Secondary Metabolites of Onygenales Fungi Exemplified by Aioliomyces pyridodomos

Article abstract of DOI:10.1021/acs.jnatprod.9b00121

Fungi from the order Onygenales include human pathogens. Although secondary metabolites are critical for pathogenic interactions, relatively little is known about Onygenales compounds. Here, we use chemical and genetic methods on Aioliomyces pyridodomos, the first representative of a candidate new family within Onygenales. We isolated 14 new bioactive metabolites, nine of which are first disclosed here. Thirty-two specialized metabolite biosynthetic gene clusters (BGCs) were identified. BGCs were correlated to some of the new compounds by heterologous expression of biosynthetic genes. Some of the compounds were found after one year of fermentation. By comparing BGCs from A. pyridodomos with those from 68 previously sequenced Onygenales fungi, we delineate a large biosynthetic potential. Most of these biosynthetic pathways are specific to Onygenales fungi and have not been found elsewhere. Family level specificity and conservation of biosynthetic gene content are evident within Onygenales. Identification of these compounds may be important to understanding pathogenic interactions.

Full text of DOI:10.1021/acs.jnatprod.9b00121

Article
pubs.acs.org/jnp
Cite This: J. Nat. Prod. XXXX, XXX, XXX−XXX
Secondary Metabolites of Onygenales Fungi Exemplified by
Aioliomyces pyridodomos
Zhenjian Lin,† Thomas B. Kakule,† Christopher A. Reilly,§ Sinem Beyhan,‡,⊥ and Eric W. Schmidt*,†
†Department of Medicinal Chemistry, University of Utah, Salt Lake City, Utah 84112, United States
‡Department of Infectious Diseases, J. Craig Venter Institute, La Jolla, California 92037, United States
⊥Department of Medicine, University of California, San Diego, La Jolla, California 92093, United States
§Department of Pharmacology & Toxicology and Center for Human Toxicology, University of Utah, Salt Lake City, Utah 84112,
United States
S
* Supporting Information
ABSTRACT: Fungi from the order Onygenales include
human pathogens. Although secondary metabolites are critical
for pathogenic interactions, relatively little is known about
Onygenales compounds. Here, we use chemical and genetic
methods on Aioliomyces pyridodomos, the first representative of
a candidate new family within Onygenales. We isolated 14
new bioactive metabolites, nine of which are first disclosed
here. Thirty-two specialized metabolite biosynthetic gene
clusters (BGCs) were identified. BGCs were correlated to
some of the new compounds by heterologous expression of
biosynthetic genes. Some of the compounds were found after
one year of fermentation. By comparing BGCs from A.
pyridodomos with those from 68 previously sequenced
Onygenales fungi, we delineate a large biosynthetic potential. Most of these biosynthetic pathways are specific to Onygenales
fungi and have not been found elsewhere. Family level specificity and conservation of biosynthetic gene content are evident
within Onygenales. Identification of these compounds may be important to understanding pathogenic interactions.
ungi produce secondary metabolites that establish and
Here, we examined Aioliomyces pyridodomos, a nonpatho-
genic Onygenales isolate from a coral reef tunicate in the
Eastern Fields of Papua New Guinea. Through genome
sequencing, we analyzed BGC content of A. pyridodomos and
compared it to the available Onygenales genomes. This
analysis revealed a large untapped diversity of Onygenales
compounds awaiting discovery from both pathogenic and
nonpathogenic isolates. Sequence analysis shows that the
number of BGCs in Onygenales is similar to that found in
Eurotiales, but encoding a different and new suite of
compounds.
To determine the chemical potential of A. pyridodomos, we
performed fermentation and chemical analysis, leading to the
discovery of 14 polyketides (1−14). Previously, we identified a
new polyketide derivative, oxazinin A (10), which exhibited
activity against transient receptor potential (TRP) channels.20
Subsequently, we described a series of unusual polyketides,
onydecalins (11−14), with anti-TRP and anti-Histoplasma
activities.21 Here, we describe further new polyketide structural
families encompassing aintennols (1−6) and aiorsellinates (7−
9). We identified some of the BGCs linked to A. pyridodomos
F
maintain infections.1 For example, Aspergillus fumigatus
(order Eurotiales) synthesizes gliotoxin as part of its repertoire
to circumvent the human immune system.2 Eurotiales fungi,
such as Aspergillus spp. and Penicillium spp., are the best-
studied producers of secondary metabolites, with thousands of
known compounds including those important in virulence. In
contrast, fungi from the order Onygenales include important
human pathogens that are challenging to treat,3,4 but much less
is known about their secondary metabolites. Several
Onygenales compounds from pathogens have been produced
via heterologous expression,5,6 but it remains unknown if these
are produced in the native host. Similarly, an Onygenales
biosynthetic gene cluster (BGC) implicated in virulence is an
iron-sequestering siderophore pathway with unknown chem-
ical products.7,8
From nonpathogenic Onygenales, many compounds have
been reported. A few examples include polyketides and
isoprenoids from marine9 and terrestrial Onygenales.10−15
Onygenales and Eurotiales strains contain a similar number of
BGCs responsible for producing secondary metabolites,
although the types of genes differ slightly between the
groups.16−19 Thus, it might be expected that both pathogenic
and nonpathogenic Onygenales provide a reservoir of new
secondary metabolites important in infection.
Received: February 8, 2019
© XXXX American Chemical Society and
American Society of Pharmacognosy
A
DOI: 10.1021/acs.jnatprod.9b00121
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Chart 1
Figure 1. Predicted PKS and NRPS BGCs in the A. pyridodomos genome.
compounds by sequence analysis and heterologous expression.
