Reappraising the structures and distribution of metabolites from black aspergilli containing uncommon 2-benzyl-4H-pyran-4-one and 2-benzylpyridin-4(1H) -one systems

Article abstract of DOI:10.1021/np200454z

To date, natural products containing 2-benzyl-4H-pyran-4-one and 2-benzylpyridin-4(1H)-one substructures have been encountered in relatively few fungi outside of the black aspergilli clade. While exploring the occurrence of these compounds among Aspergillus spp., it was determined that the structures of the unusual furopyrrols tensidols A and B (5 and 6) and JBIR-86 and JBIR-87 (9 and 10) were incorrect and should be reassigned as 2-benzyl-4H-pyran-4-ones (7, 8, 11e, and 12, respectively). The origin of the unique N-phenyl groups in the 2-benzylpyridin-4(1H)-ones nygerones A and B (1 and 2) were also examined, and it was established that N-phenylamides added to the culture medium were suitable substrates for generating these metabolites; however, this phenomenon remained limited to a single fungus in our collection (Aspergillus niger ATCC 1015). A variety of 2-benzyl-4H-pyran-4-ones and 2-benzylpyridin-4(1H)-ones were detected among the black aspergilli, but only pestalamide B (13) was found in all 11 of the tested strains. These metabolites, as well as a group of synthetic analogues, demonstrated weak antifungal activity against several Candida strains, Aspergillus flavus, and Aspergillus fumigatus.

Full text of DOI:10.1021/np200454z

ARTICLE
pubs.acs.org/jnp
Reappraising the Structures and Distribution of Metabolites from
Black Aspergilli Containing Uncommon 2-Benzyl-4H-pyran-4-one
and 2-Benzylpyridin-4(1H)-one Systems
Jon C. Henrikson,† Trevor K. Ellis,† Jarrod B. King,† and Robert H. Cichewicz*,†,‡
†Natural Products Discovery Group, Department of Chemistry and Biochemistry, Stephenson Life Sciences Research Center,
101 Stephenson Parkway, University of Oklahoma, Norman, Oklahoma 73019, United States
‡Graduate Program in Ecology and Evolutionary Biology, University of Oklahoma, Norman, Oklahoma 73019, United States
S Supporting Information
b
ABSTRACT: To date, natural products containing 2-benzyl-4H-
pyran-4-one and 2-benzylpyridin-4(1H)-one substructures have
been encountered in relatively few fungi outside of the black
aspergilli clade. While exploring the occurrence of these com-
pounds among Aspergillus spp., it was determined that the
structures of the unusual furopyrrols tensidols A and B (5 and 6)
and JBIR-86 and JBIR-87 (9 and 10) were incorrect and should
be reassigned as 2-benzyl-4H-pyran-4-ones (7, 8, 11e, and 12,
respectively). The origin of the unique N-phenyl groups in the
2-benzylpyridin-4(1H)-ones nygerones A and B (1 and 2) were
also examined, and it was established that N-phenylamides added to the culture medium were suitable substrates for generating these
metabolites; however, this phenomenon remained limited to a single fungus in our collection (Aspergillus niger ATCC 1015). A
variety of 2-benzyl-4H-pyran-4-ones and 2-benzylpyridin-4(1H)-ones were detected among the black aspergilli, but only
pestalamide B (13) was found in all 11 of the tested strains. These metabolites, as well as a group of synthetic analogues,
demonstrated weak antifungal activity against several Candida strains, Aspergillus flavus, and Aspergillus fumigatus.
reviously, we reported the discovery of the 2-benzylpyridin-
4(1H)-one-containing metabolites nygerones A and B (1, 2)
and 2-benzyl-4H-pyran-4-one metabolites among black aspergilli
and explored the origins of the N-phenyl groups in 1 and 2, as
well as investigated the biological activities of these compounds.
