Geographic variability and anti-staphylococcal activity of the chrysophaentins and their synthetic fragments

Article abstract of DOI:10.3390/md10051103

Drug-resistant Staphylococcus aureus is a continuing public health concern, both in the hospital and community settings. Antibacterial compounds that possess novel structural scaffolds and are effective against multiple S. aureus strains, including current drug-resistant ones, are needed. Previously, we have described the chrysophaentins, a family of bisdiarylbutene macrocycles from the chrysophyte alga Chrysophaeum taylori that inhibit the growth of S. aureus and methicillin-resistant S. aureus (MRSA). In this study we have analyzed the geographic variability of chrysophaentin production in C. taylori located at different sites on the island of St. John, U.S. Virgin Islands, and identified two new linear chrysophaentin analogs, E2 and E3. In addition, we have expanded the structure activity relationship through synthesis of fragments comprising conserved portions of the chrysophaentins, and determined the antimicrobial activity of natural chrysophaentins and their synthetic analogs against five diverse S. aureus strains. We find that the chrysophaentins show similar activity against all S. aureus strains, regardless of their drug sensitivity profiles. The synthetic chrysophaentin fragments indeed mimic the natural compounds in their spectrum of antibacterial activity, and therefore represent logical starting points for future medicinal chemistry studies of the natural products and their analogs.

Full text of DOI:10.3390/md10051103

Mar. Drugs 2012, 10, 1103-1125; doi:10.3390/md10051103
OPEN ACCESS
Marine Drugs
ISSN 1660-3397
www.mdpi.com/journal/marinedrugs
Article
Geographic Variability and Anti-Staphylococcal Activity of the
Chrysophaentins and Their Synthetic Fragments
Jessica L. Keffer 1, Jared T. Hammill 2, John R. Lloyd 1, Alberto Plaza 1,†, Peter Wipf 2 and
Carole A. Bewley 1,*
1
Laboratory of Bioorganic Chemistry, National Institute of Diabetes and Digestive and Kidney
Diseases, National Institutes of Health, Bethesda, MD 20892, USA;
E-Mails: kefferj@mail.nih.gov (J.L.K.); lloydj@niddk.nih.gov (J.R.L.);
a.plaza@mx.uni-saarland.de (A.P.)
2
Center for Chemical Methodologies and Library Development, University of Pittsburgh, Pittsburgh,
PA 15260, USA; E-Mails: jth25@pitt.edu (J.T.H.); pwipf@pitt.edu (P.W.)
†
Current address: Department of Pharmaceutical Biotechnology, Saarland University,
66123 Saarbrücken, Germany.
* Author to whom correspondence should be addressed; E-Mail: caroleb@mail.nih.gov;
Tel.: +1-301-594-5187; Fax: +1-301-401-4192.
Received: 15 April 2012; in revised form: 5 May 2012 / Accepted: 7 May 2012 /
Published: 22 May 2012
Abstract: Drug-resistant Staphylococcus aureus is a continuing public health concern,
both in the hospital and community settings. Antibacterial compounds that possess novel
structural scaffolds and are effective against multiple S. aureus strains, including current
drug-resistant ones, are needed. Previously, we have described the chrysophaentins, a
family of bisdiarylbutene macrocycles from the chrysophyte alga Chrysophaeum taylori
that inhibit the growth of S. aureus and methicillin-resistant S. aureus (MRSA). In this
study we have analyzed the geographic variability of chrysophaentin production in
C. taylori located at different sites on the island of St. John, U.S. Virgin Islands, and
identified two new linear chrysophaentin analogs, E2 and E3. In addition, we have
expanded the structure activity relationship through synthesis of fragments comprising
conserved portions of the chrysophaentins, and determined the antimicrobial activity of
natural chrysophaentins and their synthetic analogs against five diverse S. aureus strains.
We find that the chrysophaentins show similar activity against all S. aureus strains,
regardless of their drug sensitivity profiles. The synthetic chrysophaentin fragments indeed
mimic the natural compounds in their spectrum of antibacterial activity, and therefore

Mar. Drugs 2012, 10
represent logical starting points for future medicinal chemistry studies of the natural
1104
products and their analogs.
Keywords: chrysophaentins; Staphylococcus aureus; antibacterial
1. Introduction
Staphylococcus aureus has long been a public health concern, and is currently the leading cause of
skin, soft tissue, bloodstream, and lower respiratory tract bacterial infections [1]. Penicillin was
discovered in 1928 and first used in 1941, but by the late 1940’s was ineffective due to the rise in
penicillinase-producing S. aureus [2,3]. Methicillin was introduced into the clinic to combat
penicillin-resistant strains, but not long after its introduction, the first case of methicillin-resistant
S. aureus (MRSA) was described [4,5]. In the late 1980s and 1990s, the proportion of methicillin
resistant infections rose significantly compared to susceptible ones worldwide [6]. In the United States,
S. aureus is the leading cause of hospital-acquired infections [7]. Most of these strains are
methicillin-resistant, and in the early 2000’s, more patients in the U.S. succumbed to lethal MRSA
infections than to AIDS, tuberculosis, and hepatitis B combined [8].
The public health threat involving MRSA is not just limited to the spread of antibacterial resistance;
the movement of MRSA infections out of hospital settings and into the community is of great concern.
These community-associated MRSA (CA-MRSA) strains have become so prevalent in the United
States that almost everyone can be considered at risk for these infections [9,10]. CA-MRSA infections
present unique challenges to their treatment because they are distinct from hospital-acquired MRSA
(HA-MRSA) from an epidemiological, genetic, and clinical perspective [11,12]. CA-MRSA strains
generally have a faster growth rate than hospital acquired strains, but are less resistant to
antibacterials [13]. These bacteria typically contain a type IV staphylococcal cassette chromosome mec
type IV gene (SCC mec IV) and are capable of producing the extracellular cytotoxin Panton-Valentine
leukocidin (PVL) [14,15]. In contrast, nosocomial S. aureus typically carry a type I, II, or III
staphylococcal cassette chromosome mec. CA-MRSA is fully virulent, perhaps even more so than
HA-MRSA, and can produce other toxins that vary by geographic location [16,17]. In the United
States, there are two distinct community-associated backgrounds that comprise the majority of
CA-MRSA infections, namely USA300 and USA400. USA400 was first identified, but for reasons not
completely understood, USA300 has become the predominant infectious CA-MRSA clone [7,18,19],
currently accounting for 90–95% of CA-MRSA outpatient isolates in the US [13,20,21]. One reason
for its relative success and apparent fitness advantage may be its increased virulence. In a rat
pneumonia model, researchers have shown increased pathogenicity that correlates with increased
expression of virulence genes and increased expression of S. aureus regulatory systems [17,22].
The decline in antibacterial drug discovery programs worldwide has led to a concurrent decrease in
the number of new antibacterial compounds [23]. This decline is apparent from the decrease in the
number of antibacterial drugs (not including vaccines) approved for use from 1981–2010. During this
period, 56 drugs were approved from 1981–1990, 32 were approved in 1991–2000, and only 16 were
approved from 2001–2010 [24]. The increasing prevalence of resistance, even to drugs of last resort

Mar. Drugs 2012, 10
1105
such as vancomycin, indicates that more new compounds are still needed [25,26]. These trends,
combined with the knowledge that while incremental changes to lead compound scaffolds that result in
increased potency and spectrum generally do not provide a permanent solution to combating resistance,
suggest that new antibiotic compounds should differ structurally from earlier generation antibacterials.
Previously, we reported a suite of novel bisdiarlybutene macrocycles with antibacterial activity
isolated from the marine chrysophyte alga, Chrysophaeum taylori, from St. John, U.S. Virgin Islands [27].
We found that these compounds have inhibitory activity against three drug-resistant Gram-positive
organisms, and several drug-susceptible strains, and we developed a structure-activity relationship to
explain their potencies. We made a subsequent collection from three locations on St. John, and have
looked at geographic and temporal variability of chrysophaentin production. Furthermore, to address
supply limitations of the natural products, we are working towards a total synthesis of the
chrysophaentins, and have in hand two fragments from this effort. In this study, we have expanded the
antibacterial activity and structure activity relationship of the natural chrysophaentins and two
synthetic fragments to two additional S. aureus strains, S. aureus UAMS-1, a clinical osteomyelitis
isolate, and CA-MRSA USA300-LAC. It was important to expand our library of S. aureus strains to
include one with a more recent clinical, but still drug-susceptible, background, and a drug-resistant
strain comprising the predominant and successfully invasive CA-MRSA USA300. We investigated the
natural and synthetic chrysophaentin antibacterial activity to determine overall potency of growth
inhibition and the mode of inhibition (bacteriostatic or bactericidal).
2. Results and Discussion
2.1. Chrysophaeum taylori Collection and Chrysophaentin Identification
The original chrysophaentins came from one collection of C. taylori made in Round Bay, located on
the southern side of St. John, U.S. Virgin Islands, in July 2007. From this collection, we identified four
asymmetrically linked macrocycles, chrysophaentins A–D (1–4); one linear analog, chrysophaentin E (5);
and three symmetrically-linked macrocycles, chrysophaentins F–H (8–10) (Figure 1). The eight
chrysophaentins were remarkably similar to one another. All possess an ether linkage between C-1 and
C-14′. If present, the second ether linkage was located between either C-14 or C-16 and C-1′, and
halogen variability only occurred at C-2 and C-2′. Chrysophaentin H additionally differed in its
pentahalogenation, with replacement of the hydrogen at C-12′ with a bromine.
To obtain additional raw material, we returned to St. John in June 2009 and made three collections
at Round Bay, Long Bay, and Hawk’s Nest (Figure 2). Each sample was dried and sequentially
extracted with hexanes, chloroform, and methanol, and the chrysophaentins were found to be present
in the methanol extract. Each methanolic extract from the three new collections plus the original
collection were chromatographed by reverse-phase HPLC on a Jupiter Proteo C12 column with a
linear gradient of 50–80% methanol in 0.05% TFA-water in fifty minutes. This afforded a
chromatographic picture of the chrysophaentins present in each extract (Figure 3). The two collections
from Round Bay (Figure 3A,C), separated by two years, were remarkably similar, as was the collection
from Long Bay (Figure 3B). However, it was immediately obvious that the methanol extract from
Hawk’s Nest (Figure 3D) was different; in fact, it lacked the asymmetrically-linked chrysophaentins