Our data suggest that Onygenales is a rich source of untapped
secondary metabolites.
a draft genome with 526 scaffolds. Using the antiSMASH22
algorithm with cluster finder,23 we identified 86 putative BGCs
in the genome. The contigs containing BGCs were manually
examined to ensure that they contained the entire BGC and
that BGCs did not span multiple scaffolds. We excluded the
category “cf_putative”, which are speculative BGC calls,
leading to 32 BGCs belonging to well-defined secondary
metabolic classes. We focused on the 19 polyketide and
RESULTS AND DISCUSSION
■
Biosynthetic Pathways in A. pyridodomos. A. pyrido-
domos E110162 was sequenced with Illumina HiSeq, providing
B
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Figure 2. Onygenales contains a wealth of secondary metabolic BGCs. The purple star indicates A. pyridodomos. A phylogenetic tree (left) was
obtained by concatenating 17 conserved proteins in 69 Onygenales strains with available genome sequences and performing a maximum-likelihood
method with MEGA-X.27 Bootstrap values are indicated at the nodes of the tree. Family level taxonomy is indicated, except for the important
genera Coccidioides and Paracoccidiodes, which belong to Ajellomycetaceae and Onygenaceae, respectively. antiSMASH was used to predict BGCs,
which are enumerated in the bar chart at the right (excluding the antiSMASH category “cf_putative”).
nonribosomal peptide metabolic BGCs (Figure 1), because
these are important biosynthetic families in fungi with
extensive experimental data.24,25 Twelve BGCs contained at
least one polyketide synthase (PKS), five contained a
nonribosomal peptide synthetase (NRPS), and two contained
a PKS and an NRPS. There were no hybrid PKS-NRPS
genes26 in the genome.
Comparison with Other Fungal Genomes. We sought
to understand the A. pyridodomos genome in comparison to
those in other Onygenales. We compared the identified A.
pyridodomos BGCs to those from 68 sequenced Onygenales
strains (Table S1) available in GenBank at the time of analysis.
To obtain BGCs, we applied antiSMASH22 to the 69 genomes
to identify 1888 BGCs (excluding “cf_putative”), of which
1184 contained a PKS, an NRPS, or both. These were overlaid
on a maximum-likelihood phylogenetic tree of the 69 strains
(Figure 2). As expected, >90% of the PKS genes belong to the
vast group of fungal type I iterative PKSs.24 Families
Arthrodermataceae and Onygenaceae contain the most
abundant BGCs, while the Ajellomycetaceae family contains
relatively fewer. Ascosphaera, which is deeply branched within
the Onygenales tree, contains the fewest BGCs.
NRPS clusters, which are the most straightforward to analyze.
Using antiSMASH, we found that only 38 out of the 1184
BGCs had significant similarity (over 70% of the genes show
similarity to known clusters) to BGCs from fungi outside of
the Onygenales. These related BGCs were found in 17 strains
from the order Eurotiales and 10 strains from class
Sordariomycetes (Figure S1). Many of these 27 strains are
plant or insect pathogens. Thus, while fungal PKS and NRPS
genes share many common features, this analysis suggests that
Onygenales encodes a unique suite of compounds in
comparison to other fungi.
We asked how the Onygenales BGCs were distributed
within the order, comparing the fungal taxonomic family to the
biosynthetic Gene Cluster Family (GCF), as defined by
Medema et al.28 The concept of GCF was introduced so that
whole biosynthetic pathways could be compared.29 Using
Medema’s method,28 1184 PKS and NRPS BGCs were
classified into 103 GCFs and 119 singletons, for a total of
222 groups. We used two different methods, Cytoscape
networking30 and cluster analysis,28 to graphically analyze the
distribution of GCF in comparison to fungi at the family level
(Figures 3 and S2). Both methods revealed that the majority of
GCFs (96% of the 222) are specific to a single fungal family
within Onygenales. Seven GCFs in the network are shared by
We next sought to determine whether the Onygenales BGCs
were similar to those from other fungi. We focused on PKS and
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specific to the family level except for siderophores involved in
iron acquisition.
Overall, these results indicate that Onygenales contain a
mostly new and uncharacterized biosynthetic potential, at least
in the PKS/NRPS gene families. Because the Onygenales
include important human pathogens, characterizing this
potential should be a priority.
A limitation in this study results from our current
understanding of fungal biosynthesis. Fungal PKS genes are
often similar, yet their chemical products differ in unpredict-
able ways. These differences are caused both by the PKSs and
by combinations of different tailoring enzymes that modify
PKS products. This reinforces the need to characterize the
chemicals from Onygenales: which of the compounds are
actually produced, do they contribute to pathogenesis, and do
they represent new structural motifs?
Polyketides from A. pyridodomos. We grew A.
pyridodomos for at least 1−2 months to obtain natural products
and for up to a year to obtain further compounds. We selected
the medium ISP2 because it was one of the few that afforded
reasonable growth for this otherwise challenging isolate. While
we initially worked with extracts isolated after a few weeks of
fermentation, because of the slow growth of the fungus we left
the culture for two months. The fungus continued to grow, and
a much larger abundance of polyketides was produced.
Therefore, we left the culture for a much longer period (one
year) to see whether the yield would increase. To our surprise,
not only did the fungus keep growing, but additional
metabolites appeared after a one-year period. The new
compounds appeared likely to be nonenzymatic, but
interesting and novel, rearrangement products31 (Table 1).
Figure 3. Cytoscape analysis reveals that most Onygenales
biosynthetic pathways are specific to individual families of fungi.
We identified 1184 PKS and NRPS BGCs, which are indicated by
circles. Connections of the circles by lines indicate GCFs. The color
of each circle indicates the fungal family that contains the BGC. Thus,
the multiple colors linked by lines indicate GCFs that are distributed
among more than one fungal family. If only a single color is present,
the BGC was found only within a single family.
two different families, and only two GCFs (GCF1 and GCF3)
are more widely distributed in Onygenales. These results
suggest a family level specialized metabolite conservation in
Onygenales.
Table 1. Compound Production after Extended
Fermentation
All of the A. pyridodomos BGCs are singletons, except for
two pathways belonging to the widely distributed GCFs 1 and
3. This result reinforces the potential family level status of this
strain. We are currently working to refine this phylogenetic
analysis.
time
shaking/30 °C
static/21 °C
25 days
60 days
365 days
1, 3, 4, 7, 8, 9, 10, 11, 12
1, 3, 4, 7, 8, 9, 11, 12
1, 2, 3, 4, 5, 6, 7, 8, 9, 11, 12, 13, 14
We examined the three most widely occurring GCFs,
GCF1−3, all of which include an NRPS gene (Figure 4). Based
upon previous studies, these common GCFs are likely
responsible for the production of hydroxamate siderophores,
involved in iron acquisition.7 For example, Hwang et al.
identified GCF1 as an iron-regulated siderophore pathway
(SID1) required for host colonization,8 although the chemical
structure of the siderophore was not defined. Thus, the
biosynthetic pathways found in Onygenales are relatively
Thus, extended fermentation may provide a method to
increase chemical diversity from microbial strains. Because of
the lengthy nature of this experiment, it was performed once.
For this reason, it is worth attempting on other systems before
generalizing our result.
