Chemical constituents of the lichen, Candelaria concolor: A complete NMR and chemical degradative investigation

Article abstract of DOI:10.1080/14786410802682536

A detailed chemical and spectroscopic investigation of the terrestrial lichen Candelaria concolor has yielded several lichenic metabolites belonging to the pulvinic acid series, as well as several depside derivatives including pulvinic dilactone (1), vulpinic acid (4) and calycin (5). The chemical transformation of 1 to pulvinic acid (3) is reported for the first time, as is the conversion of atranorin (6) to 5-chloroatranorin (7) and then finally to 5,5'-dichloroatranorin (8) under very mild conditions. Also presented is the complete 1D and 2D NMR assignment for compounds 1, 3, 4, 5 and 8, including partial NMR chemical shift assignments for the unstable depside (7). Previously, these metabolites had only been partially assigned by NMR spectroscopy.

Full text of DOI:10.1080/14786410802682536

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Chemical constituents of the lichen,
Candelaria concolor: A complete NMR
and chemical degradative investigation
Daniel A. Dias a & Sylvia Urban a
a School of Applied Sciences (Discipline of Applied Chemistry),
RMIT University, Melbourne, Australia
Available online: 29 Oct 2009
To cite this article: Daniel A. Dias & Sylvia Urban (2009): Chemical constituents of the lichen,
Candelaria concolor: A complete NMR and chemical degradative investigation, Natural Product
Research: Formerly Natural Product Letters, 23:10, 925-939
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Natural Product Research
Vol. 23, No. 10, 10 July 2009, 925–939
Chemical constituents of the lichen, Candelaria concolor: A complete NMR
and chemical degradative investigation
*
Daniel A. Dias and Sylvia Urban
School of Applied Sciences (Discipline of Applied Chemistry), RMIT University, Melbourne,
Australia
(Received 10 July 2008; final version received 7 January 2009)
A detailed chemical and spectroscopic investigation of the terrestrial lichen
Candelaria concolor has yielded several lichenic metabolites belonging to the
pulvinic acid series, as well as several depside derivatives including pulvinic
dilactone (1), vulpinic acid (4) and calycin (5). The chemical transformation of 1
to pulvinic acid (3) is reported for the first time, as is the conversion of atranorin
(6) to 5-chloroatranorin (7) and then finally to 5,50-dichloroatranorin (8) under
very mild conditions. Also presented is the complete 1D and 2D NMR
assignment for compounds 1, 3, 4, 5 and 8, including partial NMR chemical
shift assignments for the unstable depside (7). Previously, these metabolites had
only been partially assigned by NMR spectroscopy.
Keywords: lichen; natural product; pulvinic acid derivatives; depside; biological
activity
1. Introduction
The remarkable symbiotic relationship of lichens is one of the most successful examples
of two or more organisms (a fungus – mycobiont and an algal partner – photobiont)
living in unison. Lichens occur in the most extreme environments on Earth, such as the
cold arctic tundra, rain forests, hot deserts, rocky coasts, as well as toxic spoil heaps,
and have evolved to battle extreme humidity, temperature and light (Purvis, 2000). With
greater than 18,500 species known worldwide, lichens produce unique classes of
secondary metabolites that are rarely encountered in other organisms and, to date, there
have been in excess of 800 new chemical entities reported from lichens (Boustie & Grube,
2005; Huneck & Yoshimura, 1996; Rundel, 1978). Structural classes range from the
more common phenolic compounds to the usnic acids, depsides, depsidones, depsones,
lactones and quinones, which are generally derived by the acetate-malonate pathway,
and the pulvinic acid derivatives, derived from the shikimic acid pathway (Boustie &
Grube, 2005; Rundel, 1978). There have been numerous reports of the pharmacological
properties of lichen substances including antibiotic, antiviral, anti-inflammatory and
*Corresponding author. Email: sylvia.urban@rmit.edu.au
ISSN 1478–6419 print/ISSN 1029–2349 online
ß 2009 Taylor & Francis
DOI: 10.1080/14786410802682536
http://www.informaworld.com

926
D.A. Dias and S. Urban
cytotoxic effects (Muller, 2001). One well-known pulvinic acid derivative is vulpinic acid
(4), which was discovered as early as 1939, and reported to display acute dyspnoea as
well as causing an increase in the rate of respiration and salivation in cats (Santesson,
1939). Progressive developments in separation and purification strategies, as well as
advancements in the sensitivity of instrumentation, have resulted in the ability to rapidly
isolate and identify lichen constituents (Culberson & Culberson, 2001). As early as
the 1900s pioneers in lichen natural products chemistry used microcrystal tests,
ultraviolet spectroscopy (UV) and paper chromatography to identify lichen metabolites.
The 1960s made more use of techniques such as infrared spectroscopy (IR), circular
dichroism (CD), and sensitive techniques such as mass spectrometry (MS), for
compound identification. It was the late 1980s which saw improvements in chromato-
graphic techniques including high pressure liquid chromatography (HPLC), gas
chromatography (GC), together with advancements in spectroscopic techniques such
as nuclear magnetic resonance (NMR) spectroscopy, single-crystal X-ray diffraction
analysis, as well as improved sensitivity in mass spectrometry techniques, coupled with
synthetic techniques to confirm absolute structures (Culberson & Culberson, 2001).
These improvements have ultimately enabled far more rapid isolation and structure
elucidation. One of the more powerful spectroscopic techniques in the identification of
organic compounds is NMR spectroscopy and presently advancements in probe
technology, such as cold flow-probes, higher field strength magnets as well as the
development of superior 2D NMR experiments have provided new opportunities to fully
characterise lichen constituents. This is particularly important for lichen metabolites,
which had previously only been partially assigned, since much less compound is now
required for complete structural assignment.
As part of the continuing activities of The Marine And Terrestrial NAtural Product
(MATNAP) research group at RMIT University (Melbourne, Victoria, Australia), which
includes the study of Australian lichens, we focused our efforts on the chemical profiling
of a yellow lichen. Recently we conducted the chemical investigation of a distinctly
bright yellow lichen for which we were only able to identify the alga component of the
lichen as belonging to a Treboxia sp. (Dias, White, & Urban, 2007). We now wish to
report the full identification of this lichen as Chrysothrix xanthina (Vain.) Kalb. This
lichen had afforded pinastric acid (2), which presented a significant structural challenge
as NMR experiments alone could not unequivocally assign the central lactone ring
fragment (Dias et al., 2007). Fortuitously, a single X-ray analysis of 2 could be obtained
to further confirm the structure of 2. As a result of this study we were prompted to
investigate another yellow lichen, identified as Candelaria concolor (Dickson) B. Stein,
collected from Merlynston, Victoria, Australia. To date, reports on the lichen
C. concolor have been restricted to biomonitoring studies (Loppi & Frati, 2006; Pis
ˇ
u´ t
& Pis t, 2006; Purvis, 2000; Ruisi et al., 2005). The crude extract of the lichen
ˇ
u´
C. concolor demonstrated significant antibacterial activity, which prompted the isolation
of several lichenic compounds, including pulvinic dilactone (1), pulvinic acid (3)
(prepared from 1), vulpinic acid (4), calycin (5), atranorin (6), and 5-chloroatranorin (7),
as well as the first reported isolation of 5,50-dichloroatranorin (8). In addition, we report
the first complete structural characterisation involving the use of 2D NMR spectroscopy
for several previously isolated compounds, including 1 and 3–8. Furthermore, the
importance of the selection of the extraction solvent, as well as the chemical degradation
of 6, is also reported.

