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Chapter 1
Natural Products
1.1
Introduction
The term ‘natural product’ means molecules of life1 but in general it refers to
distinct low molecular weight organic compounds produced by living organisms such as
bacteria, fungi, lichens, marine invertebrates, plants, insects, mammals etc2 which are
recognized for their pharmacological and biological activities. Natural products are also
known as secondary metabolites as they are not crucial for basic life processes like
growth and reproduction but assist the host organisms in their survival, facilitate
interaction and communication, help in adaption to varied environments3 and they may
have evolved to provide defence mechanism against pathogens and predators.4 They
possess diverse and complex chemical structures which are distinctive of the species or
producing organism and are formed at specific stage of the morphological development
of the host organism,5 for example during sporulation or pigment production. Secondary
metabolites are synthesized from simple intermediate products from primary
metabolism, the main precursors are acetyl Coenzyme A, shikimic acid, mevalonic
acids and amino acids.6 Their ability to bind biological targets and stimulate bioactive
effects has attracted the attention of pharmaceutical industries, natural-product chemists
and biologists to explore different natural habitats, terrestrial, aquatic and the microbial
world for the presence of natural products and for many years they have been a wealthy
source of potential drugs.7
Since primitive times many different plant species have been documented to be
used as medicines for human ailments and later many natural product compounds
purified from plants prove to be among initial lead drugs. For example, acetyl salicyclic
acid 1 (aspirin) is based upon the natural product salicin 2, isolated form the bark of the
willow tree Salix alba L.7, 8 The pain killer codeine was synthesized from the alkaloid
morphine7 3 obtained from the plant Papaver somniferum L.; digitoxin 11 from
Digitalis purpurea L. is used as a cardiotonic to ease congestive heart failure,8 and the
antimalarial drug quinine 4 has been used for years to treat fever, malaria and mouth and
throat diseases. There are a number of plant derived anti-tumor drugs, for example
paclitaxel 9 and baccatin 10 from Taxus brevifolia9 and numerous other anti-tumor
1
compounds are in clinical trials. Various plants’ secondary metabolites display
inhibitory activity against microbes for example phenols, quinone 6, coumarin 5,
catechin 7, terpenoids and essential oils possess antimicrobial effects, flavones 8 inhibit
multiple viruses including HIV and catechin 7 is also used as an anti-coagulant.10
The discovery of the antibiotics penicillin11 161 (see section 3.3) and
cephalosporin12
16
from
the
antibacterial
fungi
Penicillium
notatum
and
Cephalosporium acremonium respectively, led to the exploration of various
microorganisms in search of assorted bioactive metabolites. Fungi from the phyla
Basidiomycota and Ascomycota delivered many pharmacological active compounds,
75% were antimicrobial and many others verified to possess antiviral, cytotoxic,
antineoplastic, cardiovascular, anti-inflammatory, antitumor and immune-stimulating
metabolites.13 The bacterial group of Actinomycetes is also rich with novel metabolites,
mainly antibacterial, antiviral, antifungal and antitumor activities.14 The antibiotics
vancomycin 15 from Amycolatopsis orientalis7,15 and erythromycin 13 from
Saccharopolyspora erythraea are active against a wide range of bacteria and are used in
treatment of various infections.7 Doxorubicin 14 isolated from the Streptomyces
peucetius is used in the treatment leukaemia, bone sarcomas and lung and thyroid
cancer.7,15 A number of immune-suppressive agents such as cyclosporin 17 from the
2
fungus Tolyplocladium inflatum and fujimycin 12 from the bacterium Streptomyces
tsukubaensis
have enhanced the field of organ transplants.2 Marine natural products
also provide a rich source of biodiversity with about 15 bioactive compounds approved
from Food and Drug Administration (FDA) and many in clinical trials.14,16 Hence, a
number of surveys conducted on drug sources for the treatment of diseases like cancer
and infections revealed that 60% of approved drugs are of natural origin.17 These
include natural products directly used as drugs as well as semi-synthetic drugs derived
from or modelled on natural products.18,19
There are major revolutions in natural product research and drug design. Large
libraries of natural product analogues and structural mimics are synthesized through
combinatorial chemistry18 and are examined by ‘High Throughput Screening
Technology’ against wide range of important biological targets.19 With present-day
genomics, cost-effective DNA sequencing of microorganisms helps in annotation of
secondary metabolites gene clusters in their respective genomes. This has motivated
researchers to express the silent secondary metabolite gene clusters whose chemical
3
products are unidentified by applying different genetic engineering techniques (see
section 3.3). Biosynthetic pathways of a large number of novel metabolites have been
elucidated from their particular biosynthetic gene clusters by different molecular
biology techniques,5 such as gene knock out, RNAi silencing, cloning, homologous
recombination and heterologous gene expression in surrogate host. These studies have
broadened the knowledge of biosynthetic genes and their encoded enzymes and
proteins. These discoveries pave the way for proteins to be used as biocatalysts20 for in
vivo synthesis of complex natural products and resolve increasing demand for new drug
products.
The main classes of natural products are terpenoids, alkaloids, nonribosomal
peptides and polyketides. They are briefly introduced in the following sections.
1.2
Terpenoids
Terpenoids are made from a five carbon unit (isoprene), which exists in two
active structural forms: dimethylallyl diphosphate 18; and isopentenyl diphosphate 19.21
These simple precursors make diverse structures and about 25,000 structures22 of
terpenes are reported. The name terpenes was named after the volatile oil of pine tree,
turpentine which is composed, among others, of the terpene compound α-pinene 20.1
Terpenoids are identified by their strong aroma, chiefly in essential oils. They have a
wide range of functions: they protect plants against attack of pests and some are used as
insecticides;22 they serve as a means of communication among organisms; they function
as attractants for insects for pollination; and many play a part as mating pheromones
and reproductive hormones.21 Many terpenes have a number of bioactivities, for
example terpenes from the genus Rubia are reported to possess antitumor, antiinflammatory, antimicrobial, anti-malarial and antidiabetic effects.23 Examples of
terpenoids include camphor 21, limonene 22 and gibberelllin GA4 23.
4
1.3
Alkaloids
Alkaloids, like terpenoids, are a large and diverse class of compounds, with
more than 12,000 examples known at present.24 They contain a basic amine group in
their structure and are derived biosynthetically from amino acids. Examples are
morphine 3 and the antimalarial drug quinine 4 (see section 1.1).
1.4
Nonribosomal peptides
Microorganisms, particularly fungi and bacteria, produce an assorted group of
peptide secondary metabolites called nonribosomal peptides (NRP) formed by
multifunctional enzymes known as nonribosomal peptide synthetases (NRPS),
independent of ribosomal pathway.25 NRP are formed from a wide range of substrates
(amino acids) which include both D and L proteiogenic and non-proteiogenic amino
acids which explains the numerous complex structures present in this class of natural
products.26 The NRP serve as antibiotics, immunosuppressants, cytostatins and
siderophores.25 Nonribosomal peptide synthetases (NRPS) are made up of set of
modules, each module consists of basic set of catalytic or enzymatic units called
domains. Modules are distinct sections in nonribosomal peptides and each module
incorporates single amino acids to the growing peptide chain.28 There are three core
domains in each module; the adenylation domain (A), the peptidyl carrier protein (PCP)
and the condensation domain (C).29 The last module normally contains a termination
domain which may be a thiolesterase (TE) or a reductase (R). The adenylation domain
selects and activates an amino acid by adenyaltion with ATP, the PCP serves as a
transporter of the activated substrate between catalytic domains by binding the substrate
to a 4’phosphopantetheine cofactor with a thioester bond. The condensation (C) domain
is responsible for peptide bond formation between the amino group of one amino acid
5
and the acyl group of amino acid of the adjacent module. At the end of last module, the
peptide chain is usually terminated and released by the thiolesterase domain. Many
NRPS may contain additional tailoring domains mainly the epimerization domain, Nmethylation and glycosylation which may further modify the structure of the NRP.30
Examples of NRP include vancomycin 15 and cyclosporin 17 (see section 1.1).
1.5
Fatty acid biosynthesis
Before describing the next class of natural products, the polyketides, it is
necessary to first discuss the biosynthesis of fatty acids. The biosynthesis of fatty acids
has many similar features to polyketides’ pathway and it has always remained a model
system to study the latter.
Fatty acids are primary metabolites present in all living cells. Structurally they
are carboxylic acids with a long saturated chain. They are important sources of fuel for
an organism and when metabolized they liberate large quantities of ATP. They are
essential components of phospholipids and play a basic structural role in assembly of
the cell membrane.31 Fatty acids are synthesized from simple building blocks
particularly the two carbon containing acetate in the form of acetyl Coenzyme A (CoA)
and malonyl CoA. The former acts as a starter unit and the latter as an extender unit.32
Fatty acid synthesis involves a number of key enzymes in each step. In a general
fatty acid pathway an acetyl group from the starter acetyl CoA 24 is transferred by an
acyl transferase enzyme (AT) to the thiol group of the 4’-phosphopantetheine arm of
acyl carrier protein (ACP). This is followed by the acetyl being transferred to another
enzyme β-ketoacyl synthase (KS) at the thiol of the active site cysteine. Meanwhile the
extender malonyl CoA 25 is attached to the ACP and condensation between acetyl
thiolester 27 and malonyl thiolester 26 is catalysed by the KS in a decarboxylative
Claisen condensation with liberation of CO2 and a β-keto thiol ester 28 bound to an
ACP is formed.33 ACP serves as a ‘handle’ to carry the growing acyl chain throughout
the whole process of fatty acid synthesis (Scheme 1.1).
The β-keto thiol ester 28 is then reduced to a secondary alcohol 30 by a βketoacyl reductase (KR) followed by dehydration by a dehydratase enzyme (DH) to
from an unsaturated thiolester 31 and finally an enoyl reductase (ER) performs a final
reduction to form fully saturated thiolester 32.34 The fully saturated thiolester 32 can
6
enter another cycle of fatty acid chain extension with addition of a second malonyl
thiolester by ACP and this process continues until the fatty acid reaches its specific
length, for example a sixteen carbon chain in case of palmitic acid and an eighteen
carbon chain for stearic acid. The last cycle of the fatty acid is terminated by a
thiolesterase enzyme (TE) which liberates the free fatty acid 33 by a hydrolytic release
and a long chain carboxylic acid is formed. Thus fatty acid synthases are large
multifunctional proteins containing KS, ACP, AT, KR, DH, ER and TE, activities.
Scheme 1.1 Series of steps in fatty acid and polyketide biosynthesis.
1.6
Polyketides
Polyketides form a large group of secondary metabolites comprising of
structurally complex compounds, produced by plants, bacteria, fungi, lichens and
marine organisms.35 They have been extensively studied by natural product researchers
because of their fascinating biosynthetic pathways and wide range of important
biological and pharmaceutical properties. Polyketides have contributed about 50
approved drugs in the pharma industry.36 Some important bioactive compounds of this
7
class include the antibiotics erythromycin 13, rifamycin 38,37 monensin 42,38
doxycycline 43,
39
the antihelminthics avermectin 39,40 the antitumor compounds
geldanamycin 4041 and macbecin 41,42 some antifungal polyenes amphotericin 4543 and
primaricin 44, the immunosuppressant tacrolimus (FK506) 46,44 the cholesterol
lowering agent lovastatin 4745 and many others. Some polyketides are also used as
insecticides like spinosad 49,46 while some are toxins, for example aflatoxins 48.47
The basic concept of polyketides biosynthetic origin was presented in 1907 by
John Norman Collie who was a professor at University College London and also a
mountaineer.48 He proposed that ‘Keten’ groups (CH2=C=O) in the form of aldehydes
and ketones can condense and polymerise by simple reactions with liberation of CO2 to
form a range of complex compounds, many of them are present in plants. Hence they
were named polyketenes and subsequently ‘polyketides’.49
8
During the 1940s, when labelled acetate incorporation was established in fatty
acids,50 Robinson supported the polyketide theory presented by Collie.51 Arthur J. Birch
worked on the biosynthetic origin of many aromatic polyketides. He showed
experimentally the incorporation of radio labelled
14
C-acetic acids in alternating
labelling pattern in 6-methyl salicylic acid 51 in Penicillium ariseofuluum (Scheme 1.2).
He endorsed that β-keto chain are formed from condensation of acetate units which
folds to form aromatic polyketides.49
Scheme 1.2: Biosynthetic steps of 6-MSA 51 showing incorporation of 14C acetate units.
Similar studies also demonstrated that methyl groups in fungal polyketides are
incorporated from L-methionine.52 When techniques like Mass spectrometry (MS) and
Nuclear Magnetic Resonance (NMR) were introduced, the biosynthetic investigations
on polyketides were accelerated. Many precursor compounds, mainly acetates were
used in the form of singly or doubly labelled 13C or 2H isotopes and fed to the cultures
of polyketides producing organisms. As
13
C or 2H enriched compounds are easily
studied by NMR, they helped to understand the acetate incorporation patterns and mode
of cyclization in many polyketides.53,54
9
Polyketides are formed by a similar biosynthetic pathway to that of fatty acids.
They are formed by repetitive decarboxylative condensation of simple acetate units in
the form of acetyl CoA 24 and malonyl CoA 25 (in some cases propionate and butyrate
units) in a head to tail manner. In polyketide pathway, the first β-keto thiolester 28 is
formed by a decarboxylative Claisen condensation of acetyl 27 and malonyl thioloester
26 by the same catalytic units KS, AT and ACP (Scheme 1.1). In fungi, the polyketide
chain can also be methylated, receiving a methyl group from S-adenosyl methionine
(SAM) to form α-methyl-β-keto thiolester 29 by a unique CMeT domain not active in
the FAS pathway. The β keto chain can be further processed by the KR, DH and ER
domains.34 The cycle again continues until PKS chain reaches it specific length and the
thiolester is hydrolysed from the peptide and released by a termination domain.
In fatty acid biosynthesis, the β-keto chain is uniformly reduced from a β-keto
group to an alcohol 30, then forms an α, β- double bond in 31 by elimination of water
and at the end is fully reduced to a methylene as observed in 32 (scheme 1.1). In the
polyketide pathway the reduction process is more controlled, selective and variable.
There may not be any reduction in the poly β-keto chain of PKS chain (Scheme 1.1, 35)
and is common in aromatic polyketides. The β-keto chain may only be reduced once to
remain as a hydroxyl group, or dehydrated to remain as a enoyl group (scheme 1.1, 36),
or may be fully reduced to methylene groups to form 34 (Scheme 1.1).33 The polyketide
chain may also hold a pendant methyl or ethyl group by incorporation of methyl
malonyl or ethyl malonyl units as observed in 37 (Scheme 1.1). Polyketide biosynthesis
also sets different stereocentres during reduction of the β-keto chain (Scheme 1.1, 36
and 37). Other factors which determine the structure of the polyketides are choice of the
starter units, number of chain extensions and pattern of cyclization.31,55 The choice of all
these structural functionalities are governed by the above described enzymes (or
domains) and these are collectively called Polyketide synthases (PKS). David Hopwood
used the term ‘programming’34 for the way PKS directs different variables or structural
features in the synthesis of the diverse polyketides structures.56 These directions are
ultimately encoded with in the sequences of PKS gene clusters.
Lots of efforts have been served by researchers to understand the core of
programming of polyketide synthases and have achieved a considerable success.
Advanced genetics and molecular biology tools like sequencing, cloning, gene
expression, and enzymes purification have helped to understand the nature and structure
10
of PKS and how they catalyse different reactions or steps in biosynthesis. Many
biosynthetic pathways of polyketide metabolites and their respective gene clusters are
known.5 Hopwood and co-workers
have achieved some pioneering work in PKS
genetics and construction of cloning vectors.56 His group discovered the first PKS
genes for the antibiotic actinorhordin.53 Different degenerate primers were developed
from the initial PKS known genes and were used to screen genome libraries of many
bacteria and fungi and pave way to discovery of numerous new PKS genes on the basis
of similarities in gene sequences. After allocation of gene clusters many PKS enzymes
and proteins were purified and a number of crystal structures were solved, which also
gave awareness of different catalytic sites in enzymes. The functions of many enzymes
are proved with cell free extracts and in vitro studies. The quest for understanding the
Polyketide synthases and their programming behaviour is still in progress and many
gene clusters and organisms need to be explored for their exclusive biosynthetic and
bioactive properties.57
1.7
Types of Polyketide Synthases
PKS enzymes are divided into three main groups according to the protein
assembly and arrangement of domains with in the polypeptide. The three main classes
are type I, type II and type III PKS.53
1.7.1
Type I PKS
In type I PKS, all the multifunctional domains required for polyketide chain
elongation and β-keto group processing are located on a single large polypeptide. Type I
PKS is further categorized into three types. They are modular PKS, trans-AT modular
PKS and iterative PKS.58
1.7.1(a)
Modular PKS
In modular PKS, the different catalytic units are arranged in the form of sets of
domains, called modules. The domains present in each module performs a single chain
extension and β-keto processing and then passed it on to the next module for another
carbon chain addition and β-keto group processing. Each domain is used once during
the cycle and the linear order of the modules and their respective domains can define the
structure of the polyketide chain. Modular PKS are common in bacteria.59
11
The best studied example of modular PKS is erythromycin A 13,60 the gene
cluster of which was studied in late 1980s.61 It is produced by the Gram positive
bacteria Saccharopolyspora erythraea. Erythromycin A 13 biosynthesis consists of an
intermediate compound 6-deoxyerythronolide B 52 (6-DEB), which is a 14-membered
macrolactone ring and a putative polyketide synthase product.53,62 The polyketide
synthase
responsible
for
the
biosynthesis
of
6-DEB
52
was
named
as
deoxyerythronolide B synthase (DEBS) encoded by three genes eryAI, eryAII and
eryAIII, each 10 kb in length (scheme 1.3).59
Scheme 1.3 Biosynthesis of erythromycin by modular Type I PKS.
The erythromycin PKS (EryPKS) was among the first PKS to be sequenced
among the complex polyketides and formed a model for studying modular PKS. The
three genes encode three proteins DEBS1, DEBS2 and DEBS 3. Each DEBS protein
holds two modules and each module contains the three basic domains KS-AT-ACP with
12
different combinations of KR, DH and ER depending upon the extent of β-group
processing. The last module ends with the thiolesterase (TE) domain. In the first cycle
the AT domain of the first module in DEBS1 binds a propionyl CoA and transfers it to
the pantotheine arm of the adjacent ACP and then to the next KS domain. The AT then
binds the extender unit, a methyl malonyl CoA to the ACP. This results in formation of
a diketide by the combination of the starter and the extender unit by the KS, followed by
keto-reduction by KR to form a β-hydroxy. The ACP in module 1 then transfers the
diketide to the module 2 where it is joined by a second extender unit, followed by keto
reduction by KR in module 2. The triketide is passed to the module 3 with condensation
with a third extender unit. The β-keto group in module 3 remains unreduced because of
a non-functional KR in module 3 as apparent in C-9 in 52, in the fourth cycle the β-keto
group undergoes subsequent reduction, dehydration and enoyl reduction to form a
methylene functionality at C-7 because of presence of KR, DH and ER in module 4.
Similar keto group condensation and reduction continues in cycle 5 and 6 depending on
the domains present in the respective modules. In the last, a 15 carbon PKS chain is
released from the ACP of DEBS 3 by the action of the last TE domain in module 6 and
forms a macrolide 52 structure by the combination of C-1 to carboxylate of C-13
hydroxyl. The post PKS steps includes attachment of a 6-deoxy sugars D-desosamine at
position 5, a L-cladinose at position 3 and P450 mediated hydroxylations at C-6 and C12 to from erythromycin A 13.59
Trans-AT PKS
There are a number of modular type I PKS gene clusters reported, which lack
the regular cis-AT, a domain found in association with ACP and KS and other domains
in a standard module.63 In trans-AT modular PKS, the prescribed function of acyl
transferase is accomplished by stand-alone AT domain(s) which act in trans with all the
other modules and serves the same function of supplying acyl building blocks to all the
respective modules in the PKS. There are a number of polyketide metabolites
biosynthesized by a trans-AT modular PKS pathway for example leinamycin 55,
64
pseudomonic acid A 54 65 and lankacidin C 53. 65 Scheme 1.4 shows the gene cluster of
leinamycin (LNM) produced by Streptomyces atroolivaceus S-140.
64
The gene cluster
consists of six PKS, one NRPS module encoded by the genes lnmI and lnmJ, among
these all the six PKS modules lack the key AT domain. The AT function is provided by
13
LnmG which loads the malonyl CoA units to all the PKS modules to biosynthesize the
compound leinamycin 55 (scheme 1.4).
Scheme 1.4 Biosynthesis of leinamycin 55 by trans-modular PKS.
1.7.1(b)
Iterative Type IPKS
Type I iterative PKSs (IPKS) are made up of a single set of multifunctional
domains found in a large polypeptide. The sole set of domains carry out all cycles of
carbon chain extension and the respective β-keto chain processing, and many domains
are used repeatedly, hence the name iterative. IPKS have long been the attention of
biosynthetic investigations because the single set of enzymes can act differently in every
cycle of chain extension and processing, portraying a high level of complex
programming. IPKS are found most commonly in fungi. IPKSs are divided into three
classes according to the extent of β-keto group reduction and on the presence of KR,
DH and ER domains in the protein architecture. They are non-reducing (NR-PKS),
14
partially reducing (PR-PKS) and highly reducing (HR-PKS). This classification was
first presented by Simpson.34
Non-reducing IPKS
As the name indicates, non-reduced polyketides are formed from the
condensation of intact and non-reduced poly β-keto chain which cyclise forming
aromatic compounds with a mono or multiple ring structures. Some common nonreduced polyketides are orsellinic acid 56, emodin 57 and norsolonic acid 58.57
Orsellinic acid 56, a tetraketide, was among the early discovered fungal PKS from
Penicillium madriti in 1968.34,66 A typical NR-PKS is composed of an N-terminal
loading component, a chain extension component and a C-terminal processing
component.
The loading component is made of a starter unit-acyl transferase (SAT). It can
load acyl CoA and in many cases complex FAS or PKS elements as starter units. For
example norsolorinic acid 58 incorporates a hexanoate unit,34 dehydrocurvularin 59
accepts a HR-tetraketide as starter unit,67 zearalenone 60 uses a HR-hexaketide as a
starter unit.68 The chain extension component consists of KS, followed by AT. The AT
is specifically a malonyl loading domain. Following the AT, is the product template
domain (PT). It is believed to be involved in chain length control34 and in some reports
it also takes part in PKS chain cyclization.57 PT is followed by ACP. Some NR-PKS
may end with ACP but many possess a distinct C-terminal processing component. The
C-terminal components may end in a Claisen cylcase-thiolesterase (CLC/TE) or may
further consist of methyl transferase (CMeT), 69 additional ACPs or thiolester reductase
(R) domains. The CLC/TE is involved in chain length decision, chain release and
cyclisation via intramolecular Claisen condensation.57,34 An example of NR-PKS with
an active CMeT domain is 3-methylorcinaldehyde 61 synthase where a methyl group is
provided by SAM (S-adenosyl methionine) (Scheme 1.5).69 The PKS structure of 3methyl orcinaldehyde synthase (MOS) consists of N-terminal (NT) domain responsible
for selecting the starter unit, followed by KS, AT, PT, an ACP, C-MeT domain and a Cterminus NADPH dependent thiolester reductase (R) domain.
15
Fig 1.1 Examples of non-reduced polyketides, the complex starter units are highlighted in green in norsolorinic acid 58,
dehydrocurvularin 59 and zearalenone 60.
Scheme 1.5 Non-reduced PKS gene cluster of 3-methylorcinaldehyde 61.
Partially reducing IPKS
The protein architecture of PR-PKS consists of KS, AT, DH, a unique core
domain, followed by KR domain and at the end an ACP domain.34 PR-PKS compounds
are formed by successive condensation of acetyl starter and malonyl extender units,
forming poly β-keto chain, while β-keto processing does not necessarily occur in every
cycle; therefore they are termed as partially reduced PKS. The core domain is believed
to maintain integration and functional stability between the domains. A well-studied
PR-PKS is 6-methylsalicylic acid (6-MSA) 51.
70
The 6-MSA was originally obtained
from Penicillium patulum biosynthesized by 6-methylsalicylic synthase (MSAS).
During biosynthesis of 51, one acetate 24 and 3 malonyl extender units 25 condense in
subsequent extensions to form a tetraketide 62. The KR domain functions after the
second extension in the presence of NADPH forming an alcohol group. After the third
16
cycle, the PKS chain undergoes cyclisation and dehydration to form 6-MSA 51
(Scheme 1.6).53
Scheme 1.6 Biosynthetic pathway of 6-MSA 51.
Highly Reducing IPKS
A typical HR PKS consists of KS, AT, DH followed by a C-MeT domain in
most cases. The next domain is the ER, but many HR-PKS may not contain a functional
ER. The ER is followed by a KR and the PKS most normally terminates with an ACP
domain. With the presence of all three β-keto processing domains used iteratively, HR
PKS synthesize structures with high level of complexity and advanced programming.34
For example lovastatin 47, is a HR polyketide produced by Aspergillus terreus.
It is biosynthesized by two PKS proteins, LovB and LovF.71,105 lovB encodes lovastatin
nonaketide synthase (LNKS) and lovF encodes lovastatin diketide synthase (LDKS).
LNKS with the assistance of LovC, a trans acting ER (the ER domain in LNKS is nonfunctional) synthesize a nonaketide PKS intermediate compound Monacolin J 63.
LDKS then produces a methylated diketide 64, which is loaded on to Monacolin J 63 at
C-10 hydroxy by a specialized acyltransferase encoded by a gene lovD to form 47
(Scheme 1.7). Squalestatin S172 65 is also made by two PKS chains, a main hexaketide
and a tetraketide sidechain. Other examples of HR PKS include the longest polyketides
Fumonisin B1 68 produced by Gibberella fujikuroi
73
and T-toxin
74
69 produced by
17
maize pathogen Cochliobolus heterostrophus. Solanopyrone A 66 and alternaric acid 67
produced by the plant pathogen Alternaria solani are also produced by HR PKS. 34
Scheme 1.7 Lovastatin biosynthetic pathway.
Hybrid IPKS-NRPS
Hybrid highly reduced polyketide synthases fused with nonribosomal peptide
synthetase (HRPKS-NRPS) are an important class of synthetases often found in fungi.
The protein architecture consists of the HRPKS domains (KS, AT, DH, MT, inactive
ER, KR, ACP) and nonribosomal peptide catalytic units (C, A, T and terminal R
domain) forming a megasynthase.75 PKS-NRPS have been extensively studied in recent
years because of their intriguing iterative programming code and important biological
18
activities demonstrated by the hybrid compounds.57,34 All PKS-NRPS discovered up till
now possess non-functional ER sequence and if ER activity is required, it is provided by
a distinct trans-acting ER encoding gene homologous to LovC. This property makes
them closely related to the lovastatin 47 gene cluster and presents a common origin.76
The HRPKS synthesizes a polyketide chain from an acetyl starter unit and subsequent
malonyl extender units with methyl groups delivered from SAM by a methyltransferase.
The adenylation domain of the NRPS selects and activates an amino acid and transfers
it to the thiolation domain. The condensation domain binds the amino acid and the
polyketide chain by an amide bond. The hybrid polyketide and peptide chain is released
from the megasynthase by the terminal R domain by either of two release
mechanisms.75 It can either be released in the form of an aldehyde forming pyrrolinone
70 by a Knoevenagel condensation (Scheme 1.8, A) reported in pseurotin A 78,
isoflavipucine 79 and chaetoglobosin A 80 biosynthesis or as a tetramic acid
(pyrrolidone 71) by direct Dieckmann cyclisation (Scheme 1.8, B) detected during
tenellin 87, desmethylbassianin (DMB) 88, equisetin 81 and cyclopiazonic acid 85
biosynthesis.75,57 The hybrid polyketide-peptide compound is further modified by
tailoring enzymes encoded by genes clustered near the megasynthase.
A
B
Scheme 1.8 A, Release mechanism by a Knoevenagel condensation by reductase domain; B, release mechanism by
Dieckmann cyclisation.
The first PKS-NRPS gene cluster was identified for fusarin C 77 from Fusarium
moniliforme and Fusarium venenatum in an attempt to search for C-methyltransferase
domains.77 The fusA gene encodes the synthesis of a tetramethylated heptaketide 72
fused by an amide bond to L-homoserine 73 by the C domain to form a covalently
19
bound intermediate 74 and is probably released by an R domain to form the aldehyde
75.78 Further Knoevenagel condensation forms the putative prefusarin 76. Subsequent
modifying steps of carboxylation, epoxidation and hydroxylation forms Fusarin C 77
(Scheme 1.9).75 Other examples of hybrid PKS-NRPS includes aspyridone A 84,
xyrrolin 86, militarinone C 82 and pramanicin 83.75
Scheme 1.9 Fusarin C biosynthesis.
Figure 1.2 Examples of hybrid PKS-NRPS compounds, the NRPS part is highlighted in red.
20
1.7.2
Type II PKS
In contrast to type I PKS, the enzymatic activities for the β- keto chain
elongation and processing in type II PKSs are present in separate polypeptides, and each
domain is used iteratively.53,58 A well-studied model of type II PKS is actinorhodin 89
biosynthesis (Scheme 1.10).
Scheme 1.10 Actinorhordin pathway showing Type II PKS system.
1.7.3
Type III PKS
Type III PKSs (Scheme 1.11) were originally identified in plants but recently
have also been isolated from several bacteria. In contrast to type I and type II polyketide
biosynthesis, the β-keto chain is elongated and processed at a single multifunctional
active site in type III PKSs.53 It does not require an acyl carrier protein and distinctively
accepts acyl coenzyme A building units, for example in chalcone 90 biosynthetic
pathway.
Scheme 1.11 Chalcone synthase biosynthesis.
21
1.8
AIMS
The main objective of our research is to contribute in understanding the
programming code veiled in the iterative protein structure of the hybrid PKS-NRPS
pathways. We aimed to study the PKS-NRPS systems of three compounds; tenellin 87,
desmethylbassianin 88 and aspyridone A 84.
Tenellin 87 and desmethylbassianin 88 are hybrid polyketide-peptide
metabolites produced by two different strains of enthomopathogenic fungi B. bassiana.
The tenellin 87 pathway has been elaborately studied in the Bristol Polyketide Group
with different genetic and chemical analysis and it helped understand the basic
biosynthetic pathway of tenellin 87. We aimed to continue the biosynthetic studies by
carrying out further heterologous expression of tenellin 87 genes or its components in a
heterologous host. These include expression of PKS-NRPS encoding gene in A. oryzae
without the tailoring genes and expression of tenellin polyketide synthase alone in A.
oryzae without its NRPS counterpart. The objective was to determine the product as
well as the intricate role of each component of the megasynthase in tenellin 87
biosynthesis.
Desmethyl bassianin 88 (DMB) and tenellin 87 gene clusters have 90%
sequence identity but DMB 88 differs in structure from tenellin 87 in having an
additional carbon chain extension and a methyl group less than tenellin 87. We next
intended to study the co-expression of tenellin 87 genes together with DMB 88 tailoring
genes in different combinations in A. oryzae. This was designed to see whether the two
similar yet different biosynthetic genes are compatible to work together in one
expression system, to obtain new engineered natural products and determine the
programming role of the particular genes.
The function of trans-acting enoyl reductase enzyme in tenellin 87 is encoded
by a discrete gene, tenC. The RNAi silencing of tenC has been successfully achieved
before in the native fungus using a strong constitutive promoter.79 We planned to
perform the RNAi silencing of tenC, again, this time using an inducible promoter and
grow the silenced transformants in different growth conditions. We desired to
investigate whether we can control the level of gene silencing and obtain new
compounds reflecting varying degree of silencing.
22
The last objective was to study aspyridone 84 biosynthesis. Aspyridone A 84, a
PKS-NRPS compound is produced from a silent gene cluster in Aspergillus nidulans
and its pathway has been proposed but not proved experimentally.80 We aimed to
investigate aspyridone pathway using an effective heterologous expression system in A.
oryzae and analyse the transformants. The objective was to determine the order of
different biosynthetic steps, role of each gene in the pathway and discover potential of
the megasynthase in synthesizing new bioactive natural products. The detail description
of the aims has been separately described in each chapter with the respective analytical
methods.
23
Chapter 2
Elucidation of new compounds from different genetic studies in
Beauveria bassiana
2.1
Introduction
Beauveria bassiana belongs to a group of entomopathogenic fungi. These fungi
are parasitic to insects and kill or disable them completely. They invade the insects
initially by their microscopic spores called conidia.81 These spores attach to the insect
cuticle and the conidia swells by secretion of lytic enzymes which helps it to breach and
penetrate the outer layer of cuticle. This is followed by development of morphological
structures such as appressorium on the cuticle, infection pegs and penetrant hyphae in
the epicuticle and procuticle helping the conidia hyphae to reach the body cavity of the
insect (hoemocoel). The fungal hyphae continue to proliferate causing damage to the
host tissue and nutrient exhaustion eventually leaves the insect body dead or
destroyed.82
Figure 2.1: Insect infected by fungus Beauveria bassiana.
83
Beauveria bassiana (Balsamo) Vuillemin has a long taxonomic history.84,85
Agostino Bassi (1835) first described this fungus as the causal agent of ‘mark disease’
also known as white muscardine disease in France.86 This disease caused destruction of
silkworm larvae in Southern Europe during the 18th and 19th centuries resulting in huge
economic losses to the silk industry. Bassi discovered that microbes can act as
contagious pathogens of animals and this formed the basic fundamentals of the ‘germ
theory of disease’.87 The first taxonomic recognition of the muscardino fungus was
proposed by Balsamo-Crivelli. He recognized Bassi’s discovery by naming this fungus
24
Botrytis bassiana. Beauverie stated that the fungus should belong to an undescribed
genus and Vuillmen established in 1912 the genus as Beauveria in his honour and
Botrytis bassiana Bals.Criv as the type species.86,88
Beauveria is distributed worldwide. They are soil borne hygomycetes (having
naked spores)89 and are pathogenic towards several different orders of insects including
Lepidoptera, Coleoptera, Hemiptera, Hymenoptera and Orthoptera.85 They are easy to
culture. There are no toxic metabolites from beauveria reported to enter the food chain
or accumulating in the environment.90 These features make B. bassiana a model system
for studying entomogenetics and effective biological control of pests.
