Bahamaolides A and B, antifungal polyene polyol macrolides from the marine actinomycete streptomyces sp.

Article abstract of DOI:10.1021/np3001915

Bahamaolides A and B (1 and 2), two new 36-membered macrocyclic lactones, were isolated from the culture of the marine actinomycete Streptomyces sp. derived from a sediment sample collected at North Cat Cay in the Bahamas. The planar structures of 1 and 2, bearing a hexaenone and nine consecutive skipped hydroxy groups, were determined by 1D and 2D NMR, mass, IR, and UV spectra. The absolute configurations of the bahamaolides were established by combined multistep chemical reactions and spectroscopic analysis. Bahamaolide A displayed significant inhibitory activity against Candida albicans isocitrate lyase and antifungal activity against various pathogenic fungi.

Full text of DOI:10.1021/np3001915

Article
pubs.acs.org/jnp
Bahamaolides A and B, Antifungal Polyene Polyol Macrolides from
the Marine Actinomycete Streptomyces sp.
Dong-Gyu Kim,† Kyuho Moon,† Seong-Hwan Kim,† Seon-Hui Park,† Sunghyouk Park,† Sang Kook Lee,†
Ki-Bong Oh,‡ Jongheon Shin,† and Dong-Chan Oh*,†
†Natural Products Research Institute, College of Pharmacy, Seoul National University, 1 Gwanak-ro, Gwanak-gu, Seoul 151-742,
Republic of Korea
‡Department of Agricultural Biotechnology, College of Agriculture and Life Science, Seoul National University, 1 Gwanak-ro,
Gwanak-gu, Seoul 151-921, Republic of Korea
S
* Supporting Information
ABSTRACT: Bahamaolides A and B (1 and 2), two new 36-membered macrocyclic lactones,
were isolated from the culture of the marine actinomycete Streptomyces sp. derived from a
sediment sample collected at North Cat Cay in the Bahamas. The planar structures of 1 and 2,
bearing a hexaenone and nine consecutive skipped hydroxy groups, were determined by 1D
and 2D NMR, mass, IR, and UV spectra. The absolute configurations of the bahamaolides
were established by combined multistep chemical reactions and spectroscopic analysis.
Bahamaolide A displayed significant inhibitory activity against Candida albicans isocitrate lyase
and antifungal activity against various pathogenic fungi.
ctinobacteria are an important source of antibiotics that
led the golden era of antibiotic discovery (1940−1960),
actinomycete strains and found that a strain isolated from
a seafloor sediment sample collected off North Cat Cay
in the Bahamas produces unusual hexaenone-bearing com-
pounds as its major chemical constituents. Further inves-
tigation of these secondary metabolites led to the discovery
of two new polyene polyol macrocyclic lactones with anti-
fungal activity, bahamaolides A and B (1 and 2). Here
we report the isolation, structural determination (including
absolute configurations), and biological activity of these rare
hexaene macrolides.
A
affording clinically useful antibiotic scaffolds such as amino-
glycosides, tetracyclines, erythromycin, vancomycin, and
amphotericin.1 However, the discovery of new bioactive natural
products from this promising bacterial group has declined
due to redundant studies on heavily investigated terrestrial
actinomycetes. Marine environments are now known to harbor
unique actinobacterial populations that may produce new
secondary metabolites with useful pharmaceutical properties.2
Despite their biomedical potential, systematic studies on the
secondary metabolites of marine actinomycetes have only
recently begun.3 Owing to the focus of efforts on marine
microbial natural products over the last 15 years, such as
actinomycetes, eubacteria, cyanobacteria, and fungi, the number
of novel bioactive chemicals from marine microorganisms has
dramatically increased since 1997,4 resulting in the discovery of
drug lead compounds that are currently in clinical trials.5
Novel antibacterial compounds such as marinomycin,6
abyssomicin,7 marinopyrrole,8 and lipoxazolidinone9 have been
discovered in marine actinomycetes. Although marine actino-
mycete-derived antifungal natural products are far less common
than their antibacterial analogues, new polyketide spiroketals,
the reveromycins,10 and a new bafilomycin derivative, all with
antifungal properties, were recently isolated from marine-
derived Streptomyces strains.11 This finding indicates the
potential for discovering antifungal lead compounds in marine
actinomycetes.
RESULTS AND DISCUSSION
■
A bacterial strain CNQ343, Streptomyces sp. based on 16S
rDNA analysis, was isolated from a sediment sample collected
at the depth of 27 m off North Cat Cay in the Bahamas. The
actinomycete was cultivated in a seawater-based medium.
Time-course chemical analysis of its EtOAc extract by LC/MS
revealed the production of polyene compounds with char-
acteristic UV spectra (λmax at 384 nm) and molecular ions
[M + H]+ at m/z 709 as major chemical constituents. We then
cultivated the bacterium on a large scale (40 L) and purified
the polyene compounds, bahamaolides A and B, by the com-
bination of normal-phase and reversed-phase chromatographic
techniques.
Bahamaolide A (1) was purified as a yellow powder and was
determined to have the molecular formula C39H64O11 by
Over the course of our search for new antibiotic lead com-
pounds in marine microbes, we screened various marine-derived
Received: March 12, 2012
Published: May 10, 2012
© 2012 American Chemical Society and
American Society of Pharmacognosy
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1
1
HRFABMS and H and 13C NMR data (Table 1). The H
and 14.9. Combined analyses of 13C, DEPT, and gHMQC
NMR spectra confirmed 55 carbon-bound protons in total;
the nine protons unaccounted for in the molecular formula
were assigned as exchangeable protons. The IR absorption at
1747 cm−1, coupled with the carbon signal at δC 168.0, revealed
the existence of an ester functionality. The observation of a UV
absorption maximum at 384 nm, 12 olefinic carbons in the 13C
NMR spectrum of 1 clearly displayed polyunsaturated and
1
NMR spectrum, and corresponding protons in the H NMR
spectrum indicated that the ester is conjugated to six double
bonds. The connectivity from C-1 to C-13 was also confirmed by
the array of COSY couplings in both pyridine-d5 and CD3OD
between the 12 olefinic protons. The HMBC showed a correlation
from H-2 (δH 6.12) to carbonyl carbon C-1 (δC 168.0). Similarly,
a combination of gCOSY, gHMQC, and gHMBC experimental
data elucidated a partial structure bearing nine hydroxy groups
1
in a repeating 1,3-pattern from C-15 to C-31. All of the H and
13C NMR chemical shifts were adequately assigned by detailed
analysis of gCOSY, gHMQC, and gHMBC spectroscopic data.