In total, 14 polyketides (1−14) were isolated from A.
pyridodomos. Our strategy was to purify all of the abundant
metabolites, as Onygenales have rarely been examined
Figure 4. Gene organization of the most common NRPS GCFs shared among multiple families in Onygenales. GCF1 is composed of relatives of
the previously identified SID1.8
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chemically. We described five of these previously: onydecalins
A−D (11−14)21 and oxazinin A (10).20 Here, we describe six
further oxazinin analogues (aintennols A−F (1−6)) and three
other polyketides (aiorsellinates A−C (7−9)). Of these, 5, 6,
8, 9, 13, and 14 likely represent rearrangement products (vide
infra). The remainder of compounds represent mature
products or intermediates, as experimentally defined below.
Compounds 5, 6, 13, and 14 were isolated only after 1 year of
fermentation (Table 1).
Aintennol A (1) was isolated as a pale yellow solid with a
1
molecular formula of C22H30O2. H, 13C, COSY, and HSQC
NMR data (Table 2 and Figure 5) revealed that 1 has some
1
Table 2. H (500 MHz) and 13C NMR (125 MHz) Data of
Compounds 1 and 2 in CDCl3
1
2
no.
δC, type
δH (J in Hz)
δC, type
δH (J in Hz)
1
153.0, C
159.0, C
1-OH
7.30, brs
12.07, s
2
3
4
5
6
7
8
127.1, C
129.8, CH 6.90, s
131.7, C
135.5, C
123.0, C
126.4, CH 6.40, d (15.7)
135.8, CH 6.05, dd (15.7,
10.4)
131.7, CH 6.22, dd (14.6,
10.8)
130.1, CH 5.76, dq (14.6, 6.8) 132.4, CH 5.85, dq (10.8, 6.8)
18.2, CH3 1.83, d (6.8)
61.5, CH2 4.93, s
29.2, CH2 3.34, d (7.3)
122.3, CH 5.34, t (7.3)
133.7, C
18.1, CH3 1.76, s
26.0, CH3 1.76, s
32.4, CH2 3.21, d (7.4)
123.6, CH 5.18, t (7.4)
131.6, C
128.4, C
137.6, CH 7.17, s
130.6, C
139.6, C
118.2, C
123.6, CH 6.57, d (15.7)
139.8, CH 6.12, dd (15.7,
10.4)
131.2, CH 6.28, dd (14.7,
10.8)
Figure 5. Key COSY and HMBC correlations for compounds 1−7.
C-4, C-5, and from H-3 to C-13, C-18 located the substitutions
of the two benzene groups, the CH2OH group, and the
pentadienyl moiety on the pentasubstituted phenyl ring in 1.
The deshielded chemical shift of C-1 (δ 153.0) of the benzene
ring indicated a hydroxy group at C-1, establishing the
complete structure of 1.
9
10
11
12
13
14
15
16
17
18
19
20
21
22
Aintennol B (2) was isolated as a yellow solid with the
molecular formula C22H28O2, which is two protons lighter than
1. This was explained by the substitution of an alcohol in 1
with an aldehyde in 2. The 1H NMR data (Table 2 and Figure
5) had very similar signals to those of 1, except for the absence
of the CH2OH group in 2. Instead, a deshielded singlet proton
was observed at δ 10.10 (H-12), consistent with an aldehyde at
C-12. HMBC correlations from H-12 to C-1, C-6, and C-5
confirmed the aldehyde substitution at C-6 in the benzene
ring. Further, the IR spectrum of 1 in comparison to 2 was
consistent with the addition of a carbonyl. However, no HSQC
correlation for H-12 was observed. Proton H-12 was not
exchangeable, as overnight incubation with 10% CD3OD did
not wash out the signal. Evidence for an aldehyde was further
established in the deshielded shift of 1-OH (δ 7.30 brs in 1
versus δ 12.07 in 2), resulting from a hydrogen bond between
1-OH and 12-CHO. Finally, the UV spectrum of 2 contains a
λmax almost identical to that of albiducin B,32 also suggesting
conjugation with an aldehyde.
Aintennols C (3) and D (4) were isolated as isomeric, pale
yellow solids with the molecular formula C22H28O4. The 1D
NMR data (Table 3 and Figure 5) of 3 and 4 are similar to
those of 1. The major difference in comparison to 1 is that in 3
one of the isoprene groups was replaced with two singlet
methyl groups (δ 1.62, 1.20, H-16, H-17), a singlet oxygenated
methine group (δH 4.40, H-3, δC 77.4 C-3), an oxygenated
nonprotonated carbon (δC 84.4 C-2), and a ketone carbonyl
(δC 194.7, C-4). HMBC correlations from H3-16/H3-17 to C-
2, C-3, from H-3 to C-2, C-4, and from H-6 to C-4 supported a
4-chromanone moiety in 3, identical to that found in oxazinin
A(10). A spin system from H-11 to H-15 was determined to
result from a pentadienyl unit based upon COSY correlations.
The chemical shifts of the carbons in this pentadienyl unit are
18.6, CH3 1.85, d (6.8)
a
ND
10.10, s
31.5, CH2 3.33, d (7.3)
121.6, CH 5.31, t (7.3)
132.5, C
17.9, CH3 1.72, s
25.7, CH3 1.76, s
27.3, CH2 3.24, d (7.4)
122.6, CH 5.16, t (7.4)
133.4, C
18.2, CH3 1.69, s
26.3, CH3 1.72, s
17.9, CH3 1.71, s
25.7, CH3 1.73, s
a
ND = not detected.
structural features similar to those of oxazinin A (10). A triplet
olefinic proton (δ 5.34, H-14) exhibited a strong COSY
correlation with a methylene group (δ 3.34, H2-13) and a weak
correlation with two singlet methyl signals (δ 1.76, 6H, H3-16,
17), indicating the presence of a prenyl group in 1. A second
prenyl group in the structure can be deduced from the NMR
data (δ 5.18, H-19; δ 3.21, H2-18; δ 1.69, 1.72, H3-21,22). The
COSY spectrum of 1 contained cross-peaks between H-7/H-
8/H-9/H-10/H3-11, indicating that 1 has the same penta-
dienyl moiety as that in 10. Additionally, five nonprotonated
carbons (δ 153.0, 127.1, C-1, C-2; 131.7, 135.5, 123.0, C-3, C-
4, C-5) and an aromatic methine group (δH 6.90, δC 129.8)
were observed in the 13C NMR data, revealing a pentasub-
stituted benzene ring. An oxygenated methylene group was
observed at δH 4.93, δC 61.5, and an exchangeable proton
signal at δ7.30 was assigned to a phenolic hydroxy group.