B. bassiana produces several secondary metabolites of varied structures but the
contribution of these metabolites to pathogenesis is mostly unknown.91 Some
metabolites reported from B. bassiana include bassianin92 91, beauvericin93 93,
bassianolide91 92, beauverolide A94 94, oosporein95 95 and tenellin96 87.
Among these, bassianolide 92 has been identified in dead silkworm larvae
infected by B. bassiana.91 Beauvericin 93 is known to have mycotoxic properties.93 A
high molecular weight protein toxin Bassiacridin is stated to be isolated from B.
bassiana strain obtained from a locust.97 The Bristol polyketide group have identified
that tenellin 87 is not involved in pathogenicity.98 The present chapter involves work on
tenellin 87 and its biosynthesis.
25
Tenellin 87 is a prominent secondary metabolite of B. bassiana and its structure
was elucidated by Wat and co-workers, together with another similar compound
bassianin 91.96 Both compounds are known by their distinctive yellow colour apparent
during fermentation and in organic extracts. Tenellin 87 possesses a 5- substitiuted 2
pyridone ring with an acylated moiety at C-3.99
Tenellin 87 has been the focus of many biosynthetic studies over a period of
many years. The most distinctive reason has been that it is formed from a combination
of an amino acid and a polyketide chain.99
The key work to determine the precursors of tenellin 87 was reported by
McInnes and co-workers.100 They used [1, 2-13C2]-acetate 96, L-[methyl-13C]
methionine 97, (±)-[1-13C] phenylalanine and (±)-[2-13C] phenylalanine 98. The results
analysed by
13
C-NMR indicated that C-2, C-3 and C-7 to C-14 of tenellin 87 were
alternately enriched with doubly labelled acetate 96 and both methyl groups at C-15 and
C-16 showed enhanced peaks for labelled
13
C-methionine 97. The carboxy carbon C-1
of phenylalanine 98 forms C-4 of tenellin 87 and C-2 of phenyl alanine 98 becomes C-6
of tenellin 87. There is an intramolecular rearrangement of phenylalanine which causes
migration of the carboxy carbon adjacent to the aromatic ring forming C-4 of 87 and the
alpha carbon of phenylalanine separates to form C-6 of tenellin 87 (Scheme 2.1). They
confirmed that tenellin 87 is formed by condensation of methylated polyketide chain
having five acetates with an entity comprising all carbons from phenylalanine.100
Scheme 2.1: Incorporation of labelled acetate, methionine and phenylalanine in tenellin 87.
26
Another similar study by Leete and coworkers101 supported the intramolecular
rearrangement of phenylalanine in tenellin 87. They fed phenylalanine labelled at two
carbon positions [1, 3-13C2]. They indicated in
13
C-NMR spectrum additional satellite
peaks adjacent to singlet peaks of C-4 and C-5 due to spin spin coupling confirming
intramolecular rearrangement of the phenylalanine side chain in tenellin 87. They
argued that had there been ‘intermolecular’ movement of carboxyl group, the 13C-NMR
would only show singlets peaks for both C-5 and C-4.
Further isotope feeding on tenellin 87 was carried out by Wright et al.102 They
also used singly and doubly labelled sodium acetate, labelled methionine and three
different labelled forms DL-[carboxy-13C], DL-[α-13C] and L-[15N] phenylalanine. Their
results supported previous feeding studies by McInnes.100 Incorporation of N-labelled
phenylalanine established that it is fused to the polyketide chain with no loss of
nitrogen. In addition they also showed severely reduced incorporation of radioactive L[U-14C] tyrosine suggesting that tyrosine is not a direct precursor of tenellin 87.
Wright and coworkers proposed a route for tetramic acid formation. First,
condensation of phenylalanine 99 with the ten carbon polyketide chain 100 and then
hydroxylation of the aromatic ring by an oxygenase enzyme give quinomethine 101.
This would then undergo rearrangement of the tetramic acid to form the pyridone of
tenellin 87 (Scheme 2.2).
27
Scheme 2.2: Proposed tenellin 87 biosynthesis by Wright et al.102
Another biosynthetic proposal suggested by Cox and O’Hagan was that
phenylalanine 99 rearranges early and then condenses with a polyketide 100 to give the
six membered pyridone directly (Scheme 2.3).103 They synthesized and fed DL-[3-13C]
and [3-14C]-3-amino-2-phenylpropionic acids 102 to the cultures of B. bassiana.
However, the
13
C NMR analysis of this experiment did not show that 3-amino-2-
phenylpropionic acid 102 is a genuine intermediate in tyrosine 87 biosynthesis.
Scheme 2.3: Proposed biosynthesis by Cox and O’Hagan.103
28
On the basis of the Wright et al.102 hypothesis, Moore et al.99 synthesized acyl
tetramic acid in two isotopically labelled forms [4-13C] in 103 and [phenyl-2H5] in 104
and carried out feeding experiments with B. bassiana fermentations. They found that
these compounds were not incorporated into tenellin 87 and there was no proof for its
presence in B. bassiana extracts. They observed a single minor metabolite 105, the
purified sample of metabolite on 1H NMR showed it to be para-substituted aromatic
moiety without N-hydroxylation. They concluded that para-hydroxylated acyl tetramic
acid emerged as a late intermediate and a precursor or a reduced metabolite in tenellin
biosynthetic pathway (Scheme 2.4). They also carried out isotopic feeding of tyrosine
13
DL-[3-
C] and [1-13C] phenylalanine and indicated that both tyrosine and phenylalanine
are efficiently incorporated into tenellin 87.
By this result, in contrast to the hypothesis of Wright et al.,102 Moore and coworkers proposed that phenolic hydroxylation in tenellin 87 is introduced by tyrosine
and not at a later stage modification of the aryl ring.
Scheme 2.4: Labelled acyl tetramic acid feeding in tenellin 87 by Moore et al.99
In recent years with advancement in molecular biology and discovery of new
tools to manipulate DNA, investigations of biosynthetic routes of natural compounds
from their specific gene clusters have been very remarkable. Lately the Bristol
Polyketide Group have carried out quite a number of successful genetic studies on
polyketides genes of tenellin 87 from B. bassiana 110.25. They discovered specific
29
genes responsible for different stages and intermediate steps in tenellin biosynthesis. All
these stages reflect cryptic programming of PKS genes.
Eley et al. identified a hybrid PKS-NRPS gene cluster from the genomic DNA
of B. bassiana and showed by gene knock out experiments that this cluster is involved
in tenellin 87 biosynthesis.98 Analysis of this cluster revealed four open reading frames
(ORF) (Figure 2.2). BLAST analysis revealed that ORF1 and 2 were homologous to
cytochrome P450 enzymes. ORF3, a putative Zn dependent oxidoreductase,104 showed
high homology to enoyl-reductase (ER) enzymes. ORF4 consists of an approximately
12-kb biosynthetic gene that encodes β-ketoacyl synthase (KS), acyl transferase (AT),
dehydratase (DH), CMeT (methyltransferase), β-ketoacyl-reductase (KR) and acyl
carrier protein (ACP) domains typical of a fungal iterative type I PKS, followed by
condensation (C), adenylation (A), thiolation (T) and putative thiol (R ester reduction)
domains, characteristic of an NRPS module. A directed gene knockout (KO) experiment
confirmed ORF4 to be involved in tenellin production, and ORF4 was therefore
renamed tenS (tenellin synthetase). In this work they also proved that B. bassiana tenS
KO and WT strains are equally pathogenic towards insect larvae suggesting that tenellin
87 is not involved in insect pathogenesis of B. bassiana.98
Figure 2.2: Tenellin gene cluster identified by Eley et al.98
They proposed a pathway for the early stages of tenellin 87 biosynthesis from its
respective PKS and NRPS enzymes (Scheme 2.5). The double methylated pentaketide
106 bound to ACP is synthesised by TENS PKS and the A domain of TENS NRPS first
activates tyrosine 107 by adenylation and transfer to the thiol group of the T domain.
The polyketide and the amino acid are fused by the C domain to form N-β-ketoacyl
amino thiolester 108. The reduction domain R carries out reduction of the thiol ester
108 with the help of NADPH and release in the form of a peptide aldehyde 109. The
aldehyde can cyclise to form pre-tenellin 110, which was assumed to be a precursor of
tenellin 87 (Scheme 2.5).
30
Scheme 2.5: Tenellin 87 pathway proposed by Eley et al.98
Halo et al. expressed the tenS gene encoding tenellin synthetase (TENS), in
Aspergillus oryzae M-2-3.104 It led to the production of three new compounds, identified
as acyl tetramic acids, prototenellin A 111, prototenellin B 112 and prototenellin C 113.
Protenellin C 113 was not fully characterized due to the low concentration of the
purified compound (Table 2.1). These compounds didn’t have the methylation pattern
of tenellin 87 and the polyketide chain length in prototenellin B 112 was shorter than
tenellin 87. In addition there were double bonds between C-11 and C-12 of prototenellin
A 111 and between C-9 and C-10 of prototenellin B 112 whereas this bond is always
saturated in 87. This shows that the enoyl reductase domain present within the TENS
protein is defective and fails to carry out reduction in the first cycle of the polyketide
chain formation. These results depict that enzymes in tenS gene when expressed on its
own lose the ‘fidelity’ in the programming of the polyketide side chain of the
compound.
Halo et al. also carried out another important experiment which was coexpression of tenS with the gene encoding the enoyl reductase enzyme (ORF3) which
led to the production of single acyl tetramic acid compound, pretenellin A 114 (Table
2.1). This compound was concluded to be a genuine precursor to 87, possessing the
31
same methylation and chain length. Pretenellin A 114 has a saturated bond in the first
keto chain extension similar to tenellin 87 proving that the enoyl reduction is carried out
by ORF3 and not by ER in the TENS protein. This result showed that in the presence of
the ORF3, the tenellin gene cluster undergoes its normal pattern of bond formation and
methylation of the polyketide unit leading to correctly programmed compound
structure. Similar stand-alone enoyl-reductase encoding genes like lovC and apdC in
PKS gene clusters are reported from lovastatin 47105 and aspyridone A80 84
respectively, where ER domain present in the PKS synthase is inoperative. The ORF3
gene in tenellin synthase is known as tenC.
The first precursor compound pretenellin A 114 produced from coexpression of
TENS and TENC was different than pre-tenellin 110 hypothesized by Eley et al.98 in
being hydroxylated at C-4 (Table 2.1). Halo et al. described that the C-terminal R
domain of the NRPS after releasing thiolester 108, does not undergo a reductive
reaction at this step but must catalyse a Dieckmann cyclisation of the N-β-ketoacyl
amino thiolester 108 directly to form the tetramic acid, pretenellin A 114 (Scheme 2.6).
This concept was further supported by similar result reported by Sims and Schmidt.106
They carried out in vitro experiments, reacting purified proteins from R domains of
equisetin synthetase (EqiS) with synthetic substrate analogues. They obtained equisetin
tetramic acids and did not observe any reduced or aldehyde intermediates thus giving
evidence for Dieckmann cyclisation activity of R domains of EqiS. Tang et al. also
produced in vitro a 3-acyltetramic acid preaspyridone A 224 by incubating Aspergillus
nidulans PKS-NRPS encoding gene apdA with its enoyl reducatse encoding gene apdC
in the absence of any oxidative enzymes, proving that its R domain also actually
catalyses a Dieckmann cyclisation.107
Scheme 2.6: Release and formation of pretenellin A 114 by Dieckmann cyclase domain of TENS.
32
In another study Halo et al. revealed late stage oxidations during the
biosynthesis of the 2-pyridone tenellin 87 in B. bassiana by a combination of gene
knockout, gene silencing by antisense RNA and gene coexpression studies.108 They
concluded that the putative cytochrome P450 oxidase encoded by tenA catalyzes the
oxidative ring expansion required to convert the tetramic acid of pretenellin A 114
(Table 2.2) to the pyridone of tenellin 87. Gene knockout and silencing of tenB
produced pretenellin B 115, which confirmed that tenB catalyzes N- hydroxylation of
115 (Table 2.2) to form 87. The tenB gene encodes a rare kind of cytochrome p450
which N-hydroxylates only 2-pyridones and not tetramic acids. Another experiment
confirmed the N-hydroxylation function of tenB when (tenA + tenB + tenS) were
coexpressed in A. oryzae producing pretenellin B 115.
The above studies on the tenellin gene cluster revealed the highly programmed
nature of the PKS-NRPS proteins. After successful studies on the tenellin 87 gene
cluster, Heneghan et al.109 carried out screening of several other species of Beauveria
bassiana to search for compounds similar to tenellin 87. Their aim was to compare new
gene clusters and their sequences of proteins to that of tenellin 87. After analysing
various B. bassiana species they found and characterized the similar compound
Desmethyl bassianin 88 (DMB) from B. bassiana strain 992.
Desmethylbassianin (DMB) 88 differs from tenellin 87 in having a single
methylation in its side chain and it also has an additional chain extention. Heneghan et
al. identified the DMB gene cluster by Southern Blot by using tenS probe (Figure 2.3).
The biosynthetic gene cluster of DMB 88 had 90% sequence identity to the tenellin 87
gene cluster.109 It has the same four open reading frames dmbA, dmbB, dmbC and dmbS
(which is the PKS-NRPS).
Figure 2.3: DMB gene cluster identified by Heneghan et al.109
33
To evaluate and confirm that dmbS is responsible for DMB 88 production,109
they carried out a double silencing and knockout strategy to disrupt the dmbS gene and
the results showed total loss of production of any DMB 88 or related compound. Similar
results to tenellin 87 were obtained when dmbS was expressed in A. oryzae (M-2-3)
producing protoDMB-B 116 and protoDMB-C 117 which cannot be regarded as
precursor of DMB 88. When dmbS was co-transformed with dmbC, it gave the preDMB
A 118 (Table 2.3) giving the same methylation and chain length pattern as DMB 88. It
confirmed that in the presence of dmbC, dmbS undergo correct reduction in the
polyketide chain of the DMB compounds, show high conformity in programming and
the yield of compounds is also increased.
In both tenellin 87 and the DMB 88 gene clusters there is more than 90%
similarity between the tenellin and DMB PKS proteins but still both these compounds
were different in their PKS chain length and methylation pattern. After successful
expression studies in both individual gene clusters, it was possible to create hybrid
tenellin and dmb genes expression in Aspergillus oryzae to know which proteins is
responsible for difference in programming in closely related gene clusters.
Heneghan et al. carried out some co-expression experiments. These showed that
DMBC and TENC are interchangeable; the programme of the PKS is influenced by tenS
and dmbS. This led to the idea of ‘fidelity’- the extent to which the PKS makes
‘mistakes’. When dmbS and dmbC were co-transformed in A. oryzae, it gave preDMB
A 118, while cotransformation of dmbS with tenC again gave preDMB A 118 (Table
2.4). This indicated that tenC and dmbC had identical effects in assisting correct
programmed compounds production. Coexpression of tenS with dmbC gave pretenellin
A 114 which is identical when tenS is expressed with tenC. These results showed that
the PKS controls the programming of polyketide chain in the presence of either of the
trans-acting ER proteins.109
In the absence of the trans acting ER encoded by tenC or dmbC the PKS
displays low fidelity. But when the trans-ER is present the PKS displays high fidelity
and high productivity.
In another experiment Heneghan and coworkers created a hybrid gene consisting
of tenSPKS with dmbNRPS to know the effect of NRPS in programming.109 This swap
produced prototenellin A 111, prototenellin B 112 and prototenellin C 113 (Table 2.4).
This means that the NRPS does not have an effect in PKS programming but it only
34
functions to connect the amino acid to the polyketide chain and plays role as off-loading
mechanism for the PKS.109 This swap produced the same compounds when tenSPKS
was expressed in A. oryzae (M-2-3).104
The productive heterologous gene expression, silencing and gene knockout
studies in DMB 88 and tenellin 87 PKS-NRPS genome resolved the characteristic
function of their respective genes. The oxidative enzymes encoded by tenA, tenB and
dmbA, dmbB carry out important ring expansion and N-hydroxylation steps. The enoyl
reductase enzymes tenC and dmbC were observed to play crucial part in production of
correctly programmed precursor compounds pretenellinA 114 and preDMB 118
(Scheme 2.7). Knocking out the hybrid PKS-NRPS genes dmbS and tenS completely
eliminated the production of tenellin and DMB in their respective fungi.
Scheme 2.7: Individual domains in TENS and DMBS proteins producing precursor compounds.
Recently Fisch and colleagues revealed the catalytic role undergone by
individual domains in DMBS and TENS proteins. Their work also solved the important
queries about the methylation and chain length difference in tenellin 87 and DMB 88
structure.110
They took tenS as a host gene and exchanged its constituent domains with
domains from dmbS one at a time in each experiment and expressed it in A. oryzae with
35
tenC. They started initially by replacing the KS-AT domain of tenS by dmbS and then
the DH in a second experiment until the entire tenS PKS gene was swapped over with
domains from dmbS PKS.
Scheme 2.8: Key metabolites desmethyl pretenellin A 119, preDMB A 118 and prebassianin 120 produced in different
domain swaps between TENS and DMBS proving catalytic role of CMeT and KR domains in programming.
No change in programming was detected by including KS-AT-DH domains
from DMBS and the clones still produced pretenellin A 114. Variations in programming
36
were observed when CMeT and KR domains were included in TENS. (KS-AT-DHCMeT) domains from DMBS produced a new compound desmethyl pretenellin A 119
possessing pentaketide side chain length of pretenellin A 114 but a single methylation in
the PKS chain (Scheme 2.8). This indicated that the CMeT from DMBS brings about
single methylation pattern of DMB 88. In another swap by adding KR in the TENS host
together with DMBS (KS-AT-DH-CMeT-ER) the clone produced the hexaketide
predmbA 118, proving that KR is the chain length determining enzyme. Another swap
supporting this result was observed when only KR from DMBS was located in TENS.
This clone produced a compound prebassianin 120 possessing hexaketide chain length
of predmbA 118 but with double methylation in PKS side chain (Scheme 2.8). This
study revealed that KR and CMeT domains exhibit the major part in selecting number
of methylation and chain elongation in iterative HR-PKS NRPS.110
The potential and diversity of secondary metabolites production from B.
bassiana was further examined by treating this fungus with epigenetic modifying
chemicals.111 B. bassiana strain 110.25 fungus was grown in the presence of genetic
modifiers, 5-azacytidine (5AC) 121 and suberoyl bis-hydroxamic acid (SBHA) 122
(Scheme 2.9).
Epigenetic modifiers are small chemicals that bring about modification of gene
activation and expression without modifying its nucleotide sequence.
Histones are the main protein component of chromatin providing a framework
for the DNA around which it winds and forms a structure. In addition chromatin also
exhibits an important role in gene regulation. It affects gene expression by removing
acetyl groups from its N-acetyl lysine amino acid with the help of deacetylase enzymes.
This deacetylation makes the DNA wrap around histones more firmly leading to
compressed chromatin which makes the genes inactive. Some chemicals can act as
histone deacetylase inhibitors such as suberoyl bis-hydroxamic acid (SBHA) 122. These
inhibitors block this action of deacetylase enzymes which increase lysine acetylation
leading to activation of silenced genes.
The expression of genes in cells is also dependent on DNA methylation. This
involves addition of methyl group to cytosine or adenine of DNA nucleotide. This
methylation inactivates the expression of certain genes. 5-azacytidine (5AC) 121 is a
chemical analogue of cytidine which is a nucleoside present in DNA and RNA. 5AC
37
can bring about changes in the cell genome and its behaviour by removing methyl
groups from the DNA thus activating the silenced genes. There a number of studies in
which epigenetic modifiers stimulated silent gene clusters in fungi producing new
compounds and adding variety in the library of natural products.112
Scheme 2.9: New compounds produced by B. bassiana WT after growing with epigenetic chemicals 5AC and SBHA.
Yakasai et al. reported that B. bassiana showed three times higher production of
tenellin related compounds when B. basssian WT cultures were grown in the presence
of 5AC 121 and SBHA 122.111 They also produced new compounds 3’,4’-antipyridomacrolidin-A
123,
3’,4’-syn-prepyridomacrolidin-A
124,
3’,4’-syn-
prepyridomacrolidin-B 125, and reprogrammed compounds prototenellin A 111 and
protenellin E 126 possessing different methylation pattern than pretenellin A 114
(Scheme 2.9).
In the same study two different clones tenA-silenced strain and tenB-silenced
strain were grown in the presence of epigenetic chemicals. The tenA-silenced strain
38
produced new compounds 12-hydroxy pretenellin A 127 and 128 (both syn and anti
diastereomer), 14-hydroxy pretenellin A 129 (Scheme 2.10) and reprogrammed
compound protenellin E 126 but not any pyridone compounds. This showed that
silenced genes of tenA were not switched on by these chemicals.
Scheme 2.10 New compounds produced by tenA aRNA transformant after growing with epigenetic chemicals 5AC and
SBHA.
tenB silenced clones in the presence of 5AC 121 and SBHA 122 produced a
variety of different new compounds (10, 11-Z ) pyridovericin 130, (10, 11-Z )-syn-13hydroxy pretenellin B 131, (10, 11-Z )-anti-13-hydroxy pretenellin B 132, prototenellin
F 133, anti-13, 15-dihydroxy pretenellin B 134, syn-13-hydroxy pretenellin B 135 and
anti-13 hydroxy pretenellin B 136. In this, epigenetic modifiers overcome the silencing
factor of tenB producing tenellin 87 which is an N-hydroxylated compound. These
chemicals demonstrated the rich tendency of PKS-NRPS genes for crafting new
compounds (Scheme 2.11).
39
Scheme 2.11: New compounds produced by tenB aRNA strain after growing with epigenetic chemicals 5AC and SBHA.
2.2
Aims and Objective of the Chapter
Tenellin 87 is an example of a secondary metabolite in entomopathogenic
fungus Beauveria bassiana, the biosynthesis of which is composed of highly
programmed steps directed by an organized enzyme complex. This enzyme complex is
encoded by a hybrid gene cluster of iterative polyketide synthases-non ribosomal
peptide synthetases (PKS-NRPS).
The modern tools of biotechnology help to manipulate and study genes in fungi.
These techniques include different gene expression in host fungi and silencing or
knock-out of target genes in native organisms. The results of these techniques are
examined with a number of the latest laboratory instruments particularly Liquid
Chromatography-Mass Spectrometry. New chemical compounds are detected and their
structures are elucidated.
Different molecular genetic techniques combined with
chemistry apparatuses help to explore the relation between compounds and their
40
corresponding proteins and genes. These help plot the biosynthetic routes of natural
product compounds.
The various genetic studies on tenellin 87 and DMB 88 PKS-NRPS gene
clusters discussed in section 2.1 encouraged further transformation experiments in A.
oryzae. The objective of this Chapter is to analyse various strains prepared from
heterologous expression of tenellin 87 PKS genes and hybrid co-expression of tenellinDMB genes in A. oryzae. The hybrid tenellin and DMB genes co-expression was carried
out in quest to know the difference in programming of both tenellin and DMB gene
clusters. We also intended to analyse tenC-aRNA silenced clones of B. bassiana carried
out using amyB promoters.
All these strains were grown under standard fermentation conditions and their
organic extracts were examined using LC-MS which displays UV diode array and mass
spectrometry data of the compounds in the crude extract. Any interesting compounds
detected in these extracts were purified using preparative HPLC and the isolated pure
compounds were subjected to various structure techniques including IR, NMR, HRMS
and X-ray diffraction.
2.3.0
Results
2.3.1
Heterelogous Expression of tenS in A. oryzae M-2-3
The effective tenS gene knock out experiment in B. bassiana reported by Eley et
al.98 resulted in loss of production of tenellin 87. This generated interest to explore the
function of TENS PKS-NRPS protein. Halo et al. carried out heterologous expression
of tenS in A. oryzae M-2-3 using the pTAex3 expression system.104 Three new
compounds were produced from this experiment which were prototenellin A 111
(C21H23NO4, m/z 353), prototenellin B 112 (C18H19NO4, m/z 313) and prototenellin C
113 (C21H25NO6, m/z 387). Prototenellin C 113 was not fully characterized previously,
due to lack of material and instability during purification.104
The experiment was repeated to characterize prototenellin C 113. Spores of A.
oryzae pTAex3-tenS were first grown on plates (DPY media) for 7-10 days. A spore
solution was made using sterile deionized water and 1 ml of spore solution was added in
41
100 ml liquid culture in 500 ml Erlenmeyer flasks (10 × 100 ml) for 7 days with 200
rpm, at 25 °C. After completion of fermentation the cultures were homogenized and
extracted with ethyl acetate (section 4.11). The ethyl acetate extracts were dried and
evaporated to yield a crude extract of 70 mg, which was dissolved in HPLC methanol
(10 mg/ml). This extract was analyzed by LCMS (Method 1, section 4.4). Analytical
LCMS showed peaks of prototenellin C at Rt 29.4 mins (Figure 2.4). ES+ and ESshowed masses of 388 and 386 respectively indicating a mass of 387 corresponding to
prototenellin C 113 (Figure 2.5). This peak was purified by mass-directed preparative
LCMS. 26 Injections were made using a 20 minute program (Method 1, section 4.5).
The purified fractions of prototenellin C were collected and dried under nitrogen gas
into pale yellow solid (2.4 mg/L).
Figure 2.4: Diodearray chromatogram of tenS expression in A. oryzae producing prototenellin C.
42
Figure 2.5: ES-, ES+ and UV spectrum of prototenellin C 113.
2.3.1(a)
Characterization of Prototenellin C 113
The pure prototenellin C 113 was dissolved in deutrated methanol and 1D and
2D NMR experiments were carried out using a 500 MHz spectrometer. The 1H NMR
showed two prominent sets of doublets (Figure 2.6) which are characteristic of the parasubstituted hydroxy phenol found in tenellin and related compounds. In the 1H-1H
COSY spectrum, the two diastereotopic protons H-16a and H-16b showed correlation
with H-5 which is a typical pattern of a tetramic acid (Figure 2.7). This was further
confirmed in the HMBC spectrum where the benzylic protons at H-16 showed two and
three bonds correlation to the phenolic protons and also to the H-5 proton of the
tetramic acid ring (Figure 2.7). Three methyl signals corresponding to H-13, H-14 and
H-15 were present in the alkane region. The HMBC correlation of H-15 with C-7 and
quaternary carbon C-6 confirmed methyl H-15 to be attached to C-7 indicating that the
polyketide chain of prototenellin C possess the same methylation pattern as
prototenellin A 111. HMBC signals at H-10 showed the occurrence of two hydroxyl
groups at H-11 and H-12. The signals at H-10 and H-14 are two overlapping doublets
and singlets respectively, providing evidence that the compound is a mixture of two
diastereomers. Collective NMR data confirmed prototenellin C structure to be 113. The
molecular formula and molecular mass was confirmed by HRMS to be C21H25NO6.
43
Fig 2.6: 1H NMR spectrum of prototenellin C 113 in methanol-d4.
Figure 2.7: 1H-1H COSY (solid lines) and 1H-13C HMBC correlations (arrows) in prototenellin C 113.
2.3.2 Heterelogous Expression of tenSPKS-dmbNRPS in A. oryzae M-2-3
Prototenellin C 113 was also produced from another strain tenSPKS-dmbNRPS
cloned in A. oryzae (This transformation was done by Katherine Williams in the School
of Biological Sciences). A. oryzae clones were grown in liquid culture (10 × 100 ml)
for 7 days, 200 rpm, at 25 °C. After fermentation the cultures were homogenized and
extracted with ethyl acetate (section 4.11). The ethyl acetate extracts were dried,
evaporated (yield was 137 mg) and dissolved in HPLC methanol (to a final
44
concentration of 10 mg/ml). This extract was analyzed by LCMS. The LCMS analysis
showed peak of prototenellin C. The ES+ and ES- showed obvious peaks of 386 and
388 respectively corresponding to the 387 mass of prototenellin C. This peak was
purified by mass- directed preparative LCMS. The purified fraction of prototenellin C
were combined and dried under nitrogen gas to give a pale solid mass of 9.6 mg.
1
H NMR was carried out from this purified fraction on a 500 MHz spectrometer.
The 1H NMR showed the same proton signals as observed from the Prototenellin C 113
obtained from tenS expression clone (Figure 2.8).
Figure 2.8: Overlay of 1H NMR of prototenellin C 113 from A, A. oryzae pTAex3-tenS expression clone; B, tenSPKSdmbNRPS clone.
2.3.3
Analysis of A. oryzae dmbS–tenC Expression Clone
Different hybrid combinations of DMB 88 genes and tenellin 87 genes were
expressed in A. oryzae (This transformation was done by Mary N. Heneghan in the
School of Biological Sciences, University of Bristol) in order to investigate their mode
of programming in terms of chain length and methylation. Many different transformants
were produced. This section is an explanation of analysis of the LCMS of different
transformants and isolation of new compounds.
45
An A. oryzae dmbS–tenC transformant was grown in CMP liquid culture (10
flasks × 100 mL) at 25 °C at 200 rpm. The fermentation cultures were homogenized and
extracted as described in section 4.11. A crude extract of 111 mg was obtained and was
made into a solution of 10 mg/ml in HPLC methanol. This extract was analyzed by
analytical LCMS. A prominent peak at Rt 19 mins was observed (Figure 2.9). Mass
analysis indicated ES+ and ES- of 368 and 366 respectively (Figure 2.10). This peak
was purified with 25 preparative runs on LCMS (Method 1, section 4.5). The pure
compound was collected in a number of different tubes. They were all collected and
dried under nitrogen gas in the form of bright yellow powder (23 mg).
Figure 2.9: Diode array chromatogram of A, dmbS-tenC expression in A. oryzae; B, Aspergillus oryzae Wild type.
Figure 2.10: ES+, ES- and UV spectrum of the 19min peak.
46
2.3.3(a)
Characterization of preDMB A 118
The high resolution mass spectrum (HRMS) of this fraction gave a molecular
formula of C22H26NO4 (observed 368.1851; calculated 368.1856 for M[H]+). The pure
compound was dissolved in deuterated methanol and 1D and 2D NMR experiments
were done on the purified fraction. The 1H NMR showed two separate doublets in the
aromatic region for H-19/23 and H-20/22 and diasteorotopic protons H-17a and H-17b
characteristic of benzylic protons of a tetramic acid (Figure 2.11). The diastereotopic
protons showed COSY connection with methine proton H-5 and HMBC correlation to
the C-19/23 of the phenol ring which confirms the tetramic acid structure (Figure 2.12).
There were two methyl signals, one for terminal H-15 and second for H-16. The HMBC
and COSY correlations confirmed that the pendant methyl at position 16 is attached to
C-13 (Figure 2.12). Six olefinic protons H-7, H-8, H-9, H-10, H-11 and H-12 in the
alkene region of the spectrum confirmed three double bonds typical for the DMB 88
polyketide chain. In the COSY spectrum the olefinic protons showed couplings of H-7
to H-8, H-9 to H-10 and H-11 to H-12. The alkene proton H-7 displayed a two bond
correlation to the quaternary carbon C-6 in HMBC. All 1D and 2D NMR data
confirmed the structure to be 118 which was named preDMB A.
47
Figure 2.11: Spectrum of 1H NMR of preDMB A 118 run in methanol-d4.
Figure 2.12: 1H-1H COSY (solid lines) and 1H-13C HMBC correlations (arrows) in preDMB A 118.
2.3.4
Heterologous expression of dmbS-dmbC in A. oryzae
The clone dmbS + dmbC gene (This transformation was done by Mary N.
Heneghan in the School of Biological sciences, University of Bristol) was analyzed
having both dmbS synthases and dmb enoyl reductase encoding genes cloned in A.
oryzae. It was grown in liquid culture (10 flasks × 100 ml) at 25 °C at 200 rpm. After
completion of fermentation the cultures were homogenized and extracted with ethyl
acetate (section 4.11). The ethyl acetate extracts were dried, evaporated (yield was 363
mg) and dissolved in HPLC methanol (10 mg/ml). The LCMS analysis showed peak of
48
predmbA at Rt 19 mins. The ES+ and ES- showed masses 368 and 366 respectively
which corresponds a mass of 367 of preDMB A 118. The peak was purified by massdirected preparative LCMS. This purification was accomplished by 18 injections on 20
minutes program with Method 1 described in section 4.5. The tubes containing purified
fractions were all collected and dried. The dried preDMB A was bright yellow with
yield of 17.6 mg.
The 1H NMR showed the same proton signals as the preDMB A obtained from
dmbS+ tenC clone as illustrated in Figure 2.13.
Figure 2.13: Overlay of 1H NMR of preDMB A 118 from A, dmbS-dmbC clone; B, dmbS-tenC expression in A. oryzae.
2.3.5
Heterologous expression of tenSPKS – dmbC in A. oryzae
In the course of investigating the biosynthetic potential of the tenellin 87
polyketide synthase-nonribosomal synthatase encoding genes, we now know that TENS
in the absence of tailoring enzymes is capable of producing tetramic acids prototenellin
A 111, protenellin B 112 and protenellin C 113 possessing different methylation and
reduction pattern than tenellin 87. When tenC is expressed with tenS, the ‘correct’
precursor pretenellin A 114 is produced. These results encouraged us to further expand
49
the gene expression experiments using the tenellin gene cluster. We now aimed to
express the HRPKS of TENS without its counterpart NRPS in A. oryzae. This
expression also included the enoyl reductase encoding gene dmbC from DMB cluster.
As the NRPS module affords a peptide or an amino acid, in its absence we
expected that the ACP bound pentaketide 106 synthesized by TENSPKS can either get
hydrolysed and released in the form of an open chain pentaketide 137 (Scheme 2.12) or
it can enter another chain extension cycle by addition of a malonyl CoA 25 by the KS
forming hexaketide 138. The hexaketide 138 can offload from the TENS by cyclisation
to form a pyrone 139 (Scheme 2.12).
Scheme 2.12: Possible compounds from expression of tenSPKS+ dmbC in A. oryzae.