The repeating 1,3-hydroxy group moiety was determined
to be connected to the hexaenone through a methylene linker
(C-14, H2-14; δC 42.4; δH 2.75, 2.60) according to COSY
correlations of H-13 (δH 6.18)−H2-14 and of H2-14−H-15
(δH 4.52) and HMBC correlations from H-12 (δH 6.32) and
H-13 to C-14 and from H2-14 to C-13 (δC 133.9) and C-15
(δC 69.1). The other end of this fragment is connected to the
adjacent aliphatic methylenes at C-32 and C-33 according to
the HMBC correlation from H2-32 (δH 1.97, 1.76) and H2-33
(δH 1.31, 1.26) to C-31 (δC 73.3) and the COSY correlation
between H2-32 and H-31. The HMBC correlation from H2-33
to C-34 (δC 34.9) and strong HMBC correlations from the
polyhydroxylated features, with 12 olefinic protons between δH
7.63 and 6.12 and 10 protons attached to oxygenated carbons
1
between δH 5.08 and 4.03. Further analysis of the H NMR
spectrum of 1 identified 24 aliphatic protons in the region δH
2.75−1.26 and three methyl groups at δH 0.96, 0.96, and 0.78.
The 13C NMR and DEPT spectra divided the carbon signals
into one quaternary carbonyl at δC 168.0, 12 olefinic methines
from δC 146.0 to 121.2, 10 oxygenated methines from δC 79.3
to 65.0, 12 aliphatic methylenes from δC 48.3 to 30.9, one
aliphatic methine at δC 30.2, and three methyls at δC 20.6, 19.0,
Table 1. NMR Spectroscopic Data for Bahamaolides A (1) and B (2) in Pyridine-d5 (800 MHz)
bahamaolide A (1)
δH, mult
bahamaolide B (2)
bahamaolide A (1)
δH, mult
bahamaolide B (2)
δC, type δH, mult (J in Hz)
position
δC, type
(J in Hz)
δC, type
167.7, C
δH, mult (J in Hz)
position
δC, type
(J in Hz)
1
2
3
168.0, C
21
22
69.5, CH 4.79, m
69.1, CH 4.83, m
47.5, CH2 1.89, m
121.2, CH 6.12, d (15.0) 121.1, CH 6.14, d (15.0)
47.7, CH2 1.88, m
1.82, m
146.0, CH 7.63, dd (15.0, 146.1, CH 7.68, dd (15.0, 11.5)
11.5)
23
24
65.0, CH 5.07, m
48.3, CH2 1.84, m
1.76, m
65.4, CH 5.02, m
47.9, CH2 1.96, m
1.84, m
4
5
130.3, CH 6.46, m
130.5, CH 6.47, dd (14.5, 11.5)
142.2, CH 6.68, dd (14.5, 142.3, CH 6.73, dd (14.5, 11.5)
11.5)
25
26
65.0, CH 5.04, m
48.0, CH2 1.80, m
65.6, CH 5.01, m
47.1, CH2 2.01, m
1.94, m
6
7
132.3, CH 6.38, m
132.7, CH 6.39, m
138.8, CH 6.54, dd (14.5, 138.7, CH 6.65, dd (14.5, 11.5)
11.5)
27
28
69.7, CH 4.74, m
69.5, CH 4.72, m
45.9, CH2 2.07, m
8
132.9, CH 6.37, m
137.1, CH 6.47, m
131.9, CH 6.31, m
135.8, CH 6.32, m
133.6, CH 6.32, m
133.9, CH 6.18, m
42.4, CH2 2.75, m
2.60, m
133.5, CH 6.39, m
46.1, CH2 1.94, br d
(14.0)
9
137.0, CH 6.52, dd (14.5, 11,0)
134.0, CH 6.42, m
10
11
12
13
14
1.59, m
74.3, CH 4.41, m
44.2, CH2 1.86, m
1.83, m
1.81, m
72.9, CH 4.52, m
45.1, CH2 2.00, m
1.92, m
130.8, CH 6.89, dd (14.5, 11.0)
130.8, CH 6.29, dd (11.0, 11.0)
131.5, CH 5.76, m
29
30
37.1, CH2 3.14, m
31
32
73.3, CH 4.03, m
36.4, CH2 1.97, m
1.76, m
72.6, CH 4.06, m
36.9, CH2 2.04, m
1.79, m
2.68, ddd (13.5, 11.0,
8.5)
15
16
69.1, CH 4.52, m
44.4, CH2 2.13, m
2.10, m
69.9, CH 4.44, m
43.6, CH2 2.15, m
2.01, m
33
30.9, CH2 1.31, m
1.26, m
31.2, CH2 1.47, m
1.39, m
34
35
36
37
38
39
34.9, CH 1.75, m
35.0, CH 1.78, m
17
18
67.2, CH 4.70, m
47.6, CH2 2.25, m
2.01, m
67.3, CH 4.68, m
47.3, CH2 2.20, m
1.95, m
79.3, CH 5.08, br d (9.5) 79.6, CH 5.10, m
30.2, CH 1.88, m
30.3, CH 1.92, m
20.6, CH3 0.96, d (6.5)
19.0, CH3 0.78, d (6.5)
14.9, CH3 0.96, d (6.5)
20.4, CH3 0.96, d (6.5)
19.1, CH3 0.82, d (6.5)
14.7, CH3 0.98, d (7.0)
19
20
70.7, CH 4.52, m
45.8, CH2 1.93, m
70.9, CH 4.68, m
45.8, CH2 1.99, m
1.89, m
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Figure 1. Derivatization of 1 to yield tetraacetonides 3−7.
doublet methyl group (H3-39, δH 0.96) to C-34 and C-33 led
to the assigned C-33−C-34−C-39 connectivity. The H-34
methine proton (δH 1.75) showed a COSY correlation with
H-35 (δH 5.08), which connected the C-34 end of the fragment
to C-35. This was supported by the HMBC correlation from
H3-39 to C-35 (δC 79.3). The final two methyl groups (δC 20.6,
δH 0.96; δC 19.0, δH 0.78) were assigned as a dimethyl moiety by
their common COSY correlation with H-36 (δH 1.88) and
HMBC coupling to C-36 (δC 30.2). This isopropyl group was
connected to C-35 according to the HMBC correlation from
H3-37 and H3-38 to C-35. The hexaenone moiety accounts for
seven out of the eight degrees of unsaturation inherent in the
molecular formula, which suggests that bahamaolide A (1) must
possess a single ring. The connectivity of the ring was assigned
by long-range coupling from H-35 to C-1 and completed the
planar macrolide structure of bahamaolide A (1).