Natural Product Research
927

928
D.A. Dias and S. Urban
2. Results and discussion
2.1. Isolation of vulpinic acid (4) and calycin (5)
The yellow lichen was extracted with 3 : 1 MeOH/CH2Cl2 and then separated by flash
silica chromatography to yield vulpinic acid (4) (also known as methylpulvinic acid).
Vulpinic acid (4) has been known since the 1830s, when early Eskimos used lichens
containing the acid to poison wolves, and in Scandinavia the lichen containing this
compound was used to kill both wolves and foxes (Knight & Pattenden, 1979; Purvis,
2000). The structure of 4 has been previously confirmed by partial synthesis (Frank,
Clark, & Coker, 1950) with the total synthesis achieved in the 1950s (Frank et al., 1950;
Moore, Shelden, Deters, & Wikholm, 1969; Pattenden, Pegg, & Kenyon, 1991; Ramage
& Griffiths, 1984). There have been several reports of the single crystal X-ray structure
of 4, previously isolated from the lichen Letharia vulpina (L.) Hue, with the first crystal
structure being reported in 1985 (Brassy, Bachet, Molho, & Molho, 1985). Vulpinic acid
(4) and its crystal structure has also been reported from the New Zealand liverwort,
T. tenellus, for which 4 was the major constituent (Toyota, Omatsu, Braggins, &
Asakawa, 2004). In 2003 the isolation of 4 and its polymorph from P. ravenelii were

Natural Product Research
929
described (Duncan et al., 2003). There have been a number of attempts to
unambiguously assign the NMR data for 4, but owing to the number of quaternary
carbons present in this molecule, the unequivocal assignment of these resonances has
been difficult, as they are often not even observed. To date there have not been any
reports of the complete 1D and 2D NMR assignment of a majority of the compounds
isolated in this study including 4 (Heurtaux, Lion, Gall, & Mioskowski, 2005; Konig &
¨
Wright, 1999; Patteneden, Pegg, & Kenyon, 1991). This prompted a complete
unequivocal NMR assignment of all proton and carbon chemical shifts on the basis
of 2D NMR connectivity experiments for all compounds isolated from C. concolor.
The complete structure of 4 was determined by detailed spectroscopic analyses involving
both 1D and 2D NMR experiments and mass spectrometry. The ESI mass spectrum
displayed a m/z 321 [MꢀH]ꢀ, consistent with a molecular formula of C19H14O5 and 13
DBE. The 13C NMR spectrum of 4 showed 15 discrete signals and the DEPT and
HSQCAD NMR experiments supported the presence of one methyl, six methines and eight
quaternary carbons. The IR spectrum of 4 displayed absorptions at 3426 and 1774 cmꢀ1
,
indicative of the hydroxy and ester carbonyl moieties. The molecular formula, together with
the observation of only 15 distinct carbon signals in the 13C NMR spectrum, immediately
suggested an element of symmetry in 4 with four of the methines being equivalent. Analysis
of the 1H NMR and the gCOSY NMR spectra of 4 suggested the presence of two isolated
mono-substituted aromatic rings (the first at ꢀ 7.27 dd, J ¼ 2.0, 8.0 Hz, (H2/H6); 7.34 dd,
J ¼ 7.0, 7.5 Hz, (H3/H5) and 7.44, m, (H4) and the second at ꢀ 8.13 dd, J ¼ 1.5, 8.5 Hz, (H20/
H60); 7.41 m, (H30/H50) and 7.42 m, (H40)). The presence of a methyl ester (ꢀ 3.89 s,
1
54.4 ppm) was supported by H, 13C and HSQCAD NMR experiments and a HMBC
correlation (171.7 (C12) ppm) confirmed it to be attached to a carbonyl moiety. Important
HMBC correlations were observed from aryl protons on both rings, which ultimately
allowed for the placement of the two mono-substituted aromatic rings. In particular, the
second mono-substituted aromatic ring showed an important correlation to 105.1 (C10)
ppm, thereby confirming the position of this ring, relative to the central lactone ring moiety.
Owing to the number of quaternary carbons in the central hydroxyfuranone core and the
lack of any HMBC correlations to these carbons, due to their remoteness from the aromatic
protons, the remaining quaternary carbons at ꢀ 165.9 (C11), 160.2 (C9) and 154.8 (C8) were
assigned on the basis of a direct comparison to the carbon shifts previously assigned for
pinastric acid (2) (Dias et al., 2007).
Analysis of another fraction resulting from the same flash silica column fractionation
step yielded a mixture of vulpinic acid (4) and calycin (5). This mixture was further purified
by additional silica chromatography and resulted in the isolation of 5 as an orange oil.
Calycin (5) had been previously isolated in 1890 as a deep red compound from the lichen
˚
L. candelaris Shaer (Akermark, 1961). Compounds 4 and 5 were also isolated from the
lichen L. clorina Stein (Grover & S.T.R., 1959). The structure of 5 had initially been
reported as 2-hydroxypulvic dilactone (9). It was not until the early 1960s that the
synthesis of 5 was proposed, for which the correct structure 5 was supported by infrared
::
˚
ꢀ
and ultraviolet spectroscopy (Akermark, 1961; Kuhler, Nilsson, Sandberg,
Wachtmeister, 1984).
&
The structure of calycin (5) was again confirmed using 1D and 2D NMR as well as by
mass spectrometry. The ESI mass spectrum displayed a m/z 305 [MꢀH]ꢀ, which suggested
a molecular formula of C18H10O5 and 13 DBE. The 13C NMR spectrum of 5 contained 15
signals (nine methines (two of these being equivalent) and eight quaternary carbons (one
not being detected)), as supported by a gHSQCAD NMR experiment. The IR spectrum of