Mar. Drugs 2012, 10
1106
A–D. Instead, the linear chrysophaentin E was the most abundant, as well as two newly identified
linear chrysophaentins, termed E2 and E3 (Figure 1, compounds 6 and 7). Algae collected from all
locations contained the symmetrical chrysophaentins, albeit at much lower abundance relative to
compounds A–E.
Figure 1. Structures of the natural chrysophaentins and synthetic fragments.
Figure 2. Locations of Chrysophaeum taylori collections on St. John, U.S. Virgin Islands.
Collections were made Long Bay in 2009 (green pin), the northern tip of Round Bay in
2007 and 2009 (yellow pin), and on the north side of the island at Hawk’s Nest in 2009
(red pin). Image was created using the Google Earth software package [28].

Mar. Drugs 2012, 10
Figure 3. Geographic distribution of chrysophaentins determined by LCMS. Analytical
1107
HPLC chromatograms of MeOH extracts of four collections of Chrysophaeum taylori
eluting with a linear gradient of 50–80% MeOH in aq. 0.05%TFA in 50 min. Panels
correspond to C. taylori extracts obtained from the following collections sites and dates:
(A) Round Bay, 2007; (B) Long Bay, 2009; (C) Round Bay, 2009; (D) Hawk’s Nest, 2009.
The expansions shown correspond to the chromatograms at 28–42 min, and UV detection
at 280 nm. Peaks are colored as shown in the legend, panel A.
2.2. Identification of New Acyclic Chrysophaentins
Previously we determined the structure of acyclic chrysophaentin E (5), which has a molecular
formula of C32H26Cl4O8 as determined by HR-MS and NMR. Chrysophaentins E2 (6) and E3 (7) were
obtained in insufficient quantities for full NMR characterization. However, through analysis of the
respective fragmentation patterns observed in the HR-ESI-MS data, we were able to assign structures
for each of these new linear chrysophaentins. HR-ESI-MS showed chrysophaentin E2 to possess a
molecular formula of C32H26BrCl3O8 (m/z 720.9786 [M − H]−, calcd for C32H25BrCl3O8, 720.9798),

Mar. Drugs 2012, 10
1108
and chrysophaentin E3 to possess a molecular formula of C32H26Br2Cl2O8 (m/z 764.9282 [M − H]−,
calcd for C32H25Br2Cl2O8, 764.9293), indicating that a bromine replaced a chlorine in E2 (6), and two
bromines replaced two chlorines in E3 (7), when compared to chrysophaentin E (5). Since positions
C-2 and C-2′ were the only sites of halogen variability in the natural chrysophaentins, we reasoned that
the dibrominated compound 7 likely contained a bromine at both of these carbons. On the other hand,
chrysophaentin E2 must contain bromine at only one of these sites, with chlorine at the other. To
identify the respective locations of chlorine and bromine in 6, we compared the mass fragmentation
patterns of the three linear analogs (Figure 4). As seen in Figure 4A, the fragmentation pattern for
chrysophaentin E showed the successive loss of four chlorine atoms, giving fragment ions with clear
isotopic signatures at m/z 641 [M − H − HCl]−, 605 [M − H − 2HCl]−, 569 [M − H − 3HCl]−, and
533 [M − H − 4HCl]−. The fragmentation pattern and peak intensities for chrysophaentin E2 (6) was
very similar to 5 (Figure 4A and 4B), where successive loss of three chlorines and one bromine gave
rise to fragment ions at m/z 685 [M − H − HCl]−, 649 [M − H − 2HCl]−, 613 [M − H − 3HCl]−, and
533 [M − H − HBr − 3HCl]−. In contrast, although loss of two bromines and two chlorines
was apparent in the mass spectrum for chrysophaentin E3, the major ion for 7 appeared at
m/z 395 [M − H − HBr − HCl − C16H14O3]−, which corresponds to the fragment generated from
cleavage at the ether bond (Figure 4C). Although MS data on model halogenated biaryl ethers was not
available, our data suggest that the presence of bromine ortho to the ether bond is destabilizing as
compared to chlorine in the same position, and facilitates cleavage at the ether bond in compound 7.
Thus, the nearly identical mass spectral patterns observed for 5 and 6, and the unique fragmentation at
the ether bond in the spectrum for 7, allow assignments of the structures of the new linear
chrysophaentins E2 and E3, as shown in Figures 1 and 4.
Figure 4. HR-MS fragmentation of chrysophaentins E, E2, and E3. Fragmentation of
(A) chrysophaentin E showing individual losses of four chlorine atoms; (B) chrysophaentin
E2 showing loss of three chlorine atoms and one bromine atom; (C) the major ion in the
spectrum for chrysophaentin E3 corresponds to fragmentation at the ether bond, with
weaker ions corresponding to loss of two chlorine and two bromine atoms.

Mar. Drugs 2012, 10
1109
Figure 4. Cont.
2.3. Chrysophaentin Fragment Synthesis
While the extracts from Long Bay and Round Bay afforded more of the asymmetrical
chrysophaentins, it became apparent that a synthetic approach to chrysophaentin production was
necessary in order to fully examine the biological activity of the structural variants. Accordingly, we
devised a convergent synthetic strategy to allow for a total synthesis of the chrysophaentins as well as
the investigation of structure-activity relationships. Fragment 12, which, in principle, could be used
twice for the assembly of the diether 1, could be constructed by a key Negishi cross-coupling between
vinyl iodide 13 and benzylic zinc species 14 (Scheme 1). A chloroiodination of alkyne 15 and a
metalation of a halide derived from aldehyde 16 were envisioned to provide access to 13 and 14,
respectively. In fact, while some modification of protective groups was required, this strategy could be
readily realized.

Mar. Drugs 2012, 10
Scheme 1. Retrosynthetic approach toward chrysophaentin A (1).
1110
The commercially available benzoic acid 17 was reduced with lithium aluminumhydride (LAH)
to the benzylalcohol 18 (Scheme 2). Bromination [29] and copper-catalyzed substitution with
TMS-acetylene magnesium bromide [30] provided the alkyne 20 in 89% overall yield from 17. After
desilylation of 20 with tetrabutylammonium fluoride (TBAF), dimethyl resorcinol 15 was subjected to
iodochlorination [31,32] in an attempt to generate 13. However, the desired alkyne addition product
was formed to a minor extent, and mainly overiodination products were observed. Therefore, we
demethylated 15 with boron tribromide and installed the deactivating pivaloyl esters in 82% yield.
Iodochlorination of 21 now proceeded in excellent yield to give a 2:1 mixture of alkene isomers 22 and
23. While this ratio could be further improved in favor of the (E)-alkene 22 using solvents other than
dichloromethane, yields were considerably lower. Furthermore, we were unable to separate the mixture
at this stage, and the clean conversion in dichloromethane allowed us to carry it forward to the
coupling step.
Scheme 2. Preparation of (E)-iodoalkene segment (22).
Formylation [33,34] of dimethyl hydroquinone 24 provided aldehyde 16, which was reduced to the
alcohol 25 and brominated to give 26 in high yield (Scheme 3). Four equivalents of this bromide were
then converted to the corresponding zinc reagent. Removal of excess zinc was necessary prior to

Mar. Drugs 2012, 10
1111
combination with 22 and 23 and subsequent microwave heating in the presence of palladium acetate.
These cross-coupling conditions [35] generated satisfactory yields of the chloroalkenes 11 and 27 in a
2:1 ratio after methanolysis of the pivaloate esters with cesium carbonate and methanol. An analytically
pure sample of 11 was obtained by separation with supercritical fluid chromatography (SFC). Boron
tribromide was used to cleave the two methyl ethers on 11 to give the tetraphenol 12, which was
obtained as the pure (E)-isomer and further purified by preparative supercritical fluid chromatography
(SFC) before submission to biological testing. In a control experiment, the use of a 2:1 mixture of 11
and 27 in the deprotection reaction led to a 2:1 mixture of 12 and its (Z)-diastereomer, as expected.
The assignments of the E/Z-configurations of 12 and its (Z)-diasteromer as well as 22 and 23 were
mainly based on the available literature data [31,32].
Scheme 3. Preparation of benzyl bromide segment (26) and Negishi coupling to give
tetraphenol 12.
2.4. Antibacterial Evaluation
With the new fragments in hand, we tested whether their potencies were similar to the parent
chrysophaentins. A microbroth dilution assay was used to evaluate the antibacterial activity of these
compounds, and the minimum inhibitory concentration that led to a 50% decrease in growth (MIC50)
was determined using a model curve fit to the data. The MIC90 was determined as the lowest
concentration where there was no visible growth, and the minimum bactericidal concentration (MBC)
was determined as the lowest concentration where a plated aliquot of treated bacteria led to a three
logarithmic decrease in colony forming units (CFU) per milliliter relative to the starting inoculum.
All compounds were evaluated against three different S. aureus strains from ATCC and two diverse
clinical isolates. The strains used in this study included: S. aureus ATCC 25923, a clinical isolate from
Seattle, 1945; methicillin-resistant S. aureus ATCC BAA-41, a hospital-acquired strain isolated in
New York City in 1994; multidrug-resistant S. aureus (MDRSA) ATCC BAA-44, a hospital-acquired
strain isolated in Lisbon, Portugal, with resistance towards ampicillin, methicillin, oxacillin, penicillin,