Similar pyrone structures are also reported from other polyketide synthase
enzymes from other fungi. Tang et al. investigated the programming role of HRPKSNRPS encoding gene apdA involved in aspyridone A 84 biosynthesis in A. nidulans.107
He manipulated the megasynthase ApdA in different in vitro assays and reported
production of a number of α-pyrones. The in vitro assay of ApdA without the enoyl
reductase ApdC produced pentaketide and hexaketide pyrone 142 and 139 respectively
(Scheme 2.13, A). The NRPS module of ApdA is very specific to recognize only fully
reduced tetraketide synthesized from the ApdA PKS. This might be the reason that
unsaturated tetraketide in the absence of ApdC enters further chain extension cycles and
methylations and get offloaded by the ACP in the form of pyrones without being
processed by the NRPS module. Another tetraketide 140 and pentaketide 141 pyrones
50
without any methylations is observed in in vitro assay where methyltransferase domain
is removed from ApdA. The in vitro assay of ApdA-PKS with ApdC independent of the
NRPS module also produced a pyrone 143 with saturated linear chain similar to
aspyridone A 87.
Kennedy et al. reported two pyrones 144 and 145 when they over expressed
lovastatin polyketide synthase LNKS in Aspergillus nidulans without the enoyl
reductase LovC (Scheme 2.13, B).71, 105
A
B
Scheme 2.13: A, α-pyrones reported from different in vitro assays with PKS-NRPS megasynthase ApdA of aspyridone
84 by Tang et al.107; B, pyrones reported from LNKS, LovB from lovastatin gene cluster.
2.3.6
Analysis of tensPKS–dmbC transformants
A total of 11 transformants were analysed (this transformation was carried out
by Dr. Walid Bakeer). They were grown in the same fermentation conditions for 7 days
and extracted with ethyl acetate (see section 4.11). The crude extracts were all in range
of 5 – 10 mg per 100 ml of liquid media. Each crude extract was made 10 mg/ml in
HPLC methanol and analysed by LCMS. In seven transformants we observed two
prominent peaks at 7.5 and 8.1 minutes (Figure 2.14) in a 15 minute program (Method
51
3, section 4.4). These were not observed in the A. oryzae wild type (M-2-3) strain. The
mass spectra of these two peaks were similar to the open chain polyketide 137, we
expected from this experiment (Scheme 2.12). The peak at 7.5 minutes showed mass of
213 in ES+ and 211 in ES- and was named compound A (Figure 2.15). The peak at 8.1
minutes gave a mass of 211 in ES+ and 209 in ES- and was named compound B (Figure
2.16). Neither compound showed a strong uv absorption.
Figure 2.14: A, LCMS chromatogram of tenSPKS-dmbC producing two new compounds at 7.5 mintues and 8.1
minutes; B, Wild type A. oryzae (M-2-3).
We thought there might be new polyketide compounds and thus planned to
purify these two peaks. These compounds were produced in low titre from each 100 ml
liquid culture. We chose one transformant producing this compound and grew large
scale fermentation of three litres liquid medium. The total crude extract was 239 mg. A
50 mg/ml sample was made in HPLC methanol for purification. About 24 runs of a 20
min program as stated in Method 2 (section 4.5) were carried out. The purified
compound A with mass (MH+) 213 was 9.5 mg and the second compound B with mass
(MH+) 211 was 9 mg.
52
Figure 2.15: A, ES- of compound A; B, ES+ of compound A; C, wavelength.
Figure 2.16: A, ES- of compound B; B, ES+ of compound B; C, wavelength.
2.3.6(a)
Identification of Compound A
Compound A had a chemical formula of C11H17O4 (observed 213.1130;
calculated 213.1121 for M[H]+) on High Resolution Mass Spectrometry. The 9.5 mg of
pure compound was dissolved in 0.65 ml of deuterated chloroform. The structure of the
53
compound was elucidated using 1D and 2D 1H and 13C NMR spectroscopic analysis. A
number of 1D and 2D NMR experiments were carried out on 500 MHz NMR
Spectroscopy. The structure of the compound A 146 is given in Figure 2.17.
The carbon spectra revealed the presence of eleven carbons, which included two
carbonyl groups at δC 176.4 (C-11) and δC 170.1 (C-9), four methylene groups in the
alkane region of the spectra and one terminal methyl carbon at δC 14.4 (C-1) (Figure
2.18). The 1H-13C HSQC gave eight protonated carbons. The 1H NMR showed two
distinct doublets in the alkene region at δH 5.95 and δH 6.54 which were assigned to two
geminal methylene protons of H-10. One of the H-10 methylene protons shows 1H-13C
HMBC correlation to an alkene carbon at δC 136.1 which was assigned C-8. Both
geminal methylene protons (H-10) show HMBC connections to carbonyl carbon at δC
170.1 (C-9) and sp3 carbon at δC 45.2 (C-7). The triplet at δH 3.65 was assigned to
methine proton (H-7) (Figure 2.17) which show a 1H-13C HMBC connection to terminal
carbonyl of a carboxylic acid at δC 176.4 (C-11), carbonyl carbon at δC 170.1 (C-9) and
an alkene carbon at δC 136.1 (C-8) (Figure 2.19).
Figure 2.17: 1H NMR of compound A run in chloroform-d.
54
Figure 2.18: 13C NMR of compound A run in chloroform-d.
The methine proton at δH 3.65 (H-7) show 1H-1H COSY connections to two
geminal methylene protons at δH 2.01 and 2.55 (H-6). Both geminal methylene protons
of H-6 show 1H-1H COSY and 1H-13C HMBC connections to a methine proton at δH
4.43 (C-5) linked to an oxygen atom (Figure 2.19). The chemical shifts of δC 79.6 for C5 support its link to an oxygen atom. The HMBC correlations of this compound
provided valid evidence for a pyran structure attached to a carboxylic acid at C-7. The
methine protons at δH 4.43 (H-5) and methylene protons at δH 2.01 and 2.55 (H-6) show
1
H-1H COSY connections to two other geminal protons at δH 1.65 and 1.81 (H-4). Two
multiplets at δH 1.37 and δH 1.46 were assigned to two geminal methylene protons H-3
which display 1H-1H COSY
link to one of the methylene at δH 1.8 (H-4). The broad
signal at δH 1.35-1.39 was assigned to methylene protons H-2 showing connections to δC
27.8 (C-3) in HMBC spectrum and to terminal methyl at δH 0.92 (H-1). Both HMBC
and COSY correlations show an aliphatic chain (C1-C4) attached to the pyran at
methine proton H-5.
55
Figure 2.19: Important 1H-1H COSY and 1H-13C HMBC connections in compound A 146.
Figure 2.20: Arrangement of protons in the pyran ring of 146 around the stereo centre at C-7 and C-5.
In the pyran ring of compound 146, the spatial orientation of the protons at the
two stereocentres C-5 and C-7 was established with the help of calculating the coupling
constant of H-5 and H-7 with their adjacent methylene protons at δH 2.01 (H-6a) and
δH 2.55 (H-6b). The coupling constant of 6 Hz between H-5 and H-6b indicated that H6b is equatorial. The J value of 10 Hz between H-5 and H-6a was 10 Hz, placing H-6a
in axial position. The coupling constant between the two geminal methylene protons H6a (2.00) and H-6b (2.55) was 12 Hz, which confirmed the conformation of H-6a to be
axial and H-6b as equatorial. The methine proton H-7 displayed J value of 12 Hz
between H-6a and 9 Hz with H-6b. These values depicts that H-7 orientation is axial.
The predicted conformation for the pyran of compound 146 from J values is given in
Figure 2.20 showing H-6a, H-7 and H-5 in axial direction and H-6b to be aligned in
equatorial position. Further confirmation of these spatial arrangements of atoms came
from 1D NOE, where both H-7 and H-5 show NOE correlations to each other which can
occur when both protons are in axial position (Figure 2.21 and 2.22).
56
Figure 2.21: In 1D-NOE, on irradiating H-5, signals of H-7 is observed in close proximity.
Figure 2.22: Irradiation of H-7 in 1D NOE, signals of H-5 are observed.
2.3.6(b) Identification of compound B
The structure of compound B 147 was interpreted with the help of HRMS and
NMR analysis and by comparison to compound A 146. The chemical formula given by
HRMS was C11H14O4 (observed 233.0799; calculated 233.0784 for [M]Na+). The 9 mg
of the pure compound was dissolved in 0.65 ml of deuterated chloroform. The ESI gave
57
idea that this compound may have the same structure with two protons less than
compound A 146. It was further confirmed by different NMR experiments.
The 13C NMR displayed the presence of 11 carbons including two carbonyls at
δC 169.5 (C-9) and 171.7 (C-11), a methine carbon at δC 80.8 (C-5) attached to an
oxygen atom, a methyl group at δC 14.0 (C-1), three methylene groups and four olefinic
carbons at 125.0 (C-7), δC 128.5 (C-8), δC 133.6 (C-10) and δC 153.5 (C-6) (Figure
2.23).
Figure 2.23: 13C NMR of compound B 147 run in chloroform-d.
The 2D HSQC showed 7 protonated carbons. In the 1H NMR, the broad doublet
signal downfield at δH 7.96 was assigned to the methine (H-6) (Figure 2.24) attached to
the olefinic carbon at δC 153.5 (C-6) in the 1H- 13C HSQC. In 1H-13C HMBC spectra, H6 shows linkage to methine carbon at δC 80.8 (C-5) and to the olefinic carbon (C-7) at
δC 125.0. This confirmed the presence of a double bond between C-6 and C-7, which is
the difference in structure between compound 146 and 147. H-6 also shows HMBC
correlations with the olefin carbons, C-8 and C-11 (Figure 2.25). The two broad signals
at δH 6.79 and δH 7.19 were identified as geminal methylene protons H-10 showing
correlations to the methine protons H-5 and H-6 in 1H- 1H COSY and 1H-
13
C HMBC
connections to the olefin carbon at δC 125.0 (C-7) and to the carbonyl group at δC 169.5
(C-9) (Figure 2.25).
58
Figure 2.24: 1H NMR of compound B run in chloroform-d.
The triplet signal at δH 4.99 was assigned to the methine H-5, displaying HMBC
correlations with the methylene groups at δC 27.3 (C-3) and δC 33.1 (C-4) and to olefin
carbons at δC 125.0 (C-7) and δC 153.5 (C-6). The two multiplet signals at δH 1.71 and
δH 1.79 were assigned to geminal methylene protons (H-4) linked to C-2 and C-3 at δC
22.6 δC 27.3 respectively in HMBC. The terminal methyl signal at δH 0.92 (H-1)
showed HMBC connections to the methylene groups at δC 22.6 (C-2) and δC 27.3 (C-3).
The 1H-13C HMBC and 1H- 1H COSY shows an aliphatic chain comprising of C-1 to
C-4 attached to the methine proton H-5. From NMR analysis, this compound was
elucidated to be a carboxylic acid pyran ring attached to an aliphatic chain and possess a
double bond between C-6/C-7.
Figure 2.25: Important 1H-1H COSY and 1H-13C HMBC connections in compound B 147.
59
2.3.6(c)
Discussion
The TENS-PKS synthesizes a polyketide chain from condensation of five
acetate units in four cycles, carrying out methylation and reduction in the first cycle and
another methylation in second cycle producing an open chain pentaketide 106. The
structures of compounds 146 and 147 were different than the compounds 137 and 139,
we proposed to be produced from this transformation. We concluded that they are not
products of TENS-PKS pathway.
Both compounds 146 and 147 have not been reported before. We searched for
similar compounds in the literature and came across a number of similar compounds
(Figure 2.26). Horhant et al. isolated three paraconic acids, protolichesterinic acid 148,
lichesterinic acid 149 and roccellaric acid 150 from lichen, Cetraria islandica (L.)
Ach.113 Dahiya and Tewari characterized three plant growth factors from the fungus
Alternaria brassica, one of them was 3-carboxy-2methylene-4-pentenyl-4-butenolide
153. They reported that 153 reduces plant growth by causing chlorosis.114 The same
structure 153 was reported by Park et al.115 They named it methylenolactocin 153 (αmethylene-γ-lactone) and obtained it from a penecillium sp.24-4 (FERM P-9437). They
stated that methylenolactocin 153 possess antimicrobial activity against Gram positive
bacteria. Huneck and Höfle isolated and characterized δ-lactone structures acaranoic
acid 151 and acarenoic acid 152 from the lichen Acarospora chlorophana.116
Figure 2.26: Similar compounds to A 146 and B 147, reported in literature.
60
Seshime et al. over expressed a type III PKS gene csyB in A. oryzae (M-2-3)
under the effect α-amylase promoter and produced a novel metabolite csypyrone B1 154
which is a 3-(3-acetyl-4-hydroxy-2-oxo-2H-pyran-6-yl) proponoic acid (Scheme
2.14).117 In a [1, 2-
13
C2] feeding, csypyrone B1 154 showed incorporation of five
acetate units which confirmed it to be a product of a PKS pathway. They presented a
proposed biosynthetic pathway for csypyrone B1 154 in which a succinyl CoA
condenses with three malonyl CoA and by pyrone ring cyclization form csypyrone B1
154.
Scheme 2.14: Proposed biosynthetic pathway for csypyrone B1 154.
It seemed highly unlikely that compound A 146 and compound B 147 could be
product of the tenellin PKS. We thus re-examined wild type A. oryzae. Although peaks
for 146 and 147 could not be observed by uv, their characteristic masses could be
detected at the correct retention times using a more sensitive 60 minutes programme
with Method 5 described in section 4.4 (Figure 2.27). We proposed a hypothetical
pathway for the biosynthesis of compound 146 and 147 (Scheme 2.15).
Figure 2.27: Section of Single ion monitoring chromatogram of compound A in ES+ and ES- mode observed at 29.1
minutes in A. oryzae tenSPKS-dmbC and Wild type A. oryzae.
61
Scheme 2.15: Proposed biosynthetic pathway for compound A 146 and compound B 147.
2.3.7
tenC RNAi Silencing in B. bassiana with amyB promoter
Messenger RNA is an important type of ribonucleic acid (RNA) and it possesses
an important role in gene expression. During gene expression, the DNA molecule
transcribes the genetic information required for a protein synthesis to a single strand of
messenger RNA. This is carried out by RNA polymerase enzymes in a process called
transcription. The messenger RNA travels outside the nucleus carrying the genetic code,
which is then translated into proteins with the support of ribosomes and transfer RNA
by an organised cell process called translation.
Figure 2.28: Process of gene expression from DNA to proteins.
62
Transcription is a process which can be interrupted by RNA interference
(RNAi). RNAi is an effective tool in eukaryotes often used in recent advances in
molecular genetic techniques.118 It is used to diminish the function of a specific gene by
destroying the messenger RNA translation stage of the target gene; this can reduce, or
completely prevent, protein production. In this technique a single stranded RNA
comprising of reverse sequence or complementary sequence to that of target gene is
introduced into the cell. This single strand binds to the native single strand RNA to form
double stranded RNA. The double stranded RNA is considered as abnormal condition
by the cell and is then degraded by enzymes called dicer (Figure 2.29). The number of
native RNAs expressed into proteins becomes less and hence eliminates the gene
product. This reverse sequence dependant genetic tool is also referred to as ‘gene
silencing’. RNA silencing does not diminish the target gene but it causes reduction of
the function of the gene so it is also named as a ‘gene knock down’ technique.119
This method helps to understand the product and function of the target gene and
aids in the investigation of the biosynthetic pathways of natural products and secondary
metabolites from their gene clusters. A lot of efforts are underway to apply this gene
silencing technique in curing many diseases by destroying function of genes which are
causal factor of viral or tumour diseases in humans.
The concept of RNA interference system came into limelight among researchers
when in a number of cases of introducing homologous RNA for high gene expression
actually diminished the outcome of the native gene unexpectedly.120 In 1990, Napoli
and Jorgensen in their attempt to investigate the enzyme responsible for colouration of
petunia petals, brought forward the hypothesis of RNA silencing for the first time.121
They overexpressed chalcone synthase in the native plant, which is the putative enzyme
for colouration in violet petunias. Instead of obtaining deep violet colour, the petals of
flowers produced were white. This led to the conclusion that the original gene of the
flower was suppressed by the transgene introduced. This suppressing factor was later
discovered to be interfering RNA in the cell which diminishes the expression of the
genes.
63
Figure 2.29: RNA silencing pathway.
In the course of transcription of genes, ‘promoters’ play a vital role in initiating
conversion of the structural gene to proteins. Promoters are regulatory regions
consisting of DNA sequences and are located upstream of the gene they transcribe. The
promoter provides a binding site for RNA polymerase (the enzyme responsible for
generation of mRNA) and for transcriptional factors (proteins that initiate RNA
64
polymerase). The transcription factors are responsible for activation or suppression of
transcription.
Figure 2.30: Promoters are responsible for initiating gene expression.
Similarly in designing gene silencing procedures, promoters are a key feature
which initiates the gene silencing pathway. There are different kinds of promoters used
in gene silencing depending on the goal of an experiment. Most commonly utilized are
constitutive and inducible promoters. Constitutive promoters always direct expression
of genes and are independent of environmental or endogenous factors in cells. The
activity of inducible promoters are dependent on external stimuli such as light,
temperature and different media sources like alcohol, different nutrients including
carbon or many herbicides and antibiotics. The use of inducible promoters makes it
possible to control the level and time of expression of target genes.
Fuji reported that the starch inducible α-amylase promoter (PamyB) is an
effective promoter for heterologous expression systems of Iterative fungal polyketide
synthases in Aspergillus oryzae host.122 Halo et al. successfully expressed hybrid
polyketide synthase – nonribosomal peptide synthatase (PKS-NRPS) encoded by tenS
in A. oryzae using PamyB.104 PamyB is induced (switched on) by starch and repressed
(switched off) by glucose. The activity of genes driven by PamyB can thus be controlled
by the addition of starch or glucose to growth media.
A good example of a strong constitutive promoter is A. nidulans PgpdA
(glyceraldehyde-3-phosphate dehydrogenase promoter). Halo et al. obtained productive
results by using PgpdA to carry out RNA silencing (iRNA) for knocking down the
function of two oxidase enzymes encoded by tenA and tenB in B. bassiana.108
65
The enoyl reductase encoding gene tenC is known to be vital for correct
programming of tenellin 87 production.104 Yakasai and colleagues carried out RNAi
silencing of tenC in B. bassiana with the constitutive PgpdA promoter.111 The B. bassiana
tenC RNAi transformant not only produced WT compounds tenellin 87, pretenellin A
114 and prototenellin D 155 but also reprogrammed compounds obtained in tenS
expression in A. oryzae, prototenellin A 111, prototenellin B 112. This silencing
experiment produced a new reprogrammed compound prototenellin E 156. The
presence of WT compounds in the culture show small varied level of TenC proteins
present in the transformant.
Figure 2.31: B. bassiana WT and reprogrammed compounds produced from RNAi tenC transformants by gpdA
promoter.111
The aim of the present project is to examine tenC RNAi transformants produced
under the effect of the inducible promoter amyB in B. bassiana. Here we design an
experiment to know whether the ‘concentration’ of TenC affects programming and new
or reprogrammed compounds are produced or not. The idea was to use an ‘inducible’
promoter ‘amyB’ the expression of which is dependent on the carbon source used in
liquid media. The expression of amyB is stimulated in the presence of maltose or starch
and is repressed by glucose.
66
The construction of the PamyB/tenC silencing vector and its transformation into
B. bassiana was done by Dr. Walid Bakeer. My role was to grow the six selected
silenced clones in different carbon sources and analyse the produced compounds.
The silenced clones (A, B, C, D, E and F) were grown along with wild type B.
bassiana using three carbon sources, mannitol, maltose and glucose.
2.3.7(a)
Growth of B. bassiana transformants in TPM (mannitol)
The spores of the six silenced clones (A, B, C, D, E and F) and wild type (WT)
B. bassiana were each grown in standard tenellin production medium (TPM) (see
section 4.8). In TPM, mannitol is used as the carbon source.
WT
A
B
C
D
E
F
Figure 2.32: tenC silencing clones and WT B. bassiana grown in standard TPM media.
The cultures were grown in 100 ml of TPM in 500 ml Erlenmeyer flasks. They
were incubated at 25 °C in shakers at 150 rpm. After ten days the cultures were filtered.
All cultures had varying degrees of yellow colour, which is an indication of tenellin
compounds. The mycelia in each flask were extracted in acetone (200 ml). The acetone
extract was concentrated under vacuum to a brown aqueous extract. It was further
diluted with deionized water (200 ml) and then extracted into ethyl acetate (200 ml).
The ethyl acetate layer was separated and dried with MgSO4. The extracts were
concentrated in vacuum to give a brown solid. All crude extracts of silenced clones
were in range of 5-8 mg and WT was 10 mg. A solution of 10 mg/ml for all extracts
was made with HPLC grade methanol and analysed with LCMS with Method 3
(section 4.4). The chromatograms of all six silenced clones and wild type are given in
figure 2.34 and 2.35.
67
The wild type B. bassiana produces four major compounds which are 15hydroxy tenellin 157 at 9.3 minutes, prototenellin D 155 at 10.0 minutes, pretenellin A
114 at 10.7 minutes and tenellin 87 at 11 minutes (Figure 2.33). The silenced clones A,
B, C, E and F still produce prototenellin D 155, pretenellin A 114 and tenellin 87. Clone
D produced only compound tenellin 87 and 157 (Figure 2.34 and 2.35). These
chromatograms showed that all the silenced clones still produce more or less the wild
type compounds. We did not observe any newly programmed or reprogrammed
compounds. This suggests that under these conditions tenC is not silenced and the amyB
promoter is inactive.
Figure 2.33: Diode array chromatogram of WT B. bassiana grown in TPM (mannitol) media showing production of 15hydroxytenellin 157 at 9.3 minutes, prototenellin D 155 at 10.0 minutes, pretenellin A 114 at 10.7 minutes and tenellin
87 at 11.0 minutes.
68
Figure 2.34: Diode array chromatograms of WT B. bassiana, Clone A, Clone B, Clone C grown in mannitol.
Figure 2.35: Diode array chromatograms of WT B. bassiana, Clone D, Clone E and Clone F grown in mannitol.
2.3.7(b)
Growth of B. bassiana transformants in maltose media
The next experiment was to grow the six silenced clones A, B, C, D, E, F and
wild type B. bassiana in TPM medium but using maltose (30 g/L) as carbon source
instead of mannitol. As amyB promoter is turned on when grown with maltose medium,
we expected strong silencing of tenC.
69
WT
A
B
C
D
E
Fig 2.36: tenC silencing clones and WT B. bassiana grown in maltose.
The six clones and WT B. bassiana were grown in the same conditions for ten
days at 25 °C at 150 rpm. All cultures were pale white colour, but produced mycelia.
The cultures were then extracted in the same way first with acetone, and then
concentrated and later diluted with deionized water and in the last extracted with ethyl
acetate. All dried extracts weighed from 28-34 mg from 100 ml liquid medium. The
concentrated extracts (10 mg/ml) in HPLC methanol were analysed in LCMS. The
chromatograms of silenced clones with WT B. bassiana are given in Figure 2.37 and
2.38.
Figure 2.37: Diode array chromatograms of WT B. bassiana, Clone A, Clone B, Clone C grown in maltose.
70
Figure 2.38: Diode array chromatograms of WT B. bassiana and tenC silencing Clone D, Clone E and Clone F grown in
maltose.
In maltose media the B. bassiana WT and tenC silenced clones did not produce
tenellin related compounds strongly. Only wild type produced small peaks of 15hydroxy tenellin 157 and prototenellin D 155. Tenellin 87 at 11 minutes was not
observed, the ESI only showed mass MH+ 354 at 11.08 minutes which may be
pretenellin B 115 but the uv was not strong or convincing. All the silenced clones failed
to produce compounds, even the wild type compounds were not detected. So, this
indicates that although amyB promoter is believed to be activated when there is maltose,
if mannitol is not used as the basic carbon source B. bassiana fails to produce any
compounds.
2.3.7(c) Growth of B. bassiana transformants in glucose media
After growing B. bassiana clones in maltose, the next experiment was to grow
them in TPM medium using glucose as carbon source (20 g/L). As the amyB promoter
is turned off when grown with glucose medium, we expected to observe an absence of
or weak silencing of tenC.
71
WT
A
B
C
D
E
F
Fig 2.39: tenC silencing clones and WT B. bassiana grown in glucose.
The six clones and WT B. bassiana were grown in the same conditions for ten
days at 25 °C at 150 rpm. They all showed very faint colour which show poor
production of tenellin compounds. The cultures were extracted first with acetone, then
concentrated and diluted with deionized water and in the last extracted with ethyl
acetate. The dried extracts weighed 9-10 mg from 100 ml liquid media. The
concentrated extracts were made 10 mg/ml in HPLC methanol and analysed in LCMS.
The chromatograms of silenced clones with WT B. bassiana is given in Figures 2.40
and 2.41. In glucose medium the WT and all six silenced clones failed to produce any
tenellin related compounds.
Figure 2.40: Diode array chromatograms of WT B. bassiana and tenC silencing Clone A, Clone B and Clone C grown in
glucose.
72
Figure 2.41: Diode array chromatograms of WT B. bassiana and tenC silencing Clone D, Clone E and Clone F grown in
glucose.
2.3.7(d)
Growth of transformants in mannitol in combination with 1%
maltose
In the previous three experiments we observed that B. bassiana produces
tenellin 87 and related compounds only when mannitol is used in TPM medium. In
maltose and glucose even the WT could not produce. Here we used mannitol with 1%
maltose to see if maltose makes silencing in the presence of mannitol. The WT B.
bassiana and silenced clones were grown in the same conditions as explained in section
4.10. The crude extracts were made 10 mg/ml in HPLC methanol and analysed by
LCMS. The diode arrays of all extracts are given in Figure 2.42 and 2.43. In all
chromatograms wild type compounds, 15-hydroxy tenellin 157, prototenellin D 155,
pretenellin A 114 and tenellin 87 were produced.
73
Figure 2.42: Diode array chromatograms of WT B. bassiana and tenC silencing Clone A, Clone B and Clone C grown in
mannitol with 1% maltose.
Figure 2.43: Diode array chromatograms of WT B. bassiana and tenC silencing Clone D, Clone E and Clone F grown in
mannitol with 1% maltose.
2.3.7(e)
Growth of transformants in mannitol in combination with
1% glucose
The WT and silenced clones of B. bassiana were grown on mannitol with 1%
glucose to see the effect of glucose on production of compounds. The WT B. bassiana
and silenced clones were grown in the same conditions as explained in section 4.10. The
74
crude extracts were made up to 10 mg/ml in HPLC methanol and analysed by LCMS.
The diode arrays of all extracts are given in figure 2.44 and 2.45. Wild type compounds
157, 155, 114 and 87 are observed in silenced clone A, B and D. Clones C, E and F also
produced more and less WT compounds.
Figure 2.44: Diode array chromatograms of WT B. bassiana and tenC silencing Clone A, Clone B and Clone C grown in
mannitol with 1% glucose.
Figure 2.45: Diode array chromatograms of WT B. bassiana and Clone D, E and F grown in mannitol with 1% glucose.
75
2.3.7(f)
Growth of transformants in mannitol with combination of
5% maltose
In this experiment the WT and silenced clones of B. bassiana were grown in mannitol
with increased percentage of maltose, 5%. The crude extracts were made 10 mg/ml in
HPLC methanol and analysed by LCMS. The chromatograms of all extracts are given in
Figure 2.46 and 2.47. WT B. bassiana produces only compound protenellin D 155 and
tenellin 87 when percentage of maltose is increased. The silenced produce WT
compounds 157, 155 and 87 in different concentrations.
Figure 2.46: Diode array chromatograms of WT B. bassiana and Clone A, B and C grown in mannitol with 5 % maltose.
Figure 2.47: Diode array chromatograms of WT B. bassiana and Clone D, E and F grown in mannitol with 5 % maltose.
76
2.3.7(g)
Growth of transformants in mannitol with combination of
5% glucose
In last experiment the WT and silenced clones of B. bassiana were grown on
mannitol with increased percentage of glucose 5%. The crude extracts were made 10
mg/ml in HPLC methanol and analysed by LCMS. The chromatograms of all extracts
are given in figure 2.48 and 2.49. Wild type compounds prototenellin D 155 and tenellin
87 are produced in some clones.
Figure 2.48: Diode array chromatograms of WT B. bassiana and Clone A, B and C grown in mannitol with 5 % glucose.
Figure 2.49: Diode array chromatograms of WT B. bassiana and Clone D, E and F grown in mannitol with 5 % glucose.
77
These experiments showed that silencing a gene with a promoter whose function
is dependent on the carbon source was not successful in this case. B. bassiana will
always require mannitol in TPM media for production of its primary and secondary
metabolites.
We suggest that if we want tenC silencing in varying degrees in B. bassiana, we
may use another kind of promoter, the expression of which is not dependent on carbon
source for example, alcohol dehydrogenase PalcDH which is stimulated by glycerol or
lactose in the media.
2.4
Conclusions
Heterelogous expression of tenellin genes alone and co-expression with DMB
genes was effectively accomplished in A. oryzae (M-2-3) and their chemical products
were isolated.
The structure of prototenellin C 113 was elucidated and fully characterized.
Prototenellin C along with prototenellin A 111 and prototenellin B 112 were produced
from two clones; A. oryzae tenSPKS-NRPS and A. oryzae tenSPKS-dmbNRPS. The
TENS PKS-NRPS in absence of enoyl reductase TenC produce re-programmed or
unusual tetramic acids. The pattern of reduction, methylation and in case of 112, even
chain length was deviated than tenellin 87. This shows that TENS PKS controls the
programming of the polyketide chain while NRPS proves to possess a broader substrate
specificity to accept altered PKS chain from the TENS in case of 111, 112 and 113. The
NRPS deliberately serves its role to select and combine tyrosine with the PKS chain and
provide offloading mechanism for the chemical product. The production of 111, 112
and 113 from A. oryzae tenSPKS-dmbNRPS verifies successful development of a
Hybrid PKS-NRPS system, with proteins obtained from two different fungal strains
working together admirably. This paves way for further manipulation of PKS-NRPS
systems and exploring their enzymes for obtaining desired compounds. The yield of
prototenellin C 113 from A. oryzae tenSPKS-dmbNRPS was more (9.6 mg/L) than 113
obtained from A. oryzae tenSPKS-NRPS (2.4 mg/L), which further adds to the efficacy
of the hybrid TENS-PKS: DMB-NRPS system.
PreDMB A 118 was produced from A.oryzae dmbS –dmbC and A.oryzae dmbStenC clones. PreDMB A 118 has similar polyketide chain pattern as DMB 88. This
78
heterologous expression shows that like tenellin pathway, the DMB PKS-NRPS in the
presence of enoyl reductase DMBC, produced the correctly programmed precursor
compound preDMB A 118. The expression of DMBS with TENC again produced
preDMB A 118. This shows that the enoyl reductase does not play any role in
programming of the polyketide chain but the presence of enoyl reductase, either DMBC
or TENC, DMBS ‘does not’ loses the fidelity of programming and compounds with
correct methylation and chain length are formed.
Heterologous expression tenSPKS-dmbC in A. oryzae without the NRPS
produced two new compounds 146 and 147, but they were not the products of tenellin
pathway. We ascertained that they are native wild type A. oryzae compounds. This
demonstrates that expression of TENS PKS without NRPS is a challenging experiment
and requires a more efficient expression system.
The silencing of tenC with amyB promoter in B. bassiana was carried out with
the purpose to achieve different level of concentration of the enoyl reductase and obtain
new compounds. This experiment was unsuccessful as substituting the carbon source
(mannitol) with maltose and glucose to obtain induction and repression of silencing,
severely abrogated the production of tenellin or related compounds. We suggested that
alternative inducible promoters can be considered in future, the induction of which is
not dependant on carbon source.
79
Chapter 3
Investigation of the Role of Genes from the Aspyridone Pathway using
Heterologous Expression and Structural Elucidation of new
Compounds
3.1
Introduction
Aspergillus is a large genus of moulds which acquired its name because of its
distinctive spore bearing structure similar to an aspergillum (a holy water sprinkler). All
aspergilli have characteristic morphology consisting of a foot cell, elongated hyphae
called a conidiophore and a round vesicle bearing the asexual spores, the conidiospores
(Figure 3.1). In many species, the colour of the spores serves as identification, for
example Aspergillus niger produces black spores, Aspergillus ochraceus have yellow or
brown spores and the colour of spores from A. nidulans, A. fumigatus and A. flavus are
green.123
Because of their asexual airborne spores and their ability to grow with minimal
nutrients, aspergilli are widespread in all ecosystems.124 They mostly occur in terrestrial
habitats, soil and mostly on plant and animal debris. They are saprophytes. After they
get in contact to their food they first breakdown complex ingredients by secreting
enzymes and acids and then absorb the nutrients. The aspergilli play an important part
in decay and decomposition of organic matter driving carbon and other important
minerals back into the environment by natural recycling.125 They also provide a means
for supply of nutrients and food for a large number of other soil dwelling
microorganisms. Their ability of bio deterioration and degradation is a major problem in
spoiling foods, textiles, paper and even historic paintings.126
There are about 250 species in the genus Aspergillus. Many of the species aid in
fermentation and are important in the food industry. Since the early 20th century
Aspergillus niger has been used in the production of citric and gluconic acids and has
also been used in the pharmaceutical industry.127 Aspergillus terreus is used in the
production of synthetic polymer.128 Aspergillus oryzae is used in the production of rice
vinegars, soy sauce, alcohol beverages,130 kojic acid used in a range of Japanese food
80
and also in making koji and synthesis of flavour enhancers.131 Many aspergillus species
are important in providing commercial enzymes.132
Aspergillus species can be easily grown in the laboratory on simple organic
media and have been studied extensively by molecular biologists and in studies for
developing biotechnology tools. A. oryzae has been an effective host for the
heterologous expression of different biosynthetic gene clusters of many secondary
metabolites133 and A. niger is used as a host for construction of heterologous proteins.134
Figure 3.1: A, microscopic view of Aspergillus spores; B, A. oryzae; C, A. terreus.
