Biosynthetically intriguing chlorinated lipophilic metabolites from geographically distant tropical marine cyanobacteria

Article abstract of DOI:10.1021/jo300160e

Five new vinylchlorine-containing metabolites, the lipoamides janthielamide A and kimbeamides A-C and the ketide-extended pyranone kimbelactone A, have been isolated from collections of marine cyanobacteria made in Curacao and Papua New Guinea. Both janthielamide A and kimbeamide A exhibited moderate sodium channel blocking activity in murine Neuro-2a cells. Consistent with this activity, janthielamide A was also found to antagonize veratridine-induced sodium influx in murine cerebrocortical neurons. These lipoamides represent the newest additions to a relatively rare family of marine cyanobacterial-derived lipoamides and a new structural class of compounds exhibiting neuromodulatory activities from marine cyanobacteria.

Full text of DOI:10.1021/jo300160e

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pubs.acs.org/joc
Biosynthetically Intriguing Chlorinated Lipophilic Metabolites from
Geographically Distant Tropical Marine Cyanobacteria
Joshawna K. Nunnery,† Niclas Engene,† Tara Byrum,† Zhengyu Cao,‡ Sairam V. Jabba,‡ Alban R. Pereira,†
Teatulohi Matainaho,§ Thomas F. Murray,‡ and William H. Gerwick*,†
†Center for Marine Biotechnology and Biomedicine, Scripps Institution of Oceanography and Skaggs School of Pharmacy and
Pharmaceutical Sciences, University of California at San Diego, La Jolla, California 92093, United States
‡Department of Pharmacology, College of Medicine, Creighton University, Omaha, Nebraska 68178, United States
§Discipline of Pharmacology, School of Medicine and Health Sciences, University of Papua New Guinea, National Capital District,
Papua New Guinea
S
* Supporting Information
ABSTRACT: Five new vinylchlorine-containing metabolites,
the lipoamides janthielamide A and kimbeamides A−C and the
ketide-extended pyranone kimbelactone A, have been isolated
from collections of marine cyanobacteria made in Curaca̧ o and
Papua New Guinea. Both janthielamide A and kimbeamide A
exhibited moderate sodium channel blocking activity in murine
Neuro-2a cells. Consistent with this activity, janthielamide A
was also found to antagonize veratridine-induced sodium influx
in murine cerebrocortical neurons. These lipoamides represent
the newest additions to a relatively rare family of marine
cyanobacterial-derived lipoamides and a new structural class of
compounds exhibiting neuromodulatory activities from marine
cyanobacteria.
incorporated into marine cyanobacterial natural products,18,19 is
present in several linear lipophilic metabolites which feature
terminal or pendant vinyl chloride functionalities, including
(−)-(E)-1-chlorotridec-1-ene-6,8-diol,20 malyngamide A21 and
additional malyngamides,10 pitiamide A,22 jamaicamides,23 the
taveuniamides,24 and grenadamides B and C,25 among others.
These vinyl chloride-containing cyanobacterial compounds
exhibit a variety of biological activities, such as VGSC blocking
activity,23 cytotoxicity,10,23 toxicity,24 and insecticidal activity.25
In the present work, two independent collections of
cyanobacteria from Caribbean and Indo-Pacific locations
INTRODUCTION
■
Marine cyanobacteria are prolific producers of diverse bioactive
secondary metabolites with many exhibiting anticancer activity
largely owing to interference with tubulin (e.g., dolastatin 10,1
curacin A2) or actin (e.g., lyngbyastatin 1,3 hectochlorin,4
lyngbyabellin A,5 desmethoxymajusculamide C6) formation and
function. While freshwater and terrestrial cyanobacteria have
long been recognized as producers of potent neurotoxins, such
as anatoxin-a,7 β-methylaminoalanine,8 and saxitoxin,9 marine
cyanobacteria have only recently garnered attention for their
potential to produce neuromodulatory compounds.10−12
Metabolites exhibiting such activity, including voltage-gated
sodium channel (VGSC) activation or blocking effects and/or
intracellular calcium ion modulation, have potential therapeutic
utility in the treatment of spinal cord injury, chronic pain, CNS
disorders including stroke and epilepsy, and treatment of
cardiovascular, inflammatory, and neurodegenerative disor-
ders.13,14 With the discovery of potent VGSC activation for
antillatoxin A15 and nanomolar VGSC blocking activity for
kalkitoxin,15 two metabolites obtained from collections of
marine cyanobacteria, the capacity of these oceanic prokaryotes
to produce neuroactive compounds was realized.
1
yielded chromatographic fractions that had similar H NMR
profiles, molecular weights, and isotopic patterns. Additionally,
fractions from both extracts exhibited similar neuromodulatory
activity profiles in both a sodium channel blocking assay in
murine Neuro-2a cells and a calcium ion modulation assay in
murine neocortical neurons. At 20 μg/mL in Neuro-2a cells,
the Curaca̧ o-derived chromatographic fraction inhibited the
ouabain/veratradine-induced toxicity by 129%, while the Papua
New Guinea-derived fraction inhibited this toxicity by 86%.
Subsequently, bioassay-guided fractionation of these chromato-
graphic fractions employing the murine Neuro-2a sodium
channel blocking assay yielded a new lipoamide, janthielamide
Halogen atom incorporation is common in natural products
from the marine realm in part owing to the relatively high
concentrations of bromide, chloride, and iodide present in
seawater.16,17 Chloride, the most common seawater halide
Received: January 25, 2012
Published: April 9, 2012
© 2012 American Chemical Society
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Table 1. NMR Spectroscopic Data for Janthielamide A (1) in CDCl3 (1H at 500 MHz, 13C at 75 MHz)
a,
b
c
position
δC
δH mult (J in Hz)
COSY
HMBC
1
116.3 CH
130.1 CH
122.3 CH
144.1 CH
37.4 CH
5.88 d (7.0)
2
2, 3
1, 4
2
6.25 dd (10.4, 7.0)
1, 3
3
6.39 dd (15.4, 10.4)
5.76 dd (15.4, 7.6)
2.29 m
2, 4
1, 2, 5
4
3, 5
2, 3, 12
3, 4, 6, 7, 12
4, 5, 7, 8, 9, 12
5, 6, 8
5
4, 6, 12
5, 7
6
39.8 CH2
127.2 CH
137.5 CH
19.9 CH
2.03 m
7
5.33 dt (15.3, 6.7)
5.25 dd (15.3, 7.9)
2.14 m
6, 8, 11a/b
7, 9
8
5, 6, 7, 13
7, 8, 10, 13
8, 11, 13
8, 11, 13
10, 1′
9
8, 10a/b, 13
9, 10b, 11a
9, 10a, 11b
7, 10a
7, 10b
5
10a
10b
11a
11b
12
13
NH
1′
36.9 CH2
1.53 m
1.40 m
37.7 CH2
3.27 m
3.19 m
10, 1′
19.8 CH3
21.2 CH3
1.02 d (6.7)
0.98 d (6.7)
5.35
4, 5, 6
9
8, 10
2′, 4′
167.0 qC
118.8 CH
150.5 qC
35.2 CH3
27.2 CH3
2′
5.51 s
4′, 5′
12, 1′, 3′, 5′
3′
4′
2.14 s
2′
2′
1′, 2′, 3′, 4′, 5′
1′, 2′, 3′, 12
5′
1.82 s
a
b
c
Recorded at 75 MHz. Multiplicity deduced from HSQC. Recorded at 500 MHz.
Figure 1. Partial and full structures for janthielamide A (1), kimbeamide A (2), and kimbelactone A (5) with COSY and key HMBC correlations
shown.
A (1), from the Curaca
̧
o collection and the novel lipoamide
with CH2Cl2/MeOH (2:1) and then fractioned by silica gel
vacuum column chromatography to produce nine subfractions
(A−I). Bioassay-guided fractionation of the main bioactive
fraction (D) utilizing normal-phase column chromatography
led to the isolation of janthielamide A (1) as a yellow, optically
active amorphous solid [34.2 mg, 1.54%, [α]D 10.2 (c 0.60,
CHCl3)]. The LR-ESI-MS spectrum of 1 revealed a 3:1 ratio at
m/z 310/312 for the [M + H]+ pseudomolecular ion,
consistent with the presence of a single chlorine atom. HR-
ESI-TOFMS established the molecular formula as
C18H28ClNO, revealing that 1 contained five double bond
equivalents. The IR spectrum displayed absorptions character-
istic for NH or OH protons (3297 cm−1) and an amide
carbonyl group (1732 cm−1), while UV showed a maximum at
224 nm (log ε = 4.4) consistent with the presence of at least
kimbeamide A (2), two cis−trans isomers, kimbeamides B (3)
and C (4), and the polyketide kimbelactone A (5) from the
Papua New Guinea collections. Despite geographical distance
and morphological differences, both collections were found to
contain evolutionarily closely related cyanobacteria based on
phylogenetic analyses. Herein, we report the isolation, structure
elucidation and bioactivity of these metabolites, and a thorough
characterization of the putative marine cyanobacterial pro-
ducers.