Mar. Drugs 2012, 10
1112
erythromycin, gentamicin, tetracycline, azithromycin, amikacin, clindamycin, cephalothin, ceftriaxone,
imipenem, lincomycin, streptomycin, perfloxacin, rifampin, and neomycin [6,36]; S. aureus UAMS-1,
a clinical osteomyelitis isolate [37]; and community-associated methicillin-resistant S. aureus,
USA300-LAC [38,39].
2.4.1. Minimum Bactericidal Concentrations for Select Natural Chrysophaentins
The MIC50 for select chrysophaentins was reported previously [27]. Those qualitative antibacterial
data have been expanded here to include descriptions of the MIC90 and the MBC for the
chrysophaentins against S. aureus 25923, MRSA BAA-41, and MDRSA BAA-44. These values are
summarized in Table 1. We found chrysophaentin A to inhibit the growth of S. aureus 25923, MRSA
BAA-41 and MDRSA BAA-44 with MIC50 values on the order of 1–4 μM, and MIC90 values ranging
from 4–9 µM against all three strains. Chrysophaentin A was bacteriostatic at its MIC90, becoming
bactericidal only at concentrations four to eight times higher than the MIC90. We have found
chrysophaentin A to be the most potent compound in all of our antibacterial assays, yet chrysophaentins
F and H which bear a symmetrical bisbibenzyl ether skeleton also inhibit S. aureus 25923 and MRSA
BAA-41 with single digit micromolar MIC50s. While their MIC90s were higher than those of
chrysophaentin A (17–19 µM), the MBC for chrysophaentin F was still four-fold higher than the
MIC90. A significant reduction in potency was observed for the acyclic natural product chrysophaentin
E (5), which was also bacteriostatic at its MIC90. Chrysophaentins D (4) and G (9), both of which
contain a bromine atom on the D ring, were the least active compounds, with an 8- to13-fold increase
in MIC50s values, and an 8-fold increase in MIC90s.
Table 1. Antibacterial summary of select chrysophaentins and synthetic fragments
against five diverse strains of Staphylococcus aureus (all values are in µM; n.d. means
not determined).
CA-MRSA
S. aureus 25923
MRSA BAA-41
MDRSA BAA-44 S. aureus UAMS-1
USA300-LAC
MIC50 MIC90 MBC MIC50 MIC90 MBC MIC50 MIC90 MIC50 MIC90 MBC MIC50 MIC90 MBC
1
4
5
8
9
2.7 ± 0.9 9.2
37 ± 16 65
37 2.3 ± 1.0 4.6
n.d. 26 ± 8.5 65
74 14 ± 4.1 37
74 6.3 ± 1.9 19
n.d. 17 ± 4.3 69
n.d. 6.2 ± 1.9 17
68 11 ± 5.4 34
150 23 ± 9.9 74
37 1.8 ± 0.5
9.2
n.d.
n.d.
n.d.
n.d.
n.d.
34
5.1 ± 2.1 9.2
19 5.0 ± 2.4 9.2
19
>65
n.d.
74
n.d.
74
n.d.
n.d.
n.d.
n.d.
n.d.
49 ± 23
n.d.
65
n.d.
37
>65 41 ± 20
n.d. n.d.
65
16 ± 5.3 37
7.9 ± 2.9 19
23 ± 7.5 69
n.d.
74
12 ± 5.6
n.d.
>74 12 ± 5.1 37
n.d.
n.d.
n.d.
n.d.
34
n.d.
n.d.
n.d.
n.d.
n.d.
n.d.
n.d.
n.d.
68
10 5.9 ± 1.8 17
11 12 ± 4.3 34
12 20 ± 5.2 74
n.d.
68 13 ± 4.7
150 27 ± 9.4
20 ± 10
31 ± 13
68 18 ± 8.4 34
150 29 ± 9.9 74
74
74
150
2.4.2. Antibacterial Activity of Synthetic Chrysophaentin Fragments
We were gratified to find that compound 11 inhibited the growth of S. aureus 25923 with MIC50
and MIC90 values comparable to those for the linear and symmetrically linked natural products 5 and 8
(Table 1). At the MIC90, compound 11 was bacteriostatic, and its MBC was 68 µM. Compound 11 also
inhibited the growth of MRSA BAA-41 and MDRSA BAA-44 with low micromolar MIC50s, and an

Mar. Drugs 2012, 10
1113
MIC90 of 34 µM for both strains. Compound 12 was slightly less active with MIC50s in the range of
20–28 μM against S. aureus 25923, MRSA BAA-41, and MDRSA BAA-44. An increase to 74 µM in
MIC90 was observed for all three strains. Compound 12 was bacteriostatic at this concentration, with a
MBC of ~150 µM.
2.4.3. Antibacterial Activity against S. aureus UAMS-1 and CA-MRSA USA300-LAC
Given the notable antibacterial activity observed for diverse laboratory strains of S. aureus, we were
interested to compare the effects of several representative compounds on the clinical S. aureus strains
UAMS-1 and CA-MRSA USA300-LAC. Compounds evaluated included chrysophaentins A, D, and F,
and synthetic compounds 11 and 12, and MIC50, MIC90 and MBC values were determined for each
compound. Chrysophaentin A was again the most potent, with an MIC50 of 5 µM for both clinical
strains (Table 1), and the MIC90 was only slightly higher at 9 µM. Chrysophaentin A was still
bacteriostatic at low micromolar concentrations, and the MBC was only 2 times higher at 19 µM. By
comparison, the symmetrical chrysophaentin F (8) was slightly less active with respective MIC50 and
MIC90 values of 12 and 37 µM, and was also bacteriostatic at the MIC90. The MIC50s for compounds
11 and 12 were similar to those of 8 as was the MIC90 for compound 11, while the MIC90 for
compound 12 was higher. For both synthetic compounds, the MBC was two-fold higher than the
MIC90. Finally, chrysophaentin D (4) was the least active with MIC50s between 41 and 49 µM, and
MIC90s at 65 µM.
Growth curves for chrysophaentins A (1), D (4), and F (8), and compound 12 were graphed as
percentage growth of untreated bacteria for S. aureus UAMS-1 (Figure 5A) and CA-MRSA
USA300-LAC (Figure 5B). These curves clearly demonstrate the relationship between the structure of
the chrysophaentins and analogs, halogen composition, and anti-staphylococcal activity.
Figure 5. Dose-response curves show the antibacterial activities and reveal a
structure-activity relationship for the chrysophaentins. Chrysophaentins A (1), D (4), and F (8),
and fragment 12 were tested for their ability to inhibit the growth of Staphylococcus aureus
UAMS-1 (panel A) and CA-MRSA USA300-LAC (panel B) in a microbroth dilution assay.
Plots represent percentage bacterial growth as a function of compound concentration.

Mar. Drugs 2012, 10
1114
2.5. Discussion
C. taylori is a rare marine chrysophyte alga. It was originally discovered in the Dry Tortugas off the
southern coast of Florida [40], and has since been found in several locations worldwide. Bioactive
natural products have been discovered from collections made in two locations. The Puerto Rico
location yielded hormothamnione [41] and 6-desmethoxyhormothamnione [42], while the collections
from St. John, U.S. Virgin Islands have yielded the ten chrysophaentins, as well as hormothamnione
and 6-methoxyhormothamnione. As we have described here, analysis of methanol extracts revealed
geographic variability within the C. taylori collections from St. John. The original collection coming
from Round Bay in 2007 yielded chrysophaentins A–H. A second collection at Round Bay two years
later contained a similar profile of compounds, indicating temporal stability in the natural compound
production at this location. A 2009 collection at Long Bay contained the same eight chrysophaentins in
a similar ratio. Both Long Bay and Round Bay are on the southeastern side of the island, and each of
these collections was made in shallow protected waters, where the algae were growing on coral sand.
The 2009 collection from Hawk’s Nest on the northwestern side of the island yielded a different
profile of compounds. It was immediately obvious when comparing the chromatograms that the
difference laid in the complete absence of the asymmetrical chrysophaentins in the samples collected
at this location. Instead, the most abundant compounds included the acyclic chrysophaentin E, as well
as two newly identified linear chrysophaentins, namely chrysophaentins E2 and E3.
Synthetic efforts to obtain a steady supply of chrysophaentin A are under way. An early approach
that was employed provided two chrysophaentin fragments that we have shown exhibit similar
potencies and spectrum of activity to those of the parent chrysophaentins. Previously, we profiled the
anti-staphylococcal activity of several of the natural chrysophaentins towards common laboratory
strains of S. aureus. In this study, we sought to expand the structure-activity relationship (SAR) by
including the synthetic fragments, and to expand the number and diversity of S. aureus strains. We
tested select compounds against five S. aureus strains to measure their potency (MIC50 and MIC90) and
to determine whether the compounds were bacteriostatic or bactericidal. All chrysophaentins, natural
and synthetic, were bacteriostatic at their MIC90; but were bactericidal at concentrations ranging from
two to eight fold higher than their respective MIC90s. Furthermore, the synthetic fragments showed
similar potency to the acyclic chrysophaentin E (5).
The synthetic fragments fit perfectly into our proposed structure-activity relationship. The
tetrachlorinated macrocycle with the asymmetrical linkage between C-16 and C-1′ (chrysophaentin A)
was essential for maximum potency. Compounds with an ether bond between C-14 and C-1′, which
lead to a symmetrical macrocycle as seen in chrysophaentin F, show a decrease in overall potency. The
presence of a fifth halogen at C-12′ did not decrease the antibacterial activity, even when the additional
halogen was a bromine, indicating that the atom type at this position or even this ring is not critical to
the observed activity. The absence of a macrocyclic structure, as in chrysophaentin E and the synthetic
fragments 11 and 12, further decreased antibacterial activity. However, the presence of the
asymmetrical macrocycle did not guarantee potency as halogen diversity on the phenyl rings also
affected activity. Chrysophaentin D, which contains two bromines in place of the chlorines on the
phenol rings in 1, was the least active compound considered in this study.