P
from Aspergillus niger.1ꢀ4 Subsequent 1H NMR-guided efforts by
our group to purify additional 2-benzylpyridin-4(1H)-one and
related 2-benzyl-4H-pyran-4-one metabolites from A. niger
yielded several compounds that we presumed were structurally
similar to 1 and 2. However, during the course of dereplication
studies, a variety of natural products possessing R-pyridone,5
γ-pyrone,6 and furopyrrol7 skeletons were found in the literature,
and all of these compounds were reported to possess 1H NMR
chemical shifts that were virtually identical to the key diagnostic
resonances in 1 and 2. Fortunately, the R-pyridone-containing
variant had been previously examined by Ye and colleagues, who
successfully revised the proposed structure of aspernigrin A (3)5
as the more plausible 2-benzylpyridin-4(1H)-one shown for
compound 4.8 Our analysis of the furopyrrol-containing
molecules, which included tensidols A and B (5, 6),7 led to the
observation that these structures had arisen from a strikingly
’ RESULTS AND DISCUSSION
Structure Revision of the Furopyrrols. The tensidol family
of metabolites, which includes 5 and 6, as well as JBIR-86 (9) and
JBIR-87 (10),10 were reported from the fermentation broths of a
marine isolate of A. niger and an unidentified Aspergillus sp.,
respectively. We were initially prompted to examine the spectro-
scopic data for 5 and 6 since the unusual furopyrrol skeleton in
these structures was reportedly unprecedented.7 Focusing on 5,
we noted several unusual chemical shift assignments that we
could not reconcile. Namely, the 13C NMR data presented for
C-2 (162.0), C-3 (178.0), C-3a (119.1), C-4 (164.3), C-5
(116.0), and C-6a (168.8) were inconsistent with our expecta-
tions for carbon resonances in isolated or fused furan and pyrrole
systems (Supporting Information, Figure S1). At the same time,
we purified a small molecule from A. niger whose MS and NMR
data matched those reported for 5 (Supporting Information,
Table S1); however, de novo structure characterization studies
1
different interpretation of H and 13C NMR data that were
indistinguishable from the values reported for the 2-benzyl-4H-
pyran-4-ones carbonarone A (7)6 and pestalamide A (8),9
respectively. In this report, we address the structure revision of
the tensidols using a combination of NMR spectroscopy, ab initio
carbon chemical shift calculations, and total synthesis. We have
also analyzed the distribution of the 2-benzylpyridin-4(1H)-one
Received: June 1, 2011
Published: August 19, 2011
r
Copyright
2011 American Chemical Society and
1959
dx.doi.org/10.1021/np200454z J. Nat. Prod. 2011, 74, 1959–1964
American Society of Pharmacognosy
|

Journal of Natural Products
ARTICLE
carried out by our group suggested that γ-pyrone 7 was a more
reasonable alternative to consider. The molecular formula for 7
exhibited a low H:C ratio (0.85), which, along with the presence
of JHꢀC couplings in the HMBC data, might have played
confounding roles in previous attempts to unambiguously assign
the correct structure for this metabolite.11,12 Further inspection
of the literature demonstrated that nearly one year after the
disclosure of 5 in 2006, Zhang and colleagues reported the
alternative structure 7 from a marine isolate of Aspergillus
carbonarius, although no mention of 5 was provided in their
account.6 We propose that 2-benzyl-4H-pyran-4-one 7 repre-
sents the correct structure of this metabolite.
Figure 1. (A) Candidate structures (11aꢀe) evaluated as alternatives
during the revision of 9. (B) Mean average errors (MAE) were calculated
using ∑(δcalc ꢀ δexp)/#. A smaller MAE value represents better overall
agreement between the computational and published 13C NMR data.
(C) 1Hf13C HMBC correlations observed for synthetic 11e.
4
respectively. This would presumably necessitate the incorpora-
tion of an odd number of nitrogens in each metabolite. However,
the published 13C NMR data for 9 clearly supported the presence
of a γ-pyrone, thus eliminating candidate 11a from further
consideration (Figure 1A). In addition, the observed chemical
shifts for the methyl group in 9 (δ 52.8) suggested that it was
attached to an oxygen atom, not the N-methyl amide shown for
candidate 11b (Figure 1A). These restrictions left open the
possibilities that the revised structure contained a primary
ketimine (11c) or methyl imidate (11d) (Figure 1A), but both
candidate compounds were also rejected based on biosynthetic
considerations and evaluations of their respective predicted 13C
NMR chemical shift data (Supporting Information, Figure S2).
With no plausible nitrogen-containing candidate structures
left to consider for 9, we focused on formulating a reasonable
alternative structure from the NMR data. This approach led us to
propose 11e as a candidate structure (Figure 1A). We proceeded
to test this hypothesis using gauge-including atomic orbital based
13C NMR chemical shift calculations on both 9 and 11e. Density
functional theory calculations performed at the b3lyp/6-31+g(d,p)
level yielded a poor mean absolute error value of 16.0 ppm for 9,
but a reasonable value of 3.8 ppm for 11e (Figure 1B). Figure 2
illustrates that structure 9 exhibited several pronounced devia-
tions in the calculated carbon NMR values for positions C-2, C-3,
C-3a, C-4, C-5, and C-6a from the published data (Δδcalcꢀδexp
ꢀ46.5, ꢀ20.0, ꢀ27.0, ꢀ41.1, ꢀ25.8, and ꢀ20.6ppm, respectively).
A similar set of circumstances surrounded compound 6, which
was originally described as a methyl succinate derivative of 5.7 In
this case, a 2008 report by Ding et al. unknowingly provided
structure 89 as a reasonable revision for compound 6. Our
1
evaluation of both data sets confirmed that the H and 13C
NMR chemical shifts reported for the two compounds were
identical and that 8 should be regarded as the correct structure of
this metabolite.