Finally, HMBC correlations from H2-12 to C-1, C-6, C-5, from
H-7 to C-6, from H2-13 to C-1, C-2, C-3, from H2-18 to C-3,
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1
1
Table 3. H (500 MHz) and 13C NMR (125 MHz) Data of
Table 4. H (500 MHz) and 13C NMR (125 MHz) Data of
Compounds 3 and 4 in CD3CN
Compounds 5 and 6 in CD3CN
3
5
5
6
no.
δC, type
δH (J in Hz)
δC, type
δH (J in Hz)
no.
δC, type
δH (J in Hz)
δC, type
δH (J in Hz)
2
3
4
5
6
7
8
9
84.4, C
84.2, C
77.5, CH 4.44, s
195.1, C
118.9, C
125.6, CH 7.57, s
134.1, C
2
3
4
5
6
7
8
9
83.6, C
83.9, C
77.4, CH 4.40, s
194.7, C
76.1, CH 4.49, s
193.6, C
76.2, CH 4.46, s
193.3, C
a
a
a
ND
ND
ND
125.9, CH 7.52, s
134.0, C
147.5, C
131.1, CH 7.77, s
131.0, CH 7.78, s
a
a
ND
ND
146.7, C
126.1, C
156.2, C
156.0, C
a
a
128.8, C
ND
ND
10 157.5, C
156.7, C
10 157.4, C
157.0, C
11 126.3, CH 6.57, d (2.9)
12 137.8, CH 6.56, d (6.7)
125.0, CH 6.29, d (11.3)
133.0, CH 6.41, dd (11.9,
11.3)
128.4, CH 5.74, dd (15.0,
11.9)
133.5, CH 5.89, dq (15.5,
6.8)
18.3, CH3 1.69, d (6.7)
27.1, CH3 1.62, s
18.1, CH3 1.23, s
32.3, CH2 3.22, d (6.0)
123.3, CH 5.20, t (6.2)
133.8, C
17.8, CH3 1.66, s
26.0, CH3 1.71, s
57.3, CH2 4.53, brs
11 40.5, CH 3.60, q (5.8)
12 46.9, CH 3.20, m
13 54.7, CH 3.07, q (5.5)
14 38.2, CH2 2.59, dq (9.4, 5.8);
2.17, d (9.4)
15 12.7, CH3 0.83, d (7.0)
16 26.4, CH3 1.76, s
17 17.0, CH3 1.24, s
18 30.7, CH2 3.27, m
19 121.3, CH 5.05, t (6.2)
20 132.8, C
21 17.7, CH3 1.70, s
22 25.3, CH3 1.71, s
23 198.1, C
40.8, CH 3.60, q (5.8)
46.4, CH 3.19, m
55.7, CH 3.07, q (5.5)
38.5, CH 2.59, dq (9.4, 5.8);
2.17, d (9.4)
12.2, CH3 0.79, d (6.7)
26.1, CH3 1.76, s
16.8, CH3 1.26, s
30.8, CH2 3.27, m
121.6, CH 5.06, t (6.2)
133.1, C
17.2, CH3 1.70, s
25.4, CH3 1.71, s
197.3, C
13 132.5, CH 6.32, ddd (15.2, 6.7,
2.9)
14 132.9, CH 5.91, dq (15.1, 6.8)
15 18.4, CH3 1.84, d (6.8)
16 26.9, CH3 1.62, s
17 18.0, CH3 1.20, s
18 32.7, CH2 3.31, m
19 123.3, CH 5.20, t (6.2)
20 133.8, C
21 17.8, CH3 1.71, s
22 26.0, CH3 1.74, s
23 56.6 CH2 4.61 d (11.7), 4.55 d
(11.7)
a
ND = not detected.
a
ND = not detected.
between the benzobicyclo[3.1.1]heptanone moiety and the
prenylated benzene ring. The chemical shifts and coupling
constants for H-14 in both 5 and 6 are almost identical to
those of a synthetic compound, endo-2-isopropyl-2,3-dihydro-
1,3-methanonaphthalen-4(1H)-one, but drastically different
from exo-2-isopropyl-2,3-dihydro-1,3-methanonaphthalen-
4(1H)-one.33 This finding established an endo-benzobicyclo-
[3.1.1]heptanone moiety in 5 and 6. There was no observed
Cotton effect in the ECD spectra of 5 and 6, indicating that
both are racemic. The 1H NMR and 13C spectra of 5 and 6 are
almost identical, with only slight differences between them. In
extended incubation in methanol, 5 and 6 slowly interconvert,
indicating that the difference between the compounds is at a
labile center and not at the carbocycle. Because C-3 is racemic
in compounds 3 and 4 and the C-3 position is labile due to the
adjacent carbonyl, it is likely that 5 and 6 differ in the relative
orientation between C-3 and the carbocyclic portion. The
absence of a Cotton effect in 5 and 6 indicates that the
carbocycle is also racemic. This indicates that the formation of
the benzobicyclo[3.1.1]heptanone moiety in 5 and 6 is likely
nonenzymatic. Thus, 5 and 6 are both racemic; they are
diastereomers that differ by the relative configuration between
C-3 and the endo-benzobicyclo[3.1.1]heptanone moiety. The
absence of distance correlations between the two halves of the
molecules precluded establishment of the relative config-
urations of 5 and 6.
very similar to those found in oxazinin A, indicating the same
trans double bonds in 3, while the chemical shifts of the
protons are very different. (This difference is caused by
deshielding from an adjacent aromatic group in 10.) Because
of overlap between H-11 and H-12, a HOMODEC decoupling
experiment (Figure S56) helped to improve coupling constant
accuracy. An unusually small coupling constant was observed
between H-12 and H-13 (JH12−H13 = 6.7 Hz). The 1D NMR
data of 4 differed from those of 3 only in the chemical shifts
and J-values of the pentadienyl unit, especially for positions 11,
12, and 13 (Table 3). The coupling constant (JH11−H12 = 11.3
Hz) suggested a cis configuration of the C-11/C-12 double
bond in 4. There was no observed Cotton effect in the
electronic circular dichroism (ECD) spectrum of 3 and 4,
despite a strong dynode voltage corresponding to the UV
absorption spectra. Thus, as found for 10, 3 and 4 are racemic.
