Revision of the Phytochemistry of Eremophila sturtii and E. mitchellii

Jan 24, 2018 - Eremophila sturtii and E. mitchellii are found in the arid and temperate regions of Australia and, because of their similar appearances...
1 downloads 2 Views 1MB Size
Download PDF
Note Cite This: J. Nat. Prod. 2018, 81, 405−409

pubs.acs.org/jnp

Revision of the Phytochemistry of Eremophila sturtii and E. mitchellii Nicholas J. Sadgrove,† Julian Klepp,† Sarah V.A.-M. Legendre,† Dane Lyddiard,† Christopher J. Sumby,‡ and Ben W. Greatrex*,† †

School of Science and Technology, University of New England, Armidale 2351, Australia Department of Chemistry, School of Physical Sciences, University of Adelaide, Adelaide 5005, Australia

‡

S Supporting Information *

ABSTRACT: Eremophila sturtii and E. mitchellii are found in the arid and temperate regions of Australia and, because of their similar appearances, are often confused. Previous phytochemical investigations have described mitchellene sesquiterpenes (1−5) reported from E. mitchellii but are here demonstrated to be from E. sturtii. A previous study that described serrulatic acids (16 and 17) from a species reported as E. sturtii actually used E. mitchellii. In addition, two new C-15 modified analogues, mitchellenes F (14) and G (15), were isolated from E. sturtii. The absolute configuration of 14 was determined with the first X-ray structure of a compound with the mitchellene skeleton.

E

nation.12 However, no additional sampling was possible to confirm the species nor has a correction appeared in the literature, resulting in the continued misattribution of the mitchellenes to E. mitchellii.12−14 This was probably not the first phytochemical study to misdetermine the two species. In 2006, Liu et al.15 identified the bactericidal diterpenes serrulatic acid (16) and 3,8dihydroxyserrulatic acid (17) from E. sturtii. Serrulatanes and serrulatic acid derivatives are common to the genus, and compounds 1−8 were not reported.16,17 The plant used in the study by Liu et al. was collected from Lightning Ridge in NSW, which is 60 km east from the known range for E. sturtii and where E. mitchellii is abundant.18 The apparent phytochemical differences between the studies prompted an inspection of the voucher provided by The Downing Herbarium, Macquarie University, which revealed a habit and leaves consistent with E. mitchellii (see the Supporting Information). As part of a research program examining Australian Eremophila species,19,20 E. mitchellii and E. sturtii extracts and volatile oils have been characterized and are reported here. These results suggest the specimen used by Barnes et al.6 was E. sturtii and not E. mitchellii. Furthermore, the samples investigated by Liu et al.15 were likely E. mitchellii and not E. sturtii. New mitchellenes from E. sturtii have also been identified as part of this investigation. Chemical assignments for components of the CH2Cl2 extracts from freshly collected leaves of E. sturtii and E. mitchellii were made by NMR spectroscopy and volatile components were identified and quantified in the hydrodistilled extracts of fresh leaf material by gas chromatography−mass

remophila mitchellii Benth. is a temperate Australian shrub to small tree (2−10 m tall) endemic to central New South Wales (NSW) and distributed across the inland regions northward toward central Queensland (Qld), forming a major component of most woodlands.1 Eremophila sturtii R. Br. is a multistemmed desert shrub (1−5 m tall) distributed widely in far western NSW, neighboring South Australia (SA), and north into Qld. A disjunct distribution occurs in Central Australia around Alice Springs and Uluru, extending as far south as Maralinga in SA.1 There is some overlap between the distribution of E. mitchellii and E. sturtii, and in these locations the two species can be found growing together.1 The morphological similarities between the species mean they are easily misidentified, especially as both broad and narrow leaf variants of E. mitchellii are known.1 This is particularly true when E. sturtii reaches 5 m and takes on the appearance of a small tree (N.J.S., personal observation; see the Supporting Information for complete species morphological characterization). The genus Eremophila has proven to be a rich source of interesting natural products.2−5 In 2011, Barnes et al. reported from E. mitchellii a tetracyclic sesquiterpene lactone natural product mitchellene B (1) as well as two oxidized analogues, mitchellene A (2) and mitchellene C (3) (Figure 1).6 Also isolated were muurolane sequiterpenes named as mitchellene D (4), michellene E (5), and the known compounds 14-hydroxy6,12-muuroloadien-15-oic acid (6),7 casticin (7), and centaureidin (8).8,9 In earlier phytochemical studies of the wood and root oil of E. mitchellii, a number of eremophilanes were described, but these were not reported in the study by Barnes et al.10,11 In Barnes’ Ph.D. thesis, the specimen studied was redetermined from E. mitchellii to E. sturtii, noting the error made by the Queensland Herbarium along with highperformance liquid chromatography evidence for the redermi© 2018 American Chemical Society and American Society of Pharmacognosy

Received: July 21, 2017 Published: January 24, 2018 405

DOI: 10.1021/acs.jnatprod.7b00616 J. Nat. Prod. 2018, 81, 405−409

Journal of Natural Products

Note

Figure 1. Compounds identified by previous studies and this work in Eremophila sturtii and E. mitchellii with reassigned species identity.