In 2010, we encountered a new opportunity to further test our
assertion that the furopyrrol metabolites should be revised as
2-benzyl-4H-pyran-4-ones. We noted that Takagi et al. had
obtained what appeared to be two new furopyrrol-containing
metabolites, JBIR-86 (9) and JBIR-87 (10);10 however, both
compounds exhibited some problematic features. Among our
initial concerns were the incongruous 13C NMR chemical shifts
(Supporting Information, Figure S1) and the presence of suspect
4JHꢀC couplings within the furopyrrol ring system.10 Addition-
ally, the spectrometric analysis of 9 and 10 was reported to have
yielded HRESIMS data containing even-mass quasi-molecular
ions at m/z 244.0986 [M + H]+ and 358.1286 [M + H]+,
1960
dx.doi.org/10.1021/np200454z |J. Nat. Prod. 2011, 74, 1959–1964

Journal of Natural Products
ARTICLE
Table 1. Comparisons of the Observed 1H and 13C NMR
Data for 11e versus the Values Reported for 9
synthetic 11e
9a
position
δH
δC
δH
δC
2
8.41, s
162.0
119.3
175.0
116.9
167.4
39.7
8.43, s
162.0
119.1
175.0
117.3
167.6
40.0
3a
4
Figure 2. Deviations in the calculated versus observed 13C NMR
chemical shifts for 9 and 11e. Differences approaching zero demonstrate
good agreement between calculated versus reported 13C NMR data,
whereas large differences indicate poor agreement between calculated
and reported 13C NMR experimental data.
5
6.19, s
3.81, s
6.21, s
3.83, s
6a
7
8
133.9
129.1
129.1
127.8
163.6
52.7
134.1
129.4
129.4
128.0
163.9
52.8
9, 13
10, 12
11
7.22, m
7.35, m
7.32, m
7.24, m
7.32, m
7.32, m
Scheme 1. Synthesis of 11e
3
3-OCH3
3.85, s
3.87, s
a Data reproduced from Takagi, M.; Motohashi, K.; Hwang, J.-H.; Nagai,
A.; Kazuo, S. J. Antibiot. 2010, 63, 371ꢀ373.
and 2-benzyl-4H-pyran-4-one skeletons have been reported from
black aspergilli.14,15 Given our previous encounter with this
family of metabolites from A. niger, as well as our desire to better
understand its distribution among the members of this clade, we
prepared a focused library of 11 Aspergillus strains for further
investigation. The fungi were grown with and without epigenetic
modifiers,2ꢀ4 and the extracts examined by LC-ESIMS. By
probing extracted-ion chromatograms for each of the target
compounds (1, 2, 4, 7, 8, and 13), we were able to determine
which black aspergilli were producers of the 2-benzylpyridin-
4(1H)-one and 2-benzyl-4H-pyran-4-one metabolites under our
culture conditions (Supporting Information, Table S2). Inter-
estingly, the 2-benzylpyridin-4(1H)-one-containing compound
pestalamide B (13)9 was the only metabolite detected in all 11 of
the black aspergilli strains (note: the 13C NMR data provided for
13 by Ding et al. have been revised16). In contrast, the 2-benzyl-
4H-pyran-4-one-containing analogue of 13, compound 8, was
observed in a much more restricted group of five fungal strains,
and in some cases it was detected only under chemical epigenetic
induction conditions. The unsubstituted amide analogues of 13
and 8, metabolites 4 and 7, respectively, were also found in
relatively few fungi (two and four strains, respectively). At this
time, the role of methyl succinate conjugation of these metabo-
lites is unknown.
We were struck by the limited distribution of compound 1,
which was detected only in A. niger ATCC 1015 treated with
suberoylanilide hydroxamic acid (SAHA) (Supporting Informa-
tion, Table S2). The origin of the N-phenyl group in 1 is an
intriguing feature of this metabolite, and its biosynthetic source
was not readily apparent to us. Our original investigation of 1,1
including follow-up feeding studies involving the incorporation
of aniline in the fungal growth medium did not support the idea
that the phenyl ring of SAHA was the N-phenyl group precursor.
To further verify the role SAHA plays in the production of
nygerones, a subsequent test employing p-fluoro SAHA
(Supporting Information, Scheme S1A) was used as the eliciting
agent. The addition of p-fluoro SAHA resulted in the production
of p-fluoro nygerone B (14) from A. niger ATCC 1015
(Supporting Information, Table S1). This led us to use p-fluoro
In contrast, 11e showed promise as a revised candidate structure
with much less deviation in the predicted versus observed
13C NMR data for each of the corresponding carbon atoms
(Δδcalcꢀδexp ꢀ5.8, ꢀ2.3, 6.6, ꢀ7.1, ꢀ2.7, and ꢀ1.7 ppm,
respectively).
We further tested the prospect of revising 9 to candidate
structure 11e by engaging in its total synthesis. We prepared 11e
using a modification of the procedure reported by McCombie
et al. (Scheme 1).13 Upon purification of the product, HRESIMS
yielded a quasi-molecular ion at m/z 245.0815 [M + H]+ (calcd
for C14H13O4, 245.0814). The 1H and 13C NMR data of
synthetic 11e were remarkably consistent with each of the
chemical shift values reported for 9 (Table 1). Moreover, both
the previously published and new 1Hf13C HMBC data that we
examined showed that all but one of the observed heteronuclear
couplings (H-5fC-3) were now accounted for as two- to three-
bond correlations (Figure 1C). We are still unable to provide an
explanation for the disagreement between the ESIMS data
published by Takagi et al. and our own results. However, in light
of the evidence, we are quite confident that 11e represents the
correct structure for JBIR-86. On the basis of this finding, we also
reevaluated compound 10. Upon inspection of its 1H, 13C, and
1Hf13C HMBC NMR data (Supporting Information, Table S1
and Figure S3), reconsideration of its probable shared biosyn-
thetic origins with 11e, and co-isolation of compound 8 from the
same crude extract, we have concluded that 10 should be revised
to the 2-benzyl-4H-pyran-4-one-containing structure 12.
