most or all of the mycelia is
159

digested (ca. three to four hours), filter the culture through sterile Miracloth. Collect the flow
through containing the protoplasts into a 50 mL Falcon tube.
7. Centrifuge the flow-through at 4 °C, 800 g for 5 minutes.
8. Decant the supernatant. Resuspend the cells in 1 mL 1.0 M ST in a sterile Eppendorf tube.
Centrifuge again at 4 °C, 800 g for 5 minutes.
9. Decant the supernatant. Resuspend the pellet once again in 1 mL STC. Centrifuge again at
4 °C, 800 g for 5 minutes.
10. Resuspend the final pellet in 500 µL STC.
11. Take 10 µL of the protoplast solution and view it under a microscope to ensure protoplast
generation and abundance.
12. Add 100 µL 40% PEG solution and mix gently but thoroughly. This PEG should be freshly
filtered, and the first 1-2 mL should be discarded.
13. To 100 µl of protoplasts, add 1-10 µg of DNA (in a volume of 15 µL or less). Gently pipette
up and down. Incubate on ice for 25 minutes. Then, add 1 mL of 40% PEG solution and
incubate for 25 min at room temperature.
14. Plate the protoplasts onto plates containing solid GMM + 0.6 M KCl (or 1.0 M sucrose),
plus any necessary supplements. Incubate at 30 °C overnight. The next day, transfer plates to
a 37 °C incubator. Transformants should appear after 2-4 days.
5.4.5 Fermentation and HPLC-DAD-MS analysis

All TS knockout strains were cultured alongside wild-type A. terreus ATCC 20516 in 125 mL
Erlenmeyer flasks containing 30 mL of three different types of liquid media: GMM, PDB, and MEB
(see Materials & Methods for full recipes). Cultures were incubated for seven days at 37 °C with
shaking at 180 rpm. Following incubation, mycelia were filtered from the culture medium. Culture
media were extracted with 30 mL ethyl acetate (EA). The organic extract was dried (TurboVap LV)
160

and resuspended in 1 mL 4:1 MeOH:DMSO. 10 µL of the resuspended extract was analyzed via
HPLC-DAD-MS. The filtered mycelia were submerged in 1:1 MeOH:DCM and sonicated for one
hour. The organic extract was filtered, dried, resuspended in 25 mL ddH 2O, and extracted with 25
mL EA. The organic extract was dried, resuspended, and separately analyzed as above.
HPLC-DAD-MS spectra were acquired on a ThermoFinnigan LCQ Advantage ion trap
mass spectrometer with a reverse phase C 18 column (Alltech Prevail C 18; particle size, 3 μm; column,
2.1 x 100 mm) at a flow rate of 125 μL min
−1
. The solvent gradient was 95% MeCN–H 2O (solvent B)
in 5% MeCN–H 2O (solvent A), both of which contained 0.05% formic acid, as follows: 0% solvent
B from 0 to 5 min, 0 to 100% solvent B from 5 min to 35 min, 100% solvent B from 35 to 40 min,
100 to 0% solvent B from 40 to 45 min, and reequilibration with 0% solvent B from 45 to 50 min.
Conditions for MS included a capillary voltage of 5.0 kV, a sheath gas flow rate at 60 AUs, an
auxiliary gas flow rate at 10 AUs, and an ion transfer capillary temperature at 350 °C.

161

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165

Chapter 6: Conclusions and Future Directions

The work herein details my efforts to further our understanding of the exemplary abilities of
fungi. Through my research, I hope to have illustrated the extensive and diverse benefits that they
may offer us. During these past five years of research, I believe that I have developed a sufficient
understanding of fungi and their prospects regarding drug discovery and their industrial applications.
I am excited to observe future work in this field, particularly in several specific areas.
First, I look forward to seeing what new SMs are identified, as well as the techniques used to
discover them. This being said, a major limitation in the field of genome mining, in my view, is that
most SMs, even if discovered, will not undergo a comprehensive series of bioactivity assays. We
know through genome sequencing efforts that a vast majority of SMs remain to be discovered.
While the identification of these molecules is clearly an extremely interesting prospect, I do believe
that we have already overlooked potential applications of SMs already identified.
One particular technique that I find interesting to address this issue is resistance-gene guided
drug discovery. This method exploits the colocalization of resistance genes within BGCs, which are
usually the target of the SM encoded by the BGC itself
1,2
. For example, the sesquiterpenoid
herbicide aspterric acid was found to exhibit submicromolar potency towards dihydroxy-acid
dehydratase in the branched-chain amino acid biosynthetic pathway. This was facilitated by the
colocalization within the BGC of a gene that encodes a protein conferring resistance to the SM that
it encodes. This technique makes readily apparent the activity of the SM encoded by a given BGC,
and therefore avoids the ambiguity associated with the bioactivity of a given SM.
I also look forward to future techniques associated with the activation of SMs. A facile
approach to genome mining that avoids laborious gene targeting techniques is still critically needed.
Clearly, a high-throughput application to activate en masse the genes within BGCs is still needed.
166