RESULTS AND DISCUSSION
Collection and Isolation. A green-pigmented filamentous
cyanobacterial mat was collected by hand from Jan Thiel Bay in
■
Curaca̧ o. The cyanobacterial tissue was extracted repeatedly
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Scheme 1. (A) Fragmentation Strategy for Janthielamide A (1) and Kimbeamide A (2). (B) Preparation of Enantiomerically
Pure 2-Methyl-γ-aminobutyric Acid Standards for Determining the Absolute Configuration of Janthielamide A (1)
1
one conjugated diene or enone. The H NMR spectrum of 1
(Table 1) contained two doublet methyls (δH 0.98 and 1.02),
two singlet methyls (δH 1.82 and 2.14), a diastereotopic
deshielded methylene (δH 3.19 and 3.27), two upfield shifted
methines (δH 2.14 and 2.29), three resonances at 1−2 ppm
accounting for two methylenes, one NH proton at δH 5.35, and
seven olefinic resonances between 5 and 7 ppm. The 13C NMR
spectrum of 1 included two quaternary (δC 167.0 and 150.5),
nine methine (δC 144.1, 137.5, 130.1, 127.2, 122.3, 118.8,
116.3, 37.4 and 19.9), three methylene (δC 39.8, 37.7 and 36.9),
and four methyl carbons (δC 35.2, 27.2, 21.2 and 19.8),
accounting for all 18 carbon atoms in the molecular formula.
The quaternary and methine carbons were further assigned as
four double bonds and one carbonyl moiety according to their
chemical shifts.
The configurations of the double bonds were determined via
coupling constant data. The vicinal coupling constant for H1/
H2 was 7.0 Hz, a value consistent with a polarized Z double
bond.26 The C3−C4 and C7−C8 olefins were both assigned as
E on the basis of vicinal coupling constants of 15.4 and 15.3 Hz
for H3/H4 and H7/H8, respectively.
Regarding the absolute configuration of 1, we envisioned that
hydrolysis and ozonolysis followed by oxidative workup would
yield 2-methylsuccinic acid (6) and 2-methyl-γ-aminobutyric
acid (GABA, 7) as key degradation products containing the
stereocenters in question (C5 and C9, Scheme 1).
Enantiomerically pure (R)- and (S)-6 are commercially
available, and thus, were initially derivatized to afford methyl
or isopropyl esters; however, these proved inseparable on chiral
GCMS. Ultimately, esterification with (S)-2-(+)-octanol27
afforded dioctan-2-yl 2-methylsuccinate diastereomers (8)
that separated well on achiral GCMS. The products of
hydrolysis and ozonolysis of janthielamide A (1) were similarly
esterified with (S)-2-(+)-octanol. Comparison of the retention
times for (R)- and (S)-8 with janthielamide A derived 8 allowed
for assignment of the absolute configuration at C5 as R.
Unfortunately, in the case of 2-methyl-GABA, enantiomeri-
cally pure standards were not commercially available. This
analogue of the neurotransmitter GABA has previously been
synthesized;28 however, the starting material is a controlled
substance, and the synthetic route is complex. We envisioned a
more facile synthetic route whereby the desired standards could
be afforded via oxidation of the corresponding amino alcohol.
1
Two H spin systems were assembled by COSY (Figure 1);
fragment 1a featured an interesting vinylic gem-dimethyl
functionality, while fragment 1b contained a conjugated
diene, two methyl-substituted methine carbons, a third olefin
group, and two adjacent methylene groups. The lone carbonyl
carbon observed in the 13C NMR spectrum was assigned to
fragment 1a based on an HMBC correlation from H2′ to C1′.
Subsequently, spin systems 1a and 1b were connected through
an amide bond based on a correlation from the amide proton to
C2′ in the HMBC spectrum. Lastly, the chlorine atom was
attached to the C1 position (δC 116.3) on the basis of chemical
shift arguments and comparison with literature values for
similar systems.20,22,25
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Table 2. NMR Spectroscopic Data for Kimbeamide A (2) in C6D6 (1H at 600 MHz, 13C at 125 MHz)
a,
b
c
position
δC
δH (J in Hz)
COSY
HMBC
NOESY
1
164.1 qC
124.0 CH
140.5 CH
130.6 CH
137.5 CH
40.1 CH2
136.4 qC
114.2 CH
16.6 CH3
2
5.32 d (14.9)
3
1, 3, 4, 5
1, 2, 4, 5
2, 3, 6
4, NH
5
3
7.45 dd (14.9, 11.2)
5.90 dd (15.0, 11.2)
5.43 dt (15.0, 7.0)
2.31 d (7.0)
2, 4
3, 5
4, 6
5
4
2, 6, 9
3, 9
4
5
3, 6, 7
6
2, 3, 5, 7, 8, 9
7
8
5.53 bs
6, 7, 9
6
9
1.54 s
6, 7, 8
4, 5
2
NH
1′
2′
3′
4′a
4′b
5′a
5′b
6′
7′
8′
4.70 d (7.4)
5.0 m
1, 1′
42.4 CH
132.6 CH
130.3 CH
27.1 CH2
8′
1, 2′, 3′, 8′
3′, 4′, 8′
1′, 2′, 4′, 5′
3′, 5′, 6′
3′, 5′, 6′
3′, 4′, 6′, 7′
3′, 4′, 6′, 7′
5′, 7′
4′a/b
5.07 dd (10.5, 9.2)
5.17 dt (10.5, 7.5)
2.10 m
1′, 3′
2′, 4′a/b
3′, 4′b
3′, 4′a
4′b, 6′
4′b, 6′
5′a/b
5′a
1′
1.96 m
1′
30.8 CH2
1.76 m
1.71 m
133.3 CH
117.8 CH
21.7 CH3
5.69 m
5.67 d (13.4)
5′, 6′
4′a/b, 5′a
1.01 d (6.6)
1′
1′ 2′
NH
a
b
c
Recorded at 125 MHz. Multiplicity deduced from HSQC. Recorded at 600 MHz.
The N-protected standards were prepared as follows: the (S)-
N-acetyl-2-methyl-GABA standard (7a) was obtained from
commercially available (R)-(−)-3-bromo-2-methyl-1-propanol
(12) via an SN2 substitution of bromine with cyanide to form
the nitrile (13), followed by reduction with LiAlH4 to give (S)-
4-amino-2-methyl-1-butanol (14a). This was N- and O-
protected with acetyl groups (15a), and then the hydroxyl
group was deprotected (16a) and oxidized to the carboxylic
acid (7a) using Jones’ reagent. To obtain the (R)-N-acetyl-2-
methyl-GABA standard (7b), the amine and hydroxyl groups of
commercially available (R)-4-amino-2-methyl-1-butanol (14b)
were protected, and then the hydroxyl group was deprotected
and oxidized as described above for 7a. Both 7a and 7b were
then esterified with (S)-2-(+)-octanol. These diastereomers
(9a, 9b) were successfully resolved utilizing achiral reverse
phase LCMS, and comparison of their retention times with that
of the corresponding janthielamide A (1) derivative led us to
assign the absolute configuration at C9 as R, completing the
structure elucidation of compound 1.
δH 1.54, a doublet at δH 2.31 integrating for two protons,
annotated as a bis-allylic methylene, four resonances between 1
and 2.5 ppm accounting for two diastereotopic methylenes, an
NH proton at δH 4.70, nine resonances from 5 to 6 ppm
characteristic of olefinic protons, and a doublet of doublets at
δH 7.45 for the β proton in a conjugated dienone system.
Analysis of the 13C NMR and HSQC spectra revealed two
quaternary (δC 164.1 and 136.4), 10 methine (δC 140.5, 137.5,
133.3, 132.6, 130.6, 130.3, 124.0, 117.8, 114.2 and 42.4), three
methylene (δC 40.1, 30.8 and 27.1), and two methyl (δC 21.7
and 16.6) carbons, accounting for all 17 carbon atoms required
by the molecular formula. Two spin systems were assembled by
COSY (Figure 1); fragment 2a featured a conjugated diene
whereas fragment 2b featured two isolated alkene function-
alities separated by two methylene groups. The lone carbonyl
carbon was assigned to fragment 2a at position C1 based on
correlations in the HMBC from H2 to C1 and H3 to C1,
respectively. Spin systems 2a and 2b were connected through
an amide bond based on HMBC data including correlations
from NH to C1 and C1′ as well as from H1′ to C1. HMBC
correlations from H8 to C6, C7 and C9 allowed for placement
of a methyl substituted alkene on the distal side of fragment 2a.