Distribution of 2-Benzylpyridin-4(1H)-ones and 2-Benzyl-
4H-pyran-4-ones among Black Aspergilli. To date, the major-
ity of secondary metabolites bearing 2-benzylpyridin-4(1H)-one
1961
dx.doi.org/10.1021/np200454z |J. Nat. Prod. 2011, 74, 1959–1964

Journal of Natural Products
ARTICLE
aniline for further feeding studies (Supporting Information,
Scheme S2), but we did not observe any production of 14 when
the substrate was administered alone or with SAHA present
(Supporting Information, Table S1B and S1C). These data
indicated that aniline was not a suitable substrate for generating
the N-phenyl group in 1. We next considered the possibility that
an acetanilide precursor might be utilized in establishing the N-
phenyl group in compound 1. Accordingly, we prepared p-fluoro
acetanilide (Supporting Information, Scheme S1D and S1E)
from p-fluoro aniline and acetyl chloride and added it alone or
with SAHA to the growth medium. Under both conditions, 14
was obtained, which allowed us to purify the compound and
confirm its structure using ESIMS and 1H, 13C, and 19F 1D and
2D NMR (Supporting Information, Table S1). These data
indicate that acetanilides and SAHA are suitable precursor
substrates for generating the N-phenyl group of the nygerones.
Evaluation of Antimicrobial Activity. In light of the sparse
biological activity data reported for this family of metabolites, we
prepared a group of several naturally occurring and synthetic γ-
pyrone- and γ-pyridone-containing compounds for investigation
(1, 8, 11e, and 14). We also took this opportunity to make 4-oxo-
6-phenyl-4H-pyran-3-carboxylic acid (15) and associated ester
derivatives (16, 17) using the synthetic approach presented in
Scheme 1 (Supporting Information, Scheme S2 and Table S1).
None of the compounds exhibited any inhibitory effects against
bacteria (Acinetobacter baumannii ATCC 19606, A. baumannii
ATCC BAA-1605, Burkholderia cepacia ATCC 25608, Entero-
bacter cloacae ATCC 13047, Escherichia coli ATCC 10798, E. coli
ATCC 11775, Pseudomonas aeruginosa ATCC 10145, Klebsiella
pneumonia ATCC 33495, K. pneumonia ATCC 51503, Staphy-
lococcus epidermidis ATCC 12228, Staphylococcus aureus
ATCC 25923, and S. aureus ATCC 700787) at a concentration of
100 μg/mL.
and semipreparative Shimadzu HPLC systems with Phenomenex C18
Gemini columns.
Density Functional Theory Calculations (DFT). Full geometry
optimization and carbon chemical shift calculations for the proposed
11e and JBIR-86 (9) were performed on eight nodes of the Pentium-4
Xeon54 Quad Core Linux Cluster (running Red Hat Linux) using
Gaussian03. Geometry optimization with frequency check and GIAO
methods for determining carbon chemical shifts were performed using
the basis set b3lyp/6-31+g(d,p). DFT calculations of tetramethylsilane
(TMS) were also performed to determine the carbon chemical shift to
be used as a reference. Carbon chemical shifts were referenced by
subtracting the calculated δC of each carbon from the calculated δC of
TMS. Analysis of results was performed by comparing the calculated
chemical shifts with the published chemical shifts of JBIR-86.
Synthesis of 3-Carbomethoxy-6-benzyl-4-pyrone (11e).
The general scheme for syntheszing 11e followed the methodology
developed by McCombie et al.13 The enaminone, methyl 2-[-
(dimethylamino)methylene]-3-oxobutanonte, was synthesized as de-
scribed with the application of methyl acetoacetate as a starting material
instead of ethyl acetoacetate. Methyl acetoacetate (5.4 mL, 50 mmol)
was added to the DMFꢀMe2SO2 adduct in CH2Cl2 (75 mL) at 0 °C,
and triethylamine (10 mL) was added slowly over 3 min. The mixture
was stirred for 2 h at room temperature, washed with aqueous tartaric
acid (10% w/v, 100 mL) and H2O (100 mL ꢁ 2), and dried over
anhydrous MgSO4. The sample was gravity filtered (Whatman #1 filter
paper), and the solvent from the filtrate was evaporated under vacuum.
The crude material was subjected to silica gel flash chromatography
using a CHCl3ꢀEtOAc gradient (100:0 to 0:100). The solvent was
evaporated from the enaminone under vacuum and further purified by
flash chromatography using a hexaneꢀEtOAc gradient (90:10 to 0:100).
The solvent was dried to give the white solid product (Z)-methyl
2-((dimethylamino)methylene)-3-oxobutanoate.