Aintennols E (5) and F (6) were isolated only after 1 year of
fermentation as isomeric, pale yellow solids with the molecular
formula C22H26O4. The 1D NMR data (Table 4 and Figure 5)
of 5 and 6 are almost identical. The spectra for 5 and 6 were
similar to those for 3 and 4, with most differences at C-23 and
C-11−C-15. A ketone carbonyl (δC 198.1, C-23), three
methines (δC 40.5, C-11, 46.9, C-12, 54.7, C-13), a methylene
(δC 38.2, C-14), and a doublet methyl (δC 12.7, C-15) were
observed in the 1D NMR data of 5. COSY correlations
between H-11/H-12/H-13/H2-14/H3-15 suggested that all of
the protons for the additional sp3 carbons are in a single spin
system. HMBC correlations from H3-15 to C-11, C-12, C-13
and from H2-14 to C-12 and C-23 established a
benzobicyclo[3.1.1]heptanone moiety in the structure of 5.
HMBC correlations from H-14 to C-8 supported the link
Aiorsellinate (7) was isolated as a colorless oil with the
molecular formula C20H30O9. The presence of a benzene ring
in 7 was deduced by the interpretation of HMBC and COSY
correlations (Figure 5). The six aromatic carbons, including a
methine (C-6, δ 112.6) and five nonprotonated carbons (C-2−
5, δ 120.9, 157.0, 119.3, 157.7 and C-7 δ 135.0), indicated the
presence of a highly substituted doubly oxygenated benzene
F
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1
Table 5. H (500 MHz) and 13C NMR (125 MHz) Data of Compounds 7−9 in DMSO-d6
7
8
9
no.
δC, type
168.7, C
120.9, C
δH (J in Hz)
δC, type
δH (J in Hz)
δC, type
δH (J in Hz)
1
2
3
4
5
6
7
8
9
10
11
12
13
14
1′
167.2, C
119.8, C
167.7, C
119.1, C
157.0, C
119.3, C
157.7, C
112.6, CH
135.0, C
20.1, CH3
23.0, CH2
123.1, CH
131.3, C
156.3, C
118.4, C
156.9, C
112.0, CH
134.2, C
20.6, CH3
22.4, CH2
123.0, CH
130.1, C
24.9, CH3
15.1, CH3
62.8, CH3
60.5, CH2
155.9, C
118.0, C
156.5, C
111.7, CH
133.7, C
19.6, CH3
22.2, CH2
122.4, CH
129.7, C
25.3, CH3
17.0, CH3
61.8, CH3
67.6, CH2
6.50, s
6.43, s
6.44, s
2.12, s
3.16, d (6.7)
5.11, t (6.7)
2.19, s
3.22 d, (6.8)
5.16, t (6.8)
2.14, s
3.15, d (6.7)
5.10, t (6.5)
26.2, CH3
18.2, CH3
62.7, CH3
64.0, CH2
1.62, s
1.74, s
3.69, s
3.65, m; 3.44, m
1.60, s
1.68, s
3.63, s
3.81, m; 3.66, m
1.60, s
1.68, s
3.64, s
4.48, d (11.3); 4.11, dd (11.3, 6.9)
a
(64.3)
1′-OH
2′
4.39, t (5.2)
3.87, m
4.59, t (5.0)
5.02, m
71.3, CH
(71.2)
75.2, CH
68.3, CH
3.74, dd (8.0, 7.0)
a
2′-OH
3′
3′-OH
4′
5.00, d (5.3)
5.27, d (7.4)
a
73.5, CH (73.8)
71.0, CH
67.5, CH
70.9, CH
3.97, dd (7.5, 7.5)
4.52, d (7.5)
3.45, m
71.0, CH
69.4, CH
3.45, m
3.59, m
3.42, m
a
(71.1)
4′-OH
5′
4.35, d (5.7)
3.85, m
4.47, d (5.2)
3.36, m
69.7, CH
(69.7)
70.9, CH
63.7, CH2
69.2, CH
63.6, CH2
3.56, m
a
5′-OH
6′
4.74, d (5.2)
3.63, m; 3.43, m
4.22, d (8.0)
3.60, m; 3.36, m
63.5, CH2
3.59, m; 3.37, m
a
(63.8)
6′-OH
4.58, t (5.0)
4.34, t (5.6)
a
13C chemical shifts (obtained from HSQC spectrum) in compound 7a.
ring in the structure of 7. The HMBC correlations from H2-9
to C-3, C-4, and C-5 established the prenyl group at C-4. The
HMBC correlations from H3-8 to C-2, C-6, and C-7, from H-6
to C-2, C-4, and C-5, and from H3-14 to C-3, as well as the
chemical shifts of C-3 and C-5 established the placement of the
other groups on the benzene ring. The chemical shift of C-2
and the ester carbonyl at C-1 determined the position of the
ester and established the presence of a prenylated orsellinate
moiety. The remaining six oxygenated sp3-hybridized carbons
and five alcohol exchangeable protons (Table 5) suggested a
hexitol moiety, which was confirmed by COSY correlations
between H-1′/H-2′/H-3′ and between H-6′/H-5′/H-6′.
Finally, the HMBC correlation from H-3′ to C-1 established
an ester bond between the orsellinate moiety and hexitol
moiety. To further confirm the structure, 7 was hydrolyzed and
analyzed by LC-MS. A new product (m/z 251 [M + H]+) was
observed, which corresponded to the expected carboxylate of 7
(Figure S5). Mass spectrometric fragmentation data of 7
provided further evidence for the structure (Figure S6). In
particular, an ion at m/z 233 suggested a loss of a hexitol (182
Da) in comparison to an ion at m/z 415 ([M + H]+), while the
ion at m/z 251 suggested an addition of H2O to the ion at m/z
233. An ion at m/z 195, which is 56 Da less than the ion at m/
z 251, suggested a loss of C4H8 from the prenyl group.
except for the chemical shifts of the hexitol moiety. In
comparison to 7, the chemical shifts of protons and carbon at
C-1′ (δH 4.48, 4.11, δC 67.6) in 8 were deshielded, while in 9
those at C-2′ (δH 5.02, δC 75.2) were deshielded. This
suggested ester bond migration to C-2′ in 8 and to C-1′ in 9,
respectively. Indeed, treatment of 7 in Na2CO3 solution
overnight led to an accumulation of 8 and 9. In early time
points of the fungal fermentation (10 days), only 7 was
present, while at later points 8 and 9 accumulated. Because 7
was the major component in the extract of the fungus at all
time points, 7 is the likely enzymatic product of the pathway.