Most Aspergillus species produce a large number of secondary metabolites and
many of them are important natural product drugs. Lovastatin 47, a cholesterol lowering
drug was isolated from A. terreus,135 cholecystokinin antagonist asperlicin 157 from A.
alliaceus,136 ion channel ligands137 and anti-fungal compounds 158 are reported from
aspergillus species.138
Besides many beneficial species of Aspergillus, there are many fungi which are
causal agents of human and animal diseases. They produce a number of mycotoxins139
mainly aflatoxin 48, patulin 159 and ochratoxin 160 which deteriorate stored seeds and
also cause loss of poultry and loss of domestic animals.140 The air borne spores may
cause respiratory tract disease like asthma, hay fever and cause a number of allergies.141
Aspergillosis caused mainly by A. fumigatus may become fatal in immunosuppressed
individuals.142
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3.2
Aspergillus nidulans
Aspergillus nidulans (also known as Emericella nidulans) propagates both by
asexual spores called conidia and sexual spores called ascospores. Conidia are produced
on specialized structures called coniodiophores and ascospores are grown inside round
sexual fruiting bodies called cleistothecia.143 A. nidulans has been used as a model
organism for over fifty years, on which various research areas of mycology, eukaryotic
cell biology and genetics were initiated. For example, the parasexual cycle was first
discovered and studied in A. nidulans by Pontecorvo and was used as a means to
produce strain of fungus with desired genetic traits before the advent of modern
biotechnology techniques.144 Ronald Morris analysed the genetics of mitosis by
studying this organism.145 A. nidulans has also been used as a model to study genetic
metabolic diseases, genetic recombination,146 explain intron splicing, chromatin, DNA
repair and regulatory pathways.123 It has a defined sexual cycle which is used as a guide
to investigate reproductive mechanism in other fungi where the sexual phase is not
clearly defined.
Figure 3.2: Aspergillus nidulans strain 2.2 grown on plate.
82
The genome of A. nidulans has been sequenced.147 It shows that the organism is
distantly related to A. oryzae and A. fumigatus. The genome sequence indicates the
presence of various secondary metabolite gene clusters, of which 27 are polyketide
synthases, 14 are nonribosomal peptide synthetases, 6 fatty acid synthases, one
sesquiterpene cyclase and two dimethylallyl tryptophan synthase genes.148
3.3
Aspergillus nidulans metabolites
The role of secondary metabolites in the life cycle of a fungus is considered
mostly ambiguous but they often exhibit important bioactivities; most importantly
antitumor, antibacterial, antifungal and other vital pharmaceutical properties.5
Aspergillus nidulans is a producer of a range of biologically active metabolites,
some of which are toxic. But the numbers of secondary metabolite genes predicted
from sequencing of A. nidulans genome are more than the reported metabolites from
this fungus.
There are a number of elements which govern the formation of these
metabolites. These include environmental factors for example temperature, light, pH
and more important are the availability of different nutrients, nitrogen and carbon
sources.143 Synthesis of some metabolites is also related to fungal vegetative growth and
morphology. Recently it has been reported that G-proteins regulate growth of asexual
spores as well as mycotoxin production.149 A main reason for the number of compounds
discovered being less than predicted from the genome sequencing is that many genes
encoding the biosynthesis of metabolites are silent under normal fermentation
conditions and they require signals or stimulants to be activated and expressed in the
form of natural products.
Many efforts over a long period of time have been made to isolate metabolites
from A. nidulans and identify genes involved in the biosynthetic pathway for the
characterized metabolites. Mostly this has been achieved by targeted gene deletion
assisted by sequence comparison of the genes with those from the known library of
metabolites.150 The genes involved in the biosynthesis of a particular metabolite are
usually clustered; this helped the natural product chemists to recognize boundaries for
83
genes encoding all essential enzymes and catalysing each step in the biosynthesis of the
natural product.
The data provided by the genome sequence of A. nidulans encouraged
researchers to devise methods to activate silent gene clusters by different genetic
engineering using modern molecular tools and advanced genomics. Considerable
success has been achieved in this regard with the discovery and over expression of
global regulators of secondary metabolism genes such as laeA151 and replacing pathway
specific transcription regulators with inducing promoters.152 Moreover, manipulating
chromatin modifying proteins and epigenetic modifiers has facilitated the up-regulation
of previously silent clusters of many bioactive compounds and has unveiled hidden
biosynthetic potential present in this fungus.153 A review of recognized metabolites of
A. nidulans is explained below.
The well-known β-lactam antibiotic compound penicillin 161 is produced from
A. nidulans. It is produced from three amino-acids: L-α-aminoadipic acid, L-cysteine
and L-valine. The biosynthesis of penicillin 161 is encoded by three genes namely,
acvA, ipnA and aat.154 Understanding the genetic and molecular regulation of penicillin
161 biosynthesis in A. nidulans can help devise ways to increase production of this
antibiotic.
Sterigmatocystin 164 is a carcinogen polyketide mycotoxin, produced by about
20 species of Aspergillus including Aspergillus nidulans.155,
156, 157
There is a great
concern of contamination caused by mycotoxins in food and feed products resulting in
health issues and huge economic loss. It also causes high level of genotoxicity in liver
samples. It was necessary to study the biosynthesis of sterigmatocystin 164 on
enzymatic level and control them by molecular studies. A lot of research had been made
to investigate the metabolic pathway of strigmatocystin 164 and determine its gene
cluster. Brown et al. identified a 60 kb cluster in A. nidulans comprising of 25 genes
reported to conduct all the steps essential for sterigmatocystin 164 biosynthesis.150 In
this cluster a NADPH dependent reductase gene stcU158 and a P450 monooxygenase
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gene stcS converts the intermediate compound versicolorin A 162 to sterigmstocystin
164159 and stcP encoding a methyltransferase carry out methylation of the intermediate
demethylsterigmatocystin 163 to form sterigmatocystin 164.160
Scherland and Hertweck identified four unique prenylated quinolone alkaloids,
aspoquinolones A-D 165, 166, 167 and 168 from Aspergillus nidulans (HKI 0410) by
growing the fungus in rice medium.161 They predicted that like other alkaloids, these
quinolones might have produced from its precursor compound anthranilic acid. The
presence of anthranilate synthase (AS) like genes in A. nidulans sequence supports this
assumption.
The conidia spore pigmentation in A. nidulans was proposed to be encoded by a
polyketide synthase gene wA. Heterologous expression of this PKS in A. oryzae
produced a yellow coloured, novel heptaketide naphthopyrone compound known as
YWA1 169. This compound was regarded as the intermediate in pigmentation of mature
green spores.162, 163
Fernandez et al. isolated shamixanthone 170, emiricellin 171, dehydroaustinol
172 and austinol 173 from Aspergillus nidulans. They reported that an essential 4’phosphopantetheinyl transferase (PPTase) encoding gene cfwA is required for the
production of secondary metabolites in A. nidulans particularly polyketides and NRPS
85
compounds. Xanthones are phenolic compounds and exhibit important biological
activities including antimicrobial, antioxidant, cytotoxic and neuropharmacological
activities.164
In recent years after genome sequencing of A. nidulans revealed that there are
many more secondary metabolite clusters present as compared to the isolated
metabolites form this fungus, a lot of efforts has been served to uncover factors
responsible for the repression of metabolites expression. Bok et al. detected a gene
called cclA, similar to an orthologous gene in S. cerevisae which is used in chromatin
mediated gene silencing by DNA modifications involving methylation of lysine of
Histones. The cclA deletion mutants in A. nidulans gave production of six aromatic
compounds not observed before in A. nidulans. They were monodictyphenone 174,
emodin 57 and emodin analogs 175, 176, 177 and 178 and two anti-osteoporosis
polyketides F-9775A 179 and F-9775B 180.153
Sanchez et al. reported two more xanthone compounds from A. nidulans
variecoxanthone A 181 and epishamixanthone 182. The gene cluster of xanthose
comprise of a cluster of ten genes including a PKS gene mdpG which is also involved in
monodictophenone 174 biosynthesis. They stated that 174 and emodin 57 are precursors
86
of prenyl xanthones. They also identified three prenyl transferase genes necessary for
encoding prenyl transferase part of the xanthone structures.165
Schroek et al. reported the production of the aromatic tetraketide orsellinic acid
56, a lichen metabolite lecanoric acid 183 and the two antiosteoporosis polyketides F9775A 179 and F-9775B 180 by growing A. nidulans in conjunction with a collection of
soil dwelling bacteria actinomycetes.166 They revealed that a NRPKS encoding gene
orsA (AN7909) is required for the biosynthesis of orsellinic acid 56, lecanoric acid 183,
179 and 180. Sanchez et al. confirmed orsA to be involved in orsellinic acid production
and in AN7909 deletion mutant isolated two bioactive aromatic compounds in A.
nidulans, gerfelin 184 and diorcinol 185.167
Nielsen et al. grew A. nidulans on eight different media and observed not only a
number of known metabolites but also arugosin A 186, arugosin H 187, antibiotic
compounds violaceol I 188 and violaceol II 189 not known before from this fungus.168
They obtained 32 PKS deletion mutants and linked violaceol I 188 to the orsA gene and
biosynthesis of arugosin 186 to the monodictophenone 174 gene mdpG.
87
Szewczyk et al. reported a new metabolite, asperthecin 192 by deleting a gene
SumO in Aspergillus nidulans which encodes a regulatory protein.169 They also
identified the gene cluster for this compound by a following a number of gene deletion
strategies. The gene cluster consists of an NR-PKS gene aptA, a gene aptB which
encodes a hydrolase and a monooxygenase gene, aptC. They proposed that the aromatic
structure 190 of asperthecin 192 is synthesized from one acetyl-CoA and seven
malonyl-CoA assembled by a NR-PKS, encoded by aptA. The PKS chain 190 is
hydrolysed by AptB into 191 and AptC carries out later oxidation to form asperthecin
192 (Scheme 3.1).
Scheme 3.1: Proposed biosynthetic pathway of asperthecin 192.
Bok and Keller identified a nuclear methyltransferase protein LaeA which
globally regulates transcription of secondary metabolites in A. nidulans.151 In the laeA
over expressed mutants, Bok and collegues (during genetic profiling of the mutants)
recognized an antitumor compound terrequinone 198 not reported before in A.
nidulans.170 They related a five open reading frame gene cluster to the biosynthesis of
terrequinone 198 named tdi. The cluster consists of a mono-modular NRPS encoding
gene tdiA encoding a protein comprising an adenylation domain, a thiolation domain
and a thioesterase domain but a condensation domain typical of NRPS was lacking.
88
Bouhired et al. proposed that the gene tdiD, which encodes a putative aminotransferase
is responsible for the deamination of L-tryptophan 193 to indole pyruvic acid 194, tdiA
encodes a protein which then adenylates the pyruvic acid 194 and dimerises it to a
quinone structure 195. A presumed prenyl transferase encoded by tdiB catalyses a first
prenylation to form 196 and then a reductase (tdiC) accomplishes hydroquinone
reduction 197 before a second prenylation (tdiE) to form terriquinone 198.171
Scheme 3.2: Proposed pathway of terriquinone A.
Emericellamide, an antibiotic known previously from marine emericella species,
is formed by the fusion of a polyketide and a nonribosomal peptide. Chiang et al.
identified emericellamide A 207 and its analogues in gene deletion studies of series of
NRPS sequences during genome mining experiments in A. nidulans. They also deduced
that emericellamides are synthesized from a gene cluster comprising of four contigious
open reading frames. The HR-PKS EasB forms a carboxylic acid polyketide chain 199
and is converted to CoA thiolester 200 by EasD (CoA ligase) and loaded on to the
acyltransferase (AT) of EasC, 201 (Scheme 3.3). The polyketide is then loaded to the
EasA which is an NRPS consisting of five modules. Each module of the EasA delivers
89
an amino peptide group to the growing chain, glycine and valine being the amino acid
provided in the first two cycles of the NRPS subsequently forming 202, 203, 204 and
205. At the end of the biosynthetic cycle the linear chain 206 is released, cyclised and
assembles to form emericellamide A 207 and its analogues C 208, D 209, E 210 and F
211.17
Scheme 3.3: Proposed biosynthetic pathway of emericellamides.
Dohren reviewed the non-ribosomal peptide synthetase encoding genes in A.
nidulans and listed 27 NRPS and NRPS related genes. He reported a number of peptide
90
and aminoacid metabolites reported from A. nidulans including echinocandin 212,
emericellamide 207, triacetylfusigen 214, fusarinine 215, terriquinone 198, emerin 213
and aspyridone 84.173
Wang et al. recognized a silent gene cluster in Aspergillus nidulans which
consist of two adjacent PKS genes, one a NR-PKS (afoE) and other a HR-PKS (afoG)
in the same cluster.152 They triggered the cluster by replacing a putative transcription
activator gene (afoA) with the inducible alcohol dehydrogenase promoter alcA. This led
to the production of a new metabolite asperfuranone 219 with subsequent gene deletion
experiments they identified five genes involved in its biosynthesis (Scheme 3.4). They
proposed a biosynthetic pathway for asperfuranone 219 which shows that the HRPKS
that (afoG) synthesizes a 3, 5-dimethyloctadienone moiety 216 from acetyl-CoA, three
malonyl-CoA and two S-adenosyl methionine (SAM). The 3, 5-dimethyloctadienone
216 is loaded on to the next NR-PKS (afoE) by the SAT domain. The NR-PKS extends
216 by condensing with four malonyl CoA and a SAM to form the first intermediate
217. A gene (afoD) encoding hydroxylase enzyme carries out hydroxylation at C-3 to
91
form 218. The afoF, encoding an FAD dependant oxygenase hydroxylates C-7 and afoC
encoding a hydrolase is involved in furan ring formation. A gene AN1030.3 encoding
an oxidoreducatse catalyzes the last reduction step forming asperfuranone 219 (Scheme
3.4).174
Scheme 3.4: Proposed biosynthetic pathway of asperfuranone.
3.4
Aspyridone pathway in A. nidulans
Hertweck and co-workers reported an 11.9 kilobase (kb), putative hybrid
polyketide synthase- nonribosomal peptide synthetase encoding gene in A. nidulans
which was named apdA.80 ApdA is homologous to TenS discussed in chapter 2. They
described the PKS-NRPS to be comprised of a number of domains, which are
ketosynthase (KS), acyltransferase (AT), ketoreductase (KR), dehydratase (DH), enoyl
reductase (ER), C-methyltransferase (C-MeT), acylcarrier protein (ACP), adenylation
domain (A), condensation domain (C), peptidyl carrier protein (PCP) and a reducase
domain (R). The apdA gene is clustered with a number of putative oxidoreductase
encoding genes; two of the genes encode cytochrome P450 monooxygenases and are
92
named as apdB and apdE and one is an FAD-dependent monooxygenases called apdD.
The PKS-NRPS is bordered downstream by a putative exporter gene apdF, an activator
gene apdR and an acyl-CoA dehydrogenase encoding gene apdG. The gene also has an
additional trans acting ER domain, apdC which is homologous to tenC. Hertweck et al.
believed this cluster to be silent as A. nidulans extracts grown in different laboratory
mediums do not contain any PKS-NRPS metabolites.80 They perceived that the
sequence of the activator gene apdR was similar to a transcription factor found in
Aspergillus fumigatus. Over-expression of apdR under the influence of the inducible
alcohol dehydrogenase promoter alcAp was achieved by homologous recombination in
A. nidulans. The mutant A. nidulans strain produced two new pyridone compounds
Aspyridone A 84 and Aspyridone B 226.
Scheme 3.5: Proposed biosynthesis pathway of aspyridone by Bergmann et al.80
93
Hertweck and coworkers proposed that the PKS synthesizes a tetraketide 220
from three malonyl CoA, an acetyl CoA and two SAM (S-adenosyl-methionine). The
enoyl reductase enzyme in ApdA was believed to be inactive and this function is
catalysed by the stand alone reductase protein ApdC acting in trans. This characteristic
is similar to the gene clusters of lovastatin, tenellin and desmethyl bassianin. 104,109,105
The tetraketide 220 fuses with the tyrosine 107 catalysed by the condensation domain to
form a hybrid polyketide-peptide 221. Based on knowledge at the time, it was assumed
that a reductive release was catalysed by the reductase domain to form an aldehyde
intermediate 222 which after Knoevenagel closure forms pyrrolinone 223. The
cytochrome P450 encoded by apdB was proposed to perform the first oxidation to form
the tetramic acid 224 and the second oxidase encoding gene apdE makes the
hydroxylation of the tetramic acid forming an intermediate 225. The oxidative ring
expansion from tetramic acid to 2-pyridone is also proposed to be catalysed by the P450
enzymes to form aspyridone A 84. The last step of phenol hydroxylation of aspyridone
to form aspyridone B 226 is performed by the FAD dependent monooxygenase apdD
(scheme 3.5).
Tang et al. confirmed the biosynthesis of the aspyridone pathway by
reconstructing the function of apdA and apdC in in vitro studies and S. cerevisiae was
used as the expression host. They reported the production of preaspyridone A 224 by
incubating purified apdA and apdC in the presence of co-factors and building blocks
(Figure 3.3, A). The production of 4-hydroxy preaspyridone proves that the hybrid Ltyrosine-tetraketide thiolester 221 after its release undergo ring closing by a Dieckman
cyclisation catalysed by the last C-terminal domain of the NRPS (Figure 3.3, B) and
disproves the presence of a reductase domain and consequent aldehyde intermediate 222
(Scheme 3.5). They highlighted the flexibility of the adenylation domain towards
different aromatic amino-acids by showing incorporation of L-tryptohan, L-4fluorophenylalanine and L-phenylalanine by apdA to form 227, 228 and 229.107
94
A
B
C
Figure 3.3: A, in vivo synthesis of preaspyridone A 224; B, Dieckmann cyclisation in preaspyridone A 224; C,
incorporation of different amino acid analogues by ApdA.
3.5
Objectives of the Chapter
Heterologous expression of biosynthetic genes in a foreign host, particularly
fungi, has been an important biotechnology tool to investigate the various steps in
biosynthetic pathways of novel natural products. It allows the determination of the role
of each gene in the gene cluster and is of particular use in the case of silent gene
clusters.175 Among different fungi, Aspergillus oryzae is considered as an effective
heterologous host, mainly because of its established use in producing large amount of
proteins.176
The Bristol Polyketide Group successfully studied the activities of a number of
HRPKS-NRPS gene clusters in detail by heterologous expression of the specific gene
clusters in A. oryzae, for example fusarin C 77,77 squalestatin 65,72 tenellin 87,104
desmethyl bassianin 88.109 These heterologous expressions experiments resolved the
methylation and chain length factors and cryptic programming between two similar yet
different compounds tenellin 87 and desmethyl bassianin 88.110 This encouraged us to
revisit and study the iterative PKS-NRPS gene cluster of aspyridone 84. We planned to
95
study heterologous expression of aspyridone biosynthetic genes in A. oryzae by using a
recently reported multiple gene expression plasmid system for
transformation.
A. oryzae
176
We also aimed to investigate and determine the function of each tailoring
enzyme, particularly the role of the different oxidative-enzyme-encoding genes in the
aspyridone gene cluster by expressing the genes in different groups in A. oryzae. These
experiments would also produce intermediate compounds and the structure elucidation
of these will aid in understanding the order of biosynthetic steps in the aspyridone
pathway.
The A. oryzae transformants obtained will be analysed. The new metabolites will
be isolated by preparative mass directed LCMS and their structures will be elucidated
and characterized. The stereochemistry of any crystallised compound will be studied
under X-ray crystallography. The chemical structure of a compound in a particular gene
expression will help to deduce the function and biosynthetic potential of the genes in the
cluster.
3.6
Heterologous Expression system used in fungal transformation
The number of secondary metabolite gene clusters annotated in A. nidulans
genome is more than the natural products known from the fungus. One of the reasons is
the mass metabolic background in the native fungus due to which many gene clusters
fail to express their products. A suitable way to determine the product of a secondary
metabolite encoding gene cluster is to express it in an appropriate host by fungal
transformation.178
Heterologous expression also helps in engineering fungal genes and provides an
alternative method for high production of novel metabolites which are formed in low
titre in their native host fungus. The first successful DNA-mediated fungal
transformation was reported in 1979.179,180 Since then, fungal transformation protocols
are still evolving and have improved through years but also have their limitations.
Fungal PKS genes encode large megasynthases and appropriate hosts with an
effective expression system are required for their expression. Quite a number of
organisms have been used for fungal PKS transformation181 for example yeast, bacteria,
plants and also fungi like A. nidulans and A. oryzae. For many PKS genes A. oryzae has
96
been a favourable host. This is because it belongs from the same fungal genus and due
to similar cellular mechanism the expressed genes function properly.122 A. oryzae has
the ability to translate mRNA from eukaryotes, carry out post-translational
modifications, produce large amounts of enzymes and secretes secondary metabolites in
the medium in high amount. It is among those fungal species which are generally
regarded as safe (GRAS) and they are easily fermented under simple laboratory
conditions. The genomic analysis as well as analysing the LCMS data of A. oryzae
transformants is easy to study. A. oryzae can also splice introns present in the PKS
genes.182
The heterologous expression plasmid used in fungal transformations usually
consists of the target genes themselves, suitable promoters and appropriate terminator
sequences and gene for the selection markers. A number of selection markers are used
in transformation, such as nutritional markers argB, amdS, pyrG or niaA used in
auxotrophic condition or different antibiotic resistance markers against hygromycin B,
phleomycin or benomyl.183 In our fungal transformations we will use the arginine
auxotrophic A. oryzae strain (M-2-3) as the host organism which is unable to grow in
the absence of arginine – i.e. on minimal media. The plasmid contains the argB gene of
A. nidulans. Thus transformed cells will be able to grow on minimal medium, whereas
the un-transformed cells will be unable to survive. In a standard fungal transformation
procedure the required genomic DNA is extracted from the subject fungus, amplified
and cloned in the respective plasmid used for the heterologous expression. The spores
of the host fungus are subjected to enzymatic treatment to break down the cell wall and
liberate protoplasts. The protoplasts are incubated with transforming DNA in a medium
containing CaCl2 and other additives and then grown on medium containing selective
nutrients allowing only heterologous transformants to grow (Figure 3.4).175
97
Figure 3.4: Steps in fungal transformation.
The PKS-NRPS encoding genes, like other fungal secondary metabolite genes,
exist in clusters and a simple precursor compound is synthesized by encoding of a
megasynthase enzyme accompanied by tailoring enzymes. This provoked biologists to
devise a multiple gene expression vector which can express four genes in a single
transformation experiment. This has been recently accomplished in the Lazarus group in
the School of Biological Sciences, University of Bristol.176 The vector comprises of
three constitutive promoters, PgpdA from A. nidulans, Padh (alcohol dehydrogenase) and
Peno (enolase) to express maximum of three tailoring genes from a gene cluster. For
expression of megasynthases like PKS-NRPS it consists of inducible PamyB (promoter of
taka-amylase coding gene in A. oryzae) and TamyB, the amyB terminator. The vector
98
contains of the argB gene as a selectable marker for expression in A. oryzae auxotroph
(M-2-3). This vector is termed as pTAYAGSargPage.
We planned to use the pTAYAGSargPage vector to transform the silent gene
cluster from the A. nidulans aspyridone pathway. The strong constitutive promoters
provided in this vector will allow the PKS-NRPS and the tailoring genes to trigger and
express into proteins.
Figure 3.5: Multiple gene expression vector pTAYAGSargPage used in fungal transformation.
3.7.0
Results
3.7.1
Heterologous expression of apdA and apdC in A. oryzae (M-2-3)
From in vitro studies by Tang et al.107 and the aspyridone pathway proposed by
Bergmann et al.,80 we knew that the megasynthase HRPKS-NRPS encoded by apdA
and the enoyl reductase apdC synthesize preaspyridone A 224, the first compound in
the aspyridone pathway. This step is similar to the biosynthesis of pretenelllin A 114104
and preDMB A 118109 in B. bassiana. So, to confirm this hypothesis in vivo we carried
out heterologous expression of apdA and apdC in A. oryzae. The cloning and fungal
transformation was done by Dr. Khomaizon Pahirulzaman from the School of
Biological Sciences, University of Bristol.
The pTAYAGSargPage vector was used to combine the iterative HRPKS-NRPS
apdA and apdC genes to form the vector pTAYAargAC. Transformation of this vector
into A. oryzae produced a number of transformants (denoted A. oryzae apdAC) and the
99
incorporation and expression of the genes apdA and apdC was confirmed by qRT-PCR.
The best producing transformants were then selected for chemical analysis and
identification and purification of novel compounds by LCMS and NMR. Wild-type A.
oryzae M-2-3 was grown in parallel in all experiments as a control.
Figure 3.6: Expression vector pTAYAargAC containing apdA and apdC.
A
B
Figure 3.7: A, A. oryzae mycelia in liquid media in a flask; B, A. oryzae apdAC expression clone on CDA plate.
The A. oryzae transformant was grown first on Czapek Dox agar (CDA) (Figure
3.7, B) and later on DPY solid media for maximum sporulation. The transformant was
grown on plates for 7-10 days. The mature spores were scratched with a sterile loop and
spores were collected in deionized water. 1 ml of the spore solution was added to 100
100
ml of CMP liquid media (see section 4.9) in a 250 ml baffled Erlenmeyer flask. The
cultures were grown at 28 °C in a shaker at a speed of 200-250 rpm for 7 days. The A.
oryzae transformants mycelia grow in a form of small balls (Figure 3.7, A). After 7 days
the cultures were removed from the shaker, and the entire fermentation mixture
(mycelia with the liquid) was acidified (pH= 3) and homogenized with ethyl acetate
(section 4.11). The organic extract was separated, then concentrated under vacuum on a
rotary evaporator and then the residue was defatted. The mass of the dried crude extract
was 40 mg (from 100 ml fermentation). A solution of 10 mg/ml of the crude extract was
made with HPLC grade methanol and 20 µl was injected and analysed by a Waters
2795HT HPLC system. It measures wavelength between 200 and 400 nm with a Waters
998 diode array detector and provides an electrospray (ES) mass spectrum with Waters
ZQ spectrometer sensing masses between the ranges of 150 to 600 m/z units.
The LCMS chromatogram of A. oryzae apdAC displayed two new peaks, one
minor peak at 13.2 minutes and the second major peak at 14.0 minutes which were not
present in the A. oryzae wild type used as a control (Figure 3.8). Both the peaks showed
m/z 332 [M]H+ and a λmax of 279 nm which is the same as described for preaspyridone
224107 by Tang et al. although they reported that only a single compound was produced
in their experiments. The ESI chromatogram showed two peaks of identical masses
indicating that they are probably isomers. To confirm the production of preaspyridone
224 and determine the two isomers we purified each compound for NMR structural
determination.
The purified minor (0.61 mg) and major (60.7 mg) components were dissolved
in deuterated chloroform (CDCl3) and 1D, 2D 1H NMR and
13
C NMR spectroscopic
studies were carried out. Both compounds were identified as preaspyridone A 224 and
the chemical shifts of the NMR spectra matched with those reported by Tang et al.107
The 1D 1H and 13C chemical shifts of both minor and major components in CDCl3 were
same with only negligible differences of 0.01ppm. But NMR spectra run in dimethyl
sulfoxide-d6 discovered key differences between the structures of the minor and major
compounds. Reinjection of the pure compounds showed that they did not interconvert.
101
14.0
ZW-II-92H
A
1.5e+2
AU
1.25e+2
3.45
1.0e+2
7.5e+1
2.65
5.0e+1
6.23
13.2
2.5e+1
0.0
-0.00
2.00
4.00
6.00
8.00
10.00
12.00
28.77
26.93
14.00
16.00
18.00
20.00
22.00
24.00
26.00
28.00
ZW-II-92M
3.22
B
6.0e+1
3.45
5.0e+1
Wildtype A. oryzae
AU
4.0e+1
5.80
3.0e+1
2.0e+1
4.25
1.0e+1
27.43
2.60
0.0
Figure 3.8: A, Diode array chromatogram of A. oryzae apdAC expression clone showing two new peaks at 13.2 and
14.0 minutes; B, Diode array chromatogram of Wild type A. oryzae used as a control.
A
100
B
330 [M]H
ZW-II-92H 246 (13.218)
100
ZW-II-92H 793
(13.200)
222
330 [M]H-
ZW-II-92H 262 (14.078)
279
apdA,C-repeat
ZW-II-92H 841 (14.000)
223
8.0e-2
2.4
2.2
7.0e-2
2.0
280
1.8
6.0e-2
1.6
AU
AU
5.0e-2
1.0
%
%
1.4
1.2
4.0e-2
3.0e-2
8.0e-1
6.0e-1
2.0e-2
331
300 305 313 316 321 325
332
2.0e-1
1.0e-2
341 346 352
331
362363 366 371376
332 [M]H+
0
297302 308309 315 317 320 325
ZW-II-92H 263 (14.105)
100
332
0.0
337
344 348
355
363 366 368
332 [M]H+
%
100
357
333 341
336 342
%
0
4.0e-1
333
358
365 370 373
334
Figure 3.9: A, ES+, ES- and UV spectrum of minor isomer at 13.2 minutes; B, major isomer at 14.0 minutes observed
in A. oryzae apdAC expression clone
3.7.1(a)
Identification of minor isomer of preaspyridone A 224
The minor compound eluting at 13.2 minutes was purified in the form of pale
crystalline solid. HRESIMS gave a molecular formula of C19H26NO4 (observed
332.1852; calculated 332.1856 for M[H]+).
102
30.00
The 13C NMR indicated the presence of 19 carbons which included two carbonyl
groups at δC 175.4 (C-2) and δC 191.9 (C-6), an enol group at δC 194.4 (C-4), a
quaternary carbon at δC 99.9 (C-3), a methine carbon at δC 62.3 (C-5), two sp3 carbons
at δC 33.4 (C-7) and δC 31.5 (C-9), two methylene groups at δC 39.6 (C-8) and δC 28.7
(C-10) and three methyl carbons at δC 16.7 (C-13), δC 18.9 (C-12) and δC 10.8 (C-11).
Figure 3.10: 1H NMR of minor component of preaspyridone A 224 run in DMSO-d6.
The 1D 1H NMR contains two distinct doublets in the aromatic region at δ H 6.88
(2H, H-16, H-20) and δH 6.59 (2H, H-17, H-19) (Figure 3.10) which is characteristic of
a para- substituted phenol ring and is further corroborated by 1H-13C HMBC correlation
of both the aromatic protons with the C-OH at δc 155.7 (C-18) (Figure 3.12). The two
singlets at δH 9.17 and δH 8.93 were assigned to the para- substituted hydroxyl group at
H-18 and to H-1 attached to a nitrogen atom respectively by HMBC correlations. The
multiplet peaks at δH 2.82 were assigned to the diastereotopic protons (H-14a, H-14b)
linked to the methine proton at δH 4.08 (H-5) in 1H-1H COSY (Figure 3.12) and to a
quaternary carbon at δC 125.3 (C-15), aromatic carbon at δC 130.6 (C-16/20) and
103
carbonyl group at δC 194.4 (C-4) in 1H-13C HMBC and confirms benzylic protons
linked to the pyrrolidine ring at C-5 (Figure 3.11).
Figure 3.11: Segment of 1H-13C HMBC NMR spectrum of minor isomer of preaspyridone A 224 showing correlations of
benzylic protons H-14 with C-5, C-16, C-15 and C-4.
The diastereotopic protons (H-14a, H-14b) and the methine proton H-5 are three
different nuclei (A, B, X) coupled to each other and have separate chemical shifts. Each
of these proton signals are doublets of doublets but the signals of H-14a and H-14b have
merged into each other creating distorted peaks typical pattern of an ABX system and
the signals at H-5 have combined to a broad peak (Figure 3.10). The quartet at δH 3.53
was assigned to H-7 linked to the carbonyl carbon at δC 191.9 (C-6), the methyl carbon
at δC 17.1 (C-13) and methylene group at δC 39.6 (C-8) in HMBC. The broad multiplet
between δH 1.24-1.27 was assigned to the methylene protons H-8 and to the methine
proton H-9. The multiplet at δH 1.09 and δH 1.26 corresponds to the geminal protons of
H-10. The doublet at δH 1.03 was assigned to the methyl group at C-13 exhibiting a
HMBC connection to C-6, C-7 and C-8. The terminal multiplet at δH 0.80-0.82 was
consigned to the last methyl H-11 and methyl group H-12 showing HMBC connections
to C-10, C-9 and C-8.
104
Figure 3.12: 1H-1H COSY (solid lines) and 1H-13C HMBC correlations (arrows) in preaspyridone minor isomer 224.
Crystals of the minor component 224 were formed in chloroform and were
analyzed on a Microstar X-ray instrument. The crystal data (ccdc code 941137) gave
the relative stereochemistry of the three chiral centres in minor preaspyridone as (5R,
7S, and 9R). The methyl groups at C-7 and C-9 possess anti-configuration in the crystal
data. Bergmann et al. reported syn configuration for both methyls in aspyridone A 84 2,
4-dimethylhexanoyl side chain.80
Figure 3.13: Crystal structure of the minor isomer of preaspyridone A 224.
3.7.1(b)
Identification of major isomer of preaspyridone A 230
The major compound eluting at 14.0 minutes was purified in the form of pale
brown solid. HRESIMS gave a molecular formula of C19H25NO4Na (observed
354.1677; calculated 354.1676).
The 13C NMR revealed the presence of 19 Carbons (Figure 3.15) having similar
chemical shifts as the
13
C NMR spectra of minor preaspyridone 224. The 1H NMR of
the major component was similar to minor preaspyridone A 224 in having similar
aromatic doublets at δH 6.59 (H-17, H-19), δH 6.90 (H-16, H-20) and the singlet at δH
9.17 (H-18) which is distinctive of para- substituted phenol in preaspyridone A. The 1H
NMR showed difference in the signals of three protons H-14, H-8 and H-13 (Figure
3.16). The splitting pattern of the diastereotopic protons H-14a and H-14b in major
105
preaspyridone 230 was different because both protons appeared at the same chemical
shift δH 2.82 as a doublet coupled to methine proton H-5 with J constant of 4.5 Hz,
unlike in minor preaspyridone 224 where both H-14 protons appeared at separate ppm
(Figure 3.10). The methylene group at H-8 in major preaspyridone 230 appeared as
geminal protons resonating at two separate chemical shifts δH 1.29 and δH 1.38. The
chemical shift of the methyl H-13 was also different as it produced a doublet at δH 0.95
(Figure 3.16) as compared to δH 1.03 in the minor isomer. From the above variations in
the 1H NMR and the crystal structure of the minor preaspyridone 224, we deduced that
both the minor and major compound observed in the apdAC expression clone, are
diastereomers of preaspyridone A being epimeric at C-5 (Figure 3.13). The 1H-
13
C
HMBC showed similar correlations as observed in minor preaspyridone 224 (Figure
3.14).