To construct the pyrone ring, a solution of hexamethldisilazine
(1 mL) in THF (6 mL) was stirred at ꢀ78 °C, and n-BuLi in hexane
(3 mL of a 1.6 M solution) was added slowly over 2 min. After
the mixture cooled, a solution of the enaminone ((Z)-methyl 2-
((dimethylamino)methylene)-3-oxobutanoate) (2 mmol) and phenyl
acetyl chloride (2.4 mmol) in THF (5 mL) was added over 1 min. The
dry ice/acetone bath was removed, and the solution was allowed to
warm for 5 min. Diethyl ether (20 mL) was added to the reaction,
followed by 3 N HCl (8 mL). The solution was stirred rapidly overnight.
The following day, the organic phase was washed with saturated sodium
bicarbonate (100 mL) and H2O (100 mL ꢁ 2), then dried over
anhydrous MgSO4. The sample was gravity filtered (Whatman #1 filter
paper), and the filtrate dried under vacuum. The residue was subjected
to silica gel flash chromatography using a CH2Cl2ꢀEtOAc gradient
(100:0 to 0:100). Final purification was performed by preparative HPLC
using a MeOHꢀH2O gradient (50:50 to 100:0). The resulting product
appeared as a red-brown, amorphous solid (16 mg, 6%): 1H (CD3OD,
400 MHz) δH 8.45 (1H, s, H-2), 7.32 (3H, m, H-10, H-11, H-12), 7.24
(2H, m, H-9, H-13), 6.22 (1H, s, H-5), 3.88 (3H, s, 14-OCH3), 3.84
(2H, s, H-7); 13C (CD3OD, 100 MHz) δC 175.1 (C-4), 167.7 (C-6),
163.9 (C-14), 161.8 (C-2), 134.0 (C-8), 129.4 (C-9, C-13), 129.3 (C-10,
C-12), 128.0 (C-11), 119.7 (C-3), 117.2 (C-5), 52.8 (14-OCH3), 39.72
(C-7); HRESIMS m/z 245.0815 [M + H]+ (calcd for C14H13O4,
245.0814).
Previously, Fukuda et al.7 and Takagi et al.10 had reported that
tensidols (7, 8, 11e, and 12) were capable of potentiating the
activity of a subinhibitory concentration of miconazole (0.06 μM)
against Candida albicans in a disk diffusion assay. Our attempts to
confirm this activity in a broth microdilution assay against C.
albicans ATCC 38245 failed to detect any imidazole potentiation.
However, we did observe that compounds 11e, 16, and 17
provided complete growth inhibition against five Candida spp.
(C. albicans, ATCC 38245, Candida glabrata NRRL Y-65, Candida
krusei ATCC 27803, Candida parapsilosis ATCC 12969, and
Candida tropicalis ATCC 12968) and two nonblack aspergilli
(Aspergillus fumigatus NRRL A1100 and Aspergillus flavus NRRL
A1120) at a concentration of 100 μg/mL. At 10 μg/mL, 11e was
capable of inhibiting only A. flavus growth, whereas 16 and 17
exhibited broader ranges of activities against C. krusei, A. flavus, and
A. fumigatus at this reduced concentration.
’ EXPERIMENTAL SECTION
General Experimental Procedures. All solvents were of ACS
grade or better. NMR data were obtained on Varian 300, 400, and 500
spectrometers and processed using VNMRj software. LC-ESIMS data
were collected using a Thermo-Finnigan Surveyor LC system and a
Finnigan LCQ Deca mass analyzer. HRESIMS data were obtained by
electrospray ionization employing an Agilent 6538 UHD Accurate-Mass
Quadrupole TOF mass analyzer. Initial separations were performed by
normal-phase flash chromatography using a Biotage Isolera system.
Further isolation and purification was performed using preparative
LC-ESIMS Analysis of Extracts from Black Aspergilli Grown
under Chemical Epigenetic Induction Conditions. Freeze-
dried spores from 11 strains of black aspergilli (Aspergillus aculeatus
NRRL 5094, Aspergillus basiliensis NRRL 35542, A. carbonarius NRRL
346, A. carbonarius NRRL 369, Aspergillus ellipticus NRRL 5120,
Aspergillus ibericus NRRL 35644, Aspergillus japonicus NRRL 2053, A.
niger NRRL 326, A. niger NRRL 1015, Aspergillus parasiticus NRRL 465,
and Aspergillus tubingensis NRRL 4875) were obtained from the ARS
1962
dx.doi.org/10.1021/np200454z |J. Nat. Prod. 2011, 74, 1959–1964

Journal of Natural Products
ARTICLE
mycology collection. All strains were cryopreserved in 15% (v/v) glycerol
solution at ꢀ80 °C. Cryopreserved samples were removed and plated on
potato-dextrose agarandgrown for 14dat25°C. Hyphaefrom eachstrain
were suspended insterile H2O and usedto inoculate three1 L Erlenmeyer
flasks containing 250 mL of sterile potato-dextrose broth. Each culture
flask was treated with either 50 μM suberoylanilide hydroxamic acid
(dissolved in 50:50 DMSOꢀH2O), 50 μM 5-azacytidine (dissolved in
50:50 DMSOꢀH2O), or a bolus of 50:50 DMSOꢀH2O. Cultures were
grown for 21 d at room temperature on a rotary shaker at 150 rpm. After
21 d, 2% (wt/vol) Diaion HP-20 resin was added to the flasks, which were
further shaken for 4 h, followed by filtration of the cell mass and resin. The
cell mass and resin from each flask was extracted in 250 mL aliquots of
MeOH over a 48 h period (2ꢁ). The MeOH extracts were combined, and
solvent was removed by rotary evaporation.