Additionally, correlations from H7′ to C5′ and C6′, as well as
from H5′ and H6′ to C7′, allowed for assignment of the
remaining proton of the terminating alkene of fragment 2b. As
with janthielamide A (1), the relatively unique chemical shifts
exhibited by C8 (δC 114.2) and C7′ (δC 117.8) indicated that
chlorine atoms were appended to these vinylic carbons.
The configurations of the double bonds were determined on
the basis of vicinal coupling constants and NOE correlations.
The C2−C3, C4−C5, and C6′−C7′ olefins were assigned as E
according to coupling constants of 14.9, 15.0, and 13.4 Hz for
H2/H3, H4/H5, and H6′/H7′, respectively. The C7−C8 olefin,
in turn, was also assigned as E based on an NOE correlation
between H8 and H6. Lastly, the C2′-C3′ olefin was determined
to be Z given a vicinal coupling constant for H2′/H3′ of 10.5
Hz.
In parallel with the above study, two extracts from Papua
New Guinea collections of orange-colored cyanobacterial
consortia were combined (see the Experimental Section) and
subjected to a similar bioassay-guided protocol employing
normal-phase column chromatography and HPLC to yield
kimbeamide A (2) as an optically active pale yellow oil [1.6 mg,
0.10%, [α]D +44 (c 0.05, MeOH)]. The LR-ESI-MS of 2
revealed an isotopic pattern for the [M + H]+ pseudomolecular
ion cluster consistent with the presence of two chlorine atoms
[328/330/332 = 1:0.5:0.15]. The HR-ESI-TOFMS of 2 gave
an [M + H]+ at m/z 328.1234 (calcd for C17H24Cl2NO,
328.1229) in agreement with a molecular formula of
C17H23Cl2NO, therefore requiring 6 degrees of unsaturation.
The IR spectrum featured absorptions for NH or OH protons
(3290 cm−1) and an amide carbonyl group (1718 cm−1), while
UV showed a maximum at 250 nm (log ε = 4.0) suggesting the
presence of a conjugated dienone system.
1
The H NMR spectrum of compound 2 (Table 2) had
resonances for a methyl doublet at δH 1.01, a methyl singlet at
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Table 3. NMR spectroscopic data for kimbelactone A (5) in C6D6 (1H at 600 MHz, 13C at 125 MHz)
a
,
b
c
position
δC
δH (J in Hz)
COSY
HMBC
NOESY
17
1
165.8 qC
91.2 CH
171.7 qC
33.0 CH2
2
4.97 s
1, 3, 4, 5
3
4a
4b
5
1.84 dd (16.6, 12.1)
1.61 dd (16.8, 3.4)
4b, 5
1, 2, 3, 5, 6
2, 3, 5
6a/b
6b
4a, 5
74.2 CH
34.4 CH2
3.85 dddd (16.0, 11.9, 8.0, 4.0)
1.51 m
4a, 6a/b
5, 6b, 7
5, 6a, 7
6a/b, 8
7, 9
3, 6, 7
7
6a
6b
7
4, 5, 7, 8
4a
1.14 m
4, 5, 7, 8
4a
28.5 CH2
133.8 CH
127.2 CH
128.1 CH
133.9 CH
35.6 CH
2.12 m
5, 6, 8, 9
5, 9
6a/b
7, 12
8
8
5.46 dt (14.8, 7.4)
6.41 dd (14.8, 11.4)
6.01 t (10.9)
6, 7, 10
9
8, 10
6, 7, 8, 10
9, 11, 12
10
11
12
13
14
15
16
17
18
19
9, 11
5.16 t (10.1)
10, 12
11, 18
12, 14
13
9, 12, 13, 18
13, 14, 18
11, 12, 15, 16, 18
12, 15, 16, 18, 19
18
3.32 ddd (14.5, 11.6, 7.2)
5.51 dd (15.6, 6.6)
5.94 d (15.6)
9, 18
18, 19
12, 18
134.5 CH
128.5 CH
137.1 qC
118.5 CH
55.1 CH3
21.2 CH3
5.89 s
14, 15, 19
2, 3
2.91 s
2
1.02 d (6.7)
12
11, 12
11, 13, 14
13, 18
12.9 CH3
1.76 s
14, 15, 16
a
b
c
Recorded at 125 MHz. Multiplicity deduced from HSQC. Recorded at 600 MHz.
The absolute configuration of kimbeamide (2) at C1′ was
δH 3.32 and δH 3.85, as well as eight resonances between 5 and
6.5 ppm characteristic of olefinic protons. The 13C NMR and
HSQC spectra revealed three quaternary (δC 171.7, 165.8 and
137.1), 10 methine (δC 134.5, 133.9, 133.8, 128.5, 128.1, 127.2,
118.5, 91.2, 74.2 and 35.6), three methylene (δC 34.4, 33.0 and
28.5), one O-methyl (δC 55.1), and two aliphatic methyl (δC
21.2 and 12.9) carbons accounting for all 19 carbons required
by the molecular formula.
determined by cleaving the C2′−C3′ olefin and the amide bond
via ozonolysis followed by oxidative workup and hydrolysis to
produce alanine as a degradation product (Scheme 1). This
residue, as well as enantiomerically pure L- and D-alanine
standards, were esterified with 2-propanol and acylated with
trifluoroacetic anhydride to furnish volatile derivatives for chiral
GCMS analysis.29,30 By comparison of retention times of the
cleaved product and standards, metabolite 2 was found to
possess the S configuration at C1′, thus completing the
structure elucidation of kimbeamide A (2).
1
One large H NMR spin system (fragment 5b) featuring a
conjugated diene, a methyl-substituted methine, an oxygen-
substituted methine (based on a chemical shift of δC 74.2), and
an additional alkene moiety was assembled by COSY data
(Figure 1). A second fragment, 5a, contained a methyl-
substituted vinyl chloride functionality and was assembled on
the basis of HMBC correlations from H19 to C15 and H16 to
C19. The chemical shift of C16 at δC 118.5 allowed for
placement of the chlorine atom at this position. Fragment 5c
possessed an enone with O-Me substitution at the β carbon as
revealed by HMBC correlations from H2 to C1 and C3 and
H17 to C2 and C3. Fragment 5a was connected to the distal
side of fragment 5b based on HMBC correlations from H14 to
C15, C16, and C19 and from H16 to C14. Reciprocal HMBC
correlations from H19 to C14 and C16 further corroborated
this connection. Fragment 5c was connected to the proximal
side of fragment 5b forming a β-methoxy-substituted pyranone
ring based on HMBC correlations from H5 to C3 and from H4
to C1, C2 and C3, as well as an HMBC correlation from H2 to
C5.
As before, the geometries of the double bonds were
determined on the basis of vicinal coupling constants and
NOE correlations. The C8−C9 and C13−C14 olefins were
found to be E on the basis of large coupling constants (14.8 and
15.6 Hz, respectively), whereas the C10−C11 olefin was
determined to be Z (coupling constant for H10/H11 = ∼10.5
Hz). Lastly, the configuration of the C15−C16 olefin was
assigned as E on the basis of similarity of carbon chemical shifts
in comparison with the kimbeamide A olefin at C7−C8. The
Two cis−trans isomers of 2 were also isolated from the
orange cyanobacterial collections from Kimbe Bay. Kimbea-
mide B (3) differed from 2 in the geometry of the C4−C5
olefin (Z, vicinal coupling constant for H4/H5 = 10.9 Hz),
while kimbeamide C (4) differed from 3 in the geometry of the
C2′−C3′ olefin (E, vicinal coupling constant for H2′/H3′ = 15.5
Hz). Unfortunately, because compounds 3 and 4 were isolated
in very low yields and were unstable, we were unable to
complete the analysis of their stereochemistry. Nevertheless, we
hypothesize that C1′ of compounds 3 and 4 is S based on their
similarity to and co-occurrence with kimbeamide A (1).
Kimbelactone A (5) was isolated as an optically active pale
yellow oil [3.3 mg, 0.20%, [α]D −164.7 (c 0.17, CH2Cl2)] from
a slightly more polar chromatographic fraction of the Papua
New Guinea cyanobacterial collections. The LR-ESI-MS
spectrum of 5 revealed a 3:1 ratio at m/z 359/361 for the
[M + Na]+ pseudomolecular ion, consistent with one chlorine
atom. The HR-ESI-TOFMS established the molecular formula
as C19H25ClO3 revealing that 5 contained 7 double bond
equivalents. A UV maximum of 227 nm suggested the presence
of one or more conjugated diene or enone systems.