930
D.A. Dias and S. Urban
5 showed absorptions at 3401 and 1804 cmꢀ1, once again being suggestive of hydroxy and
1
ester carbonyl moieties. The H NMR and gCOSY NMR spectra again identified the
presence of two isolated aromatic ring systems. These included a mono-substituted
aromatic ring (ꢀ 8.19 d, J ¼ 7.5 Hz, (H20/H60); 7.47 m, (H30/H50); 7.39 dd, J ¼ 7.5, 7.5 Hz,
(H40)) and a di-substituted aromatic ring (ꢀ 7.98 d, J ¼ 7.5 Hz, (H3); 7.45, m, (H5); 7.32 dd,
J ¼ 7.5, 8.0 Hz, (H4) and 7.23 d, J ¼ 8.0 Hz, (H6)). A key HMBC correlation from ꢀ 7.98
(H3) to the carbon at 107.0 (C8) ppm unequivocally positioned the benzofuranone moiety
adjacent to the hydroxyfuranone core, whilst another HMBC correlation positioned the
other mono-substituted aromatic ring (ꢀ 8.19 (H20/H60) to the carbon at 105.9 (C11) ppm).
Positions C10 and C12 were assigned on the basis of comparisons to carbon shifts
previously assigned to 2 (Dias et al., 2007).
2.2. Identification of atranorin (6), 5-chloroatranorin (7) and the isolation of
5,50-dichloroatranorin (8)
In an effort to separate 4 and 5 by HPLC, the 3:1 MeOH/CH2Cl2 crude extract was further
sequentially partitioned using CH2Cl2 followed by MeOH. The MeOH partition was then
subjected to reversed-phase HPLC, which resulted in the isolation of atranorin (6).
Surprisingly, 4 and 5 were not detected in the MeOH partition. However, the presence of
4 and 5 were detected in the CH2Cl2 partition, as confirmed by analytical HPLC.
Atranorin (6) was first isolated in 1898 and reported to be always accompanied by 4 in the
lichen L. vulpina (Hesse, 1898; Solberg & Remedios, 1978). In addition, this ꢁ-orcinol
depside (6) has been described as being a significant taxonomic marker for various species
of lichens that possess ecological and biological properties (Elix, Whitton, & Jones, 1982;
Elix, 1993; Rundel, 1978). It was not until 1999 that the re-isolation together with the
complete 1D and 2D NMR for 6 was described (Huneck & Yoshimura, 1996; Konig &
Wright, 1999).
¨
In the present isolation of atranorin (6), the structure was confirmed on the basis of 1D
and 2D NMR experiments and mass spectrometry. The ESI mass spectrum showed a m/z
1
373 [MꢀH]ꢀ, consistent with a molecular formula of C19H18O8 and 11 DBE. The H
NMR, gHSQCAD and gHMBC NMR spectra supported the assignments of 6 as reported
in the literature (Konig & Wright, 1999). During acquisition of the NMR data for 6 it was
¨
observed that changes occurred in the 1H NMR spectrum (in particular, the disappearance
1
of the H NMR signal at ꢀ 6.41 s, (H5)) after approximately 16 h. Clearly a chemical
conversion was occurring and this could be monitored by the acquisition of 1D and 2D
NMR experiments as well as ESIMS. The NMR data acquired, together with the
supporting ESIMS data, identified the presence of a m/z 407 [MꢀH]ꢀ (molecular formula
of C19H17ClO8 and 11 DBE) and confirmed the presence of the depside 5-chloroatranorin
(7). Due to the rapid conversion of 6 to 7, only a gHSQCAD NMR experiment could be
acquired for 7 within the time it took for it to further convert to 8. This permitted the
observation of four methyls (ꢀ 2.86 s (H9); 2.09 s (H80); 2.54 s (H90); and 3.99 s (H100)) and
one deshielded methine (ꢀ 6.52 s (H50)). The methine proton at (C5) was absent as it had
now been replaced by a chlorine substituent, as confirmed by the isotopic ratio observed in
the ESIMS. The identification of 5-chloroatranorin (6) dates back to as early as the 1930s
(Pfau, 1933). It was postulated, that in the biogenesis of chlorodepsidones, a depside may
undergo halogenation and that the halogenated derivative can then undergo ring closure
by the elimination of hydrogen halides (Neelakantan, Padmasani, & Seshadri, 1964).

Natural Product Research
931
It was not until 1964 that the halogenation of 6 was reported, together with the
chlorination of lecanoric acid (10), to 5-chlorolecanoric acid (11) (Neelakantan et al., 1964;
Neelakantan, Seshadri, & Subramanian, 1962).
During the NMR data acquisition of 7 (approximately 14 h post-acquisition), it was
1
observed that the H NMR spectrum of 7 showed the disappearance of the signal at
(ꢀ 6.52 s, (H50)) and that all other observed signals, which had been evident for 7 were now
1
slightly shifted further downfield. Analysis of the H NMR, gHSQCAD and gHMBC
NMR data identified the presence of 17 carbons (one methine, four methyls and twelve
quaternary carbons (two not being detected)). The IR spectrum of 8 once again showed
absorptions at 3401 and 1735 cmꢀ1, suggestive of hydroxy and ester carbonyl moieties.
The ESIMS displayed a m/z 441 [MꢀH]ꢀ, consistent with a molecular formula of
C19H16Cl2O8 and 11 DBE. Key HMBC correlations included one from the methyl group
at ꢀ 2.65 (H90) to the carbon bearing the chlorine substituent at 118.4 (C50) ppm. This
enabled the position of the chlorine substituent to be deduced as being adjacent to the
bridging ester at (C40). The methyl group at ꢀ 2.13 (H80) also showed key HMBC
correlations to 148.8 (C40), 159.6 (C20) and 119.1 (C30) ppm. Furthermore, the methyl
group ꢀ 2.92 (H9) exhibited a three-bond HMBC correlation to the carbon containing the
second chlorine at 115.6 (C5) ppm. The hydroxy group at ꢀ 12.22 (2-OH) displayed a
two-bond HMBC correlation to 166.7 (C2) ppm and allowed the carbon (109.1 (C3) ppm)
bearing the aldehyde substituent to be positioned. Finally, the methyl ester at ꢀ 4.01 (H100)
showed a three-bond HMBC correlation to the ester carbonyl at 171.2 (C70) ppm). These
1
key HMBC correlations and similar H and 13C chemical shifts observed for 6 and 7
supported the structure of the depside, 5,50-dichloroatranorin (8) (Table 1 and Figure 1).
Table 1. 1H (500 MHz) and 13C (125 MHz) NMR data of (8) in CDCl3.
Position
ÁH, J (Hz)
ꢀC
gHMBC and CIGAR
1
2
3
4
–
–
–
–
103.6, s
166.7, s
109.1, s
ND
–
–
–
–
5
6
7
8
–
–
115.6, s
149.1, s
ND
–
–
–
–
10.38, s
2.92, s
–
193.9, d
21.5, q
111.9, s
159.6, s
119.1, s
148.8, s
118.4, s
136.6, s
171.2, s
10.5, q
19.8, q
53.1, q
–
–
C1, C5, C6
9
10
–
–
–
–
–
–
–
20
–
–
–
–
–
30
40
50
60
70
–
80
2.13, s
2.65, s
4.01, s
12.22, s
11.52, s
ND
C20, C30, C40
C10, C50, C60
C70
C1, C2, C3
C10, C20
–
90
100
2-OH
20-OH
4-OH
–
–
Note: ND indicates signal was not detected.