Coupling constants indicated that the hexitols in 7−9 were
likely to be either sorbitol or mannitol. Chemical shifts of the
hexitols in 7−9 matched those of mannitol very well, especially
C-2′ to C-6′ of 7, which were almost identical to the same
carbons found in mannitol. Analogue 7a was synthesized and
had nearly identical chemical shifts and coupling constants to
those of 7, supporting the presence of mannitol. By comparing
the experimental and calculated ECD spectra (Figure S7) of 7,
the hexitol was determined to be D-mannitol.
Biological Activities of A. pyridodomos Polyketides.
Several of the new compounds were active in assays. We first
investigated the extract because it showed activity against a
panel of transient receptor potential channels. TRPs are
important throughout the body and include several channels
important in itch and pain, which may potentially be of interest
Aiorsellinates B (8) and C (9) were constitutional isomers
of 7. The NMR data of 8 and 9 were very similar to those of 7
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in topical fungal infections.34 However, the purified com-
pounds were only modestly active. In particular, 1 was active
against TRPV3 and TRPV4 at 32 and 70 μM, respectively. The
compounds were tested for activity against Histoplasma
capsulatum. Several compounds exhibited modest activity
(Table 6), with selectivity for H. capsulatum in comparison
to other fungi.
named “ain” for its putative role in biosynthesis of 1 (Figure
S4). Onydecalins (11−14) were previously defined as
biogenetically similar to betaenones,35 the biosynthesis of
which has been well defined by the Oikawa group;36 a single
betaenone-like cluster (ond) is present in the genome (Figure
S8). Work is ongoing in our laboratories to biochemically
confirm this assignment.
Finally, aiorsellinates (7−9) originate with the single
compound 7. In this instance, the core structure appears to
be composed of orsellinic acid, the biosynthesis of which has
been well studied for decades,37 while new decorations of O-
methylation, prenylation, and esterification make the com-
pound unique. A single BGC, aio (Figure 6), contained
putative orsellinate PKS (AioS), prenyltransferase (AioP),
methyltransferase (AioM), and possible mannitol transferase
(AioT) genes. Predicted AioS is very similar to characterized
orsellinic acid synthases, except that it terminates with an acyl
carrier protein domain instead of a thioesterase. We propose
that the neighboring AioT is likely to be involved in the
transfer of the ACP-linked orsellinate group, in analogy to a
similar protein involved in trichothecene species acetylation.
Other alternatives include the possibility that mannitol could
be added as a detoxification mechanism, which may not be
encoded by AioT.
Table 6. Anti-Histoplasma Activity of Purified Compounds
compound
Histoplasma IC50 (μg/mL)
1
3
5
6
13
14
8
16
32
64
2
16
0.5
0.001
a
a
amphotericin B
itraconazole
a
Previously reported data.21
Identification of Biosynthetic Gene Clusters. Bioge-
netic hypotheses for 1−14 were proposed based upon
structural data. Compounds 1−14 are all polyketide derivatives
that are likely to result from one of the 14 PKS genes in the
genome. Aintennols (1−6) and oxazinin (10) likely result
from a single biosynthetic pathway, in which 1 or 3 is the
probable primary enzymatic product. This pathway would
require at least one prenyltransferase, at least one oxidase, and
at least one PKS with a terminal reduction domain. Indeed,
one BGC was identified to have all of these features, which
were absent in other clusters in the genome; this cluster was
To confirm these assignments, we turned to heterologous
expression in a model fungus. This route was chosen because
of the challenge of generating a new genetic model,38 especially
because we have not yet found conditions under which
sporulation occurs for this new family. Genes aioS, aioM, aioP,
and aioT were cloned into an expression vector that we
designed for high-titer polyketide expression and then
transformed either singly or in combination into the expression
Figure 6. (A) aio gene cluster and domain organization of the PKS gene. (B) The genes aioT, aioS, aioM, and aioP were overexpressed in F.
heterosporum. Incubation with precursor analogue 15 led to formation of two new compounds (16 and 17), as shown in HPLC traces at 290 nm.
H
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host, Fusarium heterosporum.39 In some cases, we have detected
>1 g/kg of heterologous polyketides using this strategy.
However, in this instance no new products could be detected.
We reasoned that the likely limiting factor would be the PKS,
as occasionally these genes do not adapt well to heterologous
systems. Therefore, we added orsellinate ethyl ester to the
medium (for better cell penetration in comparison to the free
acid), affording the predicted prenylated and methylated
products only when the correct genes were in place (Figure 6).
These derivatives were further modified by oxidation of the
isoprene group in the course of fermentation by a nonspecific
oxidase in the F. heterosporum genome, indicating that F.
heterosporum may not be a good host for heterologous
production of 1−3. By contrast, in control fermentations
consisting of wild-type F. heterosporum or a mutant containing
GFP in place of aio genes, no new products were detected
unless orsellinic acid ethyl ester was supplied. This
demonstrates that aioP and aioM are the functional orsellinate
prenyltransferase and methyltransferase, respectively.
(1.2 mg). Compounds 1, 3, 4, 7, 8, and 9 were also isolated from a 2-
month shaking culture (extracts A)21 following a similar separation
procedure.
Aintennol A (1): pale yellow solid; UV (MeOH) λmax 224, 267 nm;
IR (polyethylene) νmax 3501, 2972, 1452, 1249, 1230, 1000 cm−1; 1H
and 13C NMR (Table 2); HRESIMS m/z 325.2167 [M − H]− (calcd
for C22H30O2, 325.2168).
Aintennol B (2): yellow solid; UV (MeOH) λmax 250, 285, 371 nm;
IR (polyethylene) νmax 2960, 1692, 1641, 1450, 1301, 1261, 991, 770
1
cm−1; H and 13C NMR (Table 2); HRESIMS m/z 325.2166 [M +
H]+ (calcd for C22H28O2, 325.2168).
Aintennol C (3): pale yellow solid; UV (MeOH) λmax 229, 308, 350
nm; IR (polyethylene) νmax 3422, 2981, 1692, 1611, 1440, 1260, 1103
1
cm−1; H and 13C NMR (Table 3); HRESIMS m/z 357.2066 [M +
H]+ (calcd for C22H28O4, 357.2066).