1,3-diol chain moiety, bahamaolide A (1) was derivatized
using 2,2-dimethoxypropane to yield five tetraacetonide products
(3−7) (Figure 1).
On the basis of the 1H and 13C, COSY, HMQC, and HMBC
NMR spectra of the acetonide products (3−7), compounds 3
and 7 were used to determine the relative polyol configuration.
The NMR spectra, including the ROESY spectra of 3 and 7,
were further analyzed to assign the 1H and 13C chemical shifts,
particularly of the nine acetonide-protected hydroxy groups.
1
The H, 13C, DEPT, and HMQC NMR spectra of tetra-
acetonide 3, which has a free alcohol (δH 2.77, d, J = 3.5 Hz) at
1
C-15, showed that eight H singlet methyl groups correlated
with several methyl carbons (δC 30.5−δH 1.51; δC 30.1−δH
1.39; δC 25.1−δH 1.35; δC 24.7−δH 1.29; δC 24.5−δH 1.34; δC
24.4−δH 1.32; δC 19.5−δH 1.35; δC 19.4−δH 1.28), indicating
the presence of two syn diols and two anti diols (Table 2). As
shown in Figure 2a, H-17 and H-19 displayed a common
ROESY correlation with the H3-41 methyl group (δH 1.28),
which is bound to C-41 (δC 19.4). This methyl group and
another methyl group (C-42: δH 1.39−δC 30.1) are coupled to
the acetal carbon at δC 98.8 in the HMBC spectrum, revealing
that this acetal structure is in a chair conformation and that the
hydroxy groups at C-17 and C-19 are in the syn orientation.
Because H-21 and H-23 correlate with H3-44 (δH 1.29, δC 24.7)
and H3-45 (δH 1.35, δC 25.1), respectively, which show HMBC
correlations to the same acetal carbon (δC 100.3), the relative
orientation of these two hydroxy groups must be anti. The
relative configuration of the two alcohols at C-25 and C-27
was established as anti on the basis of the chemical shifts
of dimethyl carbons C-47 and C-48 (δC 24.5 and 24.4,
respectively). Similarly, the diol moiety at C-29 and C-31 was
found to be syn (Figure 2a).
The carbinol proton signals are sufficiently separated to
completely assign their corresponding chemical shifts in the 1H
NMR spectrum in pyridine-d5, but the olefinic protons overlap
significantly in this solvent, which makes the assignment of the
alkene geometries challenging. This problem was overcome by
changing the NMR solvent to CD3OD. Although the 1H NMR
spectrum at 900 MHz in CD3OD displayed highly congested
signals for the carbinol protons, the alkene protons were well
separated without a second-order peak (see Figure S3 and
1
Table S1). On the basis of the large vicinal H−1H coupling
constants (J = 14.5−15.5 Hz), the alkene configurations were
assigned as 2E, 4E, 6E, 8E, 10E, and 12E.
The dimethyl carbon chemical shifts in the acetonide
derivatives of a 1,3-diol indicate their relative configurations.12
When a 1,3-diol is syn, its acetonide dimethyl carbons show
two distinctive chemical shifts around δC 19 and 30, but when it
is anti, the carbons display nearly identical signals near δC 25.
To apply this rule to the relative configurations of the repeating
In contrast, the other tetraacetonide (7) was revealed to have
a free hydroxy group (δH 3.52, br s) at C-31 on the basis of
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Table 2. NMR Spectroscopic Data for Tetraacetonides 3 and 7 in Toluene-d8 (800 MHz)
tetraacetonide 3
tetraacetonide 7
tetraacetonide 3
δH, mult
tetraacetonide 7
δH, mult
δH, mult
(J in Hz)
δH, mult
(J in Hz)
position
δC, type
δC, type
position
24
δC, type
(J in Hz)
δC, type
(J in Hz)
1
2
3
166.5, C
167.2, C
42.5, CH2
1.35, m
43.5, CH2
1.38, m
120.7, CH
145.0, CH
5.93, d (15.0)
121.1, CH
146.4, CH
5.83, d (15.0)
1.31, m
4.07, m
1.36, m
1.25, m
3.99, m
1.98, m
1.39, m
3.93, m
1.44, m
1.23, m
3.70, m
1.74, m
1.58, m
1.25, m
1.36, m
4.23, m
1.30, m
1.20, m
3.99, m
0.97, m
0.78, m
3.73, m
1.62, m
1.61, m
3.70, m
1.84, m
1.71, m
1.41, m
1.09, m
1.67, m
7.51, dd (15.0,
11.5)
7.40, dd (15.0,
10.5)
25
26
62.3, CH
38.9, CH2
62.2, CH
43.9, CH2
4
5
129.8, CH
140.9, CH
6.00, m
130.2, CH
141.8, CH
5.96, m
5.97, m
6.13, dd (14.5,
11.0)
27
28
65.8, CH
42.6, CH2
64.7, CH
39.3, CH2
6
7
135.6, CH
137.3, CH
6.09, m
132.5, CH
132.4, CH
5.87, m
6.13, m
6.05, dd (14.0,
11.0)
29
30
65.7, CH
36.5, CH2
71.7, CH
43.8, CH2
8
131.3, CH
133.4, CH
131.8, CH
6.00, m
6.02, m
135.5, CH
136.3, CH
135.3, CH
6.15, m
6.10, m
6.15, m
9
31
32
70.0, CH
34.0, CH2
73.5, CH
34.8, CH2
10
5.96, dd (14.5,
11.0)
11
134.6, CH
6.11, m
138.1, CH
6.05, dd
(14.5, 11.0)
33
29.6, CH2
31.5, CH2
12
13
14
132.4, CH
132.2, CH
40.2, CH2
6.01, m
5.77, m
133.1, CH
131.9, CH
40.5, CH2
5.89, m
6.09, m
34
35
34.6, CH
79.3, CH
1.64, m
34.3, CH
78.3, CH
5.02, dd
(10.0, 2.5)