Table 1. Natural Products Isolated from E. sturtii and E. mitchellii by CH2Cl2 Extraction or by Hydrodistillation Compared with Previous Studies with Reassigned Species Identity2,11 E. sturtii a

compound

% yield

1 2 3 4 5 6 7, 8d 9 10 11 12 13 14 15 16 17

0.84 − 0.01 − 0.12 0.05 0.10 0−0.05 0.03−0.2 0−0.1 0.07 0.01−0.05 0.09 0.01 − −

E. mitchellii b

oil (n = 8)

(ref 2)

3.3 ± 2.3 − − − − − − 10.0 ± 10.1 26.7 ± 17.9 7.5 ± 8.9 − 7.1 ± 2.5 − − − −

0.10 0.04 0.01 0.04 0.03 0.19 0.07 − − − − − − − − −

c

% yield

a

− − − − − − − − − 0.01−0.07 − − − − 0.05 −

oilb (n = 3)

(ref 11)c

− − − − − − − − − 29.1 ± 18.1 − − − − − −

− − − − − − − − − − − − − − 0.03 0.04

a

% yield based upon wet weight of leaves. bOil refers to the hydrodistilled oil, and the composition is reported as a percentage of the GC-MS TIC. The composition is reported as the mean from hydrodistillation of leaves from individual specimens (E. sturtii, n = 8; E. mitchellii, n = 3). cPrevious studies quoted isolated percentages based upon dry weight. dIsolated as a mixture.

Figure 2. X-ray crystal structures of (8S)-8,14-cedranediol (12) and mitchellene F (14). Ellipsoids shown at the 50% probability level (carbon, gray; hydrogen, white; oxygen, red). Intramolecular hydrogen bonds shown with red dashed bonds [12, DO14−O8 = 2.641(3) Å, angleO14−H14−O8 = 158(4)°; 14, DO14−O15 = 2.657(2) Å, angleO14−H14−O15 = 160(3)°].

al.15 after species reassignment. Altogether, 32 components were detected in the hydrodistilled oil from E. sturtii and 28

spectrometry (Table 1). These results are shown alongside the results from the previous studies by Barnes et al.6 and Liu et 406

DOI: 10.1021/acs.jnatprod.7b00616 J. Nat. Prod. 2018, 81, 405−409

Journal of Natural Products

Note

Information). Some specimens were dominated by α-pinene and lesser amounts of bicyclogermacrene; other specimens were dominated by bicyclogermacrene and spathulenol with minor sesquiterpene alcohols such as globulol and muurolol. These results are consistent with previous studies of E. mitchellii where it was found that spathulenol, pinene, globulol, and viridiflorene were the major components of the plant oils.10 Analysis of a solvent extract from leaves of one specimen (NE102334) collected close to the location specified in Liu et al. also yielded serrulatic acid 16, confirmed by matching 13C NMR data.15 None of the mitchellenes were found in this plant, which reinforces the redetermination made by the authors of the current study. In conclusion, this study has demonstrated that the natural products previously reported from E. mitchellii are actually present in the species E. sturtii and vice versa. Furthermore, the phytochemical differences between E. sturtii and E. mitchellii identified in this study indicate that analysis of oils or extractables by GC-MS, using the mitchellenes as markers, could be used to assist future taxonomic classification of the species.