Mar. Drugs 2012, 10
1115
We have shown that subtle changes to the chrysophaentin structure and composition lead to
differences in antibacterial activity. Knowing that the macrocycle is essential for full antibacterial
potency, it is not surprising that the synthetic fragments described here have diminished activity
compared to the cyclic natural products. However, we were gratified to find that they are indeed
antimicrobial and their activities are comparable to the linear chrysophaentins. Their ease of synthesis
and antimicrobial activity provide an excellent starting point not only toward the cyclic
chrysophaentins, but also for generating more elaborate synthetic analogs.
The biosynthetic pathway leading to production of the chrysophaentins has not been characterized
and a genome sequence for any Chrysophaeum species has yet to be published. Interestingly, the most
closely related natural products include the marchantins, bazzanins, and chlorinated isoplagiochins, all
of which were isolated from different species of liverworts [43]. While these macrocyclic compounds
are reminiscent of the chrysophaentins, each family possesses defining structural characteristics that
differ relative to one another and to the chrysophaentins. The marchantins [44] are bisbibenzyl ethers
whose phenol rings are connected by an ethyl group, and thus far do not contain halogens. The
bazzanins [45,46] and chlorinated isoplagiochins [47], on the other hand are chlorinated, but are
bisbibenzyls rather than bisbibenzyl ethers. Like the marchantins and all liverwort metabolites, they
also contain ethyl linkages. The presence of a butene linkage is unique to the chrysophaentins. While
some information is known about the biosynthesis of the liverwort metabolites [48], the pathways are
incompletely characterized; moreover, parallels to chrysophaentin biosynthesis cannot be predicted as
the producing organisms are not closely related. The lack of biosynthetic information underlines the
importance of a synthetic approach that could supply large quantities of the chrysophaentins for further
biological studies.
3. Experimental Section
3.1. Biological Material
Samples of the marine algae were collected from Round Bay in July 2007, and Long Bay, Round Bay,
and Hawk’s Nest in June 2009, St. John, U.S. Virgin Islands (Collection permit VIIS-2008-SCI-0034
National Park Service). Each of the alga samples were observed as fluffy, sulfur yellow colonies
growing on coarse sand at shallow depths averaging −20 ft. The samples were identified as
Chrysophaeum taylori Lewis and Bryan as described previously [27]. Briefly, the presence of clear
stalk like structures made up of branching mucilaginous “streamers” of pear shaped, invaginated cells
were visible by light microscopy (4–10×) on freshly collected specimens, as well as those preserved in
2.5% glutaraldehyde in sterile sea water immediately after collection.
3.2. LC-MS
Lyophilized material was sequentially extracted with hexanes, CH2Cl2, and MeOH. The MeOH
extract was analyzed by reverse-phase HPLC with an MS detector (Jupiter Proteo C12, 250 × 4.6 mm,
4 µm, DAD at 220 and 280 nm, flow rate 0.5 mL/min) eluting with a linear gradient of 50–80%
methanol in 0.05% TFA-water in 50 min.

Mar. Drugs 2012, 10
1116
3.3. Mass Spectrometry and MS/MS Fragmentation
The MeOH extract was partitioned between n-BuOH–H2O (1:1), and the organic layer was
fractionated on Sephadex LH-20. Fractions containing the unknown chrysophaentins were subjected to
HR-ESI-MS and MS in-source fragmentation. High resolution accurate mass data was obtained using a
Waters LCT Premiere ESI-TOF mass spectrometer equipped with a Z-Spray electrospray ion source.
The instrument was operated in the w-mode at a nominal resolution of 10,000 and an internal standard
was used as reference. The ionization mode was negative ion and the ESI capillary voltage was −3000 V
and the cone voltage was 30 V. The collisionally activated dissociation (CAD) mass spectra were
obtained using a Waters LCT Premiere ESI-TOF mass spectrometer according to the method described
by Ren et al. [49] The ion guide 1 voltage was 65 V and the cone voltage was 30 V. The instrument
was operated in negative ion mode.
3.4. General Synthesis
All moisture sensitive reactions were performed using syringe-septum techniques under an
atmosphere of either dry N2 or dry argon unless otherwise noted. All glassware was dried in an oven at
140 °C for a minimum of 6 h or flame-dried under an atmosphere of dry nitrogen prior to use.
Reactions carried out at −78 °C employed a CO2(s)/acetone bath. Et2O and tetrahydrofuran were dried
by distillation over sodium/benzophenone under an argon atmosphere. Dry methylene chloride was
purified by filtration through an activated alumina column. All degassed solvents were prepared using
the freeze/pump/thaw method (3×). Methanol, acetonitrile, and N,N-dimethylformamide were stored
over molecular sieves (3 Å). Deuterated chloroform was stored over anhydrous potassium carbonate.
Reactions were monitored by TLC analysis (pre-coated silica gel 60 F254 plates, 250 µm layer thickness)
and visualized by using UV lamp (254 nm) or by staining with either Vaughn’s reagent (4.8 g of
(NH4)6Mo7O24·4H2O and 0.2 g of Ce(SO4)2 in 100 mL of a 3.5 N H2SO4) or a potassium permanganate
solution (1.5 g of KMnO4 and 1.5 g of K2CO3 in 100 mL of a 0.1% NaOH solution). Flash column
chromatography was performed with 40–63 µm silica gel (Silicycle). Microwave reactions were
performed on a Biotage Initiator microwave reactor. Infrared spectra were measured on a Smiths
Detection IdentifyIR FT-IR spectrometer (ATR). Unless otherwise indicated, all NMR data were
collected at room temperature in CDCl3 or (CD3)2CO on a 300, 400, 500, 600, or 700 MHz Bruker
instrument. Chemical shifts (δ) are reported in parts per million (ppm) with internal CHCl3 (δ 7.26 ppm
1
13
1
13
for H and 77.00 ppm for C), or internal acetone (δ 2.05 ppm for H and 29.85 ppm for C), as the
1
reference. H NMR data are reported as follows: chemical shift, multiplicity (s = singlet, bs = broad
singlet, d = doublet, t = triplet, q = quartet, m = multiplet, dd = doublet of doublets, dt = doublet of
triplets, td = triplet of doublets, qd = quartet of doublets, sep = septet), integration, and coupling
constant(s) (J) in Hertz (Hz).
3,5-Dimethoxy benzyl alcohol (18) [29]. To a stirred suspension of lithium aluminum hydride (13.1 g,
329 mmol, 2 equiv) in dry THF (400 mL) at 0 °C was added a solution of 3,5-dimethoxybenzoic acid
(30.0 g, 164 mmol, 1 equiv) in dry THF (400 mL) over 45 min. Upon completion of the addition, the
reaction mixture was diluted with THF (300 mL), warmed to rt, stirred for 5 h, and slowly quenched
with a saturated aqueous solution of Na/K tartrate. The resulting biphasic mixture was stirred at r.t. for