5.13, dd
(10.0, 2.5)
2.34, ddd (13.5,
3.5, 3.5)
2.69, ddd (13.5,
3.5, 3.5)
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
29.9, CH
19.9, CH3
18.5, CH3
13.7, CH3
98.8, C
1.87, m
30.6, CH
20.7, CH3
18.8, CH3
15.3, CH3
100.2, C
1.87, m
2.33, ddd (13.5,
11.0, 2.5)
1.92, ddd (13.5,
11.0, 2.5)
1.02, d (6.5)
0.76, d (6.5)
0.98, d (7.0)
1.02. d (6.5)
0.75, d (6.5)
0.95, d (7.0)
15
16
68.5, CH
41.3, CH2
4.04, m
66.0, CH
4.02, m
1.61, ddd (14.0,
5.0, 4.0)
36.5, CH2
2.24, ddd (14.0,
5.0, 4.0)
1.57, m
4.15, m
1.24, m
1.18, m
4.00, m
1.97, m
1.38, m
3.84, m
1.37, m
1.24, m
4.02, m
1.63, m
4.03, m
1.84, m
1.80, m
3.75, m
1.17, m
0.90, m
3.91, m
1.37, m
19.4, CH3
30.1, CH3
100.3, C
1.28, s
1.39, s
27.7, CH3
25.3, CH3
98.6, C
1.46, s
1.40, s
17
18
67.2, CH
35.9, CH2
64.9, CH
40.9, CH2
24.7, CH3
25.1, CH3
100.2, C
1.29, s
1.35, s
19.8, CH3
31.0, CH3
100.9, C
1.26, s
1.40, s
19
20
62.9, CH
43.1, CH2
67.5, CH
35.4, CH2
24.5, CH3
24.4, CH3
98.4, C
1.34, s
1.32, s
25.0, CH3
25.1, CH3
99.1, C
1.45, s
1.41, s
21
22
62.5, CH
39.3, CH2
65.1, CH
40.8, CH2
30.5, CH3
19.5, CH3
1.51, s
1.35, s
30.7, CH3
20.3, CH3
1.39, s
1.34, s
23
62.4, CH
62.5, CH
4.13, m
1H, 13C, COSY, HMQC, and HMBC NMR spectra. As observed
C-50 and C-51 methyl groups (δC 30.7 and 20.3). As a result
of the spectroscopic analysis of tetraacetonides 3 and 7, the
relative configuration of the polyol moiety was assigned as
15R*, 17S*, 19S*, 21S*, 23S*, 25R*, 27S*, 29S*, and 31S*.
To determine the relative configuration of the stereogenic
centers at C-34 and C-35, hetero half-filtered TOCSY
(HETLOC) and ROESY experiments were performed in
CD3OD, which avoids the overlap of the methyl groups. The
HETLOC NMR spectroscopic data showed that the H-35−
C-39 3JCH value was 6.0 Hz, which is large enough to assign an
anti relationship in oxygenated systems.13 In the case where
H-35 is anti to C-39, only one rotamer can satisfy the observed
ROESY correlations around the stereogenic centers at C-34
and C-35 (Figure 3). Therefore, the relative configurations of
C-34 and C-35 were assigned as 34S* and 35S*.
1
with 3, the analysis of H, 13C, DEPT, and HMQC spectra
connected all of the singlet methyl groups to methyl carbons
(δC 31.0−δH 1.39; δC 30.7−δH 1.39; δC 27.7−δH 1.46; δC
25.3−δH 1.40; δC 25.1−δH 1.41; δC 25.0−δH 1.45; δC 20.3−δH
1.34; δC 19.8−δH 1.26), suggesting the presence of two syn and
two anti diol relationships (Table 2). Specifically, the C-41 and
C-42 dimethyl protons (δC 27.7−δH 1.46; 25.3−δH 1.40),
which displayed HMBC correlations with the C-40 acetal
carbon at δC 100.2, coupled through space with the carbinol
protons at δC 4.03 (H-17) and 4.02 (H-15), respectively.
This finding unambiguously established an anti relationship
(Figure 2b). The syn relationship between the two hydroxy
groups at C-19 and C-21 was inferred from the chemical shifts
of dimethyl carbons C-44 (δC 31.0) and C-45 (δC 19.8), which
were assigned to this acetal ring on the basis of ROESY and
HMBC spectra. In a process similar to that used to determine
the anti diol relationship at C-15 and C-17, the relative
configuration of the diol moiety at C-23 and C-25 was
established as anti, in agreement with the chemical shifts of
dimethyl carbons C-47 and C-48 (δC 25.0 and 25.1). The two
hydroxy groups at C-27 and C-29 were determined to be syn on
the basis of two distinct carbon peaks corresponding to the
The absolute configuration of the repeating 1,3-diol moiety
in bahamaolide A (1) was determined by the application of the
modified Mosher’s method.14 The tetraacetonide product (7)
was treated with either (R)- or (S)-MTPA-Cl to yield (S)-
1
and (R)-MTPA esters 8 and 9, respectively. The H chemical
shifts around the esterified position (C-31) of 8 and 9 were
1
adequately assigned using H, COSY, HMQC, and HMBC
NMR experiments. The analysis of ΔδS−R values allowed the
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Figure 2. Key ROESY correlations of tetraacetonide (a) 3 and (b) 7 for assigning the relative configurations of the repeating 1,3-diol moiety of 1.
(Figure 5). The 1H chemical shifts around C-31 and C-35 were
assigned by 1H and COSY NMR spectroscopic analysis.
Because the 31S configuration had already been assigned, we
could utilize the distribution of ΔδS−R values for a 1,n-diol
(n odd) to establish the absolute configuration at C-35.15 The
relative configurations of 34S* and 35S* were used to assign
the absolute configuration at C-34 as S, thereby completing the
stereochemical assignment of bahamaolide A (Figure 6).