Extracts were prepared for LC-ESIMS analysis by suspending samples
in a minimal amount of MeOH and adsorbing the solution onto C18.
The dried C18 stationary phases loaded with samples were then placed in
SPE cartridges and rinsed with 5 mL of deionized H2O followed by 5 mL
of MeOH (each collected separately on a vacuum manifold). The
MeOH fractions were evaporated in preweighed vials. The vials that
contained the dried material from the MeOH fractions were weighed,
and the dried material was resuspended in a 90:10 MeOHꢀH2O
solution at a concentration of 20 mg/mL. A portion of each sample
was then transferred to a 2 mL Eppendorf tube and centrifuged at
14 000 rpm for 5 min. The supernatants were placed in vials for LC-
ESIMS analysis using a MeOHꢀH2O gradient (25:75 to 100:0).
The mass detector was set to scan a range from m/z 120ꢀ1500 in
positive mode.
Synthesis of p-Fluoro SAHA. Suberic acid monomethyl ester
(2.01 g, 10.7 mmol), p-fluororaniline (1.42 g, 12.8 mmol), and hydro-
xybenzotriazole (1.73 g, 12.8 mmol) were dissolved in 25 mL of DMF.
To this solution was added diisopropylcarbodiimide (1.62 g, 12.8
mmol), and the reaction mixture was stirred at room temperature for
3 h. The reaction was quenched by pouring the mixture over 450 mL of
ice stirred in H2O. After the ice had melted, the resulting precipitate was
filtered using vacuum-assisted filtration, followed by purification via flash
silica chromatography using a hexaneꢀEtOAC gradient. The procedure
provided the suber-p-fluoroanilic acid methyl ester in 82% yield (2.47 g).
Fresh methanolic hydroxyl amine was prepared by dissolving hydro-
xylamine hydrochloride (9.43 g, 135.6 mmol) in MeOH (25 mL)
followed by the addition of KOH (7.60 g, 135.6 mmol). The mixture
was allowed to stir under an atmosphere of nitrogen in an ice bath to aid
the precipitation of the resulting salt, which was subsequently removed
by filtration. The filtrate was transferred to a round-bottom flask, and a
solution of suber-p-fluoroanilic acid methyl ester (2.07 g, 7.36 mmol) in
10 mL of MeOH was added. This reaction mixture was stirred under
nitrogen at 50 °C for 12 h. The reaction was quenched via dilution in
MeOH with ice H2O (250 mL). The resulting aqueous solution was
neutralized with acetic acid, resulting in the precipitation of p-fluoro
SAHA. The white precipitate was filtered and evaporated under vacuum
to provide an overall product yield of 61%. Analysis of the product by
1H, 13C, and 19F NMR revealed that the p-fluoro SAHA was obtained in
high chemical purity without the need for further purification. 1H NMR
((CD3)2SO, 500 MHz) δ 10.35 (1H, br s, N-H-12), 9.91 (1H, br s, N-H-
2), 7.60 (2H, m, H13, H-17), 7.10 (2H, m, H14, H-16), 2.27 (1H, t, J =
7.5 Hz, H-9), 1.93 (2H, t, J = 7.5 Hz, H-4), 1.55 (2H, m, H-8), 1.48 (2H,
m, H-5), 1.26 (4H, m, H-6 and -7); 13C NMR ((CD3)2SO, 125 MHz) δ
171.7 (C-10), 169.8 (C-3), 159.6 (C-15), 136.3 (C-12), 121.1 (C-13,
C-17), 115.7 (C-14, C-16), 36.7 (C-9), 32.8 (C-4), 28.9 (C-6, C-7), 25.5
(C-5, C-8); 19F NMR ((CD3)2SO, 282 MHz) δ ꢀ119.96 (F-15);
HRESIMS m/z 305.1274 [M + Na]+ (calcd for C14H19FNaO3,
305.1277).