Mar. Drugs 2012, 10
1117
1 h, the organic layer was separated, the remaining aqueous layer was extracted with EtOAc (×2), and
the combined organic layers were dried (MgSO4), filtered, and concentrated under reduced pressure to
give 27.0 g (97%) of 18 as a colorless oil: Rf 0.33 (EtOAc/hexanes, 1:2); IR (CDCl3) 3390, 2937,
1
1594, 1456, 1428, 1318, 1294, 1202, 1146, 1057, 829 cm−1; H NMR (600 MHz, CDCl3) δH 6.45
(d, 2 H, J = 2.4 Hz), 6.32 (d, 1 H, J = 2.4 Hz), 4.50 (d, 2 H, J = 5.4 Hz), 3.71 (s, 6 H), 3.49 (t, 1 H,
J = 5.4 Hz); 13C NMR (150 MHz, CDCl3) δC 160.5, 143.3, 104.2, 99.2, 64.5, 55.0.
3,5-Dimethoxybenzyl bromide (19) [29]. To a stirred solution of 18 (17.5 g, 104 mmol, 1 equiv) in
CH2Cl2 (500 mL) at 0 °C was added dropwise PBr3 (12.1 mL, 125 mmol, 1.2 equiv). The reaction
mixture was slowly warmed to rt, stirred for 3 h, quenched with a saturated aqueous solution of
NaHCO3, and stirred at r.t. for 1 h. The organic layer was separated and the aqueous layer was
extracted with Et2O. The combined organic layers were dried (MgSO4), filtered, and concentrated
under reduced pressure to give 22.2 g (92%) of 19 as a white solid: Rf 0.75 (EtOAc/hexanes, 1:2);
1
IR (CDCl3) 2999, 2954, 1596, 1458, 1428, 1345, 1323, 1297, 1204, 1152, 1062, 930 cm−1; H NMR
(400 MHz, CDCl3) δH 6.54 (d, 2 H, J = 2.0 Hz), 6.39 (d, 1 H, J = 2.4 Hz), 4.42 (s, 2 H), 3.80 (s, 6 H);
13C NMR (150 MHz, CDCl3) δC 160.9, 139.7, 107.0, 100.6, 55.4, 33.6; HRMS (ESI+) m/z calcd for
C9H12O2Br 231.0021, found 231.0038.
(3-(3,5-Dimethoxyphenyl)prop-1-ynyl)trimethylsilane (20). To a stirred solution of ethynyl
trimethysilane (39.1 mL, 277 mmol, 4 equiv) in THF (120 mL) at 0 °C was added ethylmagnesium
bromide (3.16 M in Et2O; 87.6 mL, 277 mmol, 4 equiv). The reaction was warmed to rt, stirred for
30 min, and CuBr (9.93 g, 69.2 mmol, 1 equiv) was added. The reaction was stirred at r.t. for 15 min,
3,5-dimethoxybenzyl bromide (19, 16.0 g, 69.2 mmol, 1 equiv) was added, the resulting mixture was
heated to 66 °C , and stirred overnight. The reaction mixture was diluted with Et2O, slowly quenched
with brine, and extracted with Et2O (×2). The combined organic layers were washed with a saturated
aqueous solution of NH4Cl, brine, dried (MgSO4), filtered, and concentrated under reduced pressure.
The crude mixture was purified by chromatography on SiO2 (EtOAc/hexanes, 1:10) to give 17.2 g
(100%) of 20 as a pale yellow oil: Rf 0.41 (EtOAc/hexanes, 1:20); IR (neat) 2956, 2898, 2175, 1754,
1
1596, 1459, 1428, 1204, 1156, 1122, 1101, 839 cm-1; H NMR (400 MHz, CDCl3) δH 6.53 (d, 2 H,
13
J = 2.4 Hz), 6.34 (t, 1 H, J = 2.4 Hz), 3.79 (s, 6 H), 3.60 (s, 2 H), 0.19 (s, 9 H); C NMR (100 MHz,
CDCl3) δC 160.8, 138.7, 105.9, 104.0, 98.7, 87.1, 55.3, 26.3, 0.1; HRMS (ESI+) m/z calcd for
C14H21O2Si 249.1311, found 249.1287.
1,3-Dimethoxy-5-(prop-2-ynyl)benzene (15). To a stirred solution of 20 (5.37 g, 21.6 mmol, 1 equiv)
and AcOH (4.99 mL, 86.5 mmol, 4 equiv) in THF (100 mL) was added dropwise TBAF (1 M in THF;
86.5 mL, 86.5 mmol, 4 equiv). The resulting reaction mixture was stirred at r.t. for 24 h, diluted with
Et2O, washed with brine (×2), dried (MgSO4), filtered, and concentrated under reduced pressure.
The crude mixture was purified by chromatography on SiO2 (EtOAc/hexanes, 1:10) to give 3.85 g
(100%) of 15 as a colorless oil: Rf 0.36 (EtOAc/hexanes, 1:10); IR (CDCl3) 3286, 2999, 2954, 1593,
1
1457, 1428, 1344, 1323, 1288, 1204, 1154, 1064, 827; H NMR (400 MHz, CDCl3) δH 6.53 (d, 2 H,
J = 2.4 Hz), 6.35 (t, 1 H, J = 2.4 Hz), 3.79 (s, 6 H), 3.56 (d, 2 H, J = 2.8 Hz), 2.20 (t, 1 H, J = 2.8 Hz);
13C NMR (100 MHz, CDCl3) δC 160.9, 138.3, 105.9, 98.7, 81.7, 70.6, 55.3, 25.0; HRMS (EI+)
m/z calcd for C11H12O2 176.0837, found 176.0834.

Mar. Drugs 2012, 10
1118
5-(Prop-2-ynyl)-1,3-phenylene bis(2,2-dimethylpropanoate (21). To a stirred solution of 15 (6.50 g,
36.9 mmol, 1 equiv) in CH2Cl2 (1200 mL) at 0 °C was added BBr3 (1 M in CH2Cl2; 92.2 mL,
92.2 mmol, 5 equiv) over 1 h via an addition funnel. The reaction mixture was warmed to r.t. and
stirred overnight. The reaction mixture was slowly quenched with a saturated aqueous solution of
NaHCO3 (500 mL) and stirred at r.t. for 4 h. The solution was acidified with HCl, extracted with
CH2Cl2 and EtOAc, and the combined organic layers were dried (MgSO4), filtered, and concentrated
under reduced pressure. The crude mixture was dissolved in CH2Cl2 (180 mL), and triethylamine
(20.9 mL, 148 mmol, 4 equiv) and PivCl (11.6 mL, 92.2 mmol, 2.5 equiv) were added. The resulting
solution was stirred at r.t. for 90 min, diluted with brine, the organic layer was separated, and the
aqueous layer was extracted with EtOAc. The combined organic layers were dried (MgSO4), filtered,
and concentrated under reduced pressure. The crude mixture was purified by chromatography on SiO2
(EtOAc/hexanes, 1:10) to give 9.6 g (82%) of 21 as a colorless oil: Rf 0.85 (EtOAc/hexanes, 3:7);
IR (neat) 3293, 2973, 1806, 1750, 1414, 1452, 1396, 1366, 1269, 1118, 1098, 1031, 1003 cm−1;
1H NMR (300 MHz, CDCl3) δH 6.95 (d, 2 H, J = 1.8 Hz), 6.76 (t, 1 H, J = 1.8 Hz), 3.61 (d, 2 H,
J = 2.4 Hz), 2.21 (t, 1 H, J = 2.4 Hz); 13C NMR (75 MHz, CDCl3) δC 176.6, 151.5, 138.3, 118.3, 113.8,
80.7, 71.2, 39.1, 27.1, 24.5; HRMS (EI+) m/z calcd for C19H24O4 316.1675, found 316.1670.
4-Chloro-2,5-dimethoxybenzaldehyde (16) [33,34]. To a stirred solution of 2-chloro-1,4-dimethoxy
benzene (25.0 g, 0.145 mol, 1 equiv) and hexamethylene tetramine (20.5 g, 0.145 mol, 1 equiv) at r.t.
was carefully added TFA (250 mL). The resulting yellow suspension was heated to 95 °C , stirred for
5 h, and the hot brown solution was poured into a 2 L Erlenmeyer flask containing approximately
250 g of crushed ice. To the vigorously stirred mixture was slowly added solid NaHCO3 (243 g,
2.90 mol, 20 equiv) in 5–10 g portions over two hours. The resulting yellow precipitate was filtered
through a pad of celite, washed with water, and dissolved in Et2O. The organic layer was washed with
water and brine, dried (MgSO4), filtered, and concentrated under reduced pressure to yield 29.0 g
(100%) of 16 as an off-white solid: Rf 0.70 (EtOAc/hexanes, 3:7); IR (neat) 2941, 2874, 1664, 1601,
1
1575, 1497, 1478, 1461, 1389, 1269, 1213, 1023, 977 cm−1; H NMR (400 MHz, CDCl3) δH 10.35
(s, 1 H), 7.33 (s, 1 H), 7.03 (s, 1 H), 3.87 (s, 6 H); 13C NMR (100 MHz, CDCl3) δC 188.4, 156.1, 149.4,
130.4, 123.4, 114.5, 109.9, 56.5, 56.3; HRMS (EI+) m/z calcd for C9H9O3Cl 200.0240, found 200.0238.
(4-Chloro-2,5-dimethoxyphenyl)methanol (25). To a stirred solution of 16 (29.0 g, 145 mmol,
1 equiv) in absolute ethanol (550 mL) was added sodium borohydride (27.3 g, 723 mmol, 5 equiv).
The reaction mixture was stirred at r.t. for 6 h, quenched via dropwise addition of acetone, diluted with
EtOAc, washed with brine (×2), dried (MgSO4), filtered, and concentrated under reduced pressure. The
crude residue was purified by chromatography on SiO2 (EtOAc/hexanes, 1:1) to give 28.0 g (96%) of
25 as a white solid: mp 89–90 °C ; Rf 0.53 (EtOAc/hexanes, 1:1); IR (neat) 3258, 2958, 2915, 1495,
1
1461, 1392, 1204, 1061, 719 cm−1; H NMR (300 MHz, CDCl3) δH 6.90 (s, 1 H), 6.80 (s, 1 H), 4.55
13
(d, 2 H, J = 4.2 Hz), 3.77 (s, 3 H), 3.71 (s, 3 H), 3.10 (bs, 1 H); C NMR (75 MHz, CDCl3) δC 150.6,
148.7, 128.4, 120.8, 112.5, 112.4, 60.3, 56.5, 55.6; HRMS (ESI+) m/z calcd for C9H12O3Cl 203.0475,
found 203.0465.
1-(Bromomethyl)-4-chloro-2,5-dimethoxybenzene (26) [50]. To a stirred solution of 25 (5.00 g,
24.7 mmol, 1 equiv) in CH2Cl2 (125 mL) at 0 °C was added dropwise HBr (47–49% aqueous solution;
4.13 mL, 74.0 mmol, 3 equiv). The resulting solution was slowly warmed to r.t. and stirred overnight.