Bahamaolide B (2) was obtained as a yellow powder, and
Figure 3. ROESY correlations determining the relative configurations
the molecular formula was determined to be C39H64O11
of C-34 and C-35 (in CD3OD).
by HRFABMS and 1H and 13C NMR spectroscopic data
(Table 1). The general features of the NMR, UV, and mass
absolute configuration at C-31 to be assigned as S (Figure 4).
spectra of 2 were very similar to those of 1, indicating a
The relative configurations assigned above were used to assign
structurally related isomer of 1. Detailed analysis of 1H and 13C,
the absolute configuration of the polyol structure as 15R, 17S,
COSY, HSQC, HMBC, and ROESY NMR spectra revealed
19S, 21S, 23S, 25R, 27S, 29S, and 31S.
that the only difference between 1 and 2 was in the alkene
The absolute configuration of the asymmetric center at C-35
geometry at C-12. The H-12−H-13 coupling constant was
was also determined through multistep chemical derivatiza-
tions, including an MTPA esterification. We attempted a
methanolysis of 1 to obtain a free hydroxy at C-35. However,
possibly due to steric hindrance, the yield of the reaction was
very low, and prolonging the reaction resulted in the formation
of the saponification product. Consequently, bahamaolide A
was treated instead with potassium hydroxide to produce linear
saponification product 10, followed by methylation with TMS-
diazomethane to afford linear methyl ester 11. In the final step
toward determining the absolute configuration at C-35, methyl
ester 11 was derivatized using either (R)- or (S)-MTPA-Cl to
yield bis-(S)- and (R)-MTPA esters 12 and 13, respectively
11.0 Hz, characteristic of the 12Z configuration supported by
ROESY analysis.
To determine the absolute configuration of 2, the alkenes in
1 and 2 were first reduced by hydrogenation. The two hydro-
genation products (12 and 13) derived from 1 and 2 displayed
1
virtually identical H NMR spectra (Figures S27 and S28) and
CD spectra (Figure S29), which enabled us to assume an
absolute configuration for the stereogenic centers in 2 identical
with those assigned in 1.
Because polyene polyol macrolides, such as the clinically
used amphotericin B, have been reported to possess antifungal
Figure 4. ΔδS−R values of 8 and 9 in toluene-d8.
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Figure 5. Multistep chemical modification of 7 to afford bis-(S)- and (R)-MTPA esters 12 and 13.
In molar concentration, bahamaolide A’s IC50 (10.8 μM) is
lower than that of the positive control compound, 3-nitro-
propionate (IC50 20.1 μM).
Bahamaolides A and B (1 and 2) were also tested in
antibacterial assays against Gram-positive (Staphylococcus aureus
ATCC 6538p, Bacillus subtilis ATCC 6633, and Micrococcus
luteus IFO 12708) and Gram-negative bacteria (Salmonella
typhimurium ATCC 14028, Proteus vulgaris ATCC 3851, and
Escherichia coli ATCC 35270). However, these compounds did
not show significant activity in these assays.
The cytotoxicities of 1 and 2 were measured against several
human cancer cell lines, including A549 (lung cancer), HCT116
(colon cancer), MDA-MB-231 (breast cancer), and SK-HEP-1
(liver cancer). Bahamaolides A and B did not display significant
inhibitory activity even at high concentrations (100 μg/mL).
Bahamaolides A and B (1 and 2), both bearing a hexaenone
and contiguous 1,3-diol groups, are structurally new 36-mem-
bered macrocyclic lactones belonging to the hexaene macrolide
class, apparently derived from the polyketide synthase type I
pathway. Hexaene macrolides are quite rare in nature, even
though pentaene and heptaene macrolides are common.17,18 To
the best of our knowledge, the only other well-characterized
hexaene-class macrolides are dermostatins A and B, isolated from
Streptomyces viridogriseus.12,19 Compared with dermostatin A
(Figure 7), the bahamaolides have one less alkene at C-32 and
Figure 6. ΔδS−R values of 12 and 13 in pyridine-d5.
properties, the biological activities of 1 and 2 were initially
evaluated by antifungal assays with various pathogenic fungal
strains, including Aspergillus fumigatus HIC 6094, Trichophyton
rubrum IFO 9185, T. mentagrophytes IFO4 0996, and Candida
albicans ATCC 10231. Bahamaolide A (1) showed moderate
antifungal activity against all tested fungal strains with an MIC
value of 12.5 μg/mL, but bahamaolide B (2) did not display
significant inhibitory activity against any tested strains. Because
bahamaolide A exhibited cell-based antifungal activity, we then
tested the inhibitory activities of the bahamaolides against
C. albicans isocitrate lyase (ICL), which is a recently highlighted
antifungal target enzyme utilizing multiple metabolic networks
and carbon sources and thus plays an important role in fungal
pathogenesis.16 Interestingly, bahamaolide A displayed signifi-
cant ICL inhibitory activity, with an IC50 of 7.65 μg/mL
(10.8 μM), whereas amphotericin B did not inhibit ICL, even at
high concentrations (100 μg/mL). Bahamaolide B showed weak
ICL inhibitory activity, with an IC50 of 90 μg/mL (127 μM).
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94 °C for 5 min, 30 cycles of denaturation at 94 °C for 30 s, annealing
at 55 °C for 30 s, extension at 72 °C for 1 min 30 s, and additional
extension at 72 °C for 10 min. The sequencing analysis data obtained
from COSMO Co., Ltd. showed that CNQ343 was most closely
related to the partial sequence of Streptomyces chartreusis (98%
identity), identifying this strain as Streptomyces sp. The strain was
deposited in The National Center for Biotechnology Information,
United States, with an accession number of AGDE01000038.
Cultivation and Extraction. To maximize the yield of
bahamaolides A and B (1 and 2), serial cultivation procedures were
employed. The bacterium on a YPM agar plate was used to inoculate
2 mL of seawater-based YPM medium (4 g of mannitol, 2 g of yeast
extract, 2 g of peptone in 1 L of artificial seawater) in a 50 mL flask.