1
Examination of the H NMR spectrum of 5 (Table 3)
revealed a methyl doublet at δH 1.02, a methyl singlet at δH
1.76, a slightly upfield-shifted O-methyl at δH 2.91, five
resonances between 1 and 2.5 ppm accounting for six
methylene protons, two midfield shifted methine protons at
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Figure 2. Molecular-phylogenetic inference of the kimbeamide-producing strains PNG 07/14/07-6.1 (JQ388599) and PNG 07/18/07-3
(JQ396912) from Papua New Guinea as well the janthielamide-producing strain NAC 12/21/08-3 (JQ388601). The clade that includes the NP-
producing strains is highlighted with a red box. The closest related group is the genus Symploca (reference strain: PCC 8002T, GenBank acc. no.
AB039021) with a p-distance of 4.9% based on the SSU rRNA gene sequence divergence (highlighted with a gray box). The cladogram is based on
SSU (16S) rRNA gene sequences using the bayesian (MrBayes) and maximum likelihood (PhyML) methods, and the support values are indicated as
posterior probability at the nodes. The specimens are indicated as species, strain and access number in brackets. Specimens designated with (T)
represent type-strains obtained from Bergey’s Manual.36 The scale bar is indicated at 0.03 expected nucleotide substitutions per site, corrected using
the general time reversal (GTR) model.
absolute configurations at C5 and C12 of kimbelactone A (5)
remain unknown as the compound decomposed before such
studies could be undertaken.
Biological Activity. Biological evaluation of fractions
arising from extracts of these Curaca̧ o and Papua New Guinea
collections of tropical marine cyanobacteria revealed similar
neuromodulatory activity profiles. The most potent bioactive
fractions from both collections exhibited sodium channel
blocking activity in murine Neuro-2a cells at 20 μg/mL and
suppressed spontaneous calcium oscillations in murine neo-
cortical neurons at 5 μg/mL. The lipoamide janthielamide A
these latter lipoamides was prevented by the rapid decom-
position of 2-5 under either dry or organic solvent storage.
Decomposition due to autoxidation is common in polyunsa-
turated compounds such as fatty acids, and thus, it is not
surprising that these lipoamides readily decomposed despite
efforts to prevent autoxidation by cold storage in benzene.31,32
Characterization of the Cyanobacterial Collections.
The similar neuromodulatory activity profiles and related
natural product skeletal types from these two collections
suggested that the producing cyanobacteria might be closely
related, despite their geographic distance and macroscopic
morphological differences. On the basis of their growth
morphologies as well as their filament and cell morphologies,
the kimbeamide- and the janthielamide-producing strains best
fell under the taxonomic definition of the genus Symploca.
However, phylogenetic inferences of the 16S rRNA gene
sequenced from each collection revealed that these two natural
product-producing strains form a separate clade that is
evolutionarily distant (p-distance = ∼5% gene sequence
divergence) from the genus Symploca (reference strain: PCC
(1), obtained as the main bioactive constituent of the Curaca̧ o
collection, exhibited sodium channel blocking activity in murine
Neuro-2a cells with an IC50 value of 11.5 μM. Janthielamide A
was also found to antagonize veratridine-induced sodium influx
in cerebrocortical neurons with an IC50 value of 5.2 μM (95%
confidence interval = 2.67−10.1 μM; n = 6). From the Papua
New Guinea collections, pure kimbeamide A (2) was found to
exhibit sodium channel blocking activity in murine Neuro-2a
cells at 20 μg/mL; however, further biological evaluation of
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8002T, GenBank acc. no. AB039021). This clade of “tropical
marine Symploca” has yielded several important natural
products, including dolastatin 10 (e.g., strain VP377; GenBank
acc. no. AF306497),33 symplostatin 1 (e.g., strain VP642b;
GenBank acc. no. AY032933),33 and the hoiamides (strain
PNG06−65.1; GenBank acc. no. HM072001)34 (Figure 2).
Although strains of this group have been published as Symploca,
the evolutionary divergence together with the distinct
ecological habitats of these two groups suggests that this
clade of “tropical marine Symploca” needs to be described as a
new genus.
In addition, microscopic morphological and phylogenetic
analyses indicated that the cyanobacterial collections from
Kimbe Bay, Papua New Guinea, consisted of a consortium of
Moorea producens35 (formerly Lyngbya majuscula) and “tropical
marine Symploca”. However, because the biomass was primarily
composed of the finer “Symploca” filaments, and the fact that
the M. producens was only found in one of the kimbeamide-
producing specimens, it is likely that these are metabolites of
the “tropical marine Symploca” clade.
Despite macroscopic morphological differences, the putative
producing organisms were revealed by microscopic and
phylogenetic analyses to be closely related, belonging to an as
yet undescribed genus of tropical marine cyanobacteria that is
most similar to the extant genus Symploca. The similarities in
the biological activity profiles, secondary metabolite chemical
structure classes, and microscopic morphological characters all
suggested that these organisms may be evolutionarily closely
related. Ongoing investigation of the chemistry of morpholog-
ically similar and phylogenetically related organisms will
certainly continue to reveal intriguing chemotaxonomic
relationships in marine cyanobacteria.
Janthielamide A (1) and kimbeamides A−C (2−4),
presumably of a mixed PKS/NRPS biosynthetic origin, further
expand the halogenated lipoamide chemotype previously found
in marine cyanobacteria which includes pitiamide A22 and
grenadamides B and C.25 On the other hand, kimbelactone A
(5) is most closely structurally related to fuligoic acid, a
chlorinated polyene-pyrone from a myxomycete.37 It is
tempting to speculate on the biogenesis of the two different
types of vinyl chloride moieties that are featured in these new
lipoamides. With regard to kimbeamides A−C (2−4) and
kimbelactone A (5), which possess a methyl-substituted vinyl
chloride moiety, and grenadamides B and C,25 which feature a
propyl-substituted vinyl chloride, we hypothesize that this
functionality arises in a fashion similar to that of the pendant
vinyl chloride present in the jamaicamides A−C.23,38 This is
likely occurring through an hydroxymethylglutaryl CoA
synthase (HMGCS) like addition of acetate to a β-keto-
thioester intermediate, followed by chlorination, dehydration
and decarboxylation. As for the terminal vinyl chloride
functionality present in janthielamide A (1), kimbeamides A−
C (2−4), grenadamide C25 and pitiamide A,22 it is conceivable
that this moiety arises first via radical chlorination at the
penultimate carbon of a β-hydroxy ACP thioester as well as
sulfation of the β-hydroxy group. This functionally dense
system is then predicted to undergo a concerted thioester-
mediated hydrolysis, decarboxylation and sulfate elimination
similar to the mechanism shown for terminal alkene formation
in curacin A.39,40
CONCLUSION
■
The lipoamides janthielamide A and kimbeamides A−C, as well
as the pyranone kimbelactone A, were isolated from
independent collections of marine cyanobacteria from Curaca̧ o
and Papua New Guinea. All of these new cyanobacterial natural
products are characterized by multiple unsaturations and an
intriguing terminal vinyl chloride moiety. Both janthielamide A
(1) and kimbeamide A (2) exhibited modest sodium channel
blocking activity in murine Neuro-2a cells. Additionally,
compound 1 antagonized veratridine-induced sodium influx
in murine cerebrocortical neurons with an IC50 value of 5.2 μM.
Thus, these metabolites constitute new lead molecules in the
development of potential neuromodulatory agents.
Similar compounds in this structure class, such as pitiamide
A,22 grenadamide B, and grenadamide C,25 were initially
reported without characterization of their absolute config-
urations, likely due to the difficulties in determining the
chirality of remote methyl substituents and the small quantities
isolated. In the case of janthielamide A (1), hydrolysis and
ozonolysis afforded two fragments, 2-methylsuccinic acid and 2-
methyl-γ-aminobutyric acid, for which standards could be
obtained commercially or via synthesis. Following optimization,
our synthetic methodology to enantioselectively produce (R)-
and (S)-N-acetyl-2-methyl-GABA could provide a facile route
for generation of GABA analogues from the corresponding
amino alcohols for further biological evaluations of these
neurotransmitter-like compounds. Similarly, ozonolysis and
hydrolysis of kimbeamide A (2) yielded alanine, which was
readily analyzed and compared with amino acid standards.