Figure 3.14: 1H-1H COSY (solid lines) and 1H-13C HMBC correlations (arrows) in preaspyridone major isomer 230.
Figure 3.15: 13C NMR of major isomer of preaspyridone A 230 run in DMSO-d6.
106
Figure 3.16: 1H NMR of major isomer of preaspyridone A 230 run in DMSO-d6.
3.7.1(c)
Discussion on the biosynthesis of preaspyridone A 224 and 230
Heterologous expression of apdA and apdC in A. oryzae (M-2-3) produced two
diastereomers of preaspyridone A, 224 and 230, which confirms the in vitro
reconstitution of apdA and apdC function by Tang and colleagues.107 There are a
number of 3-acyl tetramic acids reported from analogous iterative HRPKS-NRPS gene
clusters from other fungi. For example, fusarin C 77, pramanicin 83, militarinone C 82,
equisetin 81, pseurotin A 78, chaetoglobosin A 80, 2-oxo-cyclopiazonic acid 85 (Figure
3.17), pretenellin A 114 and predesmethylbassianin A 118 (see chapter 1). The
biosynthetic pathway of preaspyridones is more similar to pretenellin A 114.
The minor 224 and major 230 preaspyridone A are epimers at methine proton H5. This illustrates that the adenylation domain of the NRPS is able to select D-tyrosine
231 during biosynthesis of minor preaspyridone 224 and L-tyrosine 232 to form the
major preaspyridone 230 (Scheme 3.5). This further confirms the in vitro studies by
Tang and coworkers where they reported the flexibility of the adenylation domain of
apdA to incorporate different aromatic amino acids.107 We also proposed that minor
preaspyridone 224 might have formed by reduction of 267.
107
Scheme 3.5: Incorporation of amino acids in isomers of preaspyridone A 224 and 230.
The structures of the diastereomers of preaspyridone A 224 and 230 also reflects
the exclusive potential of the enoyl reductase domain (ApdC) of aspyridone gene
cluster, particularly setting the opposite R and S stereo centres at the pendant methyls in
the 2, 4-dimethyl hexanoyl side-chain of preaspyridone A 224 and 230. In the initial
cycle of the polyketide chain biosynthesis the AT and KS domains form a diketide,
followed by methyl group transfer by CMeT domain and ketoreduction by the KR. The
dehydratase (DH) further reduces the diketide to an enol group. The enoyl reductase is
defective in the ApdA protein and the reduction is carried out by the stand alone ER
protein encoded by apdC. The ER reduces the enoyl group to a saturated bond and at
the same time sets the stereochemistry of the methyl chain, and in pre aspyridone A the
first methyl group is arranged in R configuration 233. The same steps repeat in the
second cycle by the iterative domains and the second methyl group is settled in S
configuration 234 by the ER domain (Scheme 3.6). In the last cycle a β-keto tetraketide
220 is formed.
108
Scheme 3.6: Proposed biosynthetic steps of polyketide chain in pre aspyridone A 224 and 230.
A similar stereoselective programming is observed by KR domain in (6’S,
10’S)-7, 8’-dehydrozearalenol (DHZ 236) biosynthesis, which is an intermediate of
hypothemycin 237. The HRPKS Hpm8 synthesizes a hexaketide 235 where KR
catalyzes opposite stereo reduction at C-6’ and C-10’. The hexaketide 235 is transferred
to a NRPKS which combines with three malonyl coenzyme A to form DHZ 236 and
later steps eventually form hypothemycin 237 (Scheme 3.7).184
Scheme 3.7: Stereoselective reduction during biosynthesis of hypothemycin 237.
109
Figure 3.17: 3-acyl tetramic acids reported from hybrid PKS-NRPS pathway, amino acids are highlighted in red.
3.7.2
Heterologous expression of apdACE in A. oryzae (M-2-3)
After successful expression of apdA and apdC, pTAYAGSargPage was
modified to develop another plasmid pTAYAargACE, containing apdA, apdC and
apdE, which was then transformed into A. oryzae M-2-3. The apdE gene encodes a
cytochrome P450 monooxygenase and it shares 48% protein identity to the ring
expandase P450 enzyme TenA present in B. bassiana 110.2.108 This expression was
aimed to delineate the role of P450 enzymes in aspyridone A 84 pathway. Selected
transformants were screened for production of new metabolites by LCMS.
The A. oryzae ACE expression clone was grown on DPY plates and mature
spores were inoculated in CMP liquid media for 7 days at 28 °C (see section 4.10). The
A. oryzae culture was homogenised, acidified and extracted with ethyl acetate. The
organic extract was concentrated under vacuum and defatted. The dried extract (112.6
mg from 100 ml of culture) was made to a solution of 10 mg/ml in HPLC grade
methanol and analysed by LCMS.
110
14.5
ZW-II-92K-50-70ch3cn
A
14.9
8.8
3.0e+2
2.0e+2
3.28
1.0e+2
3.72
2.58
0.0
-0.00
ZW-II-92M
6.0e+1
2.00
4.00
24.8
6.4
6.00
8.00
10.00
12.00
14.00
16.00
18.00
20.00
22.00
24.00
26.00
28.00
3.22
B
Wildtype A. oryzae
3.45
5.0e+1
4.0e+1
5.80
3.0e+1
2.0e+1
1.0e+1
4.25
2.60
27.43
0.0
Figure 3.18: A, Diode array chromatogram of A. oryzae apdACE expression clone showing four new peaks at 8.8, 14.5,
14.9 and 24.8 minutes; B, Diode arrat chromatogram ofvWild type A. oryzae used as a control.
The diode array chromatogram of A. oryzae apdACE expression clone indicated
four new peaks which were not present in the A. oryzae wild type (WT) extract. The
first peak at 8.8 minutes possessed a λmax (222, 281) which is similar to uv absorption of
the 3-acyl tetramic acid preaspyridone A 224. The ESI spectrum of this peak showed
m/z 348 [M]H+ which is 16 mass units more than that of preaspyridone A 224. The
second peak at 14.5 minutes showed mass ion of 238 [M]H+ and a longer λmax (229, 325
nm). The peak eluting at 14.9 minutes showed a uv absorption (λmax 246, 344) and ESI
(m/z 330 [M]H+), spectra corresponding to aspyridone A 84. The last peak at 24.8
minutes had a mass ion of m/z 316 [M]H+, which is 16 mass units less than
preaspyridone A 224 (m/z 332 [M]H+) and a λmax 281 nm.
The transformant was grown on large scale (100 ml × 10 flasks) and massdirected purification of the above peaks was performed on a Waters LCMS
autopurification system (see section 4.5). The dried crude extract (1303 mg/L) was used
to prepare a solution of 50 mg/ml and about 200 µl was injected in each preparative run.
The gradient use on a 30 duration programme was acetonitrile Method 5 (section 4.5)
on a C18 Phenomenex LUNA column. The purified fractions of each compound were
collected and dried under nitrogen gas. Structural elucidation was achieved using 1D
and 2D NMR spectroscopy and High Resolution Mass Spectrometry.
111
3.7.2(a)
Identification of 14-Hydroxypreaspyridone A 238
The compound eluting at 8.8 minutes was obtained in the form of waxy light
brown solid (67 mg/L). HRESIMS gave a molecular formula of C19H25NO5 (observed
370.1624; calculated 370.1625 for M[Na] +). The
13
C NMR in methanol-d4 showed
many signals coming at similar chemical shifts as preaspyridone A 224. These included
two methylene carbons at δC 41.2 (C-8) and δC 30.2 (C-10), a methine carbon at δC 68.5
(C-5), two sp3 carbons at δC 33.5 (C-9) and δC 35.5 (C-7), three methyl groups at δC 17.5
(C-13), δC 19.6 (C-12) and δC 11.4 (C-11), two aromatic carbon at δC 115.5 (C-17, C-19)
and δC 129.6 (C-16, C-20) and carbonyl group at δC 195.2 (C-6). A new signal at δC 74.8
indicated carbon attached to a hydroxyl (OH) group. The 1H NMR presented a similar
spectrum to preaspyridone A 224 spectra. A doublet at δH 4.99 attached to δC 74.8 (C14) in the 1H-13C HSQC was assigned to H-14 (Figure 3.20). The aromatic doublets at
δH 7.11 (H-16, H-20) and δH 6.65 (H-17, H-19) exhibited HMBC correlations to parasubstitited phenol at δC 158.6 (C-18), quaternary carbon at δC 130.3 (C-15) and benzylic
carbon δC 74.8 (C-14) (Figure 3.19). The doublet at δH 4.22 was assigned to methine H5 showing a 1H-1H COSY to H-14 and HMBC correlation to benzylic carbon at C-14
which confirmed para-substituted phenol attached at hydroxy-benzyl to a pyrrolidine
nucleus at methine H-5 (Figure 3.21). The quartet at δH 3.55 (H-7) was linked to C-8
and C-13 in HMBC. The multiplet spread between δH 1.29-1.40 was assigned to
methylene protons (H-8) and δH 1.09-1.33 to methylene groups (C-10) and methine at
C-9. The terminal methyl at δH 0.81 (H-11) and methyl at 0.82 (H-12) showed HMBC
connection to C-13, C-9 and C-8 confirming a similar 2,4-dimethylhexanoyl side chain
possessed by preaspyridone A 224 (Figure 3.19). All correlations confirmed the
structure to be 14-hydroxypreaspyridone A 238.
Figure 3.19: 1H-1H COSY (solid lines) and 1H-13C HMBC correlations (arrows) in 14-hydroxypreaspyridone 238.
112
Figure 3.20: 1H NMR of 14-hydroxy preaspyridone A 238 run in methanol-d4.
Figure 3.21: Segment of 1H-13C HMBC NMR spectrum of 14-hydroxy preaspyridone A 238 showing key correlations of
hydroxyl benzyl at C-14 with a methine at H-5 and aromatic proton H-16.
113
Figure 3.22: ES+ and UV spectrum of 14-hydroxy preaspyridone A 238 eluted at 8.8 mins.
3.7.2(b)
Identification of 4-hydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 239
The compound eluting at 14.5 minutes was obtained as pale white solid (63
mg/L). The HRESIMS gave a molecular formula of C13H20NO3 (observed 238.1428;
calculated 238.1437 for M[H] +). The 13C NMR in DMSO-d6 indicated the presence of
all 13 carbons consisting of two carbonyl groups at δC 177.6 (C-4) and δC 211.9 (C-7),
two methylene groups at δC 40.0 (C-9) and δC 29.7 (C-11) and three methyl groups at δC
11.1 (C-12), δC 16.7 (C-14) and δC 18.8 (C-13). The 1H NMR showed a triplet
downfield at δH 7.60 and a doublet at δH 5.92 which were assigned to H-6 and H-5
respectively (Figure 3.24), linked to an amide proton at δH 11.49 (H-1) in 1H-1H COSY
(Figure 3.23). The aromatic protons H-5 and H-6 showed 1H-13C HMBC correlations to
quaternary carbon C-3 (δC 105.9), a hydroxyl group at C-4 (δC 177.6) and carbonyl
group at δC 161.8 (C-2) verifying the structure of 2-pyridone (Figure 3.25). The quartet
at δH 4.25 was assigned to a methine proton H-8 linked to the carbonyl group at δC 211.9
(C-7), methyl group at C-14 and to a methylene group at C-9 (Figure 3.23). The methyl
group at δH 1.02 (H-14) showed HMBC connection to carbonyl at C-7 and methine at C8 which established that it is attached to C-8. The multiplets at δH 1.21 and δH 1.49 were
assigned to geminal protons at H-9 and multiplets at δH 1.09 and δH 1.25 were allocated
to geminal protons at H-11. A number of COSY and HMBC showed the structure to be
a 2, 4-dimethylhexanoyl side chain attached to 2-pyridone at C-3 and was named as 4hydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 239.
114
Figure 3.23: 1H-1H COSY (solid lines) and 1H-13C HMBC correlations (arrows) in 4-hydroxy-3-(2, 4-dimethylhexanoyl) 2pyridone 239 and crystal structure of 239.
Figure 3.24: 1H NMR of 4-hydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 239 run in DMSO-d6.
Figure 3.25: Section of 1H-13C HMBC NMR spectrum of 4-hydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 239 showing
important correlations of pyridone protons H-5 and H-6 with quaternary carbons C-2, C-4 and C-3.
115
Figure 3.26: ES+ and UV spectrum of 4-hydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 239.
3.7.2(c)
Identification of aspyridone A 84
The compound separating at 14.9 minutes (Figure 3.18) was obtained as light
brown solid. The HRESIMS gave a molecular formula of C19H24NO4 (observed
238.1692; calculated 330.1699 for M[H]+). From 1D 1H and 13C NMR and 2D NMR the
compound was identified as aspyridone A 84.80 The two aromatic doublets at δH 7.26
(H-16, H-20) and δH 6.80 (H-17, H-19) (Figure 3.28) exhibited HMBC correlations to
para-substituted phenol carbon at δC 158.3 (C-18) and quaternary carbon at δC 125.2 (C15). The singlet at δH 7.47 was assigned to methine proton H-6 which displayed 1H- 13C
HMBC connections to a hydroxyl group at δC 177.6 (C-4) and carbonyl group at C-2
(δC 163.9) (Figure 3.27). The H-6 singlet also showed HMBC connection to quaternary
carbon C-15 which confirmed 2-pyridone linked to para-substituted phenol at C-15.
The quartet at δH 4.39 was allotted to methine proton H-8 linked to methylene group at
δC 41.2 (C-9) and to methyl group at δC 17.6 (C-14). The last methyl at δH 0.87 (H-12)
showed correlations to C-10 (δC 33.7) and methylene group at δC 33.7 (C-11). The
methyl group at δH 0.90 (H-13) showed connections to C-9, C-10 and C-11 giving
116
evidence that a similar polyketide chain of preaspyridone A 224 has combined intact to
2-pyridone at C-7 (δC 214.4) to give a structure of aspyridone A 84.
Crystals of aspyridone A 84 were formed. The methyl groups at C-8 and C-10
possess anti-configuration in the crystal data, similar to preaspyridone A 224 minor
isomer and 239.
Figure 3.27: 1H-1H COSY (solid lines) and 1H-13C HMBC correlations (arrows) in aspyridone A 84 and crystal structure
of aspyridone A 84.
Figure 3.28: 1H NMR of aspyridone A 84 run in methanol-d4.
117
Figure 3.29: ES+ and UV spectrum of aspyridone A 84.
3.7.2 (d)
Identification of 18-deshydroxypreaspyridone A 240
The final compound from A. oryzae apdACE expression clone separated at 24.8
minutes (Figure 3.18). It was obtained as light brown solid (12 mg/L) and HRESIMS
showed molecular formula of C19H25NO3 (observed 338.1735; calculated 338.1726 for
M[Na]+).
The 13C NMR revealed the presence of 15 carbons and quaternary carbons were
observed in the 2D NMR. A significant difference in the 1H NMR of compound 240
was observed due to the presence of distinct multiplets in the aromatic region (Figure
3.31) in contrast to the distinct pair of doublets which are characteristic of the parasubstituted phenol present in the spectra of preaspyridone 224 and aspyridone A 84
(Figure 3.10 and 3.28). The multiplets from δH 7.16-7.22 were assigned to three
aromatic protons (H-17/19), (H-18) and (H-16/20) (Figure 3.31). The aromatic protons
H-16/20 and H-17/19 displayed HMBC correlations to quaternary carbon at δC 136.9
(C-15) and benzylic carbon at δC 38.3 (C-14) (Figure 3.32). Two set of doublets of
doublets at δH 2.98 and δH 3.07 were assigned to two diastereotopic protons at the
benzylic position, H-14 and showed connection to δC 130.7 (C-17/19), C-15 and to
methine carbon at δC 63.5 (C-5) confirming the structure to be benzene ring attached to
pyrrolidine ring at C-5 connected through benzylic carbon, C-14. The multiplet at δH
3.65 was allotted to methine proton H-7 linked to a carbonyl group at δC 196.0 (C-6),
methylene group at C-8 (δC 41.3), a methine at δC 33.5 (C-9) and a methyl group at δC
118
17.6 (C-13) (Figure 3.30). The multiplets at δH 1.33 and 1.44 were assigned to geminal
protons at H-8 correlated to C-6, C-13, C-12 (δC 19.5), C-10 (δC 30.3), C-9 (δC 33.5) and
C-7 in 1H-13C HMBC. The multiplets at δH 1.09 and δH 1.34 were assigned to a second
methylene group at H-10 linked to C-11 (δC 11.5), C-12 and C-9 in the HMBC
correlations. All the 1D and 2D NMR elucidated the structure to be 18deshydroxypreaspyridone A 240.
Figure 3.30: 1H-1H COSY (solid lines) and 1H-13C HMBC correlations (arrows) in 18-deshydroxypreaspyridone A 240.
Figure 3.31: 1H NMR of 18-deshydroxypreaspyridone A 240 run in methanol-d4.
119
Figure 3.32: Segment of 1H-13C HMBC NMR of 18-deshydroxypreaspyridone A 240.
Figure 3.33: ES+ and UV spectrum of 18-deshydroxypreaspyridone A 240.
3.7.2 (e)
Role of apdE in aspyridone pathway
Heterologous expression of hrPKS-NRPS ApdA, enoyl reductase ApdC and
cytochrome P450 enzyme ApdE in A. oryzae was accomplished using the
120
pTAYargACE expression vector. The transformant produced three new products; 14hydroxy preaspyridone 238, 4-hydroxy-3-(2,4-dimethylhexanoyl)2-pyridone 239,
aspyridone A 84 and 18-deshydroxypreaspyridone A 240. This experiment exemplified
the role of ApdE in the aspyridone pathway as the new compounds 238, 239 and 240
were not observed in the A. oryzae apdAC expression clone.
The apdE gene encodes a monooxygenase enzyme which belongs to a large
family of cytochrome P450 oxidases. Cytochrome P450 enzyme in fungi are known to
perform important bioconversions and reactions in the biosynthesis of many natural
products, for example selective olefin epoxidation and oxidation of methyl groups of
natural product compounds. They also execute hydroxylation of complex polyaromatic
hydrocarbons, steroids such as progesterone. Some plant pathogenic fungi are reported
to encode P450 enzymes.185 Cytochrome P450 contain a heme cofactor and are known
as hemo proteins. Scheme 3.8 illustrates the reaction by which Ferrous (II) ions in heme
uses molecular oxygen as the oxidant and carries out oxidation of organic substrates by
introduction of oxygen into C-H bond.
Scheme 3.8: The heme in Cytochrome P450 uses molecular oxygen for oxidation of organic substrates.
In aspyridone biosynthesis, ApdE catalyses three important reactions which are:
conversion of preaspyridone A 224 and 230 to aspyridone A 84 by oxidative ring
expansion; hydroxylation of preaspyridone A to form 238; and oxidative dephenylation
of preaspyridone A to from 239. A similar oxidative ring expansion reaction takes place
in tenellin 87 biosynthesis in B. bassiana 110.0 by a cytochrome P450 enzyme known
as TenA. It drives the formation of pretenellin B 115 from pretenellin A 114 by
oxidative ring expansion of 3-acyl tetramic acid to 2-pyridone (see chapter 2).
Prototenellin D 155 is another compound similar to 14-hydroxy preaspyridone A 238 in
tenellin pathway. It is formed by hydroxylation of pretenellin A by direct rebound
121
mechanism by an unknown oxidase enzyme in B. bassiana as it is only produced in the
native organism and not observed during heterologous expression of tenellin 87 genes
in A. oryzae. Halo and colleagues have comprehensively studied and presented a
hypothesis for the mechanism of oxidative ring expansion and benzylic hydroxylation
of tetramic acid in tenellin 87 pathway.104,108
Scheme 3.9: Biosynthesis of pretenellin B 115 and protonellin D 155 in B.bassiana.
Scheme 3.10: A, Proposed oxidative mechanisms for biosynthesis of hydroxyl tetramic acid 238; B, Proposed
mechanism for oxidative ring expansion during biosynthesis of 2-pyridone 84.
122
We proposed similar biosynthetic routes for compounds 238, 84 and 239
(Scheme 3.10, 3.11). Cytochrome P450 initiates by hydrogen atom abstraction from the
benzylic position and forms carbon-centred radical 241. The C-centred radical reacts
with an iron bound hydroxyl radical and gets hydroxylated directly by the oxygen
rebound mechanism186 to form 14-hydroxy preaspyridone 238 (Scheme 3.10, A). The
carbon centred radical 241 can seemingly also follow another route of single electron
transfer and form cyclopropyl oxy-radical 242,187 followed by another short lived
intermediate 243, which consequently leads to ring expansion and form 2-pyridone
compound aspyridone A 84 (Scheme 3.10, B).
Scheme 3.11: Proposed oxidative dephenylation for biosynthesis of 239.
A third mechanism was proposed for the biosynthesis of 4-hydroxy-3-(2,4dimethylhexanoyl) 2-pyridone 239. A peroxo-iron intermediate 244 can make
hydroxylation at the phenyl-bridge head carbon of preaspyridone A 224 which results in
formation of assumed hydroxyquinone 245 (Scheme 3.11). Further electron transfer
leads to ring expansion of the pyrrolidone ring to 2-pyridone 246 and the hydroquinone
separates causing dephenylation. The 2-pyridone rearranges and forms 239 (Scheme
3.11). A number of related natural products are known. For example Torrubiellone A
247 and Torrubiellone B 248 from the spider-pathogenic fungi Torrubiella sp. BCC
2165,188
(+)-N-deoxymilitarinone
A
249
and
militarinone
A
250
from
entomopathogenic fungi Paecilomyces farinosus,189 all feature structures which have
been hydroxylated at the carbon corresponding to the oxidation target in the proposed
mechanism, while jacaglabroside B 251190,191 features a more highly oxidised version of
the same structural feature.
123
TenA in the tenellin 87 gene cluster has a similar catalytic activity to ApdE, yet
the latter displays a broader chemical diversity as no dephenylated or similar
compounds were reported from the tenellin 87 pathway. There are a number of reported
structures which are similar to dephenylated compound 239. Piericidins A 252 and B
253 are natural insecticides isolated from Streptomyces mobaraensis.192 They exhibit
specific
inhibitory
action
for
electron
transport
system
in
mitochondria.
Sapinopyridione 254 and 255 were purified from a fungal pathogen of conifers,
Sphaeropsis sapinea.193 Atpenins 256, 257, 258 are antifungal metabolites of molds
Penicillium sp.FO-125. They are known to inhibit the succinate-ubiquinone reductase
activity of mitochondrial complex II.194 The atpenins also possess pendant methyls
arranged in anti-configuration.
124
3.7.3
Heterologous Expression of apdABC in A. oryzae
The apdB gene encodes a second cytochrome P450 monooxygenase in the
aspyridone cluster.80 The pTAYAargABC plasmid was constructed to reveal the role of
apdB. Selected transformants were grown to be analysed for production of new
metabolites. The transformants were grown on DPY medium on plates and spores were
afterwards inoculated in CMP (section 4.9.b) growth media for seven days according to
methods dercribed in section 4.10. The cultures of A. oryzae apdABC transformants
were extracted in the usual way and the crude extract was defatted with wet methanol
and hexane and then evaporated (80.5 mg from 100 ml).
A
14.0
apdA,B,C
ZW-II-92D
1.5e+2
3.32
1.0e+2
5.0e+1
2.67
0.0
-0.00
ZW-II-92M
6.0e+1
5.0e+1
4.52
2.00
4.00
13.2
6.10
6.00
8.00
10.00
12.00
29.25
14.00
16.00
18.00
20.00
22.00
24.00
26.00
28.00
3.22
B
Wildtype A. oryzae
3.45
4.0e+1
5.80
3.0e+1
2.0e+1
1.0e+1
4.25
2.60
27.43
0.0
Time
Figure 3.34: A, Diode array chromatogram of A. oryzae apdABC expression clone showing peaks of preasyridone A
minor isomer 224 at 13.2 and major preaspyridone A 230 at 14.0 minutes; B, Wild type A. oryzae used as a control.
A solution of 10 mg/ml in HPLC grade MeOH of the crude extract was injected
for analysis on LCMS. The diode array chromatogram showed two peaks at 13.2 and
14.0 minutes. They were identified as the minor and major isomers of preaspyridone A
224 and 230 which were characterised from the apdAC expression clone. We observed
an increase in the titres of both compounds in the apdABC expression clone to be 2.2
mg/L and 141.0 mg/L for the minor and major components of preaspyridone A,
respectively. This was double the amount of preaspyridone A in comparison to apdAC
expression clone where the yields of minor and major component were 0.6 mg/L and 60
125
mg/L, respectively. This expression experiment showed that the ApdB cytochrome
P450 cannot chemically act on the tetramic acid preaspyridone A, but it may act later in
the pathway.
3.7.4 Heterologous Expression of apdACEB in A. oryzae
The A. oryzae apdABC expression clone did not produce any new compounds,
compared to A. oryzae apdAC. We next planned to express the apdB gene in the
presence of the megasynthase ApdA, enoyl reductase ApdC and monooxygenase ApdE
in A. oryzae. The pTAYAGSargPage vector has a capacity to express maximum of four
genes owing to the presence of four promoters.176 Thus pTAYAGSargPage was
modified to construct a plasmid pTAYAargASP inserting all four genes apdA, apdB,
apdC and apdE (Figure 3.35). The expressions of all the genes were confirmed by qRTPCR.
Figure 3.35: (left) pTAYAargASP expression plasmid, (right) A. oryzae apdACEB expression clone on Czapek Dox
Agar.
The A. oryzae clones were initially grown on selection media and later on DPY
media (section 4.7.b) for maximum production of spores. The one week old spores were
inoculated in liquid media (CMP) at 28 °C at 200 rpm for 7 days (section 4.10). The
fermentations were extracted in the usual way and the organic layer was concentrated
under vacuum and then defatted with wet methanol and hexane. The resulting crude
126
extract was dried to a thick mass weighing 30 mg/100ml. 10 mg/ml of the crude extract
was made with HPLC methanol and 20 µl was injected in LCMS.
A
14.2
13.5
3.43
4.18
2.25
2.00
3.00
B
4.00
5.00
6.00
7.00
8.00
9.00
20.0 20.3
16.5
9.1
10.00
11.00
12.00
13.00
14.00
3.22
15.00
16.00
17.00
18.00
19.00
20.00
21.00
20.00
21.00
Wildtype A. oryzae
3.45
5.8
4.25
2.60
2.00
3.00
4.00
5.00
6.00
7.00
8.00
9.00
10.00
11.00
12.00
13.00
14.00
15.00
16.00
17.00
18.00
19.00
Figure 3.36: A, Diode array chromatogram of A. oryzae apdACEB expression clone showing production of 14hydroxypreaspyridone A 238, minor 224 and major isomer of preaspyridone A 230, aspyridone A 84 and three new
compounds at 16.5, 20.0 and 20.3 minutes; B, A. oryzae Wild type.
The LCMS chromatogram displayed a number of peaks which were not present
in the diode array chromatogram of the Wild type (WT) A. oryzae. The wavelength and
mass spectrum of first peak at 9.2 min. was identified as 14-hydroxy preaspyridone 238,
the second peak at 13.6 min. was minor isomer of preaspyridone A 224, third peak at
14.2 min. was major isomer of preaspyridone A 230 and the compound eluting at 14.5
min was aspyridone A 84 (Figure 3.36). These compounds were later confirmed by 1H
NMR.
Three new peaks were detected at 16.5, 20.0 and 20.3 min. (Figure 3.36). The
mass spectrum for the compound eluting at 16.5 min. was m/z 254 [M]H+, 16 mass units
more than the dephenylated aspyridone 239 and observed λmax of 278, 335 nm. The
peak at 20.0 minutes showed a uv absorption 228, 330 nm and ESI spectrum presented
m/z 268 [M]H+. The last peak at 20.3 min. showed a higher uv absorption (294, 376 nm)
and m/z 330 [M]H+.
For purification of all the peaks 1 litre (100 ml x 10 flasks) of the cultures of the
transformant was grown according standard fermentation conditions (section 4.10). The
127
Time
crude extract obtained after extraction and concentration was 1.13 g per liter. A
concentration of 50 mg/ml was made with HPLC grade methanol. 200 µl of the solution
was injected in Waters LCMS mass directed purification in each preparative run. The
purification was achieved with an acetonitrile and water solvent system with Method 3
(section 4.5) in a 30 min. duration programme. The purified fraction of each peak was
collected and dried under nitrogen. The last peak at 20.3 min. was not collected due to
lower titres (it was later purified in another experiment explained in section 3.7.5). The
structures of pure compounds were each studied with the help of 1D and 2D NMR
spectrometer and high resolution mass spectrum.
3.7.4 (a)
Identification
of
1, 4-dihydroxy-3-(2, 4-dimethylhexanoyl) 2-
pyridone 259
The compound eluting at 16.5 min. was obtained as light brown solid 6.2 mg/L.
The HRESIMS provided the molecular formula to be C13H19NO4 (observed 276.1218;
calculated 276.1206 for [M]Na+). The 13C NMR showed the presence of all 13 carbons,
with many chemical shifts similar to dephenylated aspyridone 239. The
13
C NMR
composed of two carbonyls groups at δC 213.6 (C-7) and δC 160.0 (C-2), a quaternary
carbon at δC 107.4 (C-3), a hydroxyl group at C-4 (δC 176.0), two methylene groups at
δC 31.0 (C-11) and δC 41.1 (C-9) and three methyl groups at δC 19.3 (C-13), δC 17.3 (C14) and δC 11.7 (C-12). A quartetet at δH 4.31 allocated to H-8 displayed 1H-1H COSY
correlation to H-14 (δH 1.11) and to the methylene carbon C-9 in the 2D 1H-13C HMBC.
The multiplets at δH 1.33 and δH 1.62 were assigned to geminal protons at H-9 linked to
C-7, C-14, C-13, C-11, C-10 and C-8 in the HMBC correlations (Figure 3.37 and 3.39).
The methine proton H-10 signals at δH 1.41 and multiplets at δH 1.18 and δH 1.36 were
consigned to the two geminal protons at H-11. These correlations verified that the
aliphatic chain is the same as in dephenylated preaspyridone 239 and no hydroxylation
has occurred on the polyketide chain. The pair of aromatic douplets at δH 5.97 and δH
7.94 were assigned to protons H-5 and H-6 respectively, of the 2-pyridone (Figure
3.38). The distance between the doublets was more than observed in the 1H NMR of
dephenylated aspyridone 239 and this was ascribed to the presence of a hydroxyl group
attached to the pyridone nitrogen. From the NMR correlations and HRMS the structure
was established to be 1, 4-dihydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 259.
128
Figure 3.37: 1H-1H COSY (solid lines) and 1H-13C HMBC correlations (arrows) in 1, 4-dihydroxy-3-(2, 4dimethylhexanoyl) 2-pyridone 259.
Figure 3.38: 1H NMR of 1, 4-dihydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 259 run in methanol-d4.
Figure 3.39: Key 1H-13C HMBC correlations in 1, 4-dihydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 259.
129
Figure 3.40: ES+ and UV spectrum of 1, 4-dihydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 259.
3.7.4 (b)
Identification of 1-methoxy, 4-hydroxy-3-(2, 4-dimethylhexanoyl) 2-
pyridone 260
The compound 260 separating at 20.0 minutes was purified in the form of light
brown solid (2.7 mg/L) and HRESIMS analysis presented a molecular formula of
C14H21NO4 (observed 290.1368; calculated 290.1362 for [M]Na+). The
13
C NMR
displayed presence of 14 Carbons. Many chemical shift values were similar to
dephenylated aspyridone A 239 as it consists of three methyl groups at δC 11.7 (C-12),
δC 17.3 (C-14) and δC 19.3 (C-13), two methylene groups at δC 31.0 (C-11) and δC 41.1
(C-9), two methine at δC 42.2 (C-8) and δC 33.5 (C-10), two carbonyls at δC 213.6 (C-7)
and δC 159.7 (C-2) and aromatic carbons at δC 100.7 (C-5) and δC 143.0 (C-6). A new
carbon signal was observed at δC 65.6 which indicated that it is attached to an oxygen
atom.
The 1H NMR consists of two sets of doublets at δH 6.02 (H-5) and δH 8.03 (H-6),
assigned to aromatic protons of 2-pyridone (Figure 3.43). The 1H NMR presented a
singlet at δH 4.01 integrating to three protons which was not observed in dephenylated
aspyridone A 239. It was attached to δC 65.6 in 1H-13C HSQC. A characteristic feature
which aided in elucidation of the structure was revealed in 1D NOESY when δH 8.03
130
(H-6) showed correlation to δH 4.01, which determined that an O-methyl is attached to
nitrogen atom at position 1 in 2-pyridone (Figure 3.41).
Figure 3.41: 1D NOESY showing connection of H-6 with methoxy group at N-1.
In 1H-13C HMBC spectrum, H-5 presented two bond and three bond correlations
with hydroxyl group at C-4 (δC 177.3) and quaternary carbon (δC 108.2) respectively
(Figure 3.44). The aromatic proton H-6 displayed connections to carbonyl group at C-2,
aromatic carbon C-5 and hydroxyl group at C-4. We didn’t observe any HMBC
correlations from H-15 to C-2 or C-6; this might be because four bond correlations are
rarely seen. The terminal methyl at δH 0.87 (H-12) was linked to methylene group at δC
31.0 (C-11) and methine at C-10 (δC 33.5) and adjacent methyl signal at δH 0.92 (H-13)
diplayed HMBC correlations to C-10, C-11 and C-9 (Figure 3.42). The geminal protons
at δH 1.31 and δH 1.61 at H-9 shows connections to carbonyl group at C-7 (δC 213.6),
methine carbon at δC 42.2 (C-8), C-10, C-11 and C-13. This confirms a 2, 4dimethylhexanoyl side chain identical to aspyridone A 84. These correlations establish
the structure to be 1-methoxy, 4-hydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 260.