After a three-day cultivation at 30 °C with shaking at 200 rpm, 10 mL
of the culture was used to inoculate 200 mL of YPM medium in a
500 mL flask with further incubation at 30 °C with shaking at 200 rpm
for three days. Finally, 30 mL of the culture in a 500 mL flask was used
to inoculate 2.8 L Fernbach flasks containing 1 L of YPM medium. In
total, 40 L of bacterial culture was incubated at 30 °C with shaking at
200 rpm. After seven days of cultivation, sterilized XAD-7 (20 g/L)
resin was added to obtain organic products, and the culture, with resin,
was shaken at 200 rpm for 2 h. The resin was filtered through
cheesecloth, washed with distilled water, and eluted with acetone. The
acetone extract was dried in vacuo to yield 2 g of extract.
Figure 7. Structure of dermostatin A.
one less methyl group at C-16. Furthermore, the absolute
configuration of the C-15 stereogenic center in bahamaolides A
and B is opposite that of dermostatin A. This inconsistency could
be due to modular variations in the biosynthetic pathways of
both dermostatins and bahamaolides, which may involve
different ketoreductases that possess opposing stereochemical
specificities.20
The structural determination of polyene polyol macrolides is
still challenging due to overlapping signals in the polyene and
polyol NMR regions; the complete NMR assignment of the
dermostatins has not yet been achieved.12,19 We overcame this
difficulty using two different NMR solvents (CD3OD, pyridine-d5)
and high-field (800 and 900 MHz) NMR spectrometers. In
addition, the absolute configuration of every stereogenic center
in bahamaolide A was unambiguously determined by multistep
chemical derivatization and subsequent spectroscopic analyses.
In biological studies, several classes of ICL inhibitors have
been reported previously, including terpenoids, indole alkaloids,
bromophenols, and hydroquinones.21 However, no compounds
belonging to the polyene polyol macrolide class, one of the
most popular antifungal agent groups, had been identified as
ICL inhibitors. Bahamaolide A is the first polyene polyol
macrolide ICL inhibitor. The discovery of bahamaolides A and
B from a marine actinomycete and the biological significance of
bahamaolide A as a C. albicans isocitrate lyase inhibitor provide
additional evidence to support the claim that marine environ-
ments are a prolific source of new bioactive chemical entities.
Isolation of Bahamaolides A and B (1 and 2). The extract was
fractionated by flash column chromatography on silica gel (100 g)
using a step elution with combinations of hexane, EtOAc, and MeOH
(hexane/EtOAc, 1:1; 100% EtOAc; EtOAc/MeOH, 10:1, 5:1, 2:1; and
100% methanol). Bahamaolides A and B eluted in the ethyl
acetate/methanol (5:1) fraction. This fraction was purified further
by semipreparative reversed-phase HPLC (Phenomenex Luna 5 μm
C8 (2) 250 × 10.0 mm column, flow rate 2 mL/min, UV 360 nm
detection) using isocratic 40% aqueous CH3CN to afford bahamaolide
A (36 mg) and bahamaolide B (6 mg) as yellow powders. Purified
bahamaolides A and B (1 and 2) eluted at 39 and 41 min in the HPLC
run, respectively.
Bahamaolide A (1): yellow powder, [α]25D −190 (c 0.01, MeOH);
UV (MeOH) λmax (log ε) 384 (5.48) nm; IR (neat) νmax 3370, 2925,
1698, 1559 cm−1; 1H and 13C NMR (pyridine-d5) see Table 1;
HRFABMS m/z 731.4341 [M + Na]+ (calcd for C39H64O11Na,
731.4346).
EXPERIMENTAL SECTION
■
Bahamaolide B (2): yellow powder, [α]25D −130 (c 0.03, MeOH);
UV (MeOH) λmax (log ε) 384 (5.45) nm; IR (neat) νmax 3354, 2927,
General Experimental Procedures. Optical rotations were
obtained using a Jasco P-1020 polarimeter with a 5 cm cell. UV
spectra were measured using a Perkin-Elmer Lambda 35 UV/vis
spectrometer. CD spectra were collected with a JACSO model J-715
CD spectrometer with a 0.5 cm cell. IR spectra were acquired using
a Thermo Nicolet iS10 IR spectrometer. Low-resolution LC/MS data
were obtained with an Agilent Technologies 6130 quadrupole mass
spectrometer coupled with an Agilent Technologies 1200 series HPLC
using a reversed-phase C18 column (Phenomenex Luna, 100 mm ×
4.60 mm). High-resolution fast-atom bombardment (HRFAB) mass
spectra were acquired with a Jeol JMS700 high-resolution mass spec-
trometer at the Korea Basic Science Institute in Daegu. NMR spectra
were obtained on Bruker 800 and 900 MHz NMR spectrometers
at the Korea Basic Science Institute in Ochang and Bruker 500 MHz
and Varian Inova 300 MHz NMR spectrometers at the College of
Pharmacy, Seoul National University.
1
1695, 1564 cm−1; For H and 13C NMR (pyridine-d5) see Table 1;
HRFABMS m/z 731.4344 [M + Na]+ (calcd for C39H64O11Na,
731.4346).
Preparation of Tetraacetonide Derivatives of 1 (3−7).
Pyridinium p-toluenesulfonate (5 mg) and distilled 2,2-dimethoxy-
propane (4 mL) were added to a solution of bahamaolide A (20 mg)
in anhydrous MeOH (200 μL) and anhydrous CH2Cl2 (2 mL) at
room temperature (rt). The mixture was stirred in the dark under
argon for 48 h, and then the reaction was quenched with saturated
NaHCO3 solution. The mixture was fractionated using a step gradient
elution system with H2O/MeOH (20%, 40%, 60%, 80%, and 100%
MeOH in H2O). The 100% MeOH fraction contained the five desired
tetraacetonide derivatives and was purified on a reversed-phase C18
column (Phenomenex Luna 5 μm C18 (2) 250 × 10.0 mm column,
flow rate 2 mL/min, UV 360 nm detection) using a solvent gradient
(50% to 100% aqueous MeOH) for 20 min and 100% MeOH after
20 min. The following five derivatives (3−7) were obtained:
tetraacetonide 3 (3.0 mg) at tR 32.0 min, tetraacetonide 4 (2.5 mg)
at tR 32.5 min, tetraacetonide 5 (0.5 mg) at tR 36.0 min, tetraacetonide
6 (5 mg) at tR 38.0 min, and tetraacetonide 7 (4 mg) at tR 41.0 min.