dextrose agar and grown for 14 d at 25 °C. Hyphae were suspended in
sterile H2O and used to inoculate a set of 20 Erlenmeyer flasks (1 L) each
containing 250 mL of sterile potato-dextrose broth. Each flask was
treated with 50 μM p-fluoro suberoylanilide hydroxamic acid (p-F-
SAHA) (dissolved in 50:50 DMSOꢀH2O). Controls were created by
inoculating four 1 L Erlenmeyer flasks containing 250 mL of sterile
potato-dextrose broth and 0.5% (v/v) of 50:50 DMSOꢀH2O. Cultures
were grown on a rotary shaker (150 rpm) for 14 d at room temperature,
and the combined culture broths were partitioned (3ꢁ) against an
equivalent volume of CH2Cl2. The organic phase was collected and
evaporated under vacuum. Treated and control extracts were analyzed
by LC-ESIMS as described above. Further fractionation was performed
on the p-F-SAHA culture extract by preparative HPLC using a
MeOHꢀH2O gradient (35:65 to 100:0) with final purification being
performed by semipreparative HPLC to yield 2 mg of light yellow,
amorphous solid: 1H NMR (CDCl3, 400 MHz) δ 9.89 (1H, br s, N-Hb-
15), 8.49 (1H, s, H-2), 7.23 (1H, m, H-11), 7.21 (2H, m, H-10, H-12),
7.10 (2H, m, H-30, H-50), 7.05 (2H, m, H-20, H-60), 6.83 (2H, m, H-9,
H-13), 6.61 (1H, s, H-5), 6.06 (1H, br s, N-Ha-15), 3.71 (2H, s, H-7);
13C NMR (CDCl3, 100 MHz) δ 177.7 (C-4), 166.0 (C-14), 164.1 (C-
40), 162.0 (C-10), 151.3 (C-6), 146.9 (C-2), 134.8 (C-8), 128.9 (C-20,
C-60, C-10, C-12), 128.5, C-9, C-13), 127.7 (C-11), 121.9 (C-5), 117.7
(C-3), 116.9 (C-30, C-50), 39.74 (C-7); 19F NMR (CDCl3, 376 MHz)
ꢀ112.36 (F-11); HRESIMS m/z 323.1192 [M + H]+ (calcd for
C19H16FN2O2, 323.1196).
3-Carboethoxy-6-phenyl-4-pyrone (16). Compound 16 was
prepared and purified on the basis of a previously reported method.13
1
The resulting product was a red crystalline solid: H NMR was in
agreement with the literature;13 HRESIMS m/z 245.0816 [M + H]+
(calcd for C14H13O4, 245.0814).
3-Carbomethoxy-6-phenyl-4-pyrone (17). Compound 17
was prepared and purified based on a previously reported method,13
with the exception that methyl acetoacetate was utilized instead of ethyl
acetoacetate. The resulting product was a red, crystalline solid: 1H NMR
(CD3OD, 500 MHz) δ 8.82 (1H, s, H-2), 7.89 (2H, m, H-8, H-12), 7.55
(2H, m H-10), 7.53 (2H, m, H-9, H-11), 6.94 (1H, s, H-5), 3.86 (3H, s,
H-14); 13C NMR (CD3OD, 125 MHz) δ 175.6 (C-4), 166.0 (C-2),
164.1 (C-6), 163.0 (C-13), 130.8 (C-9, C-10, C-11), 129.6 (C-7), 126.2
(C-8, C-12), 118.3 (C-3), 112.4 (C-5), 52.3 (C-14); HRESIMS m/z
231.0659 [M + H]+ (calcd C13H11O4, 231.0657).
3-Carboxylic-6-phenyl-4-pyrone (15). A 10 mg sample of 17
was reacted with 1 N NaOH in distilled THF for 3 h at room
temperature, then adjusted to pH 3 with 1 N HCl. The resulting solution
was partitioned against EtOAc, and the solvent removed under vacuum.
Purification was performed by preparative HPLC, yielding 6.2 mg of a
yellow, amorphous solid. 1H NMR ((CD3)2CO, 500 MHz) δ 9.88 (1H,
s, H-2), 8.04 (2H, m, H-8, H-12), 7.66 (1H, m, H-10), 7.60 (2H, m, H-9,
H-11), 6.95 (1H, s, H-5); 13C NMR ((CD3)2CO, 125 MHz) δ 193.2
(C-2), 178.5 (C-4), 167.1 (C-6), 161.7 (C-13), 132.8 (C-10), 130.4 (C-
7), 129.2, (C-9, C-11), 126.7 (C-8, C-12), 100.6 (C-3), 97.6 (C-5);
HRESIMS m/z 215.0358 [M + H]ꢀ (calcd for C12H7O4, 215.0350).
1
Pestalamide A (8). H and 13C NMR were in agreement with
published values;9 HRESIMS m/z [M + Na]+ 366.0956 (calcd for
C18H17NO6Na, 366.0954).
Biological Assays. Twelve bacteria and seven fungi were used in
the antimicrobial assays. Stock cultures of strains were delivered into
presterilized 96-well plates, and compounds prepared in DMSO were
added to each well of the 96-well plate (final DMSO concentration of
1%). Plates were shaken orbitally for 5 s, and an initial OD600 value was
recorded. Bacteria and fungi in 96-well plates were incubated for 24 and
48 h, respectively, in a humidified incubator at 37 °C. Plates were
removed and orbitally shaken, and a final OD600 value was recorded. The
final OD600 reading was subtracted from the initial OD600 reading to
obtain the change in OD600 (antimicrobial activity was determined by
Isolation and Purification of p-Fluoro Nygerone B (14). A
cryopreserved sample of A. niger ATCC 1015 was plated on potato-
1963
dx.doi.org/10.1021/np200454z |J. Nat. Prod. 2011, 74, 1959–1964

Journal of Natural Products
ARTICLE
the change in optical density). Chloramphenicol and ketoconazole were
used as antibacterial and antifungal positive controls, respectively.