131
Figure 3.42: 1H-1H COSY (solid lines) and 1H-13C HMBC correlations (arrows) in 1-methoxy, 4-hydroxy-3-(2, 4dimethylhexanoyl) 2-pyridone 260.
Figure 3.43: 1H NMR of 1-methoxy, 4-hydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 260 run in methanol-d4.
Figure 3.44: Key 1H-13C HMBC correlations in 1-methoxy, 4-hydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 260.
132
Figure 3.45: ES+ and UV spectrum of 1-methoxy, 4-hydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 260.
3.6.4 (c)
Discussion on role of apdB
The apdABCE expression clone produced a range of compounds including the
two diastereomers 224 and 230 observed when megasynthase ApdA was expressed with
enoyl reducatse ApdC and 14-hydroxy preaspyridone A 238, dephenylated aspyridone
239 and aspyridone A 84 which are products of the monooxygenase ApdE. This
illustrates that as more genes from a cluster are expressed in a heterologous experiment,
we observe novel biosynthetic potential of the specific gene cluster.
The production of 1-methoxy dephenylated pyridone 260 and N-hydroxy
dephenylated pyridone 259 ascribes a role for the cytochrome P450 enzyme ApdB, that
it is involved in N-hydroxylation of 5-dephenylated pyridone 239 to form 259 and 260.
It exhibits substrate specificity for dephenyalted pyridines only, because we didn’t
observe any N-hydroxylation in tetramic acids 224 and 230 and neither in aspyridone A
84.
In tenellin, the cytochrome P450 enzyme TenB also displays N-hydroxylation
for only pretenellinB to form tenellin 87 and TenB does not N-hydroxylates tetramic
acids pretenellin A 114 (see chapter 2). We assumed that the O-methylation in 260 is
carried out by a native enzyme present within A. oryzae as there is no enzyme for
putative methyl transferase characteristic in aspyridone cluster.80 There are a number of
reported compounds which possess N-methoxy group, for example Cordypyridone C
133
261 and Cordypyridone D 262 from the insect pathogenic fungus Cordyceps
nipponica.195 Kumarihamy et al. isolated four new N-methoxy-2-pyridinone compounds
from the plant pathogen Septoria pistaciarum, which are 17-hydroxy-N-(O-methyl)
septoriamycin A 263, 17-acetoxy-N-(O-methyl) septoriamycin A 264, 13-(S)-hydroxyN-(O-methyl) septoriamycin A 265 and 13-(R)-hydroxy-N-(O-methyl) septoriamycin A
266.196’’
3.7.5
Heterologous expression of apdACED in A. oryzae
ApdD encodes a FAD dependent monooxygenase. We planned to coexpress the
apdD gene in the presence of the megasynthase ApdA, enoyl reductase ApdC and
monooxygenase ApdE in A. oryzae to illustrate the role of ApdD in aspyridone
biosynthesis. The pTAYAargASP plasmid was used to construct pTAYAargACED
consisting of apdA, apdE, apdC and apdD and this was transformed into A. oryzae. The
transcription levels of all the genes were confirmed by qRTPCR (This transformation
was done by Dr. Khomaizon Pahirulzaman).
The selected transformant was grown on DPY plates and when mature spores
were visible after seven days, a spore solution was made in sterilised deionized water
and was inoculated in each 100 ml liquid media kept in 250 ml Erlenmeyer flask. 1
Litre of the media (10 x 100 ml flasks) was grown with constant shaking at 200 rpm at
28 °C. After seven days the fungal cultures were extracted with ethyl acetate (section
4.11). The organic layer was concentrated and defatted with hexane. Drying the crude
extract under nitrogen, gave a brown mass (1.128 g / 1000 ml). A 10 mg/ml solution
was made with HPLC grade methanol and analysed on LCMS using CH 3CN/H2O
analytical programme with a gradient (50-65%) with 30 min. duration (Figure 3.46).
134
A
16.6
17.3
3.55
24.4
4.02
4.00
3.22
10.2
6.00
8.00
10.00
12.00
14.00
16.00
18.00
20.00
22.00
24.00
26.00
28.00
B
Wildtype A. oryzae
3.45
5.80
4.25
Figure 3.46: A, Diode array chromatogram of apdACED consisting of 14-hydroxy preaspyridone A 238 at 10.2 minutes,
dephenylated pyridone 239 at 16.6 minutes, aspyridone A 84 at 17.3 minutes and a new compound at 24.4 minutes
(green); B, Diode array chromatogram of wild type A. oryzae.
The LCMS chromatogram displayed four noticeable peaks which were not
present in the wild type A. oryzae. The peak at 10.2 min. was identified as 14-hydroxy
preaspyridone 238 possessing a mass of m/z 348 in [M]H+ and λmax of (221, 283 nm);
the second peak at 16.6 min. was recognized as dephenylated pyridone 238 with a λmax
of (221, 283 nm) and a m/z of 238 in the ES+. The third peak at 17.3 min. had a uv
absorption of (246, 345 nm), characteristic of aspyridone A 84, exhibiting a molecular
ion of m/z 330 [M]H+. The last peak at 24.4 min. showed a m/z of 330 as [M]H+ and a
λmax of (294, 376 nm). This peak was observed in the apdABCE expression clone but
was not purified and characterised previously because of low titre (section 3.7.4).
The crude extract (1.13g) was prepared to a concentration of 100 mg/ml and 200
µl was injected in each run in mass directed preparative LCMS. The gradient used was
(50-65%) in acetonitrile/water (Method 6, section 4.5). Each of the above peaks were
purified, collected and dried under nitrogen. The first three compounds at 10.2, 16.6 and
17.3 min. after purification were analysed by 1D 1H NMR and confirmed to be 14hydroxy preaspyridone 238, dephenylated pyridone 239 and aspyridone A 84. The last
peak which was a new compound was studied with 1D 1H and
13
C and 2D NMR
experiments.
135
From the above experiment we couldn’t observe any precise role for ApdD.
Bergmann et al. guessed that apdD performs hydroxylation of aspyridone A 84 to form
aspyridone B 226.80 But we did not observe any peak confirming the production of
aspyridone B 226 in the expression clone. However, we observed that the presence of
ApdD in the expression clone with ApdE reduced the production of metabolites to 71.5
mg as compared to A. oryzae apdACE where total titres of metabolites were 390.6 mg.
(section 3.7.10).
3.7.5(a)
Identification of Z-5, 14-anhydropreaspyridone A 267
The compound eluting at 24.4 min. was obtained as a bright yellow solid (4 mg)
after repurification. The molecular formula in HRESIMS analysis was given as
C19H23NO4 (observed 352.1528; calculated 352.1519 for [M]Na+). The
13
C NMR
spectra consisted of three methyl groups at δC 11.6 (C-11), δC 17.5 (C-13), δC 19.5 (C12), two methylene groups at δC 30.6 (C-10) and δC 41.5 (C-8) and a methine group at
δC 33.6 (C-9), two aromatic signals at 116.9 (C-17, C-19) and 132.3 (C-16, C-20), a
quaternary signal at 126.5 (C-15) and a hydroxyl signal at C-18 (δC 159.4). A
quaternary carbon at δC 184.0 observed in 2D HMBC was assigned for C-4 and an
alkene carbon at δC 110.4 in 2D HSQC was assigned for C-14.
A structural feature of this compound was revealed in the 1H NMR by the
presence of a singlet at δH 6.52 (Figure 3.47), joined to δC 110.4 in the 1H-13C HSQC.
The signal at δH 6.52 displayed 1H-
13
C HMBC connection to 4-hydroxy of the
pyrrolidine nucleus at δC 184.0 (C-4) and to aromatic carbons at C-16/20 (Figure 3.49
and 3.50). From the above correlations the signal at δH 6.52 was assigned to H-14
attached at the benzylidene position of a tetramic acid. The aromatic doublet at δH 7.40
assigned to H-16/H-20 displayed HMBC connections to benzylidene carbon at δC 110.4
(C-14) and hydroxyl group at C-18. The second aromatic doublet at δH 6.83 assigned to
H-17 and H-19 show couplings to quaternary carbon C-15 and para hydroxyl group at
C-18 (Figure 3.49 and 3.50).
136
Figure 3.47: 1H NMR of Z-5, 14-anhydropreaspyridone A 267 run in methanol-d4.
In order to determine the orientation of the olefin between C-5 and C-14, a three
bond long range J coupling between H-14 and quaternary carbon C-4 was measured
from a 2D selective EXSIDE NMR experiment.191 The EXSIDE NMR is a 2D spectrum
and displays cross peak similar to a HMBC spectrum. The cross peak is a split in the
Carbon dimensions and this splitting gives the value of J (C-H) constant. Figure 3.48
shows cross peaks between H-14 and C-4, labelled in Hertz. The second number in each
peak is the 13C frequency in Hertz and the difference between both peaks (22974.16 Hz
-23019.82 Hz) is 45 Hz. A 15-fold J-scaling factor is used which give a value of 3 Hz
coupling constant between H-14 and C-4. The value of 3JHC for two possible E/Z
isomers of 267 by DFT calculations gave a value of 7.5 and 4.5 Hz respectively, which
further supported the EXSIDE experiment and the double bond between C-5 and C-14,
was confirmed to be a cis configuration.191 (DFT calculations were performed by Dr.
Craig Butts, School of Chemistry, University of Bristol). We failed to determine the
geometry of this compound from 1D NOESY and 1D ROESY because of exchangeable
protons between N-H and hydroxyl group at C-18.
137
Figure 3.48: Selective EXSIDE NMR spectrum showing cross peaks coupling between H-14 and C-4 to be 3 Hz in 267.
The multiplet at δH 3.85, assigned to methine at H-7 showed couplings to
pendant methyl group at C-13 and to methylene group at C-8. The geminal protons at δH
1.40 and δH 1.59 were assigned to H-8, linked to methyls at C-13, C-12 and to
methylene at C-10 and to methine at C-9. The multiplets at 0.87 and 0.90 were allocated
to terminal methyl H-11 and pendant methyl H-12 respectively displaying HMBC
correlations to C-9, C-10 and methylene C-8 (Figure 3.49). These connections showed
that this compound also possess a similar dimethylated tetraketide chain as observed in
preaspyridone 224 and 230. The above correlations decided the structure to be Z-5, 14anhydropreaspyridone A 267.
Figure 3.49: 1H-1H COSY (solid lines) and 1H-13C HMBC correlations (arrows) in Z-5, 14-anhydropreaspyridone A 267.
138
Figure 3.50: Key 1H-13C HMBC NMR correlations in Z-5, 14-anhydropreaspyridone A 267.
Figure 3.51: ES+ and UV spectrum of Z-5, 14-anhydropreaspyridone A 267.
3.7.6 Coexpression of apdG with apdACEB in A. oryzae
The apdG gene, located downstream of megasynthase apdA and exporter gene
apdR in the gene cluster, encodes an acyl dehydrogenase. Bergmann et al.80 predicted
that it assists in tetramic acid ring closing and reductive release of preaspyridone A 224
from the PKS-NRPS domains (section 3.4). However, the production of preaspyridone
139
A 224 and 230 from A. oryzae apdAC expression clone and from in vitro analysis by
Tang et al. (section 3.4), contradicts Bergmann et al. hypothesis. In order to explore the
function of ApdG, we planned to co-express apdG with apdA, apdB, apdC and apdE in
A. oryzae.
The vector pTAYAargASP carried apdABCE and can express only four genes.
To express apdG Dr. Khomaizon Pahirulzaman designed another plasmid which
contained a different selection marker, the bleomycin resistance gene, ble. The argB
gene in the original vector pTAYAGSargPage was replaced by bleomycin resistance
gene ble and the insertion of this gene was driven by PtrpC. This plasmid was used to
enclose apdG inserted next to PgpdA, this plasmid was named pTAYAGSbleG (Figure
3.52). A. oryzae (M-2-3) was co-expressed with pTAYAargASP (carrying apdABCE)
and pTAYAGSbleG (carrying apdG). The minimal media was supplemented with
antibiotic for selection of transformants.
Figure 3.52: Vector pTAYAGSbleG carrying apdG used in A.oryzae apdABCEG expression clone.
The selected transformant was grown on DPY plates for a week, spores were
prepared in sterile deionized water and then transferred to liquid media (CMP media,
section 4.9.b). One flask (100 ml media in 250 ml Erlenmeyer flask) was grown for
140
seven days at 28 °C with constant shaking at 200 rpm. The mycelia and liquid media
were homogenized and acidified before extraction and defatting in the usual way to give
crude extract weighing 62.2 mg/100 ml. A 10 mg/ml solution was made and analysed
with LCMS on a 30 minutes analytical programme using CH3CN/H2O (50-70%
gradient) (Figure 3.53).
14.4
8.9
14.9
A
6.00
21.2
17.0
6.48
7.00
8.00
9.00
10.00
11.00
12.00
13.00
14.00
15.00
16.00
17.00
18.00
19.00
20.00
21.00
22.00
23.00
Wildtype A. oryzae
B
8.28
6.00
7.00
8.00
10.02
9.00
10.00
13.27
11.00
12.00
13.00
14.00
15.00
16.00
17.00
18.00
19.00
20.00
21.00
22.00
Time
23.00
Figure 3.53: A, Diode array chromatogram of apdABCEG expression clone showing the production of 14-hydroxy
preaspyridone 238 at 8.9 minutes, diastereomers of preaspyridone A 224 and 230 at 13.9 and 14.4 minutes, aspyridone
A 84 at 14.9 minutes, N-hydroxy dephenylated pyridone 259 at 17 minutes and O methoxy dephenylated pyridone 260
at 21.2 minutes; B, Diode array chromatogram of wild type A. oryzae.
The yield of these compounds from 1 litre fungal culture was calculated by
quantification experiment (see section 3.6.10). The quantity of 14-hydroxy
preaspyridone 238 was 42.6 mg/L, major diasteomer of preaspyridone A 230 was 98
mg/L, aspyridone A 84 was 12.6 mg/L, N-hydroxy dephenylated pyridone 259 was 15.8
mg/L and
N-O methoxy dephenylated pyridone 260 was 8.2 mg/L. The above
mentioned compounds are the same as observed in the apdABCE expression clone only
they are produced in higher titres in apdABCEG expression clone (see section 3.7.4).
From the above heterologous expression we couldn’t settle a precise role for apdG only
that when it was expressed with other four genes from the cluster it displayed increase
in the production of metabolites.
141
3.7.7
Heterologous expression of apdAC and tenA in A. oryzae
The tenA gene encodes a cytochrome P450 enzyme during tenellin 87
biosynthesis in B. bassiana 110.25 and the protein encoded by apdE in the aspyridone
gene cluster displays 48 % sequence similarity to TenA. TenA catalyses the oxidative
ring expansion of the pyrrolidone in pretenellin A 114 to a 2-pyridone compound,
pretenellin B 115.108 We planned to express megasynthase apdA and enoyl reductase
encoding gene apdC with tenA in A. oryzae to determine if TenA displays broader
substrate selectivity and turnover of preaspyridone A 224 and 230 produced by apdAC
to aspyridone A 84. Earlier the Cox group achieved effective gene swaps between
tenellin 87 and DMB 88 when enoyl reductase dmbC from DMB 88 cluster and PKSNRPS tenS were co-expressed in A. oryzae, they produced a precursor compound of
tenellin 87 pathway, pretenellin A 114.109
Dr. Khomaizon Pahirulzaman constructed a vector pTAYAargACtenA carrying
PKS-NRPS coding gene apdA, enoyl reductase encoding gene apdC and ring expandase
gene tenA and expressed it in A. oryzae. One producing transformant was grown on
DPY solid media and later mature spores were inoculated in aspyridone growth media
(CMP media) after growing for 7-10 days. The fungal culture was grown in 100 ml
liquid media in 250 ml flask at 28 °C on shakers at a speed of 200 rpm for seven days.
Mature mycelia were homogenised and extracted with ethyl acetate (see section 4.11).
The organic layer was concentrated and defatted. A crude extract of 65.3 mg/100 ml
was obtained after drying. The crude extract prepared in HPLC grade methanol (10
mg/ml) was analysed on analytical LCMS programme with CH3CN/H2O having a
gradient of 55-65% in 30 minutes (Figure 3.54).
142
14.3
3.38
A
2.62
4.55
2.00
4.00
13.3
6.07
6.00
8.00
10.00
12.00
28.52
14.00
16.00
18.00
20.00
22.00
24.00
26.00
28.00
3.22
Wildtype A. oryzae
3.45
5.80
B
4.25
27.43
2.60
2.00
4.00
6.00
8.00
10.00
12.00
14.00
16.00
18.00
20.00
22.00
24.00
26.00
28.00
Time
Figure 3.54: A, Diode array chromatogram of apdACtenA expression clone showing production of minor preaspyridone
A diastereomer 224 at 13.3 min. and major preaspyridone A diastereomer 230 at 14.3; B, Diode array chromatogram
of wild type A. oryzae.
The LCMS chromatogram displayed production of major and minor diasteomers
of preaspyridone A 224 and 230 presenting a molecular ion of 332 [M]H+ and a uv
spectrum of 222, 278 nm. This experiment indicated that TenA has restricted substrate
specificity and didn’t convert preaspyridone A 224 to aspyridone A 84. But the titres of
minor component of preaspyridone A 224 (2.2 mg/L) and major preaspyridone A 230
(121 mg/L) in apdACtenA expression clone was higher than produced in apdAC
expression (see section 3.7.1). A similar effect in increase in preaspyridone A 224 and
230 was observed in apdABC expression. With presence of ApdB and TenA we didn’t
observe any new metabolites but the production of preaspyridone A 224 and 230 was
increased.
3.7.8
Heterologous expression of apdACE and tenB in A. oryzae
TenB, a cytochrome P450 protein in tenellin 87 gene cluster catalyses the N-
hydroxylation of the 2-pyridone in pretenellin B 115 and froms tenellin 87.108 In an A.
oryzae transformation we planned to express tenB with apdACE to explore the catalytic
activity of TenB and whether it N-hydroxylates the 2-pyridone compound, aspyridone A
143
84. Dr.Khomaizon Pahirulzaman modified the vector pTAYAargASP to form
pTAYAargACEtenB carrying genes apdACE and tenB.
A number of A. oryzae transformants were obtained and one was selected for
screening by LCMS. The transformant was grown on DPY media and after 7-10 days
when spores were well grown, spore solution was inoculated in CMP liquid media (1 x
100 ml) in a 250 ml flask for seven days at 28 °C (see section 4.10). The fungus culture
was extracted and defatted in the usual way. A solid mass weighing 27.4 mg/100ml was
obtained. 10 mg/ml in HPLC methanol was made from the crude extract and analysed
by LCMS with CH3CN/H2O with gradient 55-65% in 30 minutes.
15.4
4.42
4.78
A
5.02
2.00
4.00
14.5
9.5
2.78
6.00
3.22
8.00
10.00
12.00
14.00
15.9
16.00
18.00
20.00
22.00
24.00
Wildtype A. oryzae
3.45
5.80
B
4.25
2.60
Figure 3.55: A, Diode array chromatogram of A. oryzae apdACEtenB expression clone displaying production of 14hydroxy preaspyridone A 238, preaspyrdione A 230, dephenylated aspyridone 239 and aspyridone A 84; B, Diode array
chromatogram of wild type A. oryzae.
The LCMS chromatogram revealed the production of 14-hydroxy preaspyridone
A 238 (m/z 348 [M]H+, λmax 223, 281 nm), preaspyridone A 230 (m/z 332 [M]H+,
λmax223, 280 nm), dephenylated aspyridone 239 (m/z 238 [M]H+, λmax229, 325 nm ) and
aspyridone A 84 (m/z 330 [M]H+, λmax247, 344 nm). One litre (10 x 100 ml) of
apdACEtenB clone was grown for purification of compounds. The crude extract
achieved after extraction and concentration was 764 mg/L. The extract was made to a
144
concentration of 50 mg/ml and 100 µl was injected in each run of auto purification on
mass directed preparative LCMS. The solvents used were acetonitrile/water with a
gradient of 50-75% carried out in 30 min (Method 3) programme. The purified
fractions of each compound was collected and dried. All above stated compounds were
verified with 1H NMR spectroscopy. The yield of 14-hydroxy preaspyridone A 238 was
5.2 mg, preaspyridone A 230 was 11 mg, dephenylated aspyridone 239 was 28 mg and
aspyridone A 84 was 2.3 mg.
We didn’t observe any new or N-hydroxylated 2-pyridone compounds which
confirm that tenB is highly selective and didn’t take any 2-pyridone compounds from
aspyridone pathway as its substrate.
3.7.9
[1-13C] L-Tyrosine feeding
Earlier aspyridone A 84 and preaspyridone A 224 were reported from hybrid
PKS-NRPS gene cluster by Bergmann et al.80 and Tang107 and co-workers. They are
synthesized from the fusion of a tetraketide 220 and an amino acid tyrosine 107. With
hetererologous expression of apdACE genes and consequential later experiments we
achieved
novel
dephenylated
aspyridone
compound,
4-hydroxy-3-(2,
4-
dimethylhexanoyl) 2-pyridone 239. We carried out [1-13C] L-tyrosine feeding with 5dephenylated pyridone 239 producing clone apdACEtenB, firstly to determine if 5dephenylated pyridone 239 utilize L-tyrosine as its amino acid precursor and secondly
to ascertain that dephenylated 2-pyridone 239 is an oxidative breakdown product of
aspyridone pathway.
25 mg of [1-13C] L-tyrosine was dissolved in 3 ml of deionized water with 5µl of
1 molar NaOH. 1 ml of this solution was added in three flasks each containing 100 ml
of 3 days old fungal culture of A. oryzae apdACEtenB in 250 ml Erlenmeyer flask. This
was repeated on 4th and 5th day as well. One 250 ml flask containing 100 ml of media
was grown as a control with no [1-13C] feeding. The fungal cultures were grown
according to the standard fermentation conditions (see section 4.10). On the seventh day
the fungal mycelia was homogenized, acidified and extracted with ethyl acetate. The
organic layer was semi concentrated and defatted. After defatting the extract was dried
to a brown mass weighing 71.7 mg/300 ml. A 50 mg/ml solution was made with HPLC
145
grade methanol and 50µl was injected in each run of mass directed auto-purfication on
LCMS instrument. The gradient used on a CH3CN/H2O solvent was 55-60 % in 30 min.
duration. The purified fractions were collected and fully dried under nitrogen gas. The
yield of 14-hydroxy preaspyridone 238 was 2.8 mg, preaspyridone A 230 was 6.8 mg,
5-dephenylated pyridone 239 was 9.8 mg and aspyridone A 84 was 1.4 mg. Each of
these compounds was studied by 1D 13C NMR spectroscopy and compared parallel with
un-labelled pure compounds.
A
13
C feeding
No feeding
B
Figure 3.56: A, 13C NMR showing incorporation of [1-13C] L-tyrosine at C-4 in preaspyridone A 230; B, 13C NMR of
preaspyridone A 230 without isotope feeding.
The products of A. oryzae apdACEtenB expression clone are 14- hydroxy
preaspyridone A 238, preaspyridone A 230, dephenylated 2- pyridone 239 and
aspyridone A 84. In [1-13C] L-tyrosine feeding preaspyridone A 230 exhibited 30%
incorporation of L-tyrosine at C-4 (Figure 3.56) and dephenylated 2-pyridone 239
displayed 13% incorporation of [1-13C] L-tyrosine at C-4 (Figure 3.57). We didn’t
observe any incorporation of isotope labelled tyrosine in 14- hydroxy preaspyridone A
238 and aspyridone A 84. The feeding experiment confirmed that dephenylated
146
pyridone 239 derives the same PKS-NRPS biosynthetic pathway as preaspyridone A
230 and aspyridone A 84. It also confirms that the adenylation domain of the NRPS
selects the amino acid, L-tyrosine.
13
C feeding
A
No feeding
B
Figure 3.57: A, 13C NMR showing incorporation of [1-13C] L-tyrosine at C-4 in dephenylated 2-pyridone 239; B, 13C
NMR of dephenylated 2-pyridone 239 without isotope feeding.
3.7.10 Quantification of metabolites
The titres of different compounds produced in various heterologous expression
clones can vary owing to differences in metabolic and environmental conditions. For
this reason, we carried out quantification of all aspyridone metabolites by measuring
standard calibration curves. Serial dilutions of pure compounds from 30 µg/ml-1000
µg/ml were made with HPLC grade methanol and diode array chromatograms were
obtained on LCMS with acetonitrile/water using gradient 50-70% in 20 min. Integration
values of absorption peaks were recorded at the λmax value for each compound. Graphs
were plotted between integration values versus different concentration used in serial
dilution. This gave a standard calibration curve and linear equation for each compound.
147
100 ml of each A. oryzae transformant in a 250 ml flask was grown and crude extracts
were obtained according to the method given in section 4.11. LCMS was run on these
samples and peaks were first identified by standard retention time, uv spectra and mass
spectra. The yields of each compound in the crude extracts were then calculated from
their respective standard linear equation at their respective retention time and λmax value.
The values are arranged in Table 3.1.
148
Table 3.1: Quantification of metabolites in mg/L.
Transformant
238
224
230
84
260
259
239
267
240
Total
yield
apdA,C
430 mg
0
0.61
60.70
0
0
0
0
0
trace
61.3
apdA,C,E
1126 mg
183.6
0
0
119.8
0
0
72.2
0
15
390.6
apdA,B,C
805 mg
0
2.2
141.0
0
0
0
0
0
trace
143.2
apdA,C,tenA
653 mg
0
2.2
121.0
0
0
0
0
0
trace
123.2
apdA,C,E,B
300 mg
2.6
5.7
34.6
1.8
2.7
6.2
trace
4
trace
53.6
apdA,C,E,tenB
(274mg from 100ml)
2.9
0
5.3
3.2
0
0
6.9
6
0
24.3
apdA,C,E,D
448 mg
apd A,B,C,E,G
(62.2 mg from
100ml)
8.2
0
0.3
5.1
0
0
49.9
8
0
71.5
42.6
0
98
12.6
8.2
15.8
0
trace
trace
177.2
149
3.7.11 Discussion and Conclusions
Aspyridone A 84 and aspyridone B 226 were reported to be the exclusive
products of a silent hybrid PKS-NRPS gene cluster in A. nidulans.80 It was discovered
by the Hertweck group80 and they activated the cluster by overexpressing the
transcription regulator apdR. In this Chapter we studied the gene cluster of aspyridone
84 using a heterologous gene expression technique in A. oryzae via a lately devised
multi gene expression plasmid pTAYAGSargPage.176 It consists of strong constitutive
promoters for each subject gene and resulted in successful transcription of apd genes in
the heterologous host. The coexpression of apd genes in A. oryzae in an orderly scheme
disclosed advance oxidative modification and programming potential of respective
genes leading to a number of new compounds.
Heterologous expression of apdAC formed the previously known precursor
compound preaspyriodone A 224.107 In our project preaspyridone A was isolated as a
major and a minor diastereomer compounds 224 and 230, being epimers at C-5 (section
3.7.1). The formation of crystals of minor preaspyridone A isomer 224 presented antiarrangement of the pendent methyl groups of the alkane chain which is opposite to that
reported before.80 We established that aspyridone A 84 is the product of three genes
apdAC and apdE. Similar result is reported by Niehaus et al.,197 they studied the
biosynthesis of fusarin C in the fungus Fusarium fujikuroi. The fusarin gene cluster
consists of nine genes but they proved that only four genes are required for the
biosynthesis of fusarin C. So, we concluded that it is difficult to predict what a gene
actually does and how many genes are required for the biosynthesis of a single
compound.
The A. oryzae apdACE expression clone showed distinctive catalytic feature of
cytochrome P450 apdE that it performs oxidative ring expansion of preaspyridone A
224 to form aspyridone A 84, benzylic hydroxylation of 224 to form 14-hydroxy
preaspyridone A 238 and loses a phenoxide to form the novel compound, 5dephenylated pyridone 239. The apparent incorporation of [1-13C]-L-tyrosine by 5dephenylated pyridone 239 and the precursor compound preaspyridone 230 confirms
that tyrosine is the precursor amino acid and biosynthesis of 239 is linked to
preaspyridone 230. The structure of 5-dephenylated pyridone 239 and its derivatives
259 and 260 are similar to known fungal metabolites, the Atpenins 256-258. The
150
biosynthetic pathway of Atpenins is not reported and we suggest that they must be
products of a similar PKS-NRPS cluster like the aspyridone genes. The polyketide chain
in Atpenins and in 84, 224, 239, possess pendant methyls arranged in anti configuration.
The methyls are derived form S-adenosyl methionine by the CMeT domain. The ER
domain in ApdA is inactive and the stand alone enoyl reductase ApdC acting in trans
sets the stereochemistry of the carbon bearing the methyls. The ApdC remarkably sets
an opposite stereochemistry of the two methyls during the first and second cycle of the
tetraketide producing anti dimethylation pattern. The characterization of 18-deshydroxy
preaspyridone A 240 in apdACE and in trace amounts in other transformants (Table 3.1)
shows a wider specificity of the adenylation domain to incorporate other amino acids
than L-tyrosine. This is also in agreement of the in vitro studies performed by Tang and
coworkers.107
The gene apdB encodes a cytochrome P450 enzyme which didn’t accomplish
any chemical step when expressed with apdAC alone but was observed to increase the
titres of the metabolites. However, when expressed with apdACE in A. oryzae apdACEB
expression it was resolved that it catalyses N- hydroxylation of dephenylated 2 pyridone
239 to form a new compound 259. The apdACEB was an exclusive clone to form nine
different compounds (Table 3.1) owing to the presence of two cytochrome P450s but
the overall yield of this transformant was less (53.6 mg/L) as compared to apdACE (390
mg/L).
We did not perceive a role for the FAD dependent mono oxygenase apdD. As
we didn’t observe the production of aspyriodone B 226, we suspect it might have been
catalysed by a gene outside the cluster which is possible in transcription mediated
recombination where simulataneously many genes are turned on. The presence of apdD
also lowered the yield (71.5 mg/L) of metabolites in apdACED expression than
achieved in apdACE transformant (Table 3.1). The gene apdG didn’t exhibit any
catalytic role in apdACEBG expression clone but increased the titres of metabolites
produced. This might due to the fact that biosynthetic proteins work in clusters and with
more proteins in a heterologous expression there is more protein-protein interaction and
more production. But it cannot be concluded decisively without further experimental
studies. The expression of tenellin genes with apd genes showed high substrate
specificity of tenellin biosynthetic genes as we didn’t observe their particular role in the
151
transformants. The presence of tenA with apdAC increased the titres of the compounds
with a similar effect as apdB in apdABC expression clone.
Thus heterologous expressions of apd genes in suitable host, A. oryzae produced
new compounds and help understand the role of specific genes in aspyridone cluster of
A. nidulans.
152
Chapter 4
Experimental
4.1
General Chemicals and Equipment
All chemicals and reagents used were of analytical grade and obtained from
Sigma Aldrich, Fischer, Fluka analytical and BDH laboratories. All solvents used in
HPLC and purification were HPLC grade. Deionized water was used in all experiments.
Weighing balances were of Sartorius AX224. Small centrifuge for obtaining clear
sample were from AG Hanburry 22331. Sterilizations were carried out using Astell
Autoclave at 121 ºC for 15 minutes. Optical rotations were measured with an ADP 220
polarimeter at 589 nm. Melting points were determined using Electrothermal apparatus.
IR data were obtained using a Perkin–Elmer FTIR instrument.
4.2
Mass Spectrometry
Electrospray ionization (ESI) mass spectra were recorded on a VG Quattro-Mass
spectrometer or Bruker microtof mass spectrometer.
4.3
Nuclear Magnetic resonance Spectroscopy
NMR experiments were conducted on Varian VNMRS-500 spectrometer, 1H
NMR at 500 MHz and
13
C NMR at 125 MHz. Chemical shifts were recorded in parts
per million (ppm) and coupling constant (J) in Hz.
4.4
Analytical LCMS
All crude extracts were prepared to a concentration of 10 mg/ml in HPLC grade
methanol, centrifuged for 60 seconds and supernatant was placed in LCMS vials. 20 µl
of the extracts were injected and analysed on a Waters 2795HT HPLC system.
Detection was achieved by uv between 200 and 400 nm using a Waters 998 diode array
detector, and by simultaneous electrospray (ES) mass spectrometry using a Waters ZQ
spectrometer detecting between 150 and 600 m/z units. Chromatography (flow rate 1
ml/min) was achieved using a Phenomenex LUNA column (5 μ, C18, 100 Å, 4.6 × 250
mm) equipped with a Phenomenex Security Guard precolumn (Luna C5 300 Å) or
Chromatography was achieved using a Phenomenex Kinetex column (2.6 µ, C18, 100
153
Å, 4.6 x 100 mm) equipped with a Phenomenex Security Guard precolumn (Luna C5
300 Å). Solvents used were: A, HPLC grade H2O containing 0.05 % formic acid; B,
HPLC grade MeOH containing 0.045 % formic acid; and C, HPLC grade CH3CN
containing 0.045 % formic acid. The following gradients were used:
Method 1. Luna/MeOH: 0 min, 25% B; 5 min, 25% B; 51 min, 95% B; 53 min, 95%
B; 55 min, 25% B; 59 min, 25% B; 60 min, 25% B.
Method 2. Luna/MeOH: 0 min, 25% B; 13 min, 95% B; 15min, 95% B; 17 min,
25% B; 20 min, 25% B.
Method 3. Kinetex/MeOH: 0 min, 10% B; 10 min, 90% B; 12 min, 90% B; 13 min,
10% B; 15 min, 10% B.
Method 4. Kinetex/CH3CN: 0 min, 10% C; 10 min, 90% C; 12 min, 90% C; 13 min,
10% C; 15 min, 10% C.
Method 5. Luna/CH3CN: 0 min, 5% C; 5 min, 5% C; 45 min, 75% C; 46 min, 95% C;
50 min, 95% C; 55 min, 5% C; 60 min, 5% C.