Thus, the determination of absolute configuration of these
compounds was accomplished using a variety of approaches
including fragmentation and derivatization followed by chiral or
achiral GCMS or achiral LCMS analyses. The generation of
diasteromers via esterification of carboxylic acids with (S)-2-
(+)-octanol, as previously explored by our group,27 has proven
to be a robust method for determining the absolute
configurations of small chiral fragments of natural products.
Implementation of this methodology was key in the assignment
of the chiral centers at C5 and C9 in 1, which had proven
difficult to solve using standard chiral GCMS and LCMS
methodologies.
EXPERIMENTAL SECTION
■
General Experimental Procedures. NMR spectra were acquired
at 75 or 500 MHz with a 5 mm probe, at 600 MHz with a 1.7 mm
cryoprobe, or at 125 MHz with a direct observe 13C 5 mm cold probe.
NMR spectra were referenced to residual solvent H and 13C signals
1
(δH 7.26, δC 77.16 for CDCl3 and δH 7.16, δC 128.62 for C6D6). HPLC
separation was accomplished with UV detection using HPLC grade
solvent and a Phenomenex Maxil Silica column (10 μm, 10 × 500
mm). GCMS analysis was accomplished using a DSQ single
quadrupole mass spectrometer, and either a chiral Cyclocil B Alltech
capillary column (25 m × 0.25 mm) or an achiral DB-5 Agilent
capillary column (30 m × 0.25 mm). LCMS analysis was accomplished
using a Phenomenex Luna 5 μ C18(2) (250 × 4.60 mm).
Biological Material Collection and Identification. For the
Curaca
cyanobacterial mat was collected by hand at 1 m depth from a sandy
reef substrate in Jan Thiel Bay on the leeward side of Curacao (N
̧ o collection, approximately 3 L of a lime green filamentous
̧
12°07.634′, W 68°88.088′). Following collection, the cyanobacterium
was stored in 1:1 EtOH:seawater and stored at −20 °C; prior to
shipping, the samples were thawed and the supernatant was discarded.
For longer storage, EtOH was added and the samples were stored at
−20 °C until workup. A voucher specimen (NAC 12/21/08-3) is
maintained at the University of California San Diego, Scripps
Institution of Oceanography, La Jolla, CA.
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For the Papua New Guinea collections, approximately 1.5 L of
bright orange cyanobacterial puffballs (PNG 07/14/07-6) were
collected by hand at 20 m depth from at least eight sites located
throughout Kimbe Bay off the North coast of New Britain, Papua New
Guinea (S 5°26.192′, E 150°40.813′). Field notes identified the
puffballs as a consortium of Schizothrix sp. with a minor amount of
Lyngbya sp. present. A separate collection of 250 mL of the
cyanobacterium (PNG 07/18/07-3) was made at 25 m depth in
Kimbe Bay (S 5°19.588′, E 150°18.034′). The collected samples were
soaked in 1:1 EtOH:seawater in the field; the supernatant was
decanted and discarded before shipment. For longer storage, EtOH
was added, and the samples were stored at −20 °C until workup.
Voucher specimens [collection numbers PNG 07/14/07-6 (1.5 L
collection) and PNG 07/18/07-3 (250 mL collection)] are maintained
at the University of California San Diego, Scripps Institution of
Oceanography, La Jolla, CA.
no. 1869). A portion of the crude extract (2.22 g) was fractionated
using vacuum liquid chromatography (VLC) on silica gel (type H, 10−
40 μM) with a stepwise gradient of hexanes/EtOAc and EtOAc/
MeOH to give nine fractions (A−I). Fraction D, eluted with 40%
EtOAc in hexanes (307 mg), was subjected to two iterations of
normal-phase chromatography using Si Varian Bond Elut Sep-Paks.
During the first NP-SPE cartridge chromatography step, the initial
condition of 2.5% EtOAc in hexanes eluted 185 mg of material. A
second NP-SPE cartridge chromatography, employing a stepwise
gradient of 100% hexanes to 100% EtOAc, a fraction (D1F) eluting
with 5% EtOAc in hexanes gave compound 1 (34.2 mg, 1.5% of
extract). The surrounding subfractions D1E (22.4 mg) and D1G (6.2
mg) eluted with 2% and 10% EtOAc in hexanes, respectively, also
contained 1; however, these fractions contained additional impurities.
An EtOH-preserved consortium of “tropical marine Symploca” and
Moorea producens (PNG 07/14/07-6, 101.7 g dry weight) was
extracted with 2:1 CH2Cl2/MeOH six times to give 1.82 g of crude
extract (extract no. 1707), while an additional sample of “tropical
marine Symploca” (PNG 7/18/07-3, 5.97 g dry weight) was extracted
with 2:1 CH2Cl2/MeOH five times to give 0.151 g of crude extract
Morphological Characterization. Morphological characteriza-
tion was performed using an epifluorescent microscope (1000×).
Morphological comparison and putative taxonomic identification of
the cyanobacterial specimens were performed in accordance with
modern classification systems.
1
(extract no. 1708). On the basis of preliminary H NMR and LCMS
DNA Extraction, PCR Amplification, and Cloning. Algal
biomass (∼50 mg) was partly cleaned under a dissecting microscope.
The biomass was pretreated using TE (10 mM Tris; 0.1 M EDTA;
0.5% SDS; 20 μg/mL RNase)/lysozyme (1 mg/mL) at 37 °C for 30
min followed by incubation with proteinase K (0.5 mg/mL) at 50 °C
for 1 h. Genomic DNA was extracted using the Wizard Genomic DNA
Purification Kit (Promega) following the manufacturer’s specifications.
DNA concentration and purity were measured using a spectropho-
tometer. The 16S rRNA genes were PCR-amplified from isolated
DNA using the modified lineage-specific primers, CYA106F 5′-
CGGACGGGTGAGTAACGCGTGA-3′ and CSL1445R 5′-
GGTAACGACTTCGGGCGTG-3′. The PCR reaction volumes
were 25 μL containing 0.5 μL (∼50 ng) of DNA, 2.5 μL of 10 ×
PfuUltra IV reaction buffer, 0.5 μL (25 mM) of dNTP mix, 0.5 μL of
each primer (10 μM), 0.5 μL of PfuUltra IV fusion HS DNA
polymerase, and 20.5 μL dH2O. The PCR reactions were performed
using the following gradient: initial denaturation for 2 min at 95 °C, 25
cycles of amplification, followed by 20 s at 95 °C, 20 s at 55 °C, 1.5
min at 72 °C, and final elongation for 3 min at 72 °C. PCR products
were purified using a MinElute PCR Purification Kit (Qiagen) before
subcloning using the Zero Blunt TOPO PCR Cloning Kit (Invitrogen)
following the manufacturer’s specifications. Plasmid DNA was isolated
using the QIAprep Spin Miniprep Kit (Qiagen) and sequenced with
M13 primers. The 16S rRNA gene sequences are available in the
DDBJ/EMBL/GenBank databases under acc. No. JQ388601 (NAC
12/21/08-3), JQ396912 (PNG 07/18/07-3), JQ388599 (PNG 07/
14/07-6.1), and JQ388600 (PNG 07/14/07-6.2).
Phylogenetic Inference. The 16S rRNA gene sequences were
aligned with evolutionary informative cyanobacteria using the L-INS-I
algorithm in MAFFT 6.71741 and refined using the SSU secondary
structures model for Escherichia coli J0169542 without data exclusion.
The best-fitting nucleotide substitution model optimized by maximum
likelihood was selected using corrected Akaike/Bayesian Information
Criterion (AICC/BIC) in jModeltest 0.1.1.43 The evolutionary
histories of the cyanobacterial genes were inferred using Maximum
likelihood (ML) and Bayesian inference algorithms. The ML inference
was performed using GARLI 1.044 for the GTR+I+G model assuming
heterogeneous substitution rates and gamma substitution of variable
sites [proportion of invariable sites (pINV) = 0.494, shape parameter
(α) = 0.485, number of rate categories = 4] with 1000 bootstrap
replicates. Bayesian inference was conducted using MrBayes 3.145 with
four Metropolis-coupled MCMC chains (one cold and three heated)
run for 3000000 generations. The first 25% were discarded as burn-in,
and the following data set was sampled with a frequency of every 100
generations. The MCMC convergence was detected by AWTY.
Extraction and Isolation. Two of the three liters of EtOH-
analyses indicating that the extracts were virtually identical, the crude
extracts were combined and fractionated using vacuum liquid
chromatography (VLC) on silica gel (type H, 10−40 μM) with a
stepwise gradient of hexanes/EtOAc and EtOAc/MeOH to give nine
fractions (A−I). Fraction D, eluted with 40% EtOAc in hexanes (159
mg), was subjected to two iterations of normal-phase chromatography
using Si Varian Bond Elut Sep-Paks. In the first round of NP-SPE
column chromatography, the majority of the material (127 mg) eluted
under the loading condition of 30% EtOAc in hexanes. The second
round of NP-SPE cartridge chromatography utilizing a stepwise
gradient of 100% hexanes to 100% EtOAc was more successful.