Mar. Drugs 2012, 10
1119
The following morning a second batch of HBr (47–49% solution; 4.13 mL, 74.0 mmol, 3 equiv) was
added to the reaction mixture, which was stirred at r.t. for an additional 4 h, extracted with Et2O (×2),
washed with water, a saturated aqueous solution of NaHCO3, brine, dried (MgSO4), filtered, and
concentrated under reduced pressure to give 6.55 g (100%) of 26 as a white solid: Rf 0.31
(EtOAc/hexanes, 3:7); IR (neat) 2962, 2947, 2844, 1732, 1582, 1495, 1458, 1443, 1389, 1301, 1204,
1182, 1033, 882 cm−1; 1H NMR (400 MHz, CDCl3) δH 6.91–6.90 (m, 2 H), 4.50 (s, 2 H), 3.84 (s, 3 H),
3.82 (s, 3 H); 13C NMR (75 MHz, CDCl3) δC 151.3, 148.9, 125.1, 123.2, 114.6, 113.6, 56.6, 56.2, 28.2.
(E)-5-(2-Chloro-3-iodoallyl)-1,3-phenylene bis(2,2-dimethylpropanoate (22) and (Z)-5-(2-Chloro-
3-iodoallyl)-1,3-phenylene bis(2,2-dimethylpropanoate (23). To a stirred solution of 21 (9.50 g,
30.0 mmol, 1 equiv) in dry CH2Cl2 (150 mL) at 0 °C was added ICl (1 M in CH2Cl2; 30.0 mL,
30.0 mmol, 1 equiv). The reaction mixture was warmed to rt, stirred protected from light (enclosed in
aluminum foil) for 3 h, diluted with Et2O, washed with Na2SO4 (×2), brine, dried (MgSO4), filtered,
and concentrated under reduced pressure. The crude mixture was purified by chromatography on SiO2
(EtOAc/hexanes, 1:10) to give an inseparable 2:1 mixture of 22 and 23 (13.7 g, 95%) as a colorless oil:
Rf 0.54 (EtOAc/hexanes, 1:10); IR (neat) 3277, 2934, 2872, 1746, 1592, 1497, 1461, 1409, 1269,
1207, 1122, 1103, 1032, 975, 723 cm−1; HRMS (ESI+) m/z calcd for C19H24O4NaClI 501.0306, found
1
501.0308. Characteristic data for the major, desired isomer 22: H NMR (300 MHz, CDCl3) δH 6.87
13
(d, 2 H, J = 1.5 Hz), 6.80 (t, 1 H, J = 2.1 Hz), 6.62 (s, 1 H), 3.90 (s, 2 H), 1.35 (s, 18 H); C NMR
(100 MHz, CDCl3) δC 176.6, 151.6, 137.9, 135.6, 119.0, 114.2, 74.8, 44.2, 39.1, 27.1. Characteristic
signals for 23: 1H NMR (300 MHz, CDCl3) δH 7.06 (s, 1 H), 3.92 (s, 2 H).
(E)-5-(2-Chloro-4-(4-chloro-2,5-dimethoxyphenyl)but-2-enyl)benzene-1,3-diol (11). A flame-dried
microwave vial was charged with a 2:1 mixture of 22 and 23 (0.520 g, 1.08 mmol, 1 equiv), Pd(OAc)2
(0.0122 g, 0.0543 mmol, 0.05 equiv), P(o-tol)3 (0.0341 g (0.109 mmol, 0.1 equiv), and freshly
distilled, dry, degassed DMF (0.8 mL). The reaction mixture was stirred at r.t. for 10 min. In a separate
dry flask, a catalytic amount of iodine (~20 mg) and zinc (0.362 g, 5.43 mmol, 5 equiv) were heated
(bunsen burner) until a purple gas coated the interior of the flask. The flask was cooled to rt, charged
with distilled, dry, degassed DMF (1 mL), and benzyl bromide 26 (1.15 g, 4.34 mmol, 4 equiv). The
mixture was allowed to stir at r.t. under argon for 6 min. The activated organozinc reagent was filtered
through an oven dried fritted funnel under an atmosphere of argon and cannulated into the stirred
solution of 22 and 23. The resulting solution was heated in a microwave reactor (2 min, 120 °C ),
treated with a second batch of Pd(OAc)2 (0.0122 g, 0.0543 mmol, 0.05 equiv) and resubjected to the
microwave conditions (2 min, 120 °C ). The crude reaction mixture was directly purified by
chromatography on SiO2 (EtOAc/hexanes, 1:20) to give a crude yellow oil which was immediately
dissolved in MeOH/CH2Cl2 (4 mL, 2:1), and Cs2CO3 (1.67 g, 5.08 mmol, 5 equiv) was added. The
reaction mixture was stirred at r.t. for 6 h, diluted with EtOAc, and acidified with conc. HCl. The
organic layer was separated, and the acidified aqueous solution was extracted with EtOAc. The
combined organic layers were washed with brine, dried (MgSO4), filtered, and concentrated under
reduced pressure. The crude mixture was purified by chromatography on SiO2 (chloroform/acetone,
8:2) to give a 2:1 mixture of 11 and the undesired regioisomer 27 (0.192 g, 51%) as a yellow oil.
Characteristic signals for 27: 1H NMR (600 MHz (CD3)2CO) δH 8.20 (bs, 2 H), 7.02 (d, 2 H, J = 3.6 Hz),
6.45 (t, 1 H, J = 7.8 Hz), 6.26 (s, 3 H), 3.84 (app s, 5 H), 3.81 (s, 3 H), 3.50 (d, 2 H, J = 7.8 Hz);

Mar. Drugs 2012, 10
1120
13C NMR (150 MHz, CDCl3) δC 159.5, 152.3, 150.0, 141.5, 140.7, 127.7, 120.9, 115.6, 113.8, 108.1,
101.9, 101.7, 56.9, 56.5, 45.1.
An analytically pure sample of 11 for biological evaluation and characterization was obtained
via SFC chromatography using a semiprep (250 × 10 mm) silica column (Rt 5.80 min, 10 mL/min,
15% methanol, 220 nm detection): Rf 0.48 (acetone/chloroform, 2:8); IR (acetone) 3375, 3001, 2952,
1
1696, 1599, 1495, 1463, 1387, 1212, 1156, 1034, 1010 cm−1; H NMR (600 MHz (CD3)2CO) δH 8.18
(bs, 2 H), 7.02 (s, 1 H), 6.99 (s, 1 H), 6.30 (s, 2 H), 6.25 (s, 1 H), 5.88 (t, 1 H, J = 7.8 Hz), 3.84 (s, 3 H),
13
3.81 (s, 3 H), 3.73 (s, 2 H), 3.51 (d, 2 H, J = 7.8 Hz); C NMR (150 MHz, CDCl3) δC 159.5, 152.3,
150.0, 140.6, 133.7, 128.1, 128.1, 120.8, 115.5, 113.8, 108.0, 101.8, 56.9, 56.5, 40.2, 29.5; HRMS
(ES−) [M + Cl]− m/z calcd for C18H18O4Cl3 403.0271, found 403.0295.
(E)-5-(2-Chloro-4-(4-chloro-2,5-dihydroxyphenyl)but-2-enyl)benzene-1,3-diol (12). A stirred
solution of 11 (0.0500 g, 0.135 mmol, 1 equiv) in CH2Cl2 (7 mL) was enclosed in aluminum foil
and cooled to 0 °C . To the stirred solution was added dropwise BBr3 (1 M in CH2Cl2; 0.677 mL,
0.677 mmol, 5 equiv). The resulting solution was slowly warmed to r.t. and stirred overnight. The
reaction mixture was quenched with a saturated aqueous solution of NaHCO3 (8 mL) and stirred at r.t.
for an additional hour. The solution was acidified with HCl, extracted with EtOAc (×2) and the
combined organic layers were dried (MgSO4), filtered, and concentrated under reduced pressure. The
crude product was purified by chromatography on SiO2 (chloroform/acetone, 3:1) to give 12 (0.0462 g,
96%) as a slightly yellow film. Before submission for biological testing, a sample of 12 was further
purified via SFC chromatography using a semiprep (250 × 10 mm) silica column (Rt 4.68 min,
8 mL/min, 25% methanol, 220 nm detection): Rf 0.15 (acetone/chloroform, 2:8); IR (neat) 3343, 1692,
1599, 1495, 1417, 1329, 1184, 1143, 1005, 822 cm−1; 1H NMR (600 MHz (CD3)2CO) δH 8.17 (bs, 4 H),
6.86 (s, 1 H), 6.84 (s, 1 H), 6.29 (d, 2 H, J = 2.4 Hz), 6.24 (t, 1 H, J = 2.4 Hz), 5.89 (t, 1 H, J = 7.8 Hz),
13
3.69 (s, 2 H), 3.47 (d, 2 H, J = 7.8 Hz); C NMR (150 MHz, CDCl3) δC 159.5, 149.0, 146.9, 140.5,
133.8, 128.0, 127.0, 118.5 (2C), 116.7, 108.0, 101.8, 40.2, 29.5; HRMS (ES−) [2M − H]− m/z calcd for
C32H27O8Cl4 679.0460, found 679.0482.
The demethylation reaction was also performed on a 2:1 mixture of 11 and 27, resulting in a
1
2:1 mixture of 12 and its (Z)-diastereomer. Characteristic signals for the (Z)-diastereomer: H NMR
(400 MHz (CD3)2CO) δH 6.85 (s, 1 H), 6.48 (t, 1 H, J = 7.6 Hz), 3.81 (s, 2 H), 3.46 (d, 2 H, J = 7.6 Hz).
3.5. Anti-Staphylococcal Activity
Microbroth dilution assays were performed as described in the CLSI guidelines. Minimum
inhibitory concentrations (MIC50 and MIC90) were determined using 96-well microbroth dilution
assays. Antimicrobial activity of pure compounds or antibiotic standards were tested by adding serial
dilutions to wells containing Mueller Hinton II broth. An overnight growth of S. aureus ATCC 25923,
MRSA ATCC BAA-41, MDRSA BAA-44, S. aureus UAMS-1, and CA-MRSA USA300-LAC was
diluted in Mueller-Hinton II broth and added to the wells to a final concentration of 5 × 105 colony
forming units (CFU) per milliliter. Plates were incubated for 18 h at 37 °C with shaking at 200 rpm.
Absorbance at 600 nm is read on a Molecular Devices plate reader. Growth curves are plotted, and
curves were fit to a one-site model using the equation y = 100/[1 + (concentration/MIC50)], where