4.5
Preparative HPLC
Purification of compounds was generally achieved using a Waters mass-directed
autopurification system comprising of a Waters 2767 autosampler, Waters 2545 pump
system, a Phenomenex LUNA column (5µ, C18, 100 Å, 10 × 250 mm) equipped with a
Phenomenex Security Guard precolumn (Luna C5 300 Å) eluted at 4 ml/min. Solvents
used were: A, HPLC grade H2O + 0.05% formic acid; solvent B, HPLC grade MeOH +
0.045% formic acid; solvent C, HPLC grade CH3CN + 0.045% formic acid. The postcolumn flow was split (100: 1) and the minority flow was made up with solvent A to 1
ml/min for simultaneous analysis by diode array detector (Waters 2998), evaporative
light scattering (Waters 2424) and ESI mass spectrometry in positive and negative
modes (Waters Quatro Micro).
Method 1: 0 min, 25% B; 13 min, 95% B; 15 min, 95% B; 17 min, 25% B; 20 min,
25% B.
Method 2: 0 min, 40% C; 15 min, 80% C; 15.50 min, 95% C; 16.50 min, 95% C;
17 min, 40% C; 20 min, 40% C.
Method 3: 0 min, 50% C; 22 min, 75% C; 24 min, 95% C; 26 min, 95% C; 27 min,
50% C; 30 min, 50% C.
154
Method 4: 0 min, 55% C; 22 min, 60% C; 24 min, 95% C; 26 min, 95% C; 27 min,
55% C; 30 min, 55% C.
Method 5: 0 min, 55% C; 22 min, 75% C; 24 min, 95% C; 26 min, 95% C; 27 min,
55% C; 30 min, 55% C.
Method 6: 0 min, 50% C; 22 min, 65% C; 24 min, 95% C; 26 min, 95% C; 27 min,
50% C; 30 min, 50% C.
4.6
X-ray Crystallography
X-ray diffraction data were analysed on a Bruker Microstar rotating anode
diffractometer using Cu-Kα radiation (λ = 1.54178 Å). Data collections were performed
using a CCD area detector from a single crystal mounted on a glass fibre. Absorption
corrections were based on equivalent reflections using TWINABS or SADABS. The
structures were solved using direct methods using SHELXS and refined against all Fo2
data with hydrogen atoms on carbon and oxygen atoms riding in calculated positions
using SHELXL.
4.7
Solid media for growth of fungal spores
The Aspergillus oryzae transformants and were first grown on Czapek Dox
Agar (minimal media) and for further production of spores, they were inoculated and
grown on DPY solid media for 7-10 days at 25 °C. Beauveria bassiana WT and RNAi
strains of B. bassiana were grown on Potato Dextrose Agar (PDA) for 10 days at 25 °C.
4.7(a) Czapek Dox Agar
50 grams of Czapek Dox agar was dissolved on 1 litre of deionized water and
sterilized in autoclave.
4.7(b) DPY media
The spores of A. oryzae transformants were grown on plates in DPY media
(dextrin-peptone-yeast extract). It is made by the following ingredients 2% (w/v)
dextrin, 1% (w/v) polypeptone, 0.5% (w/v) yeast extract, 0.5% (w/v) potassium
dihydrogen phosphate, 0.05% (w/v) magnesium sulphate and 2.5% (w/v) agar.
155
4.7(c) PDA media
B. bassiana was grown on potato dextrose agar for growth of spores. 39 gram of
PDA was dissolved in 1 litre of deionized water and sterilized by autoclave.
4.8
Preparation of Tenellin production Media
The liquid media used for growing B. bassiana spores was Tenellin production
media (TPM). The ingredients include D-mannnitol (50 g), KNO3 (5 g), KH2PO4 (1 g),
MgSO4·7H2O (0.5 g), NaCl (0.1 g), CaCl2 (0.2 g), FeSO4·7H2O (20 mg) and mineral
ion solution (10 ml, ZnSO4·7H2O (880 mg), CuSO4·5H2O (40 mg), MnSO4·4H2O (7.5
mg), boric acid (6 mg), and (NH4)6Mo7O24·4H2O (4 mg) made up to 1 L in deionized
water) made up to 1 L in deionized water. This medium solution was divided into 100
ml each in 500 ml Erlenmeyer flask, covered with foam bung covered with aluminium
foil and sterilized by autoclaving.
4.9
Czapek-Dox minimal medium (CD)
A. oryzae tenPKS-dmbC strain was grown in CD minimal media. It was
prepared by adding 10 g peptone, 20 g glucose, and 30 g sucrose, 50 ml of solution A
(40 g NaNO3, 40 g KCl, 10 g MgSO4.7H2O, 0.2 g FeSO4.7H2O in 1 litre deionized
water) and 50 ml of solution B (20 g K2HPO4 in 1 litre deionized water) in 1 litre
deionized water. 100 ml of the media was divided in 500 ml conical Erlenmeyer flasks,
covered with bung form and alumium foil and autoclaved. The A. oryzae spores were
inoculated in flasks and were incubated for 4 days at 28 °C and shaken at 200 rpm. This
was followed by changing the media (under sterile condition) with induction medium
for production of secondary metabolites. The inducing medium was made by dissolving
20 g starch, 10 g peptone, 50 ml solution A and 50 ml solution B in 1 litre deionized
water and incubated under the same previous conditions over 5-7 days.
4.9(a) Czapek-Maltose-Polypeptone (CMP) medium A
This medium was prepared by adding 20 g maltose, 10 g polypeptone, 50 ml
solution A and 50 ml solution B in 900 ml deionized water. The medium was divided
into 100 ml each in 500 ml Erlenmeyer flask, covered with foam bung and aluminium
foil and then sterilized by autoclaving.
156
4.9(b) Czapek-Maltose-Polypeptone (CMP) medium B
The liquid medium for A. oryzae transformants studied in aspyridone
biosynthesis (chapter 3) was made by adding 30 g sucrose, 20 g maltose, 10 g
polypetone, 50 ml solution B and 50 ml solution C (60 g NaNO3, 10 g KCl, 10 g
MgSO4.7H2O and 0.2 g FeSO4.7H2O in 1 litre deionized water) in 900 ml deionized
water. The medium (100 ml) was divided in 500 ml Erlenmeyer flasks as described
previously.
4.10
Culturing and inoculation of fungal spores in liquid media
10 mL of deionized water was added on plates having 10 days old growing
spores of the fungus. The spores were made to pass into the deionized water by careful
scratching the surface with sterilized loop. The spore suspension (1 ml) from the plate
was added in each 100 mL liquid medium contained in 500 ml Erlenmeyer flask. The A.
oryzae spores were allowed to grow in the liquid culture for 7 days on shakers at 200
rpm at 25 °C- 28 °C and B. bassiana spores were grown on shakers for at least 10 days
at 25 °C at 150-200 rpm.
4.11
Extraction of A. oryzae transformants
Cells and media (1 L) were homogenized using a hand-held electric blender and
then acidified to pH 4.0 using 37% aqueous HCl. An equal volume of ethyl acetate was
added and stirred for 10 min. The resulting mixture was vacuum filtered through
Whatman no. 1 filter paper. The filtrate was transferred into a separating funnel and
shaken vigorously. The mixture was allowed to stand to separate the layers. The organic
layer was washed once with concentrated brine solution and then with deionized water.
The organic phase was dried (MgSO4), filtered and evaporated to dryness. The crude
extract was dissolved in 10 % aqueous methanol and defatted by extraction with
hexane. The methanolic layer was evaporated to dryness and then made into a solution
of 10 mg/ml in HPLC grade methanol and analysed by LCMS. For purification of
compounds the crude extract was made into a solution of 50 mg/ml in HPLC grade
methanol and 200 µl aliquots were injected in each run of mass-directed HPLC
preparative purification.
157
4.12
Extraction of B. bassiana transformants
The 10 days old cultures of B. bassiana were vacuum filtered and the mycelia
was collected and kept in equal volume of acetone overnight. The acetone layer was
separated from the mycelia by filtration with Whatman filter paper and acetone was
concentrated by vacuum to form a brown aqueous extract. It was further diluted with
deionized water and then extracted with ethyl acetate (2 × 500 ml). The ethyl acetate
layer was separated from the water layer by a separating funnel. MgSO4 was added to
absorb any trapped water droplets. The ethyl acetate was filtered and concentrated under
vacuum to obtain a semi solid crude extract. The extract was dissolved in 10 % aqueous
methanol and defatted by extraction with hexane. The methanolic layer was obtained in
separated glass vial and dried. The crude extract was prepared to a concentration of 10
mg/ml in HPLC grade methanol and analysed by LCMS.
4.13
Purification of metabolites from A. oryzae pTAex3-tenS and A. oryzae
tenSPKS-dmbNRPS
The 1 litre culture of A. oryzae pTAex3-tenS was extracted by the procedure
explained above and a crude extract of 70 mg was achieved. The extract was dissolved
in HPLC grade methanol to a concentration of 50 mg/ml and used for mass-directed
HPLC preparative purification. 100 – 200 μl of the crude solution was injected during
successive rounds of a 20 minute HPLC program (Method 1). Fractions corresponding
to prototenellin C 113 were collected and evaporated to yield 2.4 mg of pure compound.
Similar protocols were used in purifying the prototenellin C 113 from 137 mg of a crude
extract of A. oryzae tenSPKS-dmbNRPS obtained from 1 L fermentation and produced
9.6 mg of the pure compound.
4.13(a)
Characterization of prototenellin C 113109
Prototenellin C 113, pale yellow solid, 1H NMR (CD3OD, 500 MHz) δ = 1.15 (d, J =
6.4 Hz, 3H, H-13), 1.29 (s, 3H, H-14), 1.93 (s, 3H, H-15), 2.85 (m,1H, H-16a), 2.99 (m,
1H, H-16b), 3.62 (m, 1H, H-12), 3.94 (m, 1H, H-5), 6.15 (d, 1H, J = 15 Hz, H-10), 6.68
158
(d, J = 8.2 Hz, 2H, H-19, H-21), 6.71 (m, 1H, H-9), 6.98 (m, 1H, H-8), 7.03 (d, J = 8.2
Hz, 2H, H-18, H-22);
13
C NMR (CD3OD, 125 MHz), δ = 11.5 (C-15), 16.3 (C-13),
22.9 (C-14), 36.3 (C-16), 61.2 (C-5), 73.3 (C-12), 75.3 (C-11), 114.4 (C-19, C-21),
124.2 (C-9), 126.3 (C-17), 130.5 (C-18, C-22), 131.7 (C-7), 138.9 (C-8), 145.5 (C-10),
155.0 (C-20), 163.1 (C-2), 188.3 (C-6), 194.0 (C-4). The
13
C signal for C-3 was not
observed. HRMS calculated for C21H25NO6Na: 410.1580; found 410.1578 [M]Na+.
Purification of metabolites from A. oryzae dmbS –tenC and A. oryzae dmbS-
4.14
dmbC
A 1 litre fermentation media was grown for A. oryzae dmbS –tenC at 25 °C (10
flasks × 100 mL) for 7 days on shakers at 200 rpm. The fungal cultures were extracted
with protocol explained in section 4.14, which gave a dark brown crude extract of 111
mg. This was made to a solution of 50 mg/ml with HPLC grade methanol for
purification of compounds on mass- directed preparative HPLC. 100µl - 200µl of crude
extract was injected in each successful preparative run. Pure fractions having bright
yellow colour of predmB A were collected after 25 preparative runs and dried to
evaporation. 23 mg of pure compound was achieved. 363 mg of crude extract was
obtained from a 1 litre culture of A. oryzae dmbS-dmbC and 17.6 mg of pure predmb A
was produced by following a similar protocol as explained above.
4.14(a)
Characterization of predmbA 118
PreDMB A 118, brownish yellow solid; IR (neat): νmax 3277, 2961, 2922, 2876,
1648, 1594, 1514 cm-1; 1H NMR (CD3OD, 500 MHz) δ = 0.92 (t, J = 7.5 Hz, 3H, H15),
1.09 (d, J = 7.5 Hz, 3H, H-16), 1.41 (m, 2H, H-14), 2.20 (m, 1H , H-13), 2.90
(brd, 1H, H-17a), 3.00 (brd, 1H, H-17b), 4.08 (brs, 1H, H-5), 5.98 (dd, J = 7.8, 15.2 Hz,
1H, H-12), 6.27 (dd, J = 10.8, 15.2 Hz, 1H, H-11), 6.43 (dd, J = 11.9, 14.5 Hz, 1H, H9), 6.70 (d, J = 8.1 Hz, 2H, H-20, H-22), 6.83 (dd, J = 14.5, 10.8 Hz, 1H, H-10), 7.00
(d, J = 8.1 Hz, 2H, H-19, H-23), 7.21 (d, J = 15.4 Hz, 1H, H-7), 7.53 (dd, J = 11.9, 15.4,
159
Hz, 1H, H-8), 13C NMR (CD3OD, 125 MHz) δ = 10.7 (C-15), 18.7 (C-16), 29.2 (C-14),
36.2 (C-17), 38.8 (C-13), 62.8 (C-5), 114.7 (C-20, C-22), 119.7 (C-7), 126.3 (C-18),
128.7 (C-11), 128.8 (C-9), 130.4 (C-19, C-23), 144.3 (C-10), 145.2 (C-8), 147.8 (C-12),
155.9 (C-21), 173.3 (C-6), 173.7 (C-2), 195.9 (C-4). The
13
C signal for C-3 was not
observed. HRMS calculated for C22H26NO4: 368.1856; found 368.1851 [M]H+.
4.15
Purication of metabolites from A. oryzae tensPKS –dmbC
3 litres of A. oryzae tenSPKS-dmbC strain (30 flasks × 100 ml) were grown at 28
°C at 200 rpm. After extraction of fungal cultures with ethyl acetate (see section 4.14), a
crude extract of 239 mg was obtained. HPLC grade methanol was added to make a 50
mg/ml solution for mass-directed purification of compounds on preparative HPLC. The
crude extract was subjected to 24 successful preperative runs with Method 2 and two
pure metabolites were obtained. The yield of compound 146 was 9.5 mg and compound
147 was 9 mg.
4.15(a)
Characterization of Compound A 146
Light brown viscous oil, [α]22D -16.9 (c = 0.23, MeOH); IR (neat): νmax 2932, 2872,
2342, 1761, 1631, 1355, 1190 1084 cm-1; 1H NMR (CDCl3, 500 MHz) δ = 0.92 (t, J = 7
Hz, 3H, H-1), 1.36 (m, 2H, H-2), 1.37 (m, 1H, H-3a), 1.45 (m, 1H, H-3b), 1.65 (m, 1H,
H-4a), 1.81 (m, 1H, H-4b), 2.01 (ddd, J = 12, 10.5, 12 Hz 1H, H-6a), 2.55 (ddd, J = 9,
6, 12 Hz, 1H, H-6b), 3.65 (t, J = 9, 12 Hz, 1H, H-7), 4.43 (m, 1H, H-5), 5.95 (b, 1H, H10a), 6.54 (b, 1H, H-10b), 13C NMR (CDCl3, 125 MHz) δ = 14.4 (C-1), 22.9 (C-2), 27.8
(C-3), 35.5 (C-4), 36.1 (C-6), 45.2 (C-7), 79.6 (C-5), 131.8 (C-10), 136.1 (C-8),
170.1(C-9), 176.4 (C-11). HRESIMS calculated for C11H17O4: 213.1121; observed
213.1130 [M]H+.
160
4.15(b)
Characterization of compound B 147
light color viscous oil, IR (neat): νmax 2962, 2930, 2873, 1736, 1582, 1454, 1045, 878
cm-1; 1H NMR (CDCl3, 500 MHz) δ = 0.92 (t, J = 7 Hz, 3H, H-1), 1.38 (m, 2H, H-2),
1.46 (m, 2H, H-3), 1.71 (m, 1H, H-4a), 1.79 (m, 1H, H-4b), 4.99 (t, J = 6.2 Hz, 1H, H5), 6.79 (b, 1H, H-10a), 7.19 (b, 1H, H-10b), 7.96 (b, 1H, H-6); 13C NMR (CDCl3, 125
MHz) δ = 14.0 (C-1), 22.6 (C-2), 27.3 (C-3), 33.1 (C-4), 80.8 (C-5), 125.0 (C-7), 128.5
(C-8), 133.6 (C-10), 153.5 (C-6), 169.5 (C-9), 171.7 (C-11). HRESIMS calculated for
C11H14O4Na: 233.0784; observed 233.0799 [M]Na+.
4.16
Purication of metabolites from A. oryzae apdACE
A brown crude extract of 1303 mg was formed from 1 litre culture of A. oryzae
apdACE strain by growing the fungus according to standard conditions and extraction
with ethyl acetate. The crude extract was made to a solution of 50 mg/ml with HPLC
grade methanol for purification of metabolites on the preparative HPLC by following
the gradient given in Method 5. Pure fractions of corresponding metabolites were
collected and dried to evaporation.
Characterisation of metabolites from A. oryzae apdACE
4.16(a)
14- Hydroxy preaspyridone A 238
Brown viscous oil, [α]22D -158.8 (c = 1.29, MeOH); IR (neat): νmax 3317, 3020, 2964,
1651, 1214 cm-1; 1H NMR (CD3OD, 500 MHz) δ = 0.81 (m, 3H, H-11), 0.82 (m, 3H,
H-12), 0.95 (d, J = 6.5 Hz, 3H, H-13), 1.09 (m, 1H, H-10a), 1.23 (m, 1H, H-9), 1.29 (m,
1H, H-8a), 1.33 (m, 1H, H-10b), 1.40 (m, 1H, H-8b), 3.55 (q, J = 7 Hz, 1H, H-7), 4.22
(d, J = 3.5 Hz, 1H, H-5), 4.99 (d, J = 4 Hz, 1H, H-14), 6.65 (d, J = 8.5 Hz, 2H, H-17,
H-19), 7.11 (d, J = 8.5 Hz, 2H, H-16, H-20) ; 13C NMR (CD3OD, 125 MHz) δ = 11.4
161
(C-11), 17.5 (C-13), 19.6 (C-12), 30.2 (C-10), 33.5 (C-9), 35.5 (C-7), 41.2 (C-8), 68.5
(C-5), 74.8 (C-14), 115.5 (C-17, C-19), 129.6 (C-16, C-20), 130.3 (C-15), 158.6 (C18), 195.2 (C-6). The
13
C signals for C-2, C-3 and C-4 were not observed. HRESIMS
calculated for C19H25NO5Na: 370.1624; observed 370.1624 [M]Na+.
4.16(b)
4-hydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 239
Light brown solid, mp 120 °C; [α]22D -24.3 (c = 0.82, MeOH); IR (neat): νmax 3408,
3298, 2923, 2287, 1734, 1601, 1462, 1227 cm-1; 1H NMR (DMSO, 500 MHz) δ = 0.79
(m, 3H, H-12), 0.81 (m, 3H, H-13), 1.02 (d, J = 7 Hz, 3H, H-14), 1.09 (m, 1H, H-11a),
1.21(m, 1H, H-9a), 1.25 (m, 1H, H-11b), 1.33 (m, 1H, H-10), 1.49 (m, 1H, H-9b), 4.25
(q, J = 7 Hz, 1H, H-8), 5.92 (d, J = 7.5 Hz, 1H, H-5), 7.60 (t, J = 6.5, 7 Hz, 1H, H-6),
11.49 (s, 1H, H-1); 13C NMR (DMSO, 125 MHz) δ = 11.1 (C-12), 16.7 (C-14), 18.8 (C13), 29.7 (C-11), 31.7 (C-10), 39.9 (C-8), 40.0 (C-9), 99.1 (C-5), 105.9 (C-3), 142.8 (C6), 161.8 (C-2), 177.6 (C-4), 211.9 (C-7). HRESIMS calculated for C13H20NO3:
238.1437; observed 238.1428 [M]H+.
Crystals were formed by slow evaporation in methanol. Crystals size/mm3 =
0.29 × 0.25 × 0.06, formula C13H19NO3, (M =237.29): monoclinic, space group P21 (no.
4), a =
9.0939(16) Å, b =
30.695(5) Å, c =
9.1390(16) Å, β =
94.405(5)°, V =
2543.5(8) Å3, Z = 8, T = 100(2) K, μ/mm-1 = 0.713, Dcalc = 1.239 g/mm3, range for
data collection = 9.7 to 133.62°, reflections collected 60175, independent reflections
8598 (Rint = 0.0426), final R1 was 0.0294 and wR2 was 0.0759, Largest diff. peak/hole/e
Å-3 = 0.12/-0.19, flack parameter 0.03(8), ccdc code 941139.
162
Aspyridone A 8480
4.16(c)
Light brown solid, mp 178 °C; [α]22D -8.2 (c = 0.48, MeOH); IR (neat): νmax 2961,
2928, 1648, 1610, 1516, 1458, 1378, 1217, 1175, 992, 835, 588 cm-1; 1H NMR
(CD3OD, 500 MHz) δ = 0.87 (t, J = 7.5 Hz, 3H, H-12), 0.90 (d, J = 6.5 Hz, 3H, H-13),
1.13 (d, J = 6.5 Hz, 3H, H-14), 1.16 (m, 1H, H-11a), 1.33 (m, 1H, H-9a), 1.34 (m, 1H,
H-11b), 1.41 (m, 1H, H-10), 1.64 (m, 1H, H-9b), 4.39 (q, J = 6.8 Hz, 1H, H-8), 6.80 (d,
J = 8.5 Hz , 1H, H-17, H-19), 7.26 (d, J = 8.5 Hz, 1H, H-16, H-20), 7.47 (s, 1H, H-6);
13
C NMR (CD3OD, 125 MHz) δ = 11.7 (C-12), 17.6 (C-14), 19.4 (C-13), 31.0 (C-11),
33.7 (C -10), 41.2 (C-9), 41.9 (C-8), 107.0 (C-3), 116.0 (C-5), 116.15 (C-17, C-19),
125.2 (C-15), 131.5 (C-16, C-20) 140.6 (C-6), 158.3 (C-18), 163.9 (C-2), 177.6 (C-4),
214.4 (C-7). HRESIMS calculated for C19H24NO4: 330.1699; observed 330.1692
[M]Na+. The specific rotation and melting point of aspyridone A 84 are not reported
before in literature.
Crystals were formed by slow evaporation in methanol and diethyl ether. Brown
crystals,
formula
C19H23NO4,
M =329.38, monoclinic,
space
group
P21, a =
7.3372(7) Å, b = 22.647(3) Å, c = 20.618(2) Å, β = 90.112(6)°, V = 3426.0(6) Å3, Z =
8, T = 100(2) K, μ/ mm-1 = 0.727, Dcalc = 1.277 g/mm3, range for data collection = 3.9
to 132.38°, reflections collected 56007, independent reflections 5995 (Rint = 0.0792),
final R1 was 0.0452 and wR2 was 0.1139, Largest diff. peak/hole/e Å-3 = 0.28/-0.29,
flack parameter 0(10), ccdc code 941138.
4.16 (d)
18-deshydroxypreaspyridone A 240
Brown viscous oil, [α]22D -142.3 (c = 0.15, MeOH); IR (neat): νmax 3019, 2962, 2875,
1709,1656, 1600, 1214 cm-1; 1H NMR (CD3OD, 500 MHz) δ = 0.83 (m, 3H, H-11),
163
0.84 (m, 3H, H-12), 1.02 (d, J = 7 Hz, 3H, H-13), 1.09 (m, 1H, H-10a), 1.27 (m, 1H, H9), 1.34 (m, 1H, H-10b), 1.33 (m, 1H, H-8a), 1.44 (m, 1H, H-8b), 2.98 (dd, J = 5.5, 14
Hz, 1H, H-14a), 3.07 (dd, J = 4, 14 Hz, 1H, H-14b), 3.65 (q, J = 6.7 Hz, 1H, H-7), 4.11
(t, J = 4.5 Hz, 1H, H-5), 7.16 (m, 2H, H-17, H-19), 7.17 (m, 1H, H-18), 7.22 (m, 2H, H16, H-20);
13
C NMR (CD3OD, 125 MHz) δ = 11.5 (C-11), 17.6 (C-13), 19.5 (C-12),
30.3 (C-10), 33.5 (C-9), 36.0 (C-7), 38.3 (C-14), 41.3 (C-8), 63.5 (C-5), 128.1 (C-18),
129.5 (C-16, C-20), 130.7 (C-17, C-19), 136.9 (C-15), 196.0 (C-6), 197.7 (C-4). The
13
C signals for C-2 and C-3 were not observed. HRESIMS calculated for C19H25NO3Na:
338.1726; observed 338.1735 [M]Na+.
4.17
Purification of metabolites from A. oryzae apdACEB
1 litre culture (100 ml x 10 flasks) of the fungal strain A. oryzae apdACEB was grown
in CMP medium B at 28 °C for 7 days at 200 rpm. The culture was then extracted with
ethyl acetate according to standard method given in section 4.11. A crude extract of
1.13 g was obtained after vacuum concentration of the ethyl acetate layer and 50 mg/ml
solution was made with HPLC grade methanol. For purification of metabolites 200 µl of
the crude extract was injected in each successive run on the preparative HPLC
following the gradient in Method 3. The purified fractions of respective metabolites
were collected and dried.
Characterization of new metabolites from A. oryzae apdACEB
4.17(a)
Preaspyridone A (minor) 224
Pale white crystalline solid, mp 146-148 °C; [α]22D 98.4 (c = 0.51, CHCl3); IR (neat):
νmax 3262, 2962, 1693, 1650, 1589 cm-1; 1H NMR (DMSO, 500 MHz) δ = 0.80 (m, 3H,
H-12), 0.82 (m, 3H, H-11), 1.03 (d, J = 6 Hz, 3H, H-13), 1.09 (m, 1H, H-10a), 1.24 (m,
1H, H-9), 1.26 (m, 1H, H-10b), 1.27 (m, 2H, H-8), 2.82 (td, J = 4.5, 15.5 Hz, 2H, H14), 3.53 (q, J = 6.8 Hz, 1H, H-7), 4.08 (brs, 1H, H-5), 6.59 (d, J = 8 Hz, 2H, H-17, H164
19), 6.88 (d, J = 8.5 Hz, 2H, H-16, H-20), 8.93 (s, 1H, H-1), 9.17 (brs, 1H, H-18); 13C
NMR (125 MHz, DMSO) δ = 10.8 (C-11), 16.7 (C-13), 18.9 (C-12), 28.7 (C-10), 31.5
(C-9), 33.4 (C-7), 35.4 (C-14), 39.6 (C-8), 62.3 (C-5), 99.9 (C-3), 114.7 (C-17, C-19),
125.3 (C-15), 130.6 (C-16, C-20), 155.7 (C-18), 175.4 (C-2), 191.9 (C-6), 194.4 (C-4).
HRESIMS calculated for C19H26NO4: 332.1856; observed 332.1852 [M]H+.
Crystals were formed by slow evaporation in methanol. Formula C19H25NO4,
M =331.40, monoclinic, space group P21, a = 8.3357(13) Å, b = 11.294(2) Å, c =
9.9646(17) Å, α = 90.00°, β = 103.551(8)°, V = 912.0(3) Å3, Z = 2, T = 100(2) K, m/
mm-1 = 0.683, Dcalc = 1.207 g/mm3, range for data collection = 9.12 to 132.5°,
reflections
collected
10439,
independent
reflections
3021(
Rint =
0.0533),
final R1 was 0.0467 and wR2 was 0.1226, Largest diff. peak/hole/e Å-3 = 0.28/-0.29,
flack parameter 0.2 (2), ccdc code 941137.
4.17(b)
Preaspyridone A (major diastereomer) 230107
Brown viscous oil, [α]22D -166.3 (c = 0.95, CHCl3) ; IR (neat): νmax 3020, 2964, 1653,
1602, 1214 cm-1. 1H NMR (500 MHz, DMSO) δ = 0.80 (m, 3H, H-12), 0.79 (m, 3H, H11), 0.95 (d, J = 7 Hz, 3H, H-13), 1.07 (m, 1H, H-10a), 1.28 (m, 1H, H-9), 1.29 (m, 1H,
H-10b), 1.29 (m, 1H, H-8a), 1.38 (m, 1H, H-8b), 2.82 (d, J = 4.5 Hz, 2H, H-14), 3.51
(q, J = 6.6 Hz, 1H, H-7), 4.08 (brs, 1H, H-5), 6.59 (d, J = 8.5 Hz, 2H, H-17, H-19),
6.90 (d, J = 8 Hz, 2H, H-16, H-20), 8.92 (s, 1H, H-1), 9.17 (brs, 1H, H-18); 13C NMR
(125 MHz, DMSO), 10.9 (C-11), 17.1 (C-13), 18.9 (C-12), 28.6 (C-10), 31.7 (C-9),
33.5 (C-7), 35.7 (C-14), 39.1 (C-8), 62.4 (C-5), 99.8 (C-3), 114.7 (C-17, C-19), 125.4
(C-15), 130.6 (C-16, C-20), 155.9 (C-18), 175.6 (C-2), 191.9 (C-6), 194.3 (C-4).
HRESIMS calculated for C19H26NO4Na: 354.1676; observed 354.1677 [M]Na+.
165
4.17 (c)
1, 4-dihydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 259
Brown viscous oil, [α]22D -14.3 (c = 0.20, MeOH) ; IR (neat): νmax 3104, 2928, 1731,
1635, 1611, 1453, 1200, 751 cm-1; 1H NMR (CD3OD, 500 MHz) δ = 0.87 (t, J = 7.2 Hz,
3H, H-12), 0.92 (d, J = 7 Hz, 3H, H-13), 1.11 (d, J = 6.5 Hz, 3H, H-14), 1.18 (m, 1H,
H-11a), 1.33 (m, 1H, H-9a), 1.36 (m, 1H, H-11b), 1.41 (m, 1H, H-10), 1.62 (m, 1H, H9b), 4.31 (q, J = 7 Hz, 1H, H-8), 5.97 (d, J = 8 Hz , 1H, H-5), 7.94 (d, J = 7.5 Hz, 1H,
H-6); 13C NMR (CD3OD, 125 MHz) δ = 11.7 (C-12), 17.3 (C-14), 19.3 (C-13), 31.0 (C11), 33.5 (C -10), 41.1 (C-9), 42.1 (C-8), 98.9 (C-5), 107.4 (C-3), 142.3 (C-6),160.2 (C2), 176.0 (C-4), 213.6 (C-7). HRESIMS calculated for C13H19NO4Na: 276.1206;
observed 276.1218 [M]Na+.
4.17(d)
1-methoxy, 4-hydroxy-3-(2, 4-dimethylhexanoyl) 2-pyridone 260
Brown viscous oil, [α]22D -32.5 (c = 0.12, MeOH) ; IR (neat): νmax 2961, 2930, 1661,
1611, 1465, 1388, 975 cm-1; 1H NMR (DMSO, 500 MHz) δ = 0.80 (t, J = 7.2 Hz, 3H,
H-12), 0.84 (d, J = 6.5 Hz, 3H, H-13), 1.03 (d, J = 7 Hz, 3H, H-14), 1.13 (m, 1H, H11a), 1.25 (m, 1H, H-9a), 1.25 (m, 1H, H-11b), 1.37 (m, 1H, H-10), 1.49 (m, 1H, H-9b),
3.93 (s, 3H, H-15), 4.15 (q, J = 7 Hz, 1H, H-8), 5.99 (d, J = 7.5 Hz , 1H, H-5), 8.24 (d,
J = 7.5 Hz, 1H, H-6); 13C NMR (DMSO, 125 MHz) δ = 11.1 (C-12), 16.5 (C-14), 18.6
(C-13), 29.4 (C-11), 31.8 (C-10), 39.6 (C-9), 39.9 (C-8), 64.4 (C-15), 98.4 (C-5), 146.2
(C-6), 109.9 (C-3), 159.9 (C-2), 177.3 (C-4), 214.5 (C-7). HRESIMS calculated for
C14H21NO4Na: 290.1362; observed 290.1368 [M]Na+.
4.18
Purification of metabolites from A. oryzae apdACED
The A. oryzae apdACED was grown in 1 litre CMP medium B (10 flasks ×100 ml) at
28°C for 7 days with constant shaking at 200 rpm. The fungal cultures were then
166
extracted with ethyl acetate following the protocol explainedin section xx. A dried crude
extract of 1.128 g was obtained which was made to a concentration of 50 mg/ml with
HPLC grade methanol for purification of metabolites on preparative HPLC with
gradient given in Method 6. The pure fractions of metabolites were collected and dried.
Characterization of new metabolites from A. oryzae apdACED
4.18(a)
Z-5, 14-anhydropreaspyridone A 267
Bright yellow solid, [α]22D 15.1° (c = 0.13, MeOH); IR (neat): νmax 3667, 3190, 2961,
2876, 2366, 1691, 1584 cm-1;
1
H NMR (DMSO, 500 MHz) δ = 0.79 (m, 3H, H-11),
0.82 (m, 3H, H-12), 1.05 (d, J = 6.5 Hz, 3H, H-13), 1.08 (m, 1H, H-10a), 1.32 (m, 1H,
H-9), 1.32 (m, 1H, H-10b), 1.38 (m, 1H, H-8a), 1.47 (m, 1H, H-8b), 3.75 (q, J = 6.6 Hz,
1H, H-7), 6.37 (s, 1H, H-14), 6.78 (d, J = 8.5 Hz, 2H, H-17, H-19), 7.46 (d, J = 8.5 Hz,
2H, H-16, H-20); 13C NMR (DMSO, 125 MHz) δ = 10.9 (C-11), 16.8 (C-13), 18.8 (C12), 28.7 (C-10), 31.5 (C-9), 40.4 (C-8), 110.4 (C-14), 115.6 (C-17, C-19), 124.8 (C15), 131.5 (C-16, C-20), 158.8 (C-18), 184.0 in CD3OD (C-4). The 13C signals for C-2,
C-3, C-4, C-5, C-6 and C-7 were not observed. HRESIMS calculated for C19H23NO4Na:
352.1519; observed 352.1528 [M]Na+.
167
Chapter 5
Summary and Future Perspective
Iterative HRPKS-NRPS produce a wide array of diverse, bioactive and
biosynthetically intriguing compounds. The programming rules embedded in the PKSNRPS enzymes have been under investigation for many years and still provide
undiscovered horizons for researchers.