Prioritization of BGCs is also needed, and, for reasons stated above, resistance-gene guided methods
may prove useful towards this end.
Another direction of research that I look forward to witnessing regards the biological
upcycling of plastics. I am proud of the progress that we’ve made regarding the upcycling of
polyethylenes, polystyrenes, and more. The major advancement, in my eyes, is the finding that we
can use substrates derived from plastics to grow fungi. The diverse products that originate from
fungi detailed in the introduction and throughout this thesis should be able to be produced from
these same substrates. Thus, the enormous array of products (SMs, proteins, dyes, biocontrol agents,
organic acids, products used in food & cosmetic industries, etc.) can reasonably be produced from
plastics through the actions of fungi.
Our specific plans regarding plastic upcycling via fungi are extensive and still growing. We
are currently aiming to produce a wide catalog of products from plastic-derived substrates. First, we
are looking to manufacture industrially relevant proteins, such as proteases, glucoamylases,
polymerases, and DNases using these methods. We are further looking to produces dyes, such as the
red, orange, and yellow dyes, derived from the azaphilone class of SMs produced by Monascus spp.
Furthermore, we have begun construction of a strain of A. nidulans expressing a gene encoding an
indigoidine synthase, which should enable production of indigo dyes. We are also expanding the
number of biocontrol agents that we can produce from plastics; other relevant species, such as
Trichoderma spp., should be able to be produced from the methods detailed above. Lastly, we are in
the process of developing a heavily engineered “superstrain” of A. nidulans, exploiting the genetic
machinery detailed in strain LO10050 in Chapter 2. We should be able to readily express essentially
any gene that we would like by incorporating it into the genome of this strain.
Taken together, my results described here expand our current scientific understanding of
fungi. I effectively employ genetic engineering and synthetic biology approaches to shape the fungal
167

phenotype. We revealed complex regulatory mechanisms regarding SM biosynthesis. We also
identified the BGC responsible for terrecyclic acid A and quadrone biosynthesis. Lastly, and in my
opinion most importantly, we leverage fungal biosynthesis to rapidly and efficiently upcycle post-
consumer plastics to high-value natural products. I am proud of the work that I have done, and I
look forward to witnessing the progress that future scientists in this field will make.

168

References
1. Tang, X., Li, J., Millán-Aguiñaga, N., Zhang, J. J., O’Neill, E. C., Ugalde, J. A., Jensen, P. R.,
Mantovani, S. M., & Moore, B. S. (2015). Identification of Thiotetronic Acid Antibiotic
Biosynthetic Pathways by Target-directed Genome Mining. ACS Chemical Biology, 10(12),
2841–2849. https://doi.org/10.1021/acschembio.5b00658

2. Yan, Y., Liu, Q., Zang, X., Yuan, S., Bat-Erdene, U., Nguyen, C., Gan, J., Zhou, J., Jacobsen,
S. E., & Tang, Y. (2018). Resistance-gene-directed discovery of a natural-product herbicide
with a new mode of action. Nature, 559(7714), 415–418. https://doi.org/10.1038/s41586-
018-0319-4

Abstract (if available)

Abstract Fungi represent one of the most versatile groups of organisms in existence. They and their fermentation products have proven to be instrumental toward the advancement of medicine, agriculture, and biotechnology. As evidence of their importance, even their transient absence in ecological systems would lead to catastrophic environmental failure. What is more, most applications in society that fungi confer simply result from wild-type strains. We are fortunate to exist in the post-genomic era, which enables us to dramatically manipulate the behavior of fungal strains through synthetic biology and genetic engineering approaches. The work detailed herein describes approaches that I have taken to exploit the abilities of fungi to the fullest.

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Creator Rabot, Christian Anthony (author)

Core Title Harnessing synthetic biology for drug discovery & waste valorization

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Publication Date 12/06/2022

Defense Date 10/27/2022

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