The molecular formulas of the five derivatives were each confirmed as
being C51H80O11 by ESIMS analysis ([M + Na]+ m/z at 891.5). For
1H and 13C NMR data of 3 and 7, see Table 2.
Bacterial Identification. The bacterium (strain CNQ343) was
isolated from a sediment sample collected at the depth of 27 m from
North Cat Cay in the Bahamas in 2000. The chromosomal DNA from
Streptomyces sp. CNQ343 was extracted using a G-spin genomic DNA
extraction kit, and the isolated DNA was amplified by PCR using
universal primers 27F and 1492R, corresponding to position 27 in the
forward direction and 1492 in the reverse direction of the E. coli 16S
rDNA sequence. For PCR, the reaction elements included 5 μL of the
template DNA, 5 μL of 10× Taq buffer, 1 μL of 10 mM mixed dATP,
dTTP, dGTP, and dCTP, 1 μL of sense-27F primer (10 pmol/μL),
1 μL of antisense-1492R primer (10 pmol/μL), and 5 U of Taq DNA
polymerase, which were prepared in a final volume of 50 μL. The PCR
thermal conditions were set as follows: preheating for denaturation at
MTPA Esterification of Tetraacetonide 7. Tetraacetonide 7
(4.0 mg) was separated into two equal portions and dried completely
under high vacuum. Freshly distilled anhydrous pyridine (1 mL) and a
catalytic amount of dimethylaminopyridine were added to each
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portion. The reaction mixtures were stirred at room temperature for
10 min. Either (R)- or (S)-α-methoxy-α-(trifluoromethyl)phenylacetyl
chloride (MTPA) (30 μL) was added to each vial. The reactions were
quenched with MeOH after 1 h. The products were purified on a
reversed-phase C18 column (Phenomenex Luna 5 μm C18 (2) 250 ×
10.0 mm column, flow rate 2 mL/min, UV 360 nm detection) using a
gradient system (40% to 100% aqueous MeOH) for 35 min and 100%
MeOH after 35 min. (S)-MTPA ester 8 (1.2 mg) and (R)-MTPA ester
9 (1.2 mg) eluted at 64.5 and 64.2 min, respectively. The molecular
formulas of the two derivatives (8 and 9) were each confirmed as
being C61H87F3O13 by ESIMS analysis ([M + Na]+ m/z at 1107.5).
The ΔδS−R values around the stereogenic centers of MTPA esters 8
and 9 were assigned by 1H NMR, COSY, HMQC, and HMBC NMR
experiments.
Saponification of Tetraacetonide 7. KOH (7%) was added to a
room-temperature solution of tetraacetonide 7 (7.0 mg) in anhydrous
MeOH (2 mL).22 The solution was stirred at reflux for 24 h and
quenched with saturated NH4Cl solution. The reaction mixture was
fractionated using a step gradient elution system with H2O/MeOH
(20%, 40%, 60%, 80%, and 100% MeOH in H2O). The 100 MeOH
fraction contained the desired product and was dried in vacuo. It was
purified on a reversed-phase C18 column (Phenomenex Luna 5 μm C18
(2) 250 × 10.0 mm column, flow rate 2 mL/min, UV 360 nm
detection) using a gradient system (50% to 100% aqueous MeOH) for
20 min and 100% MeOH after 20 min. Saponification product 10
(5.0 mg) eluted at 30.0 min. The molecular formula of 10 was
confirmed as being C51H82O12 by ESIMS analysis ([M + Na]+ m/z at
909.5).
Hydrogenation of 1 and 2. Bahamaolides A and B (1 and 2),
each in a separate vial, were suspended with a 10% Pd/C catalyst in
absolute EtOH (1.5 mL). The reaction mixtures were hydrogenated
under 1 atm of H2 at rt. Each product was filtered through a
polytetrafluoroethylene syringe filter and dried in vacuo. The molecular
formulas of the resultant products 14 and 15 (derived from
bahamaolides A and B, respectively) were each confirmed to be
C39H76O11 by ESIMS analysis ([M + H]+ m/z at 721.5).
Hydrogenation product 14: CD (2.1 × 104 M, MeOH), λmax (Δε)
1
314 (+0.10), 362 (+0.40), 397 (+0.28), 430 (+0.70) nm; H NMR
(CD3OD, 500 MHz) δ 5.33 (1H, m), 4.68 (1H, dd, J = 8.0, 3.5 Hz),
4.07 (2H, m), 4.01 (2H, m), 3.95 (1H, m), 3.89 (1H, m), 3.81 (1H,
m), 3.66 (1H, m), 2.34 (2H, m), 2.18 (2H, m), 2.02 (1H, m), 1.91
(1H, m), 1.77 (1H, m), 1.69−1.50 (13H, m), 1.50−1.39 (3H, m),
1.39−1.24 (25H, m), 0.90 (3H, d, J = 6.5 Hz), 0.90 (3H, d, J = 6.5
Hz), 0.86 (3H, d, J = 6.5 Hz).
Hydrogenation product 15: CD (2.1 × 104 M, MeOH), λmax (Δε)
1
314 (+0.10), 362 (+0.43), 397 (+0.32), 430 (+0.72) nm; H NMR
(CD3OD, 500 MHz) δ 5.33 (1H, m), 4.68 (1H, dd, J = 8.0, 3.5 Hz),
4.07 (2H, m), 4.01 (2H, m), 3.95 (1H, m), 3.89 (1H, m), 3.81 (1H,
m), 3.66 (1H, m), 2.34 (2H, m), 2.18 (2H, m), 2.02 (1H, m), 1.91
(1H, m), 1.77 (1H, m), 1.69−1.50 (13H, m), 1.50−1.39 (3H, m),
1.39−1.24 (25H, m), 0.90 (3H, d, J = 6.5 Hz), 0.90 (3H, d, J = 6.5 Hz),
0.86 (3H, d, J = 6.5 Hz).
Antifungal Activity Assay. YPD medium (1% yeast extract, 2%
peptone, 2% dextrose) was used for cultivation of Candida albicans.
After incubation for 48 h at 28 °C, cells were harvested by centrifuga-
tion and then washed twice with sterile distilled water. A. fumigatus,
T. rubrum, and T. mentagrophytes were plated on potato dextrose agar
and incubated for 2 weeks at 28 °C. Spores were washed three times
with sterile distilled water and resuspended in potato dextrose broth
(PDB, Difco) to prepare an initial inoculum size of 105 spores/mL.