932
D.A. Dias and S. Urban
HO
H
CH
O
3
CH
CH
3
O
OH
O
O
OH
Cl
CH
3
O
3
HO
H
CH
O
3
CH
3
O
OH
O
O
OH
Cl
CH
3
O
CH
3
HO
H
CH
O
3
CH
3
O
OH
O
OH
O
CH
Cl
3
O
CH
3
Figure 1. Chemical conversion of 6 to 7 and then finally to 8 in CDCl3 in situ (mild conditions).
The first report of 8 appeared in 1964, where the halogenation of depsides was
described (Neelakantan et al., 1964). This study reported the halogenation of 6 to 7 and
then 7 to 8, using Cl2 (2 moles). Methanolysis of 8 produced two fission products,
5-chlorohaematommate (12) and methyl-5-chloro-ꢁ-orcinol carboxylate (13), which were
then used to deduce the structure of the chlorinated depside. At the time,
5,50-dichloroatranorin (8) had not been isolated or fully characterised. This represents
the first isolation of 8 along with its detailed chemical characterisation.
2.3. Isolation of pulvinic dilactone (1) and the conversion to pulvinic acid (3)
The identification of 1 was previously reported in 1883, while the methanolysis of 1 to 3
had been reported in the late 1950s (Frank et al., 1950). HPLC profiling led us to believe
that 4 and 5 were in fact the methanolysis products resulting from the initial extraction
step. In an effort to confirm this, a separate extraction of C. concolor was carried out using
1
CH2CH2, resulting in almost 95% pure (on the basis of H NMR spectroscopy) pulvinic
dilactone (1). In terms of the chemical profiling of lichens, selection of the extraction
solvent is crucial in correctly determining the ‘true’ lichen constituents present and should
be ideally performed using a selection of solvents.
Pulvinic dilactone (1) was isolated as yellow–green oil. The ESI mass spectrum showed
the presence of m/z 291 [MþH]þ, consistent with a molecular formula of C18H10O4 and 14
DBE. The 13C NMR spectrum of 1 contained seven discrete signals. The highly

Natural Product Research
933
symmetrical nature of 1 enabled the presence of 10 methines and 8 quaternary carbons to
be concluded, as supported by a gHSQCAD NMR experiment. The 1H NMR and gCOSY
NMR experiments identified the presence of two ortho-coupled aromatic methines
(ꢀ 8.04 d, J ¼ 8.0 Hz, (H5/H50/H9/H90)) and (ꢀ 7.50 dd, J ¼ 7.0, 7.5 Hz (H6/H60/H8/H80))
that in turn showed a COSY correlation to an aromatic methine at ꢀ 7.44 dd, J ¼ 7.0,
6.5 Hz, (H7/H70). The only key HMBC correlation was that observed from the protons at
ꢀ 8.04, (H5/H50/H9/H90) to 101.4 (C2/C20) ppm, which enabled the positioning of these
aromatic protons as being adjacent to the furanone. Positions C1, C10 and C3, consistent
with ester carbons, were assigned on the basis of carbon chemical shift comparisons to
those previously assigned in compounds 2, 4 and 5.
The conversion of 1 to pulvinic acid (3) was carried out, and to the best of our
knowledge, no previous allocation of the complete 1H and 13C NMR assignments of 3 has
been previously reported. The first report of pulvinic acid (3) dates back to 1894 (Volhard,
1894) including a report of the conversion of 1 to 3 by acid hydrolysis (Frank et al., 1950).
The synthesis of 3 was carried out in CDCl3, where a solution of permethylated pulvinic
acid, treated with iodotrimethylsilane, upon sealing the NMR tube under N2 was heated to
55ꢁC (Pattenden et al., 1991). After 3 days the reaction mixture was hydrolysed with
1
MeOH to afford 3, which was characterised by H and 13C NMR, but the complete
assignment of the NMR data as well as the structure remained unassigned (Pattenden
et al., 1991).
In the present study, pulvinic dilactone (1) was heated with aqueous acetone for
15 min, resulting in the isolation of 3 (in 495% purity based on 1H NMR spectroscopy).
The structure of 3 was confirmed by 1D and 2D NMR experiments, as well as mass
spectrometry, and the structure found to be similar to 4, differing only in the presence of a
carboxylic acid at C12. The negative mode ESI mass spectrum showed the presence of a
m/z 307 [MꢀH]ꢀ (molecular formula of C18H12O5 and 13 DBE). The IR spectrum of 3
displayed absorptions at 3307 (broad) and 1611 cmꢀ1, suggestive of carboxylic and ester
carbonyl moieties. The 1H NMR spectrum, together with the gCOSY NMR identified the
presence of two isolated mono-substituted aromatic rings (ꢀ 7.33 m (H4); 7.29, m (H3/H5)
and 7.19 d, J ¼ 7.0 Hz, (H2/H6) and ꢀ 8.14 d, J ¼ 7.0 Hz (H20/H60); 7.30, m (H30/H50) and
7.12, dd, J ¼ 7.0, 7.5 Hz (H40)). Diagnostic HMBC correlations were observed from aryl
protons on both rings (ꢀ 8.14 (H20/H60) to 95.7 (C10) ppm) and (ꢀ 7.19 (H2/H6) to 118.3
(C7) ppm), which enabled them to be positioned accordingly. As HMBC correlations were
not observed for the exchangeable protons, the remaining quaternary carbon shifts were
assigned by comparison to those previously assigned in 2, 4 and 5, for which 2 was
assigned by a single crystal X-ray analysis (Dias et al., 2007). Figure 2 displays the
conversion of 1 to 4 and 5, as well as the conversion of 1 to 3.
Compounds 1, 3–5 and 8 were all found to display insignificant activity against the
P388 murine leukaemia cell line (IC50412,500 ng mLꢀ1 when tested at 1 mg mLꢀ1).
Pulvinic dilactone (1) moderately inhibited the growth of Bacillus subtilis, while
compounds 3–5 and 8 only showed slight inhibition against B. subtilis. Compounds 6
and 7 were unable to be tested due to their instability and ultimate conversion to 8.
The medicinal properties of lichens dates back to folklore. Hippocrates recommended
Usnea barbata for uterine trouble, and in the 15th century, Lobaria pulmonaria was used
for the treatment of catarrhal haemoptysis and pulmonary tuberculosis (Vartia, 1973). In
Finnish folklore, lichens have been used to treat athlete’s foot and oral doses have been
used to treat coughs, usually taken in the form of so-called ‘lichen milk’ (Vartia, 1973). The
antibacterial activity of lichen acids has been reported as early as the 1940s, where it was