Mar. Drugs 2012, 10
1121
MIC50 is the concentration at which the growth of bacterial cultures are reduced by 50%. MIC90 values
were visually calculated by determining the lowest concentration of test compound that allowed no
visible growth (no difference of absorption between treated samples and blank controls).
Bacteriostatic (BS) or bactericidal (BC) properties were determined by plating onto a fresh TSA
plate an aliquot from each well that showed no visible growth starting at the MIC90. Plates were
incubated for 24 h at 37 °C , and colonies were counted. Bactericidal concentrations were defined as
those that resulted in a three logarithmic decrease in the number of viable bacteria relative to the
starting inoculum, and the lowest bactericidal concentration was defined as the MBC.
4. Conclusions
The rare marine chrysophyte alga C. taylori has yielded a number of biologically active natural
products. While the occurrence of C. taylori has been documented at a growing number of locations
globally, thus far only collections obtained from the southeastern side of St. John appear to contain the
full suite of chrysophaentins. The geographical variability described here, along with temporal and
seasonal changes in the abundance of C. taylori, limited our ability to obtain additional material for
study. Nevertheless, early synthetic efforts toward the synthesis of chrysophaentin A in particular have
provided fragments of the natural products that have proven useful for biological studies. Here we
have expanded the antimicrobial studies of several chrysophaentins and two synthetic fragments to
include drug-susceptible and drug-resistant S. aureus strains that are of current clinical interest.
Although approximately half the size of the cyclic chrysophaentins, we have shown that the potencies
and spectrum of antimicrobial activity of the synthetic fragments are comparable to those of the parent
chrysophaentins, in particular to the acyclic compound chrysophaentin E. Moreover, both the natural
products and synthetic fragments are equally effective against both drug-resistant and drug-susceptible
S. aureus strains. In summary, our results show promise for the notion of synthesizing equally potent
chrysophaentin analogs that will be more readily available, and provide new data on the antimicrobial
spectrum of this novel class of antibiotics.
Acknowledgments
We thank the United States National Park Service for research collecting permits for the U.S.
Virgin Islands, P. Dunman for clinical S. aureus strains, and M. Bewley for assistance with collections.
We also thank Boehringer Ingelheim Pharmaceuticals Inc. for financial support (P.W.). This work was
supported in part by the National Institutes of Health Intramural Research Program (NIDDK), and the
Intramural AIDS Targeted Antiviral Program, Office of the Director, NIH (C.A.B.). J.L.K. is a
recipient of a predoctoral Intramural AIDS Research Fellowship, Office of the Director, NIH.
References
1. Diekema, D.J.; Pfaller, M.A.; Schmitz, F.J.; Smayevsky, J.; Bell, J.; Jones, R.N.; Beach, M.
Survey of infections due to Staphylococcus species: Frequency of occurrence and antimicrobial
susceptibility of isolates collected in the United States, Canada, Latin America, Europe, and the
Western Pacific region for the SENTRY Antimicrobial Surveillance Program, 1997–1999. Clin.
Infect. Dis. 2001, 32 (Suppl. 2), S114–S132.

Mar. Drugs 2012, 10
1122
2. Kirby, W.M. Extraction of a highly potent penicillin inactivator from penicillin resistant
Staphylococci. Science 1944, 99, 452–453.
3. Barber, M.; Rozwadowska-Dowzenko, M. Infection by penicillin-resistant staphylococci. Lancet
1948, 2, 641–644.
4. Jevons, M.P. “Celbenin”-resistant Staphylococci. Br. Med. J. 1961, 1, 124–125.
5. Stewart, G.T.; Holt, R.J. Evolution of natural resistance to the newer penicillins. Br. Med. J. 1963,
1, 308–311.
6. Mato, R.; Sanches, S.I.; Venditti, M.; Platt, D.J.; Brown, A.; Chung, M.; de Lencastre, H. Spread
of the multiresistant Iberian clone of methicillin-resistant Staphylococcus aureus (MRSA) to Italy
and Scotland. Microb. Drug Resist. 1998, 4, 107–112.
7. Klevens, R.M.; Morrison, M.A.; Nadle, J.; Petit, S.; Gershman, K.; Ray, S.; Harrison, L.H.;
Lynfield, R.; Dumyati, G.; Townes, J.M.; et al. Invasive methicillin-resistant Staphylococcus
aureus infections in the United States. J. Am. Med. Assoc. 2007, 298, 1763–1771.
8. Bordon, J.; Master, R.N.; Clark, R.B.; Duvvuri, P.; Karlowsky, J.A.; Ayesu, K.; Klotchko, A.;
Kapoor, R.; Ramirez, J. Methicillin-resistant Staphylococcus aureus resistance to non-beta-lactam
antimicrobials in the United States from 1996 to 2008. Diagn. Microbiol. Infect. Dis. 2010, 67,
395–398.
9. Moran, G.J.; Krishnadasan, A.; Gorwitz, R.J.; Fosheim, G.E.; McDougal, L.K.; Carey, R.B.;
Talan, D.A. Methicillin-resistant S. aureus infections among patients in the emergency department.
N. Engl. J. Med. 2006, 355, 666–674.
10. Miller, L.G.; Diep, B.A. Clinical practice: Colonization, fomites, and virulence: Rethinking the
pathogenesis of community-associated methicillin-resistant Staphylococcus aureus infection.
Clin. Infect. Dis. 2008, 46, 752–760.
11. Baba, T.; Takeuchi, F.; Kuroda, M.; Yuzawa, H.; Aoki, K.; Oguchi, A.; Nagai, Y.; Iwama, N.;
Asano, K.; Naimi, T.; et al. Genome and virulence determinants of high virulence
community-acquired MRSA. Lancet 2002, 359, 1819–1827.
12. Daum, R.S.; Ito, T.; Hiramatsu, K.; Hussain, F.; Mongkolrattanothai, K.; Jamklang, M.;
Boyle-Vavra, S. A novel methicillin-resistance cassette in community-acquired methicillin-resistant
Staphylococcus aureus isolates of diverse genetic backgrounds. J. Infect. Dis. 2002, 186,
1344–1347.
13. Okuma, K.; Iwakawa, K.; Turnidge, J.D.; Grubb, W.B.; Bell, J.M.; O’Brien, F.G.; Coombs, G.W.;
Pearman, J.W.; Tenover, F.C.; Kapi, M.; et al. Dissemination of new methicillin-resistant
Staphylococcus aureus clones in the community. J. Clin. Microbiol. 2002, 40, 4289–4294.
14. Vandenesch, F.; Naimi, T.; Enright, M.C.; Lina, G.; Nimmo, G.R.; Heffernan, H.; Liassine, N.;
Bes, M.; Greenland, T.; Reverdy, M.E.; et al. Community-acquired methicillin-resistant
Staphylococcus aureus carrying Panton-Valentine leukocidin genes: Worldwide emergence.
Emerg. Infect. Dis. 2003, 9, 978–984.
15. Lina, G.; Piemont, Y.; Godail-Gamot, F.; Bes, M.; Peter, M.O.; Gauduchon, V.; Vandenesch, F.;
Etienne, J. Involvement of Panton-Valentine leukocidin-producing Staphylococcus aureus in
primary skin infections and pneumonia. Clin. Infect. Dis. 1999, 29, 1128–1132.