Aintennol D (4): pale yellow solid; UV (MeOH) λmax 228, 265,
297, 343 nm; IR (polyethylene) νmax 3421, 2982, 1692, 1602, 1433,
1261, 1104 cm−1; 1H and 13C NMR (Table 3); HRESIMS m/z
357.2067 [M + H]+ (calcd for C22H28O4, 357.2066).
Aintennol E (5): yellow solid; UV (MeOH) λmax 205, 242, 266, 354
nm; IR (polyethylene) νmax 2972, 1691, 1600, 1454, 1212, 1110 cm−1;
1H and 13C NMR (Table 4); HRESIMS m/z 355.1907 [M + H]+
(calcd for C22H26O4, 355.1909).
Aintennol F (6): yellow solid; UV (MeOH) λmax 205, 242, 266, 354
nm; IR (polyethylene) νmax 2972, 1700, 1600, 1453, 1212, 1111 cm−1;
1H and 13C NMR (Table 4); HRESIMS m/z 355.1908 [M + H]+
(calcd for C22H26O4, 355.1909).
Aiorsellinate A (7): pale yellow solid; UV (MeOH) λmax 199, 251
nm; ECD (0.5 mg/mL, methanol), λmax (Δε) 290 (−0.8), 254 (1.23)
nm; IR (polyethylene) νmax 3390, 1712, 1598, 1463, 1272, 1080 cm−1;
1H and 13C NMR (Table 5); HRESIMS m/z 415.1969 [M + H]+
(calcd for C20H30O9, 415.1963).
Aiorsellinate B (8): pale yellow solid; UV (MeOH) λmax 199, 251
nm; IR (polyethylene) νmax 3391, 1712, 1598, 1462, 1272, 1081 cm−1;
1H and 13C NMR (Table 5); HRESIMS m/z 415.1970 [M + H]+
(calcd for C20H30O9, 415.1963).
Aiorsellinate C (9): pale yellow solid; UV (MeOH) λmax 199, 251
nm; IR (polyethylene) νmax 3390, 1712, 1599, 1461, 1272, 1082 cm−1;
1H and 13C NMR (Table 5); HRESIMS m/z 415.1969 [M + H]+
(calcd for C20H30O9, 415.1963).
Sequencing. The mycelium of A. pyridodomos was collected by
centrifugation and then extracted as previously described.41 The
resulting DNA samples were repurified with the Genomic DNA Clean
& Concentrator kit (Zymo Research). Illumina libraries were
prepared as ∼350 bp inserts from genome DNA. Libraries were
sequenced using an Illumina HiSeq 2000 sequencer in 101 bp/125 bp
paired-end runs. Raw fasta files were trimmed by sickle and assembled
using IDBA_ud42 (--mink = 20 --maxk = 100 --step = 20
--inner_mink = 10 --inner_step = 5 --prefix = 3 --min_count = 2
--min_support = 1 --num_threads = 0 ==seed_kmer = 30
--min_contig = 200 --similar = 0.95 --max_mismatch = 3 --min_pairs
= 3). The genome was annotated using Augustus43 de novo gene
prediction parameters, with ‘Coccidioides_immitis’ as the model
organism. The resulting gff output file was converted to a GenBank
file and submitted to the RAST44−46 online server for gene
identification.
CONCLUSION
■
We describe 14 natural products that are so far unique to a
new family of Onygenales fungi represented by A. pyridodomos.
We tentatively assigned their BGCs and confirmed one of the
assignments by heterologous expression. A large number of
uncharacterized natural products and BGCs await analysis in
the order Onygenales, which is a key group of pathogenic
fungi.
EXPERIMENTAL SECTION
■
General Experimental Procedures. Circular dichroism spectra
were obtained on a Jasco J720A spectropolarograph. IR spectra were
measured on a Thermo Fisher iS50 Fourier transform IR
spectrometer. NMR data were collected using either a Varian
INOVA 500 (1H 500 MHz, 13C 126 MHz) NMR spectrometer
with a 3 mm Nalorac MDBG probe or a Varian INOVA 600 (1H 600
MHz, 13C 150 MHz) NMR spectrometer equipped with a 5 mm
1H[13C,15N] triple resonance cold probe with a z-axis gradient,
utilizing residual solvent signals for referencing.40 High-resolution
mass spectra (HRMS) were obtained using a Bruker APEXII FTICR
mass spectrometer equipped with an actively shielded 9.4 T
superconducting magnet (Magnex Scientific Ltd.), an external Bruker
APOLLO ESI source, and a Synrad 50W CO2 CW laser. Supelco
Discover HS (4.6 × 150 mm) and semipreparative (10 × 150 mm)
C18 (5 μm) columns were used for analytical and semipreparative
HPLC, respectively, as conducted on a Hitachi Elite Lachrom System
equipped with a diode array L-2455 detector. Unless stated otherwise,
all reagents and solvents were purchased from commercial suppliers,
and all reactions were carried out under anhydrous conditions with an
argon atmosphere. Yields were calculated by HPLC chromatography
1
or H NMR spectroscopy.
Fermentation and Extraction. A. pyridodomos strain was grown
and extracted as previously described.21
Purification. The extract B21 (600 mg) from a 1-year-old culture
was separated into seven fractions (Fr1−Fr7) on a C18 column using
step-gradient elution of MeOH in H2O (20%, 40%, 60%, 70%, 80%,
90%, and 100%). Fr3 was further subjected to C18 HPLC (35%
CH3CN in H2O) to yield compounds 7 (2.5 mg), 8 (1.0 mg), and 9
(1.1 mg). Fr6 was purified by C18 HPLC (85% CH3CN in H2O) to
yield compound 1 (4.2 mg). Fr7 was purified by C18 HPLC (89%
CH3CN in H2O) to yield compound 2 (1.0 mg). Fr5 was fractionated
using a gradient HPLC method (50% CH3CN in H2O to 80%
CH3CN in H2O over 20 min) to yield five subfractions (Fr5-1−Fr5-
5). Fr5-3 was purified by C18 HPLC (63% CH3CN in H2O) to yield
compound 3 (8.0 mg) and 4 (7.0 mg). Fr5-4 was purified by C18
HPLC (66% CH3CN in H2O) to yield compounds 5 (1.0 mg) and 6
GCF Analysis. The Augustus43 annotation output was converted
to a GenBank file and used as the input for a standalone version of
antiSMASH 4.0.22 All BGCs were separated into individual GenBank
files. Initial pairwise similarity matrices between every BGC pair were
calculated using MultiGeneBlast28 with the parameters (-distancekb
10 -minpercid 50 -minseqcov 80) and then normalized as follows: (1)
The “total score” (MultiGeneBlast) between the query BGC and itself
was set as 1.0; (2) the “total score” between query BGC and target
BGC was normalized to the total score between query itself in step 1.