(16) The 13C NMR chemical shift data published by Ding et al.9 in
Table 4 for pestalamide B were found to be incorrect. It appears as
though the 13C NMR shift data for pestalamide A in Table 3 were
superimposed onto Table 4. A list of the corrected chemical shifts is
found in Table S1 of the Supporting Information.
’ ASSOCIATED CONTENT
Supporting Information. Predicted (ACD) 13C NMR of
S
b
furan and pyrrol ring systems (Figure S1) and partial structures
of 11c and 11d (Figure S2), comparison of HMBC correlations
of JBIR-87 (10 and 12) (Figure S3), tables of 1H and 13C NMR
chemical shifts of 1, 2, 5/7, 6/8, 9/11e, 10/12, 13, 14, 15, and 17
(Table S1), LC-ESIMS analysis of black aspergilli (Table S2),
antimicrobial assay results (Table S3), schemes of feeding studies
for the production of 14 (Scheme S1AꢀE), and synthesis of 15,
16, and 17 (Scheme S2) are available free of charge via the
Internet at http://pubs.acs.org.
’ AUTHOR INFORMATION
Corresponding Author
*Tel: +1 (405) 325-6969. E-mail: rhcichewicz@ou.edu.
’ ACKNOWLEDGMENT
We wish to acknowledge the National Institutes of Health
(1RO1GM092219-01 and 1RO1AI085161-01) for financial sup-
port. DFT calculations was performed at the OU Supercomput-
ing Center for Education & Research (OSCER) at the University
of Oklahoma. We also thank J. Hayle for his assistance and
training with Gaussian03.
’ REFERENCES
(1) Henrikson, J. C.; Hoover, A. R.; Joyner, P. M.; Cichewicz, R. H.
Org. Biomol. Chem. 2009, 7, 435–438.
(2) Fisch, K. M.; Gillaspy, A. F.; Gipson, M.; Henrikson, J. C.;
Hoover, A. R.; Jackson, L.; Najar, F. Z.; Wagele, H.; Cichewicz, R. H.
J. Ind. Microbiol. Biotechnol. 2009, 36, 1199–1213.
(3) Williams, R. B.; Henrikson, J. C.; Hoover, A. R.; Lee, A. E.;
Cichewicz, R. H. Org. Biomol. Chem. 2008, 6, 1895–1897.
(4) Cichewicz, R. H.; Henrikson, J. C.; Wang, X.; Branscum, K. M.
Manual of Industrial Microbiology and Biotechnology, 3rd ed.; American
Society for Microbiology: Washington DC, 2009; pp 78ꢀ95.
(5) Hiort, J.; Maksimenka, K.; Reichert, M.; Peroviꢀc-Ottstadt, S.;
Lin, W. H.; Wray, V.; Steube, K.; Schaumann, K.; Weber, H.; Proksch, P.;
Ebel, R.; M€uller, W. E. G.; Bringmann, G. J. Nat. Prod. 2004,
67, 1532–1543.
(6) Zhang, Y. P.; Zhu, T. J.; Fang, Y. C.; Liu, H. B.; Gu, Q. Q.; Zhu,
W. M. J. Anitbiot. 2007, 60, 153–157.
(7) Fukuda, T.; Hasegawa, Y.; Hagimori, K.; Yamaguchi, Y.; Masuma,
R.; Tomoda, H.; Omura, S. J. Antibiot. 2006, 59, 480–485.
(8) Ye, Y. H.; Zhu, H. L.; Song, Y. C.; Liu, J. Y.; Tan, R. X. J. Nat. Prod.
2005, 68, 1106–1108.
(9) Ding, G.; Jiang, L. H.; Guo, L. D.; Chen, X. L.; Zhang, H.; Che,
Y. S. J. Nat. Prod. 2008, 71, 1861–1865.
(10) Takagi, M.; Motohashi, K.; Hwang, J.-H.; Nagai, A.; Shin-ya, K.
J. Antibiot. 2010, 63, 371–373.
(11) White, K. N.; Amagata, T.; Oliver, A. G.; Tenney, K.; Wenzel,
P. J.; Crews, P. J. Org. Chem. 2008, 73, 8719–8722.
(12) Choi, H.; Engene, N.; Smith, J. E.; Preskitt, L. B.; Gerwick,
W. H. J. Nat. Prod. 2010, 73, 517–522.
(13) McCombie, S. W.; Metz, W. A.; Nazareno, D.; Shankar, B. B.;
Tagat, J. J. Org. Chem. 1991, 56, 4963–4967.
(14) Nielsen, K. F.; Mogensen, J. M.; Johansen, M.; Larsen, T. O.;
Frisvad, J. C. Anal. Bioanal. Chem. 2009, 395, 1225–1242.
(15) Abarca, M. L.; Accensi, F.; Cano, J.; Caba~nes, F. J. Antonie van
Leeuwenhoek 2004, 86, 33–49.
1964
dx.doi.org/10.1021/np200454z |J. Nat. Prod. 2011, 74, 1959–1964