components in the hydrodistilled oil from E. mitchellii. Only those structures confirmed by NMR spectroscopic interpretation are presented in Table 1, and the full list assigned by their MS fragmentation pattern and retention index can be found in the Supporting Information. The major components found in this study of E. sturtii match those reported by Barnes et al. originally reported as E. mitchellii with minor differences. Compounds 1, 3, and 5−8 reported by Barnes et al. were similarly isolated in the current study. Mitchellene A (2) was not observed in the CH2Cl2 extracts or the hydrodistilled oils of E. sturtii in the present work. In addition, the known sesquiterpenes ngaione (9),21,22 myomontanone (10),23 bicyclogermacrene (11), (8S)-8,14cedranediol (12),24,25 and 8,14-cedranoxide (13)24 were isolated. Cedranediol 12 was assigned on the basis of the 2D NMR spectroscopy, and crystals of 12 suitable for X-ray analysis were obtained, confirming the assigned structure of the molecule (Figure 2).24 Two new sesquiterpenes related to 1 were also isolated that have been named mitchellene F (14) and mitchellene G (15). Serrulatic acids 16 and 17, previously reported by Liu et al.,15 were not observed in E. sturtii in the current study. High-resolution mass spectrometry of 14 produced a sodiated molecular ion at m/z 259.1666 [M + Na]+, consistent with a molecular formula of C15H24O2 for the parent molecule. Comparison of the 1H NMR spectrum of 14 to 1 showed similarities including the characteristic δΗ 2.96 ppm signal attributed to C-6 and indicated that 14 had four more protons than 1. The vinylic signal was shifted upfield because of the absence of the lactone ring, and an AB quartet for the C-15 methylene was observed at δΗ 4.06 and 4.10 ppm. The 13C NMR spectrum lacked the signal for the lactone carbonyl seen in 1 and showed an additional signal at δC 68.6 ppm. The presence of one unsaturated bond was inferred from resonances at δC 128.6 and 137.3 ppm, and these resonances had a HMBC correlation with the methylene signal at δΗ 4.06 and 4.10 ppm. The absolute configuration was assigned by single-crystal X-ray analysis and is opposite to that drawn by Barnes et al. (Figure 2). Previously, the absolute configuration of a diastereomer of the putative precursor 6 had been determined, but only the relative configuration of 6 was assigned when it was isolated from E. virgata.26 Compound 14 was also synthesized from 1 using LiAlH4 in THF in 54% yield. The NMR spectra and optical rotations for the synthetic material matched 14 found in the plant, thereby determining the absolute configuration of 1 and confirming the relative configuration assigned by Barnes et al.6 High-resolution electron impact mass spectrometry of 15 produced a molecular ion at m/z 220.1834 [M]+, consistent with a molecular formula of C15H24O. Comparison to the 1H NMR spectrum of 14 indicated 15 lacked the C-15 hydroxy group. The AB quartet was replaced with a 3H singlet at δΗ 1.75 ppm, broadened because of long-range coupling, which showed a correlation with the alkene resonances at δC 133.1 and 125.8 ppm in the HMBC spectrum. Excluding the C-15 center and the neighboring alkene, there was less than 0.1 ppm difference in the 1H NMR chemical shifts for the two tricyclic structures 14 and 15, which strongly suggested that the configurations for the two compounds are the same. The leaves of E. mitchellii produced lower yields of hydrodistilled oils than E. sturtii, at 0.1−0.2% w/w of fresh leaves. Furthermore, E. mitchellii displayed chemovariation, with no apparent geographical relationship (see the Supporting

■

EXPERIMENTAL SECTION

General Experimental Procedures. Optical rotations were recorded on a Rudolph Research Analytical Autopol 1 polarimeter. IR spectra were recorded on a PerkinElmer Spectrum Two spectrometer. NMR spectra were recorded at 298 K on a 500 MHz Bruker Avance III spectrometer and referenced to solvent signals (CDCl3, δH at 7.26 ppm, δC at 77.0 ppm). Liquid chromatography− mass spectrometry (LC-MS) data were recorded using a Varian mass spectrometer. GC-MS analyses were performed using an Agilent Technologies 7890A GC-system coupled with an Agilent 5975C mass selective detector (triple-Axis detector) using a HP-5MS Agilent column (30 m × 250 μm × 0.25 μm). The initial oven temperature was 50 °C (no hold) with heating at 5 °C per minute, then at 280 °C the temperature was held for 5 min. MS data were acquired at −70 eV using a mass scan range of m/z 30−400. Components were identified by comparison to the NIST11 library and confirmed by matching arithmetic indices calculated relative to n-alkanes, to those published by Adams27 or listed in the NIST Chemistry WebBook Database.28 Plant Material. Voucher specimens were deposited at the NCW Beadle Herbarium, at the University of New England, Armidale, NSW, Australia. The leaves of E. mitchellii were collected from three different locations in NSW (Cobar, Nyngan, and Collarenebri); accession numbers, respectively NE102351, NE100294, and NE102334. The E. sturtii specimens were collected from just north of Currawinya National Park (Qld), south across western NSW to Broken Hill. Accession numbers are NE102349, NE102350, NE100286, NE102352, NE102353, NE102354, NE100061, and NE100065. Hydrodistillation. Leaves were frozen, crushed, then removed from the twig. Crushed leaves were placed into a 5 L round bottomed flask with 2.5 L of deionized distilled water. Leaves were heated for 3 h using a 6 L electric mantle, and the steam/oil mix was condensed and collected in a 500 mL separating funnel. Oil yields ranged from 0.21 to 0.51% w/w of fresh leaves from E. sturtii and 0.1−0.2% w/w for E. mitchellii. Oils were separated from the hydrosol and stored away from light at 4 °C until used. Extraction and Isolation. The leaves of E. sturtii (27.5 g) were extracted with CH2Cl2 (2 × 400 mL) and methanol (2 × 400 mL). All the extraction layers were combined then concentrated under reduced pressure to yield a dark brown gum (5.23 g). Column chromatography with a gradient from 1:9 EtOAc-hexanes to 100% EtOAc gave three fractions (A, B, and C in order of elution). Fraction A was analyzed by GC-MS and NMR spectroscopy and discarded, as this fraction comprised the same components as detected in the hydrodistilled oils. Fraction B was further purified by column chromatography with a gradient from 1:9 EtOAc-hexanes up to 1:4 EtOAc-hexanes to give 407