934
D.A. Dias and S. Urban
O
O
O
H3CO
O
HO
OH
O
MeOH
and
O
O
O
O
O
O
(1)
(4)
(5)
80% aqueous acetone (heat, 70°C, 15 min)
HO
O
OH
O
O
(3)
Figure 2. Chemical conversion of 1 to 4 and 5 using MeOH as the extraction solvent and the
preparation of 3 from 1, highlighting the importance of extraction solvent selection.
demonstrated that compounds 4–6 displayed antibiotic activity against Gram-positive
bacteria (Krogg, 1954). Around the same time, it was concluded that 1 displayed inotropic
effects in the same order of concentration as that necessary for the cardiac glycosides,
ouabain and digitoxin (Giarman, 1949). In 1991 it was demonstrated that 4 inhibited the
growth of B. subtilis (MIC of 15.8 mg mLꢀ1) (Nadir, Rashan, Ayoub, & Awni, 1992). Other
reported activities for 4 include anti-inflammatory, inhibition of influenza RNA viruses,
analgesic, local anaesthetic activity and toxicity effects (Rao & Prabhakar, 1988; Foden,
McCormick, & O’Mant, 1975; Rashan, Ayoub, Al-Omar, & Al-Khayatt, 1990). In
another study, 4 displayed activity against the fungus U. violacea, M. microspora and
E. repens, and the bacteria B. megaterium, whereas 6 displayed activity only against
E. repens (Konig & Wright, 1999).
¨
3. Experimental
3.1. General experimental procedures
All organic solvents used were analytical reagent (AR or GR), UV spectroscopic or
HPLC grades with milli-Q water also being used. IR spectra were recorded as a film
using a NaCl disk on a Perkin-Elmer, Spectrum One FTIR spectrometer. UV/vis spectra
were recorded on a Varian CARY 50 Bio spectrophotometer, using ethanol. In addition,
a UV profile was obtained from the HPLC (PDA detection) by extraction of the 2D
1
contour plot. H (500 MHz) and 13C (125 MHz) NMR spectra were acquired in CDCl3
or DMSO-d6 on a 500 MHz Varian INOVA spectrometer with referencing to solvent
signals (ꢀ 7.26, 77.0 ppm and ꢀ 2.50, 39.5 ppm, respectively). 1D and 2D NMR
experiments included gCOSY, gHSQCAD gHMBC and constant time inverse detection
gradient accordian rescaled (CIGAR) experiments. ESI mass spectra were obtained on a
Micromass Platform II mass spectrometer equipped with a LC-10AD Shimadzu solvent
delivery module (50% CH3CN/H2O at a flow rate of 0.1 mL minꢀ1) in both the positive

Natural Product Research
935
and negative ionisation modes, using cone voltages between 20 and 30 V. HRESIMS
was carried out on an Agilent 6200 Series TOF system (ESI operation conditions of
8 L minꢀ1 N2, 350ꢁ drying gas temperature and 4000 V capillary voltage), equipped with
an Agilent 1200 Series LC solvent delivery module (100% MeOH at a flow rate of
0.3 mL minꢀ1) in the negative ionisation mode (in all cases the instruments were
calibrated using the ‘Agilent Tuning Mix’ using purine as the reference compound). TLC
was performed on pre-coated aluminium backed silica TLC plates (MerckTM silica gel 60
F254) using the solvent system 65 : 25 : 4 (CHCl3 : MeOH : H2O) or 100% (EtOAc),
visualised at 254 and 365 nm and further developed using A: iodine vapour and B: a
ninhydrin dip consisting of 0.3 g ninhydrin in 100 mL of n-butanol and 3 mL acetic acid.
Silica flash chromatography was carried out with MerckTM; silica gel (60 mesh) using
nitrogen and a 50% stepwise solvent elution from 100% hexane to 100% CH2Cl2 to
100% EtOAc and finally to 100% MeOH. All analytical HPLC analyses were performed
on a Dionex P680 solvent delivery system equipped with a PDA100 UV detector
(operated using ‘Chromeleon’ software). Analytical HPLC analyses were run using either
a gradient method (0 min 10% CH3CN/H2O; 2 min 10% CH3CN/H2O; 14 min 75%
CH3CN/H2O; 24 min 75% CH3CN/H2O; 26 min 100% CH3CN; 30 min 100% CH3CN;
32 min 10% CH3CN/H2O and 40 min 10% CH3CN/H2O) or an isocratic method (100%
˚
CH3CN) on a Phenomenex Prodigy ODS (3) C18 100A 250 ꢂ 4.6 (5 m) and on a
˚
Phenomenex Luna ODS (3) C18 100A 250 ꢂ 4.6 (5 m) column at a flow rate of
1.0 mL minꢀ1. All semi-preparative HPLC was carried out on a Varian Prostar 210
(Solvent Delivery Module) equipped with a Varian Prostar 335 PDA detector using
STAR LC WS Version 6.0 software using an isocratic method (100% CH3CN) and a
˚
Phenomenex Prodigy ODS (3) 100A C18 250 ꢂ 10 (5 m) column at a flow rate of
3.5 mL minꢀ1
.
3.2. Biological evaluation and details of assays
Extracts of C. concolor (Dickson) B. Stein were evaluated against a P388 Murine
Leukaemia cell line (antitumour assay), as well as against a number of bacteria and fungi
(antimicrobial assays) at the University of Canterbury, Christchurch, New Zealand. Only
moderate antitumour activity was observed for the crude C. concolor extract
(IC50 31,776 ng mLꢀ1 at 10 mg mLꢀ1). The C. concolor extract displayed significant
antibacterial activity against B. subtilis. No activity was observed against Eschericha coli,
Pseudomonas aeruginosa, Candida albicans, Trichophyton mentagrophytes or Cladosporium
resinae. For further details on the antitumour and antimicrobial assays, please see Dias
et al. (2007).
3.3. Lichen material
The yellow lichen was collected on 19 August 2007 near Merlynston Railway Station,
Victoria, Australia. The lichen and bark were chiselled off a eucalyptus tree and a small
(2 ꢂ 2 cm2) sample sent to Emeritus Prof. John A. Elix (Australian National University,
Canberra, ACT, Australia) for taxonomic identification and HPLC profiling. A voucher
specimen designated the code 2007-04 is deposited at the School of Applied Sciences
(Discipline of Applied Chemistry), RMIT University.