We mixed 90 μL of PDB (104 cells/mL) with 10 μL of test compound
solutions (bahamaolides A and B) in 5% DMSO in each well of a
96-well plate. A culture of DMSO (0.5%) was used as a solvent
control, and a culture supplemented with amphotericin B was used as a
positive control. Culture plates were incubated at 28 °C for 48−72 h.
The MIC values were determined at the lowest concentration where
test compounds inhibited fungal growth.
ICL Activity Assay. ICL enzyme activity was determined using
Dixon and Kornberg’s method.23 The basic concept of this method is
to spectrophotometrically measure the formation of glyoxylate
phenylhydrazone at 324 nm in the presence of phenylhydrazine and
isocitrate. A 1 mL aliquot of the reaction mixture contained 20 mM
sodium phosphate buffer (pH 7.0), 1.27 mM threo-DL (+) isocitrate,
3.75 mM MgCl2, 4.1 mM phenylhydrazine, and 2.5 μg/mL purified
ICL.24 The reaction was performed at 37 °C for 30 min with and
without a prescribed concentration of the inhibitor dissolved in
DMSO (final concentration, 1%). The effect of the inhibitor on ICL
was calculated as a percentage, relative to solvent-treated control, and
the IC50 values were calculated using nonlinear regression analysis
(percent inhibition versus concentration). Protein concentration was
determined by the method of Bradford25 using Bio-Rad protein assay
kit (Bio-Rad) and bovine serum albumin as a standard. The ICL
inhibitor 3-nitropropionate was used as a positive control.
Methylation of Saponification Product 10. The purified
saponification product (10, 5.0 mg) was dried under high vacuum
and dissolved in anhydrous MeOH (2 mL). A solution of 2.0 M TMS-
diazomethane in tetrahydrofuran (3 mL) was added to the solution.
The solution was stirred for 2 h, and the reaction was quenched by
adding acetic acid. The mixture was fractionated using a step gradient
elution system with H2O/MeOH (20%, 40%, 60%, 80%, and 100%
MeOH in H2O). The 100% MeOH fraction contained methylated
product 11 and was purified on a reversed-phase C18 column
(Phenomenex Luna 5 μm C18 (2) 250 × 10.0 mm column, flow rate
2 mL/min, UV 360 nm detection) using a gradient system (50% to
100% aqueous MeOH) for 20 min and 100% MeOH after 20 min.
Methyl ester 11 (3.0 mg) eluted at 31.5 min in this HPLC run. The
molecular formula of 11 was confirmed as being C52H84O12 by ESIMS
analysis ([M + Na]+ m/z at 923.5).
1
Methyl ester 11: H NMR (DMSO-d6, 300 MHz) δ 7.29 (1H, dd,
J = 15.0, 10.5 Hz), 6.80 (1H, dd, J = 15.0, 10.5 Hz), 6.55 (1H, dd, J =
15.0, 10.0 Hz), 6.51−6.26 (6H, m), 6.19 (1H, dd, J = 15.0, 10.5 Hz),
5.96 (1H, d, J = 15.0 Hz), 5.77 (1H, m), 5.75 (1H, m), 5.23 (1H, m),
4.25 (1H, m), 4.07 (1H, m), 4.04−3.77 (8H, m), 3.65 (3H, s), 2.86
(1H, m), 2.26 (1H, m), 1.67−1.37 (8H, m), 1.33 (3H, s), 1.31 (3H, s),
1.30−1.11 (10H, m), 1.26 (3H, s), 1.25 (3H, s), 1.22 (3H, s), 1.22
(3H, s), 1.22 (3H, s), 1.21 (3H, s), 1.02 (1H, m), 0.94 (1H, m), 0.85
(3H, d, J = 6.5 Hz), 0.78 (3H, d, J = 7.0 Hz), 0.75 (3H, d, J = 7.0 Hz),
0.52 (1H, m), 0.43 (1H, m).
MTPA Esterification of Methyl Ester 11. Methyl ester 11 (3.0
mg) was separated into two equal portions and placed into separate
vials. The same procedure described for the MTPA esterification of
tetraacetonide 7 was performed for the esterification of 11 with either
(R)- or (S)-MTPA chloride and yielded bis-(S)- and bis-(R)-MTPA
esters 12 and 13, respectively. The reactions were monitored by LC/
MS and quenched with MeOH after 3 h. Bis-MTPA esters 12 and 13
were purified on a reversed-phase C18 column (Phenomenex Luna
5 μm C18 (2) 250 × 10.0 mm column, flow rate 2 mL/min, UV 360 nm
detection) using a gradient system (40% to 100% aqueous MeOH
(0.1% formic acid)) for 35 min and 100% MeOH (0.1% formic acid)
from 35 to 70 min. Bis-(S)-MTPA ester 12 (0.2 mg) and bis-(R)-
MTPA ester 13 (0.5 mg) eluted at 62 and 63 min, respectively.
The molecular formulas of 12 and 13 were each confirmed as being
C72H98F6O16 by ESIMS analysis ([M + Na]+ m/z at 1355.6). The
ΔδS−R values around the stereogenic centers of the two derivatives
ASSOCIATED CONTENT
■
S
* Supporting Information
NMR spectra of 1 and 2 and their derivatives are available free
of charge via the Internet at http://pubs.acs.org.
AUTHOR INFORMATION
■
Corresponding Author
*Tel: 82 2 880 2491. Fax: 82 2 762 8322. E-mail: dongchanoh@
snu.ac.kr.
Notes
1
were assigned by H and COSY NMR experiments.
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS
■
We specially thank Prof. W. Fenical and Dr. P. R. Jensen at the
Scripps Institution of Oceanography for providing the microbial
strain. We would like to thank the Basic Science Research
Institutes in Daegu and Ochang, Korea, for mass spectrometric
data and for high-field NMR experiments. We appreciate Mr.
M. Bae’s help in taking the photos of the bacterial strain. This
work was supported by the National Research Foundation of
Korea Grant funded by the Korean Government (MEST)
(NRF-2010-0010141 and NRF-2010-0020428). D.C.O. is a
Howard Hughes Medical Institute International Early Career
Scientist.
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