BGC pairs with a final pairwise similarity matrix (normalized score)
over 0.6 were selected and visualized in Cytoscape.30 Each GCF was
manually checked by examining the MultiGeneBlast alignment.
I
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Construction of Phylogenetic Tree. Proteins encoded in
Onygenales genomes were obtained using the Augustus annotation
output. Seventeen widespread genes present in a single copy in all
genomes were identified by performing a blastp search using A.
pyridodomos against 68 other Onygenales genomes. Hits were defined
as identity > 80%, query coverage > 0.8, and subject coverage > 0.8.
Genes that met those criteria and that were present in all 68
Onygenales genomes were selected and used for phylogenetic
analysis. Orthologous genes were aligned using t-Coffee47 (-mode
mcoffee -output = msf, fasta_aln). To remove poorly aligned regions,
the resulting alignment was subsequently trimmed with trimAl v1.448
with a gap threshold of 0.4. A concatenation of alignment of all
conserved proteins was done by PerlScript catfasta2phyml.pl (https://
github.com/nylander/catfasta2phyml). The maximum likelihood
trees were constructed using MEGA-X using the Jukes−Cantor
model and 500 bootstrap replicates to assess node support.
72 h time point of the culture. The culture was harvested and
extracted with EtOAc. The products were analyzed by HPLC-DAD,
LC-MS, and NMR, resulting in the identification of two new
compounds, 16 and 17. Compound 16: 1H NMR (500 MHz, CDCl3)
δH 6.27 (1H, s); 6.24 (1H, s); 4.36 (2H, q, J = 7.1 Hz); 3.78 (3H, s);
2.25 (3H, s); 1.36 (3H, t, J = 7.1 Hz); ESIMS m/z 211 [M + H]+.
Compound 17: 1H NMR (500 MHz, CDCl3) δH 11.87 (1H, s); 6.28
(1H, s); 4.64 (1H, t, J = 8.8 Hz); 4.40 (2H, q, J = 7.1 Hz); 3.04 (2H,
m); 2.43 (3H, s); 1.43 (3H, t, J = 7.1 Hz); 1.36 (3H, s); 1.23 (3H, s);
ESIMS m/z 281 [M + H]+.
ECD Calculation. ECD calculation was performed as previously
described.21
ASSOCIATED CONTENT
* Supporting Information
■
S
The Supporting Information is available free of charge on the
ACS Publications website at DOI: 10.1021/acs.jnat-
prod.9b00121.
Bioactivity Screening. Assays were performed as previously
described.21
Synthesis of Compound 7a. (See Scheme S1.) Ethyl orsellinate
(P1) was prepared as reported by Chen et al.49 Ethyl 4-(benzyloxy)-
2-hydroxy-6-methylbenzoate (P3) was prepared as described by Zhu
et al.50 Product P3 (300 mg) was treated with LiOH (96 mg) in
MeOH−H2O (1:2; 10 mL). The solution was refluxed for 5 h to yield
product P4 (270 mg). A mixture of product P4 (260 mg), thionyl
chloride (2 mL), and dimethylformamide (DMF) (100 μL) in dry
CH2Cl2 (150 mL) was heated to reflux for 6 h, and then the extra
thionyl chloride was removed under vacuum to yield product P5 (290
mg, oil). 1,2:5,6-Bis-O-(1-methylethylidene)-D-mannitol (262 mg) in
tetrahydrofuran (THF) (100 mL) was pretreated with NaH (40 mg,
60% in mineral oil) for 1 h and added to the flask containing product
P5 (210 mg). The resulting mixture was stirred for 2 h to yield
product P6 (350 mg). Product P6 (300 mg) was dissolved in 100 mL
of EtOH, 10% palladium on carbon (30 mg) was added, and the
mixture was treated with hydrogen gas (1 atm) for 3 h to yield
product P7 (200 mg). Product P7 (42 mg) was dissolved in dry DMF
(50 mL), and NaH (4 mg, 60% in mineral oil) was added. The
mixture was stirred at 40 °C for 2 h and cooled to room temperature.
3,3-Dimethylallyl bromide (14.7 mg, 12 μL) was added, and the
mixture was stirred at 60 °C overnight. The product was precipitated
by adding H2O (100 mL), filtered, and washed with H2O to yield a
white solid, which was further treated with 6 M HCl (10 mL) and
purified by C18 HPLC (20% MeCN in H2O) to yield compound 7a
Additional information (PDF)
AUTHOR INFORMATION
Corresponding Author
*Tel: 801-585-5234. Fax: 801-581-7087. E-mail: ews1@utah.
■
edu.
ORCID
Christopher A. Reilly: 0000-0002-5006-1982
Eric W. Schmidt: 0000-0001-5839-694X
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
This work was funded by NIH R01GM107557 and
R35GM122521 to E.W.S. and C.R. and NIH R00AI112691
and R01AI137418 to S.B. We thank the University of Utah
Center for High Performance Computing for access to
computational resources.
1
(2.1 mg): H NMR (600 MHz, DMSO-d6) δH 9.59 (1H, s, 5-OH);
REFERENCES
■
6.34 (1H, s, H-6); 5.16 (1H, d, J = 7.6 Hz, H-3′); 4.95 (1H, t, H-10);
4.83 (1H, d, 2′-OH); 4.57 (1H, d, 5′-OH); 4.43 (1H, t, 6′-OH); 4.30
(1H, t, 1′-OH); 4.22 (1H, d, 4′-OH); 3.75 (1H, m, H-2′); 3.75 (1H,
m, H-5′); 3.38 (1H, m, H-4′); 3.63 (3H, s, H-14); 3.62 (1H, m, H-
1′a); 3.33 (1H, m, H-1’b); 3.59 (1H, m, H-6′a); 3.38 (1H, m H-6′b);
3.17 (2H, d, H-9); 2.06 (3H, s, H-8); 1.55 (1H, m, H-11); 1.69 (3H,
s, H-12); 1.61 (3H,s, H-13); ESIMS m/z 415 [M + H]+.
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