Subfraction D1E (24.7 mg), eluted with 8% EtOAc in hexanes, was
then fractionated using normal-phase HPLC with an isocratic
condition of 18.5% EtOAc in hexanes at 6 mL/min on a Phenomenex
Maxil 10 silica (500 × 10.0 mm, 10 μ) column to yield 2 (peak
centered at 19.5 min, 1.6 mg, 0.10% of extract), 3 (peak centered at
15.5 min, 1.4 mg, 0.09% of extract) 4 (peak centered at 21 min, 0.8
mg, 0.05% of extract) and 5 (peak centered at 33 min, 0.7 mg, 0.04%
of extract. Subfraction D1F (14.4 mg), eluted with 11% EtOAc in
hexanes, was then fractionated using normal-phase HPLC with an
isocratic condition of 30% EtOAc in hexanes at 6 mL/min on a
Phenomenex Maxil 10 silica (500 × 10.0 mm, 10 μ) column to yield 5
(peak centered at 17.5 min, 2.6 mg, 0.16% of extract).
Janthielamide A (1): 34.2 mg (1.54%); yellow amorphous solid;
[α]24D +10.2 (c 0.60, CHCl3); UV (MeCN) λmax (log ε) 224 (4.4) nm;
IR νmax (film) 3297, 2961, 2925, 1732, 1667, 1540, 1452, 1376, 1259,
1
1180, 973, 851, 755, 716 cm−1; H and 13C NMR data, see Table 1;
ESIMS m/z 310 (100% rel abund) [M + H]+; HRESI-TOFMS m/z
[M + H]+ 310.1934 (calcd for C18H29ClNO, 310.1932).
Kimbeamide A (2): 1.6 mg (0.10%); pale yellow oil; [α]24D +44.0 (c
0.05, MeOH); UV (MeOH) λmax (log ε) 250 (4.0) nm; IR νmax (film)
3290, 2926, 2855, 1718, 1660, 1541, 1449, 1373, 1255, 1140, 1115,
1078, 1001, 936 cm−1; 1H and 13C NMR data, see Table 2; ESIMS m/
z 328 (28% rel abund) [M + H]+, 373 (100% rel abund), 655 (5% rel
abund) [2 M + H]+, 677 (14% rel abund) [2 M + Na]+, 700 (6% rel
abund) [2 M + 2 Na]+; HRESI-TOFMS m/z [M + H]+ 328.1234
(calcd for C17H24Cl2NO, 328.1229).
Kimbeamide B (3): 1.4 mg (0.09%) pale yellow oil [α]23D +33.4 (c
0.17, CH2Cl2); UV (obtained from UV trace during LCMS analysis)
1
λmax 246; H and 13C NMR data see the Supporting Information;
ESIMS m/z 328 (13% rel abund) [M + H]+, 373 (100% rel abund).
Kimbeamide C (4): 0.8 mg (0.05%); pale yellow oil; UV (obtained
1
from UV trace during LCMS analysis) λmax 258; H NMR data, see
Table S1 in Supporting Information; ESIMS m/z 328 (23% rel abund)
[M + H]+, 350 (11% rel abund) [M + Na]+, 373 (100% rel abund),
677 (47% rel abund) [2 M + Na]+.
preserved biomass (884.6 g, dry weight) of the Curaca
“tropical marine Symploca” (NAC 12/21/08-3) were extracted with
̧
o collection of
Kimbelactone A (5): 3.3 mg (0.20%); pale yellow oil; [α]23D −164.7
(c 0.17, CH2Cl2); UV (MeCN) λmax (log ε) 227 (4.2) nm; 1H and 13
C
2:1 CH2Cl2/MeOH eight times to give 2.77 g of crude extract (extract
NMR data, see Table 3; ESIMS m/z 301 (2% rel abund) [M − Cl]+,
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337 (<1% rel abund) [M + H]+, 359 (9% rel abund) [M + Na]+, 695
(100% rel abund) [2 M + Na]+; HRESI-TOFMS m/z [M + Na]+
359.1383 (calcd for C19H25ClNaO3, 359.1384).
dried over Na2SO4, and evaporated to dryness under N2 gas to give
(S)-4-hydroxy-3-methylbutanenitrile (13). This residue was resus-
pended in THF (3.5 mL) and added dropwise to a solution of LiAlH4
(0.271 g, 7.40 mmol, 5.25 equiv) in THF (3.7 mL) at 0 °C. The
reaction was allowed to warm to rt and stirred 19 h. The reaction was
cooled to 0 °C and quenched with H2O (3 mL) and a few drops of
10% NaOH. After the mixture was stirred 30 min, a small amount of
Na2SO3 was added, and the mixture was stirred for 10 min and then
filtered over qualitative paper #1. The filtrate was dried under N2 to
yield (S)-4-amino-2-methyl-1-butanol [14a, HR-ESI-TOFMS m/z
104.1072 [M + H]+ (calcd for C5H14NO, 104.1070)]. The N-acetyl
and (S)-2-octanol esterified (S)-2-methyl-GABA standard [9a, HR-
ESI-TOFMS m/z 294.2042 [M + Na]+ (calcd for C15H29NO3Na,
294.2040)] was obtained as described above for the derivatized (R)-2-
methyl-GABA standard from (S)-4-amino-2-methyl-1-butanol (14a).
The standards and natural product derived fragment were resuspended
in MeCN at a concentration of 1 mg/mL and subjected to analysis by
LCMS. At a 0.4 mL/min flow rate, the run conditions were as follows:
60% acidified water (0.1% formic acid in 99.9% H2O):40% MeCN
held for 15 min, then ramped to 35% acidified water:65% MeCN over
90 min, then ramped to 100% MeCN over 10 min and held for 15
min, then ramped back to 60% acidified water:40% MeCN over 10
min and held 10 min. Co-injection of the derivatized (R)-2-methyl-
GABA standard with the derivatized fragment from 1 gave one peak at
71.47 min, while coinjection of the derivatized (S)-2-methyl-GABA
standard with the derivatized fragment from 1 gave two peaks at 70.47
min (S) and 71.51 min (janthielamide A). Thus, the absolute
configuration at C9 was assigned as R.
Stereoanalysis of C-1′ in Kimbeamide A (2). Kimbeamide A (2)
was ozonized (0.2 mg of 2 in 400 μL CH2Cl2, 5 min, −78 °C) with
oxidative workup, dried under a stream of N2 gas, and hydrolyzed (1
mL of 6 N HCl, 110 °C, 16 h). The hydrolysate was dried under a
constant stream of N2 gas and treated with 150 μL of acetyl chloride
and 500 μL of IPA and heated at 110 °C for 1 h to generate the
isopropyl ester of alanine. The excess reagent was evaporated under a
constant stream of N2 gas, and the dried residue was derivatized with
trifluoroacetic anhydride (400 μL) in CH2Cl2 (400 μL) at 110 °C for
15 min. The N-(trifluoroaceto)isopropyl ester of the alanine residue
was solubilized in EtOAc. Both L- and D-alanine (0.5 mg each) were
derivatized using the same conditions as described for the ozonized
hydrolysate of 2 to furnish standards. All samples were analyzed by
chiral GCMS under identical conditions; the initial oven temperature
of 50 °C was held for 3 min, increased to 90 °C at a rate of 1 °C/min,
and held for 10 min. Co-injection of the derivatized L-alanine standard
with the derivatized fragment from 2 gave one peak at 18.82 min,
while coinjection of the D-alanine standard with the derivatized
fragment from 2 gave two peaks at 16.87 min (D-alanine) and 18.82
min (kimbeamide A).
Sodium Channel Activation and Blocking Assay. Neuro-2a
cells were added to 96-well plates at 3.0 × 105 cells/mL of RPMI 1640
medium with 10% FBS and 1% penicillin/streptomycin. The cells, in a
volume of 200 μL per well, were incubated (37 °C, 5% CO2)
overnight to allow recovery before treatment with compounds.