Mar. Drugs 2012, 10
1123
16. Moellering, R.C., Jr.; Abbott, G.F.; Ferraro, M.J. Case records of the Massachusetts General
Hospital. Case 2–2011. A 30-year-old woman with shock after treatment of a furuncle. N. Engl. J.
Med. 2011, 364, 266–275.
17. Deleo, F.R.; Otto, M.; Kreiswirth, B.N.; Chambers, H.F. Community-associated meticillin-resistant
Staphylococcus aureus. Lancet 2010, 375, 1557–1568.
18. Abi-Hanna, P.; Frank, A.L.; Quinn, J.P.; Kelkar, S.; Schreckenberger, P.C.; Hayden, M.K.;
Marcinak, J.F. Clonal features of community-acquired methicillin-resistant Staphylococcus
aureus in children. Clin. Infect. Dis. 2000, 30, 630–631.
19. Tenover, F.C.; McDougal, L.K.; Goering, R.V.; Killgore, G.; Projan, S.J.; Patel, J.B.;
Dunman, P.M. Characterization of a strain of community-associated methicillin-resistant
Staphylococcus aureus widely disseminated in the United States. J. Clin. Microbiol. 2006, 44,
108–118.
20. Diep, B.A.; Stone, G.G.; Basuino, L.; Graber, C.J.; Miller, A.; des Etages, S.A.; Jones, A.;
Palazzolo-Ballance, A.M.; Perdreau-Remington, F.; Sensabaugh, G.F.; et al. The arginine
catabolic mobile element and staphylococcal chromosomal cassette mec linkage: Convergence of
virulence and resistance in the USA300 clone of methicillin-resistant Staphylococcus aureus.
J. Infect. Dis. 2008, 197, 1523–1530.
21. Gorwitz, R.J. Understanding the success of methicillin-resistant Staphylococcus aureus strains
causing epidemic disease in the community. J. Infect. Dis. 2008, 197, 179–182.
22. Montgomery, C.P.; Boyle-Vavra, S.; Adem, P.V.; Lee, J.C.; Husain, A.N.; Clasen, J.; Daum, R.S.
Comparison of virulence in community-associated methicillin-resistant Staphylococcus aureus
pulsotypes USA300 and USA400 in a rat model of pneumonia. J. Infect. Dis. 2008, 198, 561–570.
23. Payne, D.J.; Gwynn, M.N.; Holmes, D.J.; Pompliano, D.L. Drugs for bad bugs: Confronting the
challenges of antibacterial discovery. Nat. Rev. Drug Discov. 2007, 6, 29–40.
24. Newman, D.J.; Cragg, G.M. Natural products as sources of new drugs over the 30 years from
1981 to 2010. J. Nat. Prod. 2012, 75, 311–335.
25. Appelbaum, P.C. Reduced glycopeptide susceptibility in methicillin-resistant Staphylococcus
aureus (MRSA). Int. J. Antimicrob. Agents 2007, 30, 398–408.
26. Zetola, N.; Francis, J.S.; Nuermberger, E.L.; Bishai, W.R. Community-acquired meticillin-resistant
Staphylococcus aureus: An emerging threat. Lancet Infect. Dis. 2005, 5, 275–286.
27. Plaza, A.; Keffer, J.L.; Bifulco, G.; Lloyd, J.R.; Bewley, C.A. Chrysophaentins A–H, antibacterial
bisdiarylbutene macrocycles that inhibit the bacterial cell division protein FtsZ. J. Am. Chem. Soc.
2010, 132, 9069–9077.
28. Google Earth. Image: U.S. Geological Survey, Image copyright: 2012 Digital Globe,
GeoEye, Google. Available online: http://www.google.com/earth/index.html (accessed on
16 February 2012).
29. Kozlowski, M.C.; Dugan, E.C.; DiVirgilio, E.S.; Maksimenka, K.; Bringmann, G. Asymmetric
total synthesis of nigerone and ent-nigerone: Enantioselective oxidative biaryl coupling of highly
hindered naphthols. Adv. Synth. Catal. 2007, 349, 583–594.
30. Gopalsamuthiram, V.; Wulff, W.D. A new convergent strategy for the synthesis of calixarenes via
a triple annulation of Fischer carbene complexes. J. Am. Chem. Soc. 2004, 126, 13936–13937.

Mar. Drugs 2012, 10
1124
31. Bellina, F.; Colzi, F.; Mannina, L.; Rossi, R.; Viel, S. Reaction of alkynes with iodine
monochloride revisited. J. Org. Chem. 2003, 68, 10175–10177.
32. Uemura, S.; Okazaki, H.; Onoe, A.; Okano, M. Chlorination and chloroiodination of acetylenes
with copper(II) chloride. J. Chem. Soc. Perkin Trans. 1 1977, 6, 676–680.
33. Wright, B.J.D.; Hartung, J.; Peng, F.; van de Water, R.; Liu, H.B.; Tan, Q.H.; Chou, T.C.;
Danishefsky, S.J. Synthesis of pluraflavin A “aglycone”. J. Am. Chem. Soc. 2008, 130,
16786–16790.
34. Bloomer, J.L.; Stagliano, K.W.; Gazzillo, J.A. Preparation of functionalized juglone acetates and
juglones via 1,4-dimethoxynaphthalene derivatives: Synthesis of anthraquinones related to rhein
and aloe emodin. J. Org. Chem. 1993, 58, 7906–7912.
35. Oswald, C.L.; Carrillo-Marquez, T.; Caggiano, L.; Jackson, R.F.W. Negishi cross-coupling
reactions of alpha-amino acid-derived organozinc reagents and aromatic bromides. Tetrahedron
2008, 64, 681–687.
36. Sanches, I.S.; Ramirez, M.; Troni, H.; Abecassis, M.; Padua, M.; Tomasz, A.; de Lencastre, H.
Evidence for the geographic spread of a methicillin-resistant Staphylococcus aureus clone between
Portugal and Spain. J. Clin. Microbiol. 1995, 33, 1243–1246.
37. Gillaspy, A.F.; Hickmon, S.G.; Skinner, R.A.; Thomas, J.R.; Nelson, C.L.; Smeltzer, M.S. Role of
the accessory gene regulator (agr) in pathogenesis of staphylococcal osteomyelitis. Infect. Immun.
1995, 63, 3373–3380.
38. McDougal, L.K.; Steward, C.D.; Killgore, G.E.; Chaitram, J.M.; McAllister, S.K.; Tenover, F.C.
Pulsed-field gel electrophoresis typing of oxacillin-resistant Staphylococcus aureus isolates from
the United States: Establishing a national database. J. Clin. Microbiol. 2003, 41, 5113–5120.
39. Burlak, C.; Hammer, C.H.; Robinson, M.A.; Whitney, A.R.; McGavin, M.J.; Kreiswirth, B.N.;
Deleo, F.R. Global analysis of community-associated methicillin-resistant Staphylococcus aureus
exoproteins reveals molecules produced in vitro and during infection. Cell Microbiol. 2007, 9,
1172–1190.
40. Bryan, H.F.; Lewis, I.F. A new protophyte from the Dry Tortugas. Am. J. Bot. 1941, 28, 343–348.
41. Gerwick, W.H.; Lopez, A.; van Duyne, G.D.; Clardy, J.; Ortiz, W.; Baez, A. Hormothamnione, a
novel cytotoxic styrylchromone from the marine cyanophyte Hormothamnion enteromorphoides
grunow. Tetrahedron Lett. 1986, 27, 1979–1982.
42. Gerwick, W.H. 6-Desmethoxyhormothamnione, a new cytotoxic styrylchromone from the marine
cryptophyte Chrysophaeum taylori. J. Nat. Prod. 1989, 52, 252–256.
43. Harrowven, D.C.; Kostiuk, S.L. Macrocylic bisbibenzyl natural products and their chemical
synthesis. Nat. Prod. Rep. 2012, 29, 223–242.
44. Asakawa, Y.; Toyota, M.; Matsuda, R.; Takikawa, K.; Takemoto, T. Distribution of novel cyclic
bisbibenzyls in Marchantia and Riccardia species. Phytochemistry 1983, 22, 1413–1415.
45. Scher, J.M.; Zapp, J.; Schmidt, A.; Becker, H. Bazzanins L–R, chlorinated macrocyclic
bisbibenzyls from the liverwort Lepidozia incurvata. Phytochemistry 2003, 64, 791–796.
46. Martini, U.; Zapp, J.; Becker, H. Lignans from the liverwort Bazzania trilobata. Phytochemistry
1998, 49, 1139–1146.

Mar. Drugs 2012, 10
1125
47. Hashimoto, T.; Irita, H.; Takaoka, S.; Tanaka, M.; Asakawa, Y. New chlorinated cyclic
bis(bibenzyls) from the liverworts Herbertus sakuraii and Mastigophora diclados. Tetrahedron
2000, 56, 3153–3159.
48. Friederich, S.; Maier, U.H.; Deus-Neumann, B.; Asakawa, Y.; Zenk, M.H. Biosynthesis of cyclic
bis(bibenzyls) in Marchantia polymorpha. Phytochemistry 1999, 50, 589–598.
49. Ren, D.; Pipes, G.D.; Hambly, D.; Bondarenko, P.V.; Treuheit, M.J.; Gadgil, H.S. Top-down
N-terminal sequencing of immunoglobulin subunits with electrospray ionization time of flight
mass spectrometry. Anal. Biochem. 2009, 384, 42–48.
50. Bloomer, J.L.; Gazzillo, J.A. An efficient route to 3-chlorojuglones. Tetrahedron Lett. 1989, 30,
1201–1204.
Samples Availability: Available from the authors.
© 2012 by the authors; licensee MDPI, Basel, Switzerland. This article is an open access article
distributed under the terms and conditions of the Creative Commons Attribution license
(http://creativecommons.org/licenses/by/3.0/).