936
D.A. Dias and S. Urban
3.4. Extraction and isolation
In the first extraction of C. concolor, the lichen, together with the bark (170 g) was
extracted with 3 : 1 MeOH/CH2Cl2 (750 mL). The crude extract was decanted and
concentrated under reduced pressure. Approximately half of the crude extract was then
subjected to a flash silica column (50% stepwise elution from petroleum spirits to CH2Cl2
and finally to EtOAc) with the third fraction yielding pure vulpinic acid (4) (7.2 mg,
0.004%, based on mass of lichen and bark extracted). In addition, calycin (5) was
identified to be present in a mixture with 4. This mixture was subjected to further flash
silica chromatography using 100% EtOAc as the eluent, affording pure calycin (5) (1.5 mg
0.0008%, based on mass of lichen and bark extracted).
In a further effort to separate 4 and 5 by HPLC, the 3:1 MeOH/CH2Cl2 crude extract
was sequentially partitioned using CH2Cl2 followed by MeOH. The MeOH partition was
then subjected to reversed-phase HPLC using isocratic conditions (100% CH3CN with
detection at 215 and 254 nm) to yield atranorin (6) as a pale yellow oil (1.1 mg, 0.006%,
based on mass of lichen and bark extracted). It was immediately noted that atranorin (6)
rapidly converted to 7 and finally to a stable pale yellow oil, 8, over a period of 1–2 days in
CDCl3 in situ.
In a second extraction of the lichen, C. concolor, together with the bark (55 g), was
extracted with 100% CH2Cl2 (50 mL). The crude extract was decanted and concentrated
under reduced pressure. The entire extract was then subjected to flash silica
chromatography using 100% EtOAc as the eluent, to afford pure 1 (201.3 mg, 0.4%,
based on mass of lichen and bark extracted). In this instance, the purity of 1 was assessed
1
on the basis of the H NMR analysis.
The chemical conversion of 1 to 3 was demonstrated by heating 1 (75 mg) in 80%
aqueous acetone (100 mL) for 15 min, resulting in 3 (495% purity as assessed by 1H NMR
spectroscopy) (65.2 mg, 82.0% yield).
Pulvinic dilactone (1) [3,6-diphenylfuro[3,2-b]furan-2,5-dione]; isolated as a stable yellow–
green oil; IR (film) ꢂmax 3401, 2917, 1816, 1660, 1632 cmꢀ1; UV (EtOH) ꢃmax 205, 231, 285
and 375 nm (" ¼ 20900, 11100, 12000 and 6000, respectively); 1H NMR (500 MHz, CDCl3)
ꢀ 8.04, d, J ¼ 8.0 Hz (H5/H50/H9/H90); 7.50, dd, J ¼ 7.0, 7.5 Hz (H6/H60/H8/H80); 7.44, dd,
13
J ¼ 7.0, 6.5 Hz (H7/H70); C (125 MHz, CDCl3) 165.7 (C3/C30); 157.1 (C1/C10); 130.1
(C7/C70); 129.2 (C6/C60/C8/C80); 128.2 (C5/C50/C9/C90); 126.4 (C4/C40); 101.4 (C2/C20);
ESIMS (þve) (25 V) m/z 291 [MþH]þ.
Pulvinic acid (3) [(E)-2-(3-hydroxy-5-oxo-4-phenylfuran-2(5H)-ylidene)-2-phenylacetic
acid]; isolated as a dark brown oil; IR (film) ꢂmax 3307, 2917, 1739, 1611, 1578, 1563,
1398, 1146 cmꢀ1; UV (EtOH) ꢃmax 200, 260 and 365 nm (" ¼ 16,500, 17,000 and 8600,
1
respectively); H NMR (500 MHz, DMSO-d6) ꢀ 8.14, d, J ¼ 7.0 Hz (H20/H60); 7.33, m
(H4); 7.30, m (H30/H50); 7.29, m (H3/H5); 7.19, d, J ¼ 7.0 Hz (H2/H6); 7.12, dd, J ¼ 7.0,
7.5 Hz (H40), 9-OH and 12-OH not detected; 13C (125 MHz, DMSO-d6) 172.1 (C12); 169.3
(C11); 167.2 (C9); 153.1 (C8); 136.7 (C1); 133.3 (C10); 130.8 (C2/C6); 128.5 (C30/C50); 127.9
(C3/C5); 127.6 (C4); 125.8 (C20/C60); 125.7 (C40); 118.3 (C7); 95.7 (C10); ESIMS (ꢀve)
(25 V) m/z 307 [MꢀH]ꢀ.
Vulpinic acid (4) [(E)-methyl 2-(3-hydroxy-5-oxo-4-phenylfuran-2(5H)-ylidene)-2-pheny-
lacetate]; isolated as a stable, bright yellow oil; IR (film) ꢂmax 3426, 2919, 2850, 1774, 1677,
1612, 1598, 1440, 1289, 1279, 1065 cmꢀ1; UV (EtOH) ꢃmax 205, 290 and 375 nm (" ¼ 20400,
5500 and 7000, respectively); 1H NMR (500 MHz, CDCl3) ꢀ 9-OH not detected; 8.13, dd,