Compounds were dissolved in DMSO to a stock concentration of
10 mg/mL. Working solutions of the compounds were made in RPMI
1640 medium without FBS, with a volume of 10 μL added to each well
to give a final compound concentration of 20 μg/mL. An equal volume
of RPMI 1640 medium without FBS was added to 16 wells designated
as negative controls for each plate. Brevetoxin-2, at a final
concentration of 0.435 μg/mL (486 nM), was used as the positive
control for the sodium channel activating assay (e.g., 100% activation =
complete cell death) and tetrodotoxin, at a final concentration of
0.0435 μg/mL (136 nM), was the positive control for the blocking
assay (e.g., 100% blocking = no cell death), as previously tested in our
laboratory. Eight wells were used for each treatment. A mixture of
ouabain, veratridine and HCl/PBS was applied to the bottom half of
each plate to cause sodium overload to varying degrees for the
blocking and activating assays. For the blocking assay, a solution of 5
mM ouabain/0.35 mM veratridine/0.75 mM HCl/PBS was added,
and for the activating assay a solution of 5 mM ouabain/0.15 mM
Stereoanalysis of C5 in Janthielamide A (1). Janthielamide A (1)
was hydrolyzed (2.8 mg of 1 in 1 mL of 6 N HCl, 110 °C, 15 h), dried
under a stream of N2 gas, and then ozonized (1.5 mL CH2Cl2, 20 min,
−78 °C) with oxidative workup (5 drops of 30% H2O2). The product
was dried and then divided into two parts. The ozonized hydrosylate
(0.8 mg) was treated with 300 μL of (S)-(+)-2-octanol and 150 μL of
acetyl chloride and heated at 110 °C for 4 h to generate the dioctan-2-
yl 2-methylsuccinate diastereomer. The excess reagent was evaporated
under a constant stream of N2 gas, and the dried residue was
resuspended in CH2Cl2. Both (R)-(+)-methylsuccinic acid (2.1 mg)
and (S)-(−)-methylsuccinic acid (1.3 mg) were derivatized using the
same conditions as described for the reaction product of 1 to furnish
the standards. All samples were analyzed by achiral GCMS under
identical conditions; the initial oven temperature of 40 °C was held for
1 min, then the temperature was increased to 200 °C at a rate of 4.0
°C/min and held for 20 min. Co-injection of the derivatized (R)-
methyl succinic acid standard with the derivatized fragment from 1
gave one peak at 46.06 min, while coinjection of the (S)-methyl
succinic acid standard with the derivatized fragment from 1 gave two
peaks at 44.93 min (S) and 46.05 min (janthielamide A). Thus, the
absolute configuration at C5 was assigned as R.
Stereoanalysis of C9 in Janthielamide A (1). The desired fragment
for stereoanalysis of C9, N-acetyl-protected and (S)-2-octanol
esterified 2-methyl-γ-aminobutyric acid (GABA), was obtained in
addition to the derivatized 2-methyl-succinic acid under the conditions
employed for stereoanalysis of C5 from the initial aliquot of 1 {HR-
ESI-TOFMS m/z 294.2035 [M + Na]+ (calcd C15H29NO3Na,
294.2040)}. The derivatized (R)-2-methyl-GABA standard was
obtained via oxidation of N-acetyl protected (R)-4-amino-2-methyl-
1-butanol (16b). To (R)-4-amino-2-methyl-1-butanol (14b, 100 mg,
106.4 μL) was added 4-dimethylaminopyridine (11.8 mg, 0.097 mmol,
0.1 equiv), pyridine (3.34 mL) and acetic anhydride (3.34 mL). The
reaction mixture was stirred 15 h at rt and then dried under a stream of
N2 gas to afford the diprotected (amine and alcohol) fragment (15b).
The residue was resuspended in 2:1 1,4-dioxane/H2O (8.66 mL), and
LiOH·H2O (407 mg, 9.7 mmol, 10 equiv) was added. After 1.5 h,
solvents were removed under vacuum, and the residue was
resuspended in H2O (2 mL), acidified to pH 1 with 6 N HCl, and
extracted with EtOAc (3 × 3 mL). The combined organic layer was
dried over Na2SO4 and evaporated to dryness under a stream of
nitrogen to give the N-acetyl-protected fragment (16b). To N-acetyl-
protected (R)-4-amino-2-methyl-1-butanol (16b) in acetone (1 mL)
at 0 °C was added dropwise Jones’ reagent (400 μL, prepared by
dissolving 2.7 g CrO3 in 2.3 mL of concd H2SO4 and diluting to 10 mL
at 0 °C). After 1 h at 0 °C, 2-propanol (800 μL) was added, and the
mixture was stirred for 15 min. A small volume of H2O was added, and
the organic solvents were removed via a stream of nitrogen. A small
volume of acetonitrile was added, the solution was vortexed for 5 s, the
organic layer was removed, and the process was repeated two
additional times until the chromium residue appeared dry. The
combined acetonitrile layers were dried under N2 gas to give N-
protected 2-methyl-GABA (7b). (S)-2-Octanol (200 μL) and acetyl
chloride (100 μL) were added to the N-protected 2-methyl-GABA,
and the reaction was sealed and warmed to 110 °C. After 4 h, the
reaction was cooled and then dried under N2 gas, thus affording the N-
acetyl and (S)-2-octanol esterified (R)-2-methyl-GABA standard [9b,
HR-ESI-TOFMS m/z 294.2039 [M + Na]+ (calcd for C15H29NO3Na,
294.2040)].
(S)-4-Amino-2-methyl-1-butanol was obtained from (R)-(−)-3-
bromo-2-methyl-1-propanol (12) via an SN2 reaction in which the
bromine was replaced with a nitrile, followed by reduction with
LiAlH4. To sodium cyanide (0.076 g, 1.55 mmol, 1.1 equiv) were
added DMSO (2.24 mL) and (R)-(−)-3-bromo-2-methyl-1-propanol
(12, 148 μL, 1.41 mmol). After 24 h, the reaction was quenched with
H2O (2.2 mL) and extracted with EtOAc (3 × 3 mL). The aqueous
layer was acidified to pH 3 with 10% H2SO4 and extracted with EtOAc
(3 × 3 mL). The organic layers were combined, washed with brine,
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veratridine/1.75 mM HCl/PBS was added. A solution of PBS/5 mM
HCl without ouabain or veratridine was added to the top half of each
plate to give the general toxicity of the test compounds unrelated to
sodium channel regulation. The final volume for each well was 230 μL.
Plates were incubated for approximately 16 h before staining with
MTT for the activating assay. For the blocking assay, plates were
incubated for approximately 24 h before MTT staining. The plates
were read at 570 and 630 nm. IC50 determinations were the result of
three technical replicates at each concentration and thus represent a
single IC50 measurement. Under these conditions, tetrodotoxin gave
an IC50 = 9.7 nM. Concentration−response graphs were generated
using GraphPad Prism.
(11) Grindberg, R. V.; Shuman, C. F.; Sorrels, C. M.; Wingerd, J.;
Gerwick, W. H. In Modern Alkaloids: Structure, Isolation, Synthesis and
Biology; Fattorusso, E., Taglialatela-Scafati, O., Eds.; Wiley-VCH
Verlag GmbH: Weinheim, Germany, 2008; pp 139−170.
(12) Arao
́ ́
z, R.; Molgo, J.; Tandeau de Marsac, N. Toxicon 2010, 56,
813−828.
(13) Denac, H.; Mevissen, M.; Scholtysik, G. Naunyn-Schmiedeberg’s
Arch. Pharmacol. 2000, 362, 453−479.
(14) Eisenhut, M.; Wallace, H. Pflugers Arch.−Eur. J. Physiol. 2011,
461, 401−421.
(15) Berman, F. W.; Gerwick, W. H.; Murray, T. F. Toxicon 1999, 37,
1645−1648.
(16) Butler, A.; Walker, J. V. Chem. Rev. 1993, 93, 1937−1944.
(17) Gribble, G. W. Prog. Chem. Org. Nat. Prod. 2010, 91, 1−613.
(18) Neumann, C. S.; Fujimori, D. G.; Walsh, C. T. Chem. Biol. 2008,
15, 99−109.
(19) A MarinLit search of chlorine-containing compounds revealed
166 unique compounds, while a search for bromine-containing
compounds yielded 58 distinct compounds.
(20) Mynderse, J. S.; Moore, R. E. Phytochemistry 1978, 17, 1325−
1326.
ASSOCIATED CONTENT
■
S
* Supporting Information
1
Tabulated NMR spectroscopic data for 3 and tabulated H
1
NMR data for 2−4; H, 13C, gCOSY, gHMBC, and gHSQC
NMR spectra for 1−3 and 5; NOESY spectra for 2, 3, and 5;
1H spectrum for 4; GCMS and LCMS chromatograms from
stereochemical analyses; CD spectra for 5; macro- and
microscopic images of the specimens; experimental data for
neocortical neuron assays; dose−response curves for janthiela-
mide A neuromodulatory activity; graphical data for neocortical
neuron assays. This material is available free of charge via the
Internet at http://pubs.acs.org.