DOI: 10.1021/acs.jnatprod.7b00616 J. Nat. Prod. 2018, 81, 405−409

Journal of Natural Products

Note

Table 2. NMR Assignments for 14 and 15 with HMBC Correlations mitchellene F (14) position

δC

1 2

35.1 30.3

3

29.0

4 5 6 7 8 9 10 11 12 13 14 15

40.3 48.4 45.7 136.4 126.2 20.6 37.8 19.2 44.6 11.2 76.6 68.5

OH

mitchellene G (15)

δH (J in Hz) 1.66, 1.54, 1.14, 1.83, 0.93, 1.26, 1.61, 2.96,

m dddd (13.6, 3.5, 3.5, 3.5) dddd (13.6, 12.6, 12.6, 3.8) m dddd (12.6, 12.0, 12.0, 3.5) dddd (12.0, 11.7, 11.7, 3.3) m dd (8.1, 7.0)

5.94, dd (6.0, 1.4) 1.85−1.88, m, 2H 1.71, m 0.90, d (6.7), 3H 1.42, ddq (11.1, 6.8, 4.8) 1.01, d (6.8), 3H 4.25, dd (7.0, 4.9) 4.10, d (11.6) 4.06, d (11.6) 2.91, br s, 2H

HMBC 11 1, 3, 1, 3, 1, 2, 1, 2, 3, 5, 3, 4, 4, 5, 6, 7, 1, 1, 3, 4, 4, 6, 6,

4, 10, 12 4, 11 4 4, 5, 12 10, 12, 13, 14 7, 9, 10, 12 7, 8, 14, 15

9, 15 8, 5, 10 2, 4, 5, 9 2, 10 4, 5, 13, 14 12, 14 5 7, 8 7, 8

δC

δH (J in Hz)

35.1 30.7

1.63−1.69, m 1.53, dddd (13.9, 3.2, 3.2, 3.2) 1.11, dddd (12.6, 12.6, 12.6, 3.7) 1.79, m 0.92, m 1.19, m 1.57, m 2.84, br dd (6.8, 9.5)

2, 1, 1, 1, 1, 3, 3, 4,

5, 3, 3, 2, 2, 5, 4, 5,

5.72 br d (7.1) 1.85−1.74, m, 2H 1.71, m 0.88, d (6.9), 3H 1.32, ddq (11.2, 6.7, 4.5) 1.00, d (6.7), 3H 4.15, ddd (6.9, 6.8, 4.5) 1.75, s, 3H

6, 1, 1, 1, 4, 4, 4, 6,

9, 10, 15 5, 7, 8 2, 4, 5, 9 2, 10 5, 13, 14 12, 13 5 7, 8

28.8 40.3 48.3 47.2 133.1 125.8 20.8 37.9 19.3 45.0 11.3 75.3 23.9

HMBC 9, 10, 11 4, 10 4, 11 4, 5 4, 5 10, 12, 13 6, 9, 10 7, 8

1.25, d (6.9)