In this research we isolated and characterized new compounds produced from
different genetically modified transformants (particularly from heterologous expression
and gene silencing). The structures of the isolated compounds gave clues to the
biosynthetic potential of the PKS-NRPS enzymes and also the tailoring enzymes present
in the PKS-NRPS clusters.
We characterized two new putative A. oryzae wild type compounds 146 and 147
from A. oryzae tenSPKS-dmbC but couldn’t determine the chemical product of
TENSPKS acting without its NRPS moiety. The tenC silencing in B. bassiana by the
carbon inducible promoter PamyB failed to deliver any outcome because B. bassiana
didn’t produce tenellin compounds when we changed the carbon source in TPM media
from mannitol to maltose and glucose.
The investigation of the aspyridone pathway of A. nidulans and heterologous
expression of apd genes in A. oryzae gave insight into the unique catalytic capabilities
of the cytochrome P450 present in the apd cluster and the biosynthetic potential of the
PKS-NRPS. The cytochrome P450 ApdE has oxidative ring expanding and unusual
dephenylation activity. ApdB catalyses N-hydroxylation in dephenylated 2-pyridones.
The production of 18-deshydroxy preaspyridone 240 displays wider amino acid
selectivity besides tyrosine. The compounds 239, 259, 260 and 267 show more diverse
chemical activities for apd PKS-NRPS enzymes as compared to tenellin and desmethyl
bassianin PKS-NRPS gene clusters. The crystal structures of 224, 84 and 239 gave
knowledge about the relative stereochemistry of their structures. The enoyl reductase
ApdC showed a significant property for setting opposite stereochemistry for the
dimethyls in different cycles of the polyketide chain of aspyridone.
168
The ApdC, ApdE and ApdB enzymes should be further studied in vitro to
explore their catalytic potential and may contribute to be used as biocatalysts in
biosynthetic reactions.
169
References
1.
J. Clayden, N. Greeves, S. Warren and P. Wothers, Organic Chemistry, Oxford
University Press, 2001, 1413-1447.
2.
J. Davies and K. S. Ryan, ACS Chem. Biol., 2012, 7, 252−259.
3.
D. O. Kennedy and E. L. Wightman, Adv. Nutr., 2011, 2, 32–50.
4.
R. A. Maplestone, M. J. Stone and D. H. Williams, Gene, 1992, 115, 151-157.
5.
N. P. Keller, G. Turner and J. W. Bennett, Nat. Rev. MicroBiol., 2005, 3, 937 947.
6.
P. M. Dewick, Medicinal Natural Products, John Wiley and Sons ltd., 2nd
edition, 2002, 7-34.
7.
D. A. Dias, S. Urban and U. Roessner, Metabolites, 2012, 2, 303-336.
8.
A. D. Marderosian and J. A. Beutler, The Review of Natural Products, Seattle,
WA, USA, 2002, 13–43.
9.
P. M. Dewick, Medicinal Natural Products: A Biosynthentic Approach, John
Wiley and Son, West Sussex, UK, 2002.
10.
M. M. Cowan, Clin. Microbiol. Rev., 1999, 12, 564–582.
11.
J. Mann, Murder, Magic, and Medicine, Oxford University Press, New York,
USA, 1994, 164–170.
12.
Y. W. Chin, M. J. Balunas, H. B. Chai and A. D. Kinghorn, The AAPS J., 2006,
8, 239-253.
13.
J. K. Zjawiony, J. Nat. Prod., 2004, 67, 300–310.
14.
C. J. Schulze, W. M. Bray, M. H. Woerhmann, J. Stuart, R. S. Lokey and R.
G. Linington, Chem. Biol., 2013, 20, 285–295.
15.
M. S. Butler, J. Nat. Prod., 2004, 67, 2141–2153.
16.
W. H. Gerwick and B. S. Moore, Chem. Biol., 2012, 19, 85-98.
17.
G. M. Cragg, D. J. Newman and K. M. Snader, J. Nat. Prod., 1997, 60, 52-60.
18.
D. J. Newman, G. M. Cragg and K. M. Snader, J. Nat. Prod., 2003, 66, 10221037.
19.
D. J. Newman and G. M. Cragg, J. Nat. Prod., 2007, 70, 461-477.
170
20.
S. K. Piasecki, C. A. Taylor, J. F. Detelich, J. Liu, J. Zheng, A. Komsoukaniants,
D. R. Siegel and A. T. Keatinge-Clay, Chem. Biol., 2011, 18, 1331–1340.
21.
J. C. Saccheittini and C.D. Poulter, Science, 1997, 277, 1788-1789.
22.
J. Gershenzon1 and N. Dudareva, Nat. Chem. Biol., 2007, 3, 408-414.
23.
K. Xu, P. Wang, B. Yuan, Y. Cheng, Q. Li and H. Lei, Chem. Cent. J., 2013, 7,
1-11.
24.
R. H. F. Manske. The Alkaloids Chemistry and Physiology, New York,
Academic Press, 1965.
25.
K. E. Bushley and B. G. Turgeon, BMC Evol Biol., 2010, 10, 1-23.
26.
R. Finking and M. A. Marahiel, Annu. Rev. Microbiol., 2004, 58, 453–488.
27.
K. H. Kim, Y. Cho, M. La Rota, R. A. Cramer, C. B. Lawrence, Mol. Plant.
Pathol., 2007, 8, 23-39.
28.
M. A. Marahiel, Chem. Biol., 1997, 4, 561–567.
29.
T. Stachelhaus and M. A. Marahiel, J. Biol. Chem., 1995, 270, 6163–6169.
30.
C. T. Walsh, H. Chen, T. A. Keating, B. K. Hubbard, H. C. Losey, L. Luo, C. G.
Marshall, D. A. Miller and H. M. Patel, Curr. Opin. Chem. Biol., 2001, 5, 525
534.
31.
D. O’Hagan, The Polyketides Metabolites, Ellis Harwood, Chichester, 1991.
32.
S. J. Wakil, J. K. Stoops and V. C. Joshi, Ann. Rev. Biochem., 1983, 52, 537759.
33.
D. A. Hopwood and D. H. Sherman, Annu. Rev. Genet., 1990, 24, 37-66.
34.
R. J. Cox, Org. Biomol. Chem., 2007, 5, 2010–2026.
35.
D. O’Hagan, Nat. Prod. Rep., 1992, 9, 447-479.
36.
C. Sheridan, Nat. Biotechnol., 2012, 30, 385-387.
37.
E. A. Campbell, N. Korzheva, A. Mustaev, K. Murakami, S. Nair, A. Goldfarb,
S. A. Darst, Cell, 2001, 104, 901–912.
38.
D. Lowicki and A. Huczynski, Biomed. Res. Int., 2013, 2013, 1-14.
39.
H. Gjonnaess and E. Holten, Acta Obstet. Gynecol. Scand., 1978, 57, 137–139.
40.
T. Pitterna, J. Cassayre, O. F. Huter, P. M. J. Jung, P. Maienfisch, F. M.
Kessabi, L. Quaranta, H. Tobler, Bioorg. Med. Chem., 2009, 17, 4085–4095.
171
41.
T. W. Schulte, S. Akinaga, S. Soga, W. Sullivan, B. Stensgard, D. Toft, L. M.
Neckers, Cell. Stress. Chaperon., 1998, 3, 100–108.
42.
M. Muroi, M. Izawa, Y. Kosai and M. Asai, J. Antibiot., 1980, 33, 205–212.
43.
N. Khan, B. Rawlings, P. Caffrey, Biotechnol Lett., 2011, 33, 1121–1126.
44.
T. Kino, H. Hatanaka, M. Hashimoto, M. Nishiyama, T. Goto, M. Okuhara, M.
Kohsaka, H. Aoki, H. Imanaka, J. Antibiot. (Tokyo)., 1987, 40, 1249-1255.
45.
R. N. Moore , G. Bigam , J. K. Chan , A. M. Hogg , T. T. Nakashima , J. C.
Vederas , J. Am. Chem. Soc., 1985, 107, 3694–3701.
46.
G. D. Crouse, T. C. Sparks, J. Schoonover, J. Gifford, J. Dripps, T. Bruce, L. L.
Larson, J. Garlich, C. Hatton, R. L. Hill, T. V. Worden, J. G. Martynow, Pest.
Manag. Sci., 2001, 57, 177–185.
47.
R. Singh, D. P. Hsieh, Arch. Biochem. Biophys., 1977, 178, 285-292.
48.
J. N. Collie, J. Chem. Soc. Trans., 1907, 91, 1806-1813.
49.
A. J. Birch, Science, 1967, 156, 202-206.
50.
D. Rittenberg and Konrad Bloch, J. Biol. Chem., 1945, 160, 417-424.
51.
R. Robinson, J. Roy. Soc. Arts., 1948, 96, 795.
52.
A. J. Birch, P. Elliott, A. R. Penfold, Aust. J. Chem.,1954, 7, 169.
53.
J. Staunton and K. J. Weissman, Nat. Prod. Rep., 2001, 18, 380–416.
54.
M. J. Garson and J. Staunton, Chem. Soc. Rev., 1979, 8, 539-561.
55.
D. A. Hopwood, Chem. Rev., 1997, 97, 2465-2497.
56.
R. Bentley and J. W. Bennet, Annu. Rev. Microbiol., 1999, 53, 411-446.
57.
Y. H. Chooi and Y. Tang, J. Org. Chem., 2012, 77, 9933−9953.
58.
S. J. Moss, C. J. Martin and B. Wilkinson, Nat. Prod. Rep., 2004, 21, 575 – 593.
59.
L. Katz, Chem. Rev., 1997, 97, 2557-2575.
60.
J. Cortes, S. F. Haydock, G. A. Roberts, D. J. Bevitt and P. F. Leadlay,
Nature, 1990, 348, 176-178.
61.
R. Stanzak, P. Matsushima, R. H. Baltz, R. N. B. Rao, J. Biotechnol., 1986,
4, 229- 232.
172
62.
C. Khosla, Chem. Rev., 1997, 97, 2577-2590.
63.
T. Nguyen, K. Ishida, H. J. Kodama, E. Dittmann, C. Gurgui, T. Hochmuth, S.
Taudien, M. Platzer, C. Hertweck and J. Piel, Nature Biotechnol., 2008, 26,
225-233.
64.
Y. Q. Cheng, G. L. Tang and B. Shen, Proc. Natl. Acad. Sci. U.S.A, 2003, 100,
3149–3154.
65.
A. Kassem El-Sayed, J. Hothersall, S. M. Cooper, E. Stephens, T. J. Simpson
and C. M. Thomas, Chem. Biol., 2003, 10, 419–430.
66.
G. M. Gaucher and M.G. Shepherd, Biochem. Biophys. Res. Commun., 1968, 32,
664-671.
67.
K. Arai, B. J. Rawlings, Y. Yoshizawa and J. C. Vederas, J. Am. Chem.Soc.,
1989, 111, 3391–3399.
68.
S.V. Pathre, P.V. Khadikar and C. J. Mirocha, Appl. Environ. Microbiol., 1989,
55, 1955–1966.
69.
A. M. Bailey, R. J. Cox, K. Harley, C. M. Lazarus, T.
Skellam, Chem. Commun., 2007, 4053–4055.
70.
J. Beck, S. Ripka, A. Siegner, E. Schiltz and E. Schweizer, Eur. J. Biochem.,
1990, 192, 487–498.
71.
L. Hendrickson, C. R. Davis, C. Roach, D. K. Nguyen, T. Aldrich, P. C. McAda
and C. D. Reeves, Chem. Biol., 1999, 6, 429-439.
72.
R. J. Cox, F. Glod, D. Hurley, C. M. Lazarus, T. P. Nicholson, B. A. M. Rudd,
T. J. Simpson, B. Wilkinson and Y. Zhang, Chem. Commun., 2004, 20, 22602261.
73.
R. H. Proctor, A. E. Desjardins, R.D. Plattner and T. M. Hohn, Fungal Genet.
Biol., 1999, 27, 100–112.
74.
G. Wang, M. S. Rose, B. G. Turgeon and O. C. Yoder, Plant Cell, 1996, 8,
2139–2150.
75.
D. Boettger and C. Hertweck, ChemBioChem, 2013, 14, 28–42.
76.
K. E. Bushley, B. G. Turgeon, BMC Evol. Biol., 2010, 10, 26.
77.
Z. S. Song, R. J. Cox, C. M. Lazarus and T. J. Simpson, ChemBioChem, 2004, 5,
1196–1203.
78.
D. O. Rees, N. Bushby, R. J. Cox, J. R. Harding, T. J. Simpson and C. L. Willis,
ChemBioChem, 2007, 8, 46–50.
J.
Simpson
and
E.
173
79.
A. A. Yakasai, J. Davison, Z. Wasil, L. M. Halo, C. P. Butts, C. M.
Lazarus, A. M. Bailey, T. J. Simpson and R. J. Cox, J. Am. Chem. Soc., 2011,
133, 10990–10998.
80.
S. Bergmann, J. Schumann, K. Scherlach, C. Lange, A. A. Brakhage, C.
Hertweck, Nat. Chem. Biol., 2007, 3, 213–217.
81.
F. E. Vega, F. Posada, M. C. Aime, M. P. Ripoll, F. Infante, S. A. Rehner, Biol.
Control, 2008, 46, 72–82.
82.
A. E. Hajek, R. J. St. Leger, Annu. Rev. Entomol., 1994, 39, 293–322.
83.
Beauveria bassiana photo by Milos Villaris.
84.
J. V. Bell, Mycoses (ed. Cantwell, G. E.), Insect Diseases, New York, Marcel
Dekker Inc., 1974.
85.
L. Zengzhi, L. Chunru, H. Bo and F. Meizhen, Chinese Sci. Bull., 2001, 46,
751–753.
86.
S. A. Rehner, E. Buckley, Mycologia, 2005, 97, 84–98.
87.
G. C. Ainsworth, Nature, 1956, 177, 255–257.
88.
D. M. Macleod, Can. J. Bot., 1954, 32, 818–890.
89.
P. H. Dunn and B. J. Mechalas, J. Invertebr. Pathol., 1963, 5, 451-459.
90.
A. Vey, R. E. Hoagland and T. M. Butt, Progress, Problems and Potential,
CABI Publishing, Oxford, UK, 2001, 311-346.
91.
Y. Xu, R. Orozco, E. M. K. Wijeratne , P. E. Artiles , A. A. L. Gunatilaka, S. P.
Stock, I. Molnar, Fungal Genet Biol., 2009, 46, 353–364.
92.
R. C. F. Jones, A. K. Choudhury, C. E. Dawson, C. Lumley, and V. McKee,
Arkivoc., 2012, 7, 12-24.
93.
Q. Wang and L. Xu, Molecules, 2012, 17, 2367-2377.
94.
M. Kuzma, A. Jegorov, P. Kacer and V. Havlicek, J. Mass Spectrom., 2001, 36,
1108–1115.
95.
S. H. Elbasyouni, L. C. Vining, Can. J. Biochem., 1966, 44, 557-565.
96.
C. K. Wat, A. G. McInnes and D. G. Smith, J. L. C. Wright and L.C .Vining,
Can. J. Chem., 1977, 55, 4090-4098.
97.
E. Q. Moraga and A. Vey, Mycol Res., 2004, 108, 441–452.
174
98.
K. L. Eley, L. M. Halo, Z. Song, H. Powles, R. J. Cox, A. M. Bailey, C. M.
Lazarus, T. J. Simpson, ChemBioChem, 2007, 8, 289–297.
99.
M. C. Moore, R. J. Cox, G. R. Duffin and D. O’Hagan, Tetrahedron, 1998, 54,
9195-9206.
100.
A. G. McInnes, D. G. Smith, J. A. Walter, L. C. Vining and J. L. C. Wright, J.
Chem. Soc. Chem. Commun., 1974, 282–284.
101.
E. Leete, N. Kowanko, R. A. Newmark, L. C. Vining, A. G. McInnes and J. L.
C. Wright. Tetrahedron Lett., 1975, 4103-4106.
102.
J. L. C. Wright, L. C. Vining, A. G. McInnes, D. G. Smith and J. A. Walter,
Can. J. Biochem. Cell B., 1977, 55, 678-685.
103.
R. J. Cox and D. O'Hagan, J. Chem. Soc. Perkin Trans.1, 1991, 2537-2540.
104.
L. M. Halo, J. W. Marshall, A. A. Yakasai, Z. Song, C. P. Butts, M. P. Crump,
M. Heneghan, A. M. Bailey, T. J. Simpson, C. M. Lazarus and R. J. Cox,
ChemBioChem, 2008, 9, 585 – 594.
105.
J. Kennedy, K. Auclair, S. G. Kendrew, C. Park, J. C. Vederas, C. R.
Hutchinson, Science, 1999, 284, 1368–1372.
106.
J. W. Sims and E. W. Schmidt, J. Am. Chem. Soc., 2008, 130, 11149–11155.
107.
W. Xu, X. Cai, M. E. Jung and Y. Tang, J. Am. Chem. Soc., 2010, 132,
13604–13607.
108.
L. M. Halo, M. N. Heneghan, A. A. Yakasai, Z. Song, K. Williams, A. M.
Bailey, R. J. Cox, C. M. Lazarus and T. J. Simpson, J. Am. Chem. Soc., 2008,
130, 17988–17996.
109.
M. N. Heneghan, A. A. Yakasai, K. Williams, K. A. Kadir, Z. Wasil, W.
Bakeer, K. M. Fisch, A. M. Bailey, T. J. Simpson, R. J. Cox and C. M. Lazarus,
Chem. Sci., 2011, 2, 972-979.
110.
K. M. Fisch, W. Bakeer, A. A. Yakasai, Z. Song, J. Pedrick, Z. Wasil, A. M.
Bailey, C. M. Lazarus, T. J. Simpson and R. J. Cox, J. Am. Chem. Soc., 2011,
133, 16635–16641.
111.
A. A. Yakasai, J. Davison, Z. Wasil, L. M. Halo, C. P. Butts, C. M.
Lazarus, A. M. Bailey, T. J. Simpson and R. J. Cox, J. Am. Chem. Soc., 2011,
133, 10990–10998.
112.
R. B. Williams, J. C. Henrikson, A. R. Hoover, A. E. Lee and R. H. Cichewicz,
Org. Biomol. Chem., 2008, 6, 1895–1897.
113.
D. Horhant, A. L. Lamer, J. Boustie, P. Uriac and N. Gouault, Tetrahedron
Lett., 2007, 48, 6031–6033.
175
114.
J. S. Dahiya and J. P. Tewari, Phytochemistry, 1991, 30, 2825-2828.
115.
B. K. Park, M. Nakagawa, A. Hirota and M. Nakayama, Agric. Biol. Chem.,
1987, 51, 3443-3444.
116.
S. Huneck and G. Hofle, 1980, Phytochemistry, 19, 2713-2715.
117.
Y. Seshime, P. R. Juvvadi, K. Kitamoto, Y. Ebizuka, I. Fujii, Bioorg. Med.
Chem., 2010, 18, 4542–4546.
118.
H. Nakayashiki and Q. B. Nguyen, Curr. Opin. Microbiol., 2008, 11, 494–502.
119.
H. Nakayashiki, FEBS Lett., 2005, 579, 5950–5957.
120.
G. L. Sen and H. M. Blau, FASEB J., 2006, 20, 1293–1299.
121.
C. Napoli, C. Lemieux and R. Jorgensen, 1990, Plant Cell, 2, 279-289.
122.
I. Fujii, Nat. Prod. Rep., 2009, 26, 155–169.
123.
J. W. Bennett, Aspergillus, Molecular Biology and Genomics, 2010, 1-17.
124.
Kanaani, Hussein, Hargreaves, Megan, Ristovski, Zoran, Morawska, Lidia,
Atmos. Environ., 2008, 42, 7141-7154.
125.
G. C. Carroll and D. T. Wicklow, The Fungal Community; Its Organization and
Role in the Ecosystem, New York, Marcel Dekker, Inc., 1992.
126.
O. Ciferri, Appl. Environ. Microbiol., 1999, 65, 879–885.
127.
D. M. Geiser, R. A. Samson, J. Varga, A. Rokas, and S. M. Witiak, Aspergillus
in the Genomic Era, Netherlands, Wageningen Academic Pubs., 2008, 17–32.
128.
C. T. Calam, A. E. Oxford and H. Raistrick, Biochem. J., 1939, 33, 1488–1495.
130.
A. Rokas, Trends genet., 2009, 25, 60-63.
131.
G. J. G. Ruijter, C. P. Kubicek and J. Vissler, The Mycota Vol. X. Industrial
Applications, Heidelberg, Springer-Verlag, 2002, 213–230.
132.
J. W. Bennett, MDD., 2001, 4, 47–51.
133.
K. Sakai, H. Kinoshita, T. Nihira, Appl. Microbiol. Biotechnol., 2012, 93, 20112022.
134.
D. B. Archer and G. Turner, The Mycota XIII, Berlin, Springer-Verlag, 2006,
75–96.
135.
A. W. Alberts, Am. J. Cardiol., 1998, 62, 10–15.
176
136.
M. A. Goetz, M. Lopez, R. L. Monaghan, R. S. Chang, V. J. Lotti, T. B. Chen, J.
Antibiot., 1985, 38, 1633-1637.
137.
D. M. Oddon, E. Diatloff and S. K. Roberts, BBA-Biomembranes, 2007, 1768,
2466–2477.
138.
A. S. Awaad, A. J. Nabilah, M. E. Zain, Phytother. Res., 2012, 26, 1872-1877.
139.
J. W. Bennett and M. Klich, Clin. Microbiol. Rev., 2003, 16, 497–516.
140.
L. Goldblatt, Aflatoxin: Scientific Background, Control and Implications, New
York, Academic Press, 1969.
141.
R. K. Bush, J. M. Portnoy, A. Saxon, A. I. Terr and R. A. Wood, J. Allergy
Clin., 2006, 117, 326–333.
142.
J. P. Debeaupuis, J. Sarfati, V. Chazalet and J. P. Latge, Infect. Immun., 1997,
65, 3080–3085.
143.
A. M. Calvo, R. A. Wilson, J. W. Bok and N. P. Keller, Microbiol. Mol. Biol.
Rev., 2002, 66, 447–459.
144.
G. Pontecorvo, Annu. Rev. Microbiol., 1956, 10, 393-400.
145.
N. R. Morris, S. A. Osmani, D. B. Engle and J. H. Doonan, BioEssays, 1989, 10,
196-201.
146.
C. Scazzocchio, Aspergillus: a multifaceted
Microbiology, Amsterdam, Elsevier, 2009.
147.
J. E. Galagan, S. E. Calvo, C. Cuomo, L. J. Ma, J. R. Wortman, S. Batzoglou, S.
I. Lee, M. Basturkmen, C. C. Spevak, J. Clutterbuck, V. Kapitonov, J. Jurka, C.
Scazzocchio, M. Farman, J. Butler, S. Purcell, S. Harris, G. H. Braus, O. Draht,
S. Busch, C. DEnfert, C. Bouchier, G. H. Goldman, D. B. Pedersen, S. G. Jones,
J. H. Doonan, J. Yu, K. Vienken, A. Pain, M. Freitag, E. U. Selker, D. B.
Archer, M. A. Penalva, B. R. Oakley, M. Momany, T. Tanaka, T. Kumagai, K.
Asai, M. Machida, W. C. Nierman, D. W. Denning, M. Caddick, M. Hynes, M.
Paoletti, R. Fischer, B. Miller, P. Dyer, M. S. Sachs, S. A. Osmani and B. W.
Birren, 2005, Nature, 438, 1105-1115.
148.
W. C. Nierman, A. Pain, M. J. Anderson, J. R. Wortman, H. S. Kim, J. Arroyo,
M. Berriman, K. Abe, D. B. Archer, C. Bermejo, J. Bennett, P. Bowyer, D.
Chen, M. Collins, R. Coulsen, R. Davies, P. S. Dyer, M. Farman, N. Fedorova1,
N. Fedorova1, T. V. Feldblyum, R. Fischer, N. Fosker, A. Fraser, J. L. Garcıa1,
M. J. Garcıal, A. Goble, G. H. Goldman, K. Gomi, S. G. Jones, R. Gwilliam, B.
Haas, H. Haas, D. Harris, H. Horiuchi, J. Huang, S. Humphray, J. Jimenez, N.
Keller, H. Khouri1, K. Kitamoto, T. Kobayashi, S. Konzack, R. Kulkarni, T.
Kumagai, A. Lafton, J. P. Latge, W. Li, A. Lord, C. Lu, W. H. Majoros, G. S.
May, B. L. Miller, Y. Mohamoud, M. Molina, M. Monod, I. Mouyna, S.
genus.
Encyclopedia
of
177
Mulligan, L. Murphy, S. O’Neil, I. Paulsen, M. A. Penalva, M. Pertea, C. Price,
B. L. Pritchard, M. A. Quail, E. Rabbinowitsch, N. Rawlins, M. A. Rajandream,
U. Reichard, H. Renauld, G. D. Robson, S. Rodriguez de Cordoba, J. M.
Rodrıguez-Pen, C. M. Ronning, S. Rutter, S. L. Salzberg, M. Sanchez, J. C. S.
Ferrero, D. Saunders, K. Seeger, R. Squares, S. Squares, M. Takeuchi, F.
Tekaia, G. Turner, C. R. Vazquez de Aldana, J. Weidman, O. White, J.
Woodward, J. H. Yu, C. Fraser, J. E. Galagan, K. Asai, M. Machida, N. Hall, B.
Barrell and D. W. Denning, 2005, Nature, 438, 1151-1156.
149.
J. H. Yu and N. Keller, Annu. Rev. Phytopathol., 2005, 43, 437–458.
150.
D. W. Brown, J. H. Yu, H. S. Kelkar, M. Fernandes, T. C. Nesbitt, N. P. Keller,
T. H. Adams, and T. J. Leonard, Proc. Natl. Acad. Sci. U S A., 1996, 93, 1418–
1422.
151.
J. W. Bok and N. P. Keller, Eukaryot. Cell., 2004, 3, 527–535.
152.
Y. M. Chiang, E. Szewczyk, A. D. Davidson, N. Keller, B. R. Oakley and C. C.
C. Wang, J. Am. Chem. Soc., 2009, 131, 2965–2970.
153.
J. W. Bok, Y. M. Chiang, E. Szewczyk, Y. R. Dominguez, A. D. Davidson, J. F.
Sanchez, H. C. Lo, K. Watanabe, J. Stra.uss, B. R. Oakley, C. C.C. Wang and N.
P. Keller, 2009, Nat. Chem. Biol., 5, 462-464.
154.
A. A. Brakhage, FEMS Microbiol. Lett., 1997, 148, 1-10.
155.
K. R. Branch, J. W. Bennett and D. Bhatnagar, Fungal Genet. Newsl., 1993, 40,
20-21.
156.
Schroeder and Kelton, Appl. Microbiol., 1975, 30, 589-591.
157.
J. D. Hajjar, J. W. Bennett, D. Bhatnagar, and R. Bahu, Mycol. Res., 1989, 94,
548-551.
158.
N. P. Keller, N. J. Kantz and T. H. Adams, Appl. Environ. Microbiol., 1994, 60,
1444-1450.
159.
N. P. Keller, S. Segner, D. Bhatnagar and T. H. Adams, Appl. Environ.
Microbiol., 1995, 61, 3628-3632.
160.
H. S. Kelkar, N. P. Keller and T. H. Adams, Applied and Environmental
Microbiology, 1996, 62, 4296–4298.
161.
K. Scherlach and C. Hertweck, Org. Biomol. Chem., 2006, 4, 3517–3520.
162.
A. Watanabe, I. Fuji, U. Sankawa, M. E. Mayorga, W. E. Timberlake and Y.
Ebizuka, Tetrahedron Lett., 1999, 40, 91-94.
163.
I. Fujii, A. Watanabe, U. Sankawa and Y. Ebizuka, Chem. Biol., 2001, 8, 189197.
178
164.
H. R. El-Seedi, M. A. El-Barbary, D. M. H El-Ghorab, L. Bohlin, A. K. BorgKarlson, U. Goransson, R. Verpoorte, Curr. Med. Chem., 2010, 17, 854–901.
165.
J. F. Sanchez, R. Entwistle, J. H. Hung, J. Yaegashi, S. Jain, Y. M. Chiang, C. C.
C. Wang and B. R. Oakley, J. Am. Chem. Soc., 2011, 133, 4010–4017.
166.
V. Schroeckh, K. Scherlach, H. W. Nutzmann, E. Shelestd, W. S. Heck, J.
Schuemann, K. Martine, C. Hertweck, and A. A. Brakhage, Proc. Natl. Acad.
Sci. U.S.A., 2009, 106, 14558–14563.
167.
J. F. Sanchez, Y. M. Chiang, E. Szewczyk, A. D. Davidson, M. Ahuja, C. E.
Oakley, J. W. Bok, N. Keller, B. R. Oakley and C. C. C. Wang, Mol.
Biosyst., 2010, 6, 587–593.
168.
M. L. Nielsen, J. B. Nielsen, C. Rank, M. L. Klejnstrup, D. K. Holm, K. H.
Brogaard, B. G. Hansen, J. C. Frisvad, T. O. Larsen and U. H. Mortensen,
FEMS Microbiol. Lett., 2011, 321, 157–166.
169.
E. Szewczyk, Y. M. Chiang, C. E. Oakley, A. D. Davidson, C. C. C. Wang and
B. R. Oakley, Appl. Environ. Microbiol., 2008, 74, 7607–7612.
170.
J. W. Bok, D. Hoffmeister, L. A. Maggio-Hall, R. Murillo, J. D. Glasner and N.
P. Keller, 2006, Chem. Biol., 13, 31–37.
171.
S. Bouhired, M. Weber, A. Kempf-Sontag, N. P. Keller, D. Hovmeister, Fungal
Genet. Biol., 2007, 44, 1134–1145.
172.
Y. Chiang, E. Szewczyk, T. Nayak, A. D. Davidson, J. F. Sanchez, H. C. Lo,
W. Y. Ho, H. Simityan, E. Kuo, A. Praseuth, K. Watanabe, B. R. Oakley and C.
C. Wang, 2008, Chem. Biol., 15, 527–532.
173.
H. V. Dohren, Fungal Genet. Biol., 2009, 46, 45–52.
174.
J. Davisona, A. AlFahad, M. Cai, Z. Song, S. Y. Yehia, C. M. Lazarus, A. M.
Bailey, T. J. Simpson and R. J. Cox, Proc. Natl. Acad. Sci. U.S.A., 2012, 109,
7642-7647.
175.
D. Lubertozzi and J. D. Keasling, Biotech. Adv., 2009, 27, 53-75.
176.
A. K. Pahirulzaman, K. Williams and C. M. Lazarus, Method. Enzymol., 2012,
517, 241–260.
178.
J. Schumann and C. Hertweck, J. Biotech., 2006, 124, 690-703.
179.
M. E. Case, M. Schweizer, S.R. Kushner and N. H. Giles, Proc. Natl. Acad. Sci.
U.S.A., 1979, 76, 2563–5259.
180.
M. J. Hynes, J. Genet., 1996, 75, 297-311.
179
181.
B. A. Pfeifer and C. Khosla, Microbiol. Mol. Biol. Rev., 2001, 65, 106-118.
182.
J. Punya, A. Tachaleat, S. Wattanachaisaereekul, R. Haritakun,
Boonlarppradab, S. Cheevadhanarak, Fungal Genet. Biol., 50, 55-62.
183.
Talbot, Molecular and Cellular Biology of Filamentous Fungi, Oxford
University Press, 2001.
184.
H. Zhou, Z. Gao, K. Qiao, J. Wang, J. C. Vederas and Y. Tang, Nat. Chem.
Biol., 2012, 8, 331-333.
185.
H. J. M. Van den Brink, R. F. M. Van Gorcom, C. A. M. J. J. Van den
Hondel and P. J. Punt, Fungal Genet. Biol., 1998, 23, 1–17.
186.
F. Ogliaro, N. Harris, S. Cohen, M. Filatov, S. P. De Visser and S. Shaik, J. Am.
Chem. Soc., 2000, 122, 8977-8989.
187.
V. W. Bowry, J. Lusztyk and K. U. Ingold, J. Am. Chem. Soc., 1991, 113, 56875698.
188.
M. Isaka, P. Chinthanom, S. Supothina, P. Tobwor and N. L. Hywel-Jones, J.
Nat. Prod., 2010, 73, 2057–2060.
189.
Y. Cheng, B. Schneider, U. Riese, B. Schubert, Z. Li and M. Hamburger, J. Nat.
Prod. 2006, 69, 436-438.
190.
M. S. Gachet, O. Kunert, M. Kaiser, R. Brun, R. A. Munoz, R. Bauer and W.
Schuhly, J. Nat. Prod., 2010, 73, 553–556.
191.
Z. Wasil, K. A. K. Pahirulzaman, C. Butts, T. J. Simpson, C. M. Lazarus and R.
J. Cox, Chem. Sci., 2013, 4, 3845-3856.
192.
Y. Kimura, N. Takahashi and S. Tamura, Agric. Biol. Chem., 1969, 33, 15071516.
193.
A. Evidente, M. Fiore, G. Bruno, L. Sparapano, A. Motta, Phytochemistry,
2006, 67, 1019–1028.
194.
H. Miyadera, K. Shiomi, H. Ui, Y. Yamaguchi, R. Masuma, H. Tomoda, H.
Miyoshi, A. Osanai, K. Kita and S. Omura, Proc. Natl. Acad. Sci. U.S.A., 2003,
100 , 473–477.
195.
M. Isaka and M. Tanticharoen, J. Org. Chem., 2001, 66, 4803-4808.
196.
M. Kumarihamy, S. I. Khan, M. Jacob, B. L. Tekwani, S. O. Duke, D. Ferreira
and N. P. D. Nanayakkara, J. Nat. Prod., 2012, 75, 883−889.
197.
E. Niehaus, K. Kleigrewe, P. Wiemann, L. Studt, C. M. K. Sieber, L. R.
Connolly, M. Freitag, U. Guldener, B. Tudzynski and H. U. Hump, Chem. Biol.,
2013, 20, 1055–1066.
C.
180