Natural Product Research
937
J ¼ 1.5, 8.5 Hz (H20/H60); 7.44, m (H4); 7.42, m (H40); 7.41, m (H30/H50); 7.34, dd, J ¼ 7.0,
7.5 Hz (H3/H5); 7.27, dd, J ¼ 2.0, 8.0 Hz (H2/H6), 3.89, s (13-OCH3); 13C (125 MHz,
CDCl3) 171.7 (C12); 165.9 (C11); 160.2 (C9); 154.8 (C8); 131.9 (C1); 129.9 (C2/C6); 128.9
(C10); 128.6 (C4); 128.4 (C30/C50); 128.3 (C40); 128.1 (C3/C5); 127.8 (C20/C60); 115.8 (C7);
105.1 (C10); 54.4 (13-OCH3); ESIMS (ꢀve) (25 V) m/z 321 [MꢀH]ꢀ.
Calycin (5) [(E)-3-(3-hydroxy-5-oxo-4-phenylfuran-2(5H)-ylidene)benzofuran-2(3H)-one];
isolated as a stable orange oil; IR (film) ꢂmax 3401, 2922, 2852, 1804, 1709, 1641, 1473,
1442, 1343, 1040 cmꢀ1; UV (EtOH) ꢃmax 202, 265, 365 and 455 nm (" ¼ 11,200, 10,500,
1
6100 and 3000, respectively); H NMR (500 MHz, CDCl3) ꢀ 12.60, br s (10-OH); 8.19, d,
J ¼ 7.5 Hz (H20/H60); 7.98, d, J ¼ 7.5 Hz (H3); 7.47, m (H30/H50); 7.45, m (H5); 7.39, dd,
J ¼ 7.5, 7.5 Hz (H40); 7.32, dd, J ¼ 7.5, 8.0 Hz (H4); 7.23, d, J ¼ 8 Hz (H6); 13C (125 MHz,
CDCl3) 173.1 (C7); 165.4 (C12); 160.0 (C10); 153.6 (C1); 152.8 (C9); 131.4 (C5); 129.1
(C4’); 128.6 (C30/C50); 128.3 (C10); 128.0 (C20/C60); 125.8 (C3 and C4); 121.6 (C2); 111.1
(C6); 105.9 (C11), 107.0 (C8); ESIMS (ꢀve) (25 V) m/z 305 [MꢀH]ꢀ.
Atranorin (6) [3-hydroxy-4-(methoxycarbonyl)-2,5-dimethylphenyl 3-formyl-2,4-dihy-
droxy-6-methylbenzoate]; isolated as an unstable colourless oil which rapidly converted
1
to 7 and then finally to the stable compound 8; H NMR (500 MHz, CDCl3) 12.55, br s
(OH); 12.50, br s (OH); 11.95, br s (OH); ꢀ 10.36, s (H8); 6.52, s (H50); 6.41, s (H5); 3.99, s
(10-OCH3); 2.69, s (9-CH3); 2.55, s (90-CH3); 2.09, s (80-CH3); 13C (125 MHz, CDCl3)
ꢀ 193.8 (C8); 172.7 (C70); 163.2 (C20); 152.6 (C6); 152.0 (C40); 140.4 (C60); 117.1 (C30); 116.1
(C50); 112.9 (C5); 110.9 (C10); 103.2 (C1); 52.7 (100-OCH3); 25.6 (9-CH3); 24.2 (90-CH3); 9.7
(80-CH3); ND (C2); ND (C3); ND (C4); ND (C7); ESIMS (ꢀve) (25 V) m/z 373 [MꢀH]ꢀ.
5-chloroatranorin (7) [3-hydroxy-4-(methoxycarbonyl)-2,5-dimethylphenyl 3-chloro-5-
formyl-4,6-dihydroxy-2-methylbenzoate]; isolated as an unstable colourless oil resulting
from chemical degradation of 6; partial identification 1H NMR (500 MHz, CDCl3)
ꢀ 12.33, br s (OH); 11.96, br s (OH); ꢀ10.36, s (H8); 6.52, s (H50); 3.99, s (10-OCH3); 2.86, s
(9-CH3); 2.54, s (90-CH3); 2.09, s (80-CH3) other OH not detected; partial identification
13
based on gHSQCAD; C (125 MHz, CDCl3) ꢀ 193.9 (C8); 116.2 (C50); 52.6 (100-OCH3);
24.3 (90-CH3); 21.3 (9-CH3); 9.9 (80-CH3); with all remaining quaternary carbons not
detected; ESIMS (ꢀve) (25 V) m/z 407 [MꢀH]ꢀ.
5,50-dichloroatranorin (8) [2-chloro-5-hydroxy-4-(methoxycarbonyl)-3,6-dimethylphenyl
3-chloro-5-formyl-4,6-dihydroxy-2-methylbenzoate]; isolated as a stable colourless oil as
the final product from the degradation of (6); IR (film) ꢂmax 3401, 2924, 2853, 1735, 1648,
1452, 1302, 1249, 1073 cmꢀ1; UV (MeOH) ꢃmax 285 and 375 nm (" ¼ 4600 and 1500,
1
respectively); H NMR (500 MHz, CDCl3) and 13C (125 MHz, CDCl3) NMR data see
Table 1; ESIMS (ꢀve) (25 V) m/z 441 [MꢀH]ꢀ; HRESIMS m/z 441.0143 ([MꢀH]ꢀ;
calculated for C19H15Cl2O8, 441.0222).
Acknowledgements
The Marine And Terrestrial NAtural Product (MATNAP) Research Group would like to thank
Emeritus Prof. John A. Elix (Department of Chemistry, Australian National University, Canberra,
Australia) for the identification of the C. xanthina (Vain.) Kalb and the C. concolor (Dickson) B.
Stein lichens, HPLC profiling of the lichen extracts and for his invaluable knowledge of Australian
lichens; Ms Gill Ellis (University of Canterbury, Christchurch, New Zealand) for biological testing;
Ms Sally Duck (School of Chemistry, Faculty of Science, Monash University) for high resolution