(21) Cardellina, J. H., II; Marner, F. J.; Moore, R. E. J. Am. Chem. Soc.
1979, 101, 240−242.
(22) Nagle, D. G.; Park, P. U.; Paul, V. J. Tetrahedron Lett. 1997, 38,
6969−6972.
(23) Edwards, D. J.; Marquez, B. L.; Nogle, L. M.; McPhail, K.;
Goeger, D. E.; Roberts, M. A.; Gerwick, W. H. Chem. Biol. 2004, 11,
817−833.
AUTHOR INFORMATION
Corresponding Author
*Tel: (858) 534-0578. Fax: (858) 534-0576. E-mail:
wgerwick@ucsd.edu.
■
(24) Williamson, R. T.; Singh, I. P.; Gerwick, W. H. Tetrahedron
2004, 60, 7025−7033.
(25) Jimen
́
ez, J. I.; Vansach, T.; Yoshida, W. Y.; Sakamoto, B.;
Porzgen, P.; Horgen, F. D. J. Nat. Prod. 2009, 72, 1573−1578.
̈
(26) Crews, P.; Rodríguez, J.; Jaspars, M. Organic Structure Analysis,
2nd ed.; Oxford University Press: New York, 2010.
(27) Pereira, A. R.; Cao, Z.; Engene, N.; Soria-Mercado, I. E.;
Murray, T. F.; Gerwick, W. H. Org. Lett. 2010, 12, 4490−4493.
(28) Duke, R. K.; Chebib, M.; Hibbs, D. E.; Mewett, K. N.; Johnston,
G. A. R. Tetrahedron: Asymmetry 2004, 15, 1745−1751.
(29) Smith, G. G.; Wonnacott, D. M. Anal. Biochem. 1980, 109, 414−
420.
ACKNOWLEDGMENTS
We thank the governments of Curaca
■
̧
o and Papua New Guinea
for permission to collect the cyanobacterial specimens. We
thank Y. Su (UCSD Chemistry and Biochemistry Mass
Spectrometry Facility) for HRMS data and J. Gerwick and R.
C. Coates for assistance with collection of source material. The
500 MHz NMR 13C XSens cold probe was supported by NSF
CHE-0741968. This work was supported by NIH TW006634,
NIH NS053398 and NIH DA007315.
(30) Mehner, C.; Muller, D.; Krick, A.; Kehraus, S.; Loser, R.;
̈
̈
Gutschow, M.; Maier, A.; Fiebig, H. H.; Brun, R.; Konig, G. M. Eur. J.
̈
̈
Org. Chem. 2008, 1732−1739.
(31) Gardner, H. W. Free Radical Biol. Med. 1989, 7, 65−86.
(32) Porter, N. A.; Caldwell, S. E.; Mills, K. A. Lipids 1995, 30, 277−
REFERENCES
■
(1) Pettit, G. R.; Kamano, Y.; Herald, C. L.; Tuinman, A. A.;
Boettner, F. E.; Kizu, H.; Schmidt, J. M.; Baczynskyj, L.; Tomer, K. B.;
Bontems, R. J. J. Am. Chem. Soc. 1987, 109, 6883−6885.
(2) Gerwick, W. H.; Proteau, P. J.; Nagle, D. G.; Hamel, E.; Blokhin,
A.; Slate, D. L. J. Org. Chem. 1994, 59, 1243−1245.
(3) Harrigan, G. G.; Yoshida, W. Y.; Moore, R. E.; Nagle, D. G.; Park,
P. U.; Biggs, J.; Paul, V. J.; Mooberry, S. L.; Corbett, T. H.; Valeriote,
F. A. J. Nat. Prod. 1998, 61, 1221−1225.
(4) Marquez, B. L.; Watts, K. S.; Yokochi, A.; Roberts, M. A.;
Verdier-Pinard, P.; Jimenez, J. I.; Hamel, E.; Scheuer, P. J.; Gerwick,
W. H. J. Nat. Prod. 2002, 65, 866−871.
(5) Luesch, H.; Yoshida, W. Y.; Moore, R. E.; Paul, V. J.; Mooberry,
S. L. J. Nat. Prod. 2000, 63, 611−615.
(6) Simmons, T. L.; Nogle, L. M.; Media, J.; Valeriote, F. A.;
Mooberry, S. L.; Gerwick, W. H. J. Nat. Prod. 2009, 72, 1011−1016.
(7) Devlin, J. P.; Edwards, O. E.; Gorham, P. R.; Hunter, N. R.; Pike,
R. K.; Stavric, B. Can. J. Chem. 1977, 55, 1367−1371.
(8) Cox, P. A.; Banack, S. A.; Murch, S. J.; Rasmussen, U.; Tien, G.;
Bidigare, R. R.; Metcalf, J. S.; Morrison, L. F.; Codd, G. A.; Bergman,
B. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 5074−5078.
290.
(33) Luesch, H.; Moore, R. E.; Paul, V. J.; Mooberry, S. L.; Corbett,
T. H. J. Nat. Prod. 2001, 64, 907−910.
(34) Choi, H.; Pereira, A. R.; Cao, Z.; Shuman, C. F.; Engene, N.;
Byrum, T.; Matainaho, T.; Murray, T. F.; Mangoni, A.; Gerwick, W. H.
J. Nat. Prod. 2010, 73, 1411−1421.
(35) Engene, N.; Rottacker, E. C.; Kast
̌
́
ovsky, J.; Byrum, T.; Choi, H.;
Ellisman, M. H.; Komarek, J.; Gerwick W. H. Int. J. Syst. Evol.
́
Microbiol. [Online early access]. DOI:10.1099/ijs.0.033761-0. Pub-
lished Online: July 1, 2011. http://ijs.sgmjournals.org (accessed Jan 9,
2012).
(36) Castenholz, R. W.; Rippka, R.; Herdman, M. In Bergey’s Manual
of Systematic Bacteriology, Boone, D. R., Castenholz, R. W., Eds.;
Springer: New York, 2001; Vol. 1, pp 473−599.
(37) Shintani, A.; Ohtsuki, T.; Yamamoto, Y.; Hakamatsuka, T.;
Kawahara, N.; Goda, Y.; Ishibashi, M. Tetrahedron Lett. 2009, 50,
3189−3190.
(38) Gu, L.; Wang, B.; Kulkarni, A.; Geders, T. W.; Grindberg, R. V.;
Gerwick, L.; Hakansson, K.; Wipf, P.; Smith, J. L.; Gerwick, W. H.;
̊
Sherman, D. H. Nature 2009, 459, 731−735.
(9) Alam, M.; Shimizu, Y.; Ikawa, M.; Sasner, J. J. J. Environ. Sci.
Health, Part A: Environ. Sci. Eng. 1978, 13, 493−499.
(10) Gerwick, W. H.; Tan, L. T.; Sitachitta, N. Alkaloids, Chem. Biol.
2001, 57, 75−184.
(39) Gu, L.; Wang, B.; Kulkarni, A.; Gehret, J. J.; Lloyd, K. R.;
Gerwick, L.; Gerwick, W. H.; Wipf, P.; Hakansson, K.; Smith, J. L.;
̊
Sherman, D. H. J. Am. Chem. Soc. 2009, 131, 16033−16035.
4207
dx.doi.org/10.1021/jo300160e | J. Org. Chem. 2012, 77, 4198−4208

The Journal of Organic Chemistry
Featured Article
(40) Gehret, J. J.; Gu, L.; Gerwick, W. H.; Wipf, P.; Sherman, D. H.;
Smith, J. L. J. Biol. Chem. 2011, 286, 14445−14454.
(41) Katoh, K.; Toh, H. Brief Bioinform. 2008, 9, 286−298.
(42) Cannone, J. J.; Subramanian, S.; Schnare, M. N.; Collett, J. R.;
D’Souza, L. M.; Du, Y.; Feng, B.; Lin, N.; Madabusi, L. V.; Muller, K.
̈
M.; Pande, N.; Schang, Z.; Yu, N.; Gutell, R. R. BMC Bioinform. 2002,
3, 1471−2105.
(43) Posada, D. Mol. Biol. Evol. 2008, 25, 1253−1256.
(44) Zwickl, D. J. Genetic Algorithm Approaches for the
Phylogenetic Analysis of Large Biological Sequence Datasets under
the Maximum Likelihood Criterion. Ph.D. Thesis, The University of
Texas at Austin, 2006.
(45) Ronquist, F.; Huelsenbeck, J. P. Bioinformatics (Oxford) 2003,
12, 1572−1574.
4208
dx.doi.org/10.1021/jo300160e | J. Org. Chem. 2012, 77, 4198−4208