938
D.A. Dias and S. Urban
mass spectrometry analyses, and the assistance of Mr Alick Dias in the collection of the lichen
C. concolor (Dickson) B. Stein.
References
˚
Akermark, B. (1961). Studies on the chemistry of lichens XIV. Structure of calycin. Acta Chemica
Scandinavica, 15, 1695–1700.
Boustie, J., & Grube, M. (2005). Lichens – a promising source of bioactive secondary metabolites.
Plant Genetic Resources, 3, 273–287.
Brassy, P.C., Bachet, B., Molho, L., & Molho, D. (1985). ꢄ-(hydroxy-3-oxo-5-phenyl-4-2H-
´ ´ ´ ´
furylidene-2)-phenylacetate de meethyle, acide vulpinique, C19H14O5. Acta Crystallographica,
C41, 781–783.
Culberson, C.F., & Culberson, L.C. (2001). Future directions in lichen chemistry. The Bryologist,
104, 230–234.
Dias, D., White, J.M., & Urban, S. (2007). Pinastric acid revisited: A complete NMR and X-ray
structure assignment. Natural Product Research, 21, 366–376.
Duncan, C.J.G., Cuendet, M., Fronczek, F.R., Pezzuto, J.M., Mehta, R.G., Hamann, M.T., &
Ross, S.A. (2003). Chemical and biological investigation of the fungus Pulveroboletus ravenelii.
Journal of Natural Products, 66, 103–107.
Elix, J.A. (1993). Progress in the genetic delimitation of Parmelia sensu lato lichens (Ascomycotina:
Parmeliaceae) and a synoptic key to Parmeliaceae. Bryologist, 96, 359–838.
Elix, J.A., Whitton, A.A., & Jones, A.J. (1982). Triterpenes from the lichen genus Physcia.
Australian Journal of Chemistry, 35, 641–647.
Foden, F.R., McCormick, J., & O’Mant, D.M. (1975). Vulpinic acids as potential antiinflammatory
agents. 1. Vulpinic acids with substituents in the aromatic rings. Journal of Medicinal
Chemistry, 18, 199–203.
Frank, R.L., Clark, G.R., & Coker, J.N. (1950). The synthesis of vulpinic acid from polyporic acid.
Journal of the American Chemical Society, 72, 1824–1826.
Giarman, N.J. (1949). Antibiotic lactones and synthetic analogs II. Cardiotonic effects on the
isolated frog heart. Journal of Pharmacology and Experimental Therapeutics, 96, 119–134.
Grover, P.K., & Seshadri, T.R. (1959). Chemical investigation of Indian lichens: Part XXIII –
Imperfect lichens. Journal of Scientific and Indutrial Research, 18B, 238–240.
Hesse, O. (1898). Contribution to the knowledge of the lichens and their characteristic components.
Journal fuer Praktische Chemie (Leipzig), 57(2), 232–318.
Heurtaux, B., Lion, C., Le Gall, T., & Mioskowski, C. (2005). Uncatalyzed reaction of silyl ketene
acetals with oxalyl chloride: A straightforward preparation of symmetrical pulvinic acids. The
Journal of Organic Chemistry, 70, 1474–1477.
Huneck, S., & Yoshimura, Y. (1996). Identification of lichen substances. Berlin, Heidelberg,
New York: Springer.
Knight, D.W. & Pattenden, G. (1979). Syntheses of permethylated derivatives of pinastric acid and
gomphidic acid, pulvinic acid pigments of lichen and fungi. Journal of the Chemical Society,
Perkin Transactions 1: Organic and Bio-Organic Chemistry (1972–1999), 84–88.
1
Konig, G.M., & Wright, A.D. (1999). H and 13C-NMR and biological activity investigations of
¨
four lichen-derived compounds. Phytochemical Analysis, 10, 279–284.
Krogg, H. (1954). Determination of the antibiotic effect of lichen acids. Det Kongelige Norske
Videnskabers Selskabs Forhandlinger, 27, 1–3.
::
0
ꢀ
Kuhler, T., Nilsson, M., Sandberg, U., & Wachtmeister, C.A. (1984). Synthesis of 2,2 -
dimethoxypulvic acid and enolic structure of its demethylation product, 2-hydroxycalycin.
Finnish Chemical Letters, (4–5), 112–115.
Loppi, S., & Frati, L. (2006). Lichen diversity and lichen transplants as monitors of air pollution in a
rural area of central Italy. Environmental Monitoring and Assessment, 114, 361–375.

Natural Product Research
939
Moore, W.H., Shelden, H.R., Deters, D.W., & Wikholm, R.J. (1969). Rearrangements of
azidoquinones V. Stereoselective acid-catalyzed rearrangements of azidoquinones to
ꢅ-cyanoalkylidine-(cyanoarylidine-)Áꢄ,ꢁ-butenolides. Journal of the American Chemical
Society, 92, 1675–1681.
Muller, K. (2001). Pharmaceutically relevant metabolites from lichens. Applied Microbiology and
Biotechnology, 56, 9–6.
Nadir, M.T. Rashan, L.J. Ayoub, M.T. & Awni, L.T. (1992). Antibacterial and antiproliferative
activities of vulpinic acids in vitro. Farmaco, 47, 643–647.
Neelakantan, S., Padmasani, R., & Seshadri, T.R. (1964). Halogenation of the depsides, lecanoric
acid & atranorin. Indian Journal of Chemistry, 2, 478–484.
Neelakantan, S., Seshadri, T.R., & Subramanian, S.S. (1962). Chemical investigation of Indian
lichens – XXVI. Tetrahedron, 18, 597–604.
Pattenden, G., Pegg, N.A., & Kenyon, R.W. (1991). Synthesis of grevillins, novel pyrandione
pigments of fungi. Biogenetic interrelationships between grevillins, pulvinic acids, terphenyl-
quinones and xylerythrins. Journal of the Chemical Society, Perkin Transactions, 2363–2372.
St. Pfau, A. (1933). Zur Kenntnis der Flechtenbestandteile III. Synthese der Hamatommsaure.
¨
¨
Helvetica Chimica Acta, 16, 282–286.
t, I., & Pis t, P. (2006). Changes of epiphytic lichens in the surroundings of magnesite factories
near Jels
Pis
ˇ
u
´
ˇ
u´
ˇ
´
ava (Se Slovakia) in the period 1973–2004. Ekologia, 25, 176–187.
Purvis, W. (2000). Lichens (pp. 1–112). Washington D.C.: The Natural History Museum.
Ramage, R., & Griffiths, G.J. (1984). Dioxolanes as synthetic intermediates. Part 3. Biomimetic
synthesis of pulvinic acids. Journal of the Chemical Society, Perkin Transactions, 1547–1552.
Rao, A.V.N., & Prabhakar, M.C. (1988). Pharmacological actions of vulpinic acid, a lichen
metabolite. Indian Journal of Chemistry, 20(2), 143–151.
Rashan, L.J., Ayoub, M.T., Al-Omar, L., & Al-Khayatt, R. (1990). Vulpinic acids inhibit influenza
(RNA) viruses but not herpes (DNA) viruses. World Journal of Microbiology & Biotechnology,
6, 155–158.
Ruisi, S., Zucconi, L., Fornasier, F., Paoli, L., Frati, L., & Loppi, S. (2005). Mapping environmental
effects of agriculture with epiphytic lichens. Israel Journal of Plant Sciences, 53, 115–124.
Rundel, P.W. (1978). The ecological role of secondary metabolites. Biochemical Systematics and
Ecology, 6, 157–170.
Santesson, C.G. (1939). Toxicity of letharia vulpina. Upsala Lakareforenings forhandlingar, 45, 1–8.
¨
¨
¨
Solberg, Y., & Remedios, G. (1978). Studies on the chemistry of lichens XVIII. Chemical
investigation of the species letharia vulpina (L.) Hue and new derivatives of vulpinic acid.
Journal of Biosciences, 33C, 449–451.
Toyota, M., Omatsu, I., Braggins, J., & Asakawa, Y. (2004). New humulane-type sesquiterpenes
from the liverworts Tylimanthus tenellus and Marchantia emarginata subsp. tosana. Chemical
and Pharmaceutical Bulletin, 52, 481–484.
Vartia, K.O. (1973). Antibiotics in lichens. The lichens (pp. 547–561). New York: Academic Press Inc.
Volhard, J. (1894). Synthesis and constitution of vulpic acid. Justus Liebigs Annalen der Chemie, 282,
1–21.
Zopf, W. (1985). Zur Kenntnis der Flechtenstoffe. 1 Mitteilung. Liebig’s Annalen, 284, 107.
¨