three fractions (B-1, B-2, and B-3, in order of elution). Fraction B-1 contained mitchellene B (1, 232 mg, 0.84% w/w), which was confirmed by 1H and 13C NMR spectroscopic analysis. Fraction B-2 was further purified by column chromatography with 2:3 EtOActoluene to give mitchellene F (14, 25 mg, 0.09% w/w), mitchellene G (15, 3 mg, 0.01% w/w), 8,14-cedranediol (12, 20 mg, 0.07% w/w), and mitchellene C (3, 2 mg, 0.01% w/w). Fraction C was further purified by column chromatography with 1:9 MeOH−CH2Cl2 to give two additional fractions C-1 and C-2. C-1 (28 mg, 0.10% w/w) contained a mixture of the flavonoids (7 and 8), which was verified by 13 C NMR spectroscopy. C-2 (48 mg, 0.17% w/w) contained a 3:7 mixture of 14-hydroxy-6,12-muuroloadien-15-oic acid (6)/mitchellene E (5), which was verified by 13C NMR spectroscopic analysis. Mitchellene F (14). Colorless crystals (CH2Cl2); mp 142−144 °C; [α]23D − 110 (c 0.2, CH2Cl2); IR (neat) νmax 3199, 2950, 2903, 1716, 1696, 1377, 1105, 982, 851, 831 cm−1; see Table 2 for NMR data; EIMS m/z 218.2 [M − H2O]+ (100), 187 (49), 161 (44), 160 (47), 145 (60), 119 (49), 105 (63), 93 (51), 91 (82), 79 (56); HRESIMS m/z 259.1666 [M + Na]+ (calcd for C15H24O2Na, 259.1674). Mitchellene G (15). Colorless oil; IR (neat) νmax 3404, 2257, 1660, 1048, 1024, 993, 826, 763 cm−1; see Table 2 for NMR data; EIMS m/z 220 (44), 202 (64), 187 (66), 162 (100), 145 (71), 132 (29), 119 (81), 105 (61), 93 (66), 91 (58), 77 (39), 67 (14), 55 (31); HREIMS m/z 220.1834 (calcd for C15H24O, 220.1827). Synthesis of 14 from 1. To a stirred solution of LiAlH4 (25 mg, 0.70 mmol) in dry THF (2.5 mL) under N2 cooled to 0 °C was added a solution of mitchellene B (80 mg, 0.35 mmol) in dry THF (2.5 mL). After 10 min at 0 °C the mixture was heated at 50 °C for 5 h then cooled to ambient temperature and diluted with Et2O (5 mL) and H2O (5 mL). The layers were separated, and the aqueous phase was extracted with additional Et2O (2 × 5 mL). The combined organic layers were dried (MgSO4) and concentrated, and the residue was purified by flash chromatography (EtOAc-hexanes 4:6), yielding 14 as a white solid that was recrystallized by slow evaporation from CH2Cl2hexanes giving colorless needles (42 mg, 54%). The optical rotation and 1H and 13C NMR spectra matched data for compound 14. Single-Crystal X-ray Crystallography of Compounds 12 and 14. Single crystals were mounted in paratone-N oil on a plastic loop, and X-ray diffraction data were collected at 150(2) K on an Oxford Xcalibur single-crystal diffractometer using Mo Kα radiation (12) or 130(2) K on an Agilent SuperNova single-crystal diffractometer using Cu Kα radiation (14). The data sets were corrected for absorption using a multiscan method, and the structures were solved by direct

methods using SHELXS-2014 and refined by full-matrix least-squares on F2 by SHELXL-2014,29−31 interfaced through the programs XSeed32 or Olex2.33 All non-hydrogen atoms were refined anisotropically, and hydrogen atoms were included as invariants at geometrically estimated positions. Full data for the structure determination have been deposited with the Cambridge Crystallographic Data Centre as CCDC 1530800 (12) and 1530801 (14). Copies of this information may be obtained free of charge from The Director, CCDC, 12 Union Street, Cambridge CB2 1EZ, U.K. (fax, + 44-1223-336-033; e-mail, [email protected]). Crystal Data for 12. C15H26O2, F.w. 238.36, orthorhombic, P212121, a 7.8987(3), b 11.4701(7), c 15.1331(7) Å, V 1371.04(12) Å3 Z = 4, Dcalc = 1.155 Mg/m3, μ 0.074 mm−1, F(000) 528, crystal size 0.56 × 0.34 × 0.24 mm3, θ range for data collection 3.41−28.44°, Reflns coll. 22156, Obs. reflns 2338, Rint 0.0724, GoF 1.057, R1 [I > 2σ(I)] 0.0571, wR2 (all data) 0.1303, Flack 0.1(10), largest diff. peak and hole 0.301 and −0.187 e·Å−3. The absolute structure of this compound (12) has been determined.24 When this enantiomer is chosen, the Flack parameter is 0.1 for the refined structure and 0.9 for the inverted structure, and the slope in a Bijvoet plot is positive (P2(true) = 0.549), suggesting the correct enantiomer.34 Crystal Data for 14. C15H24O2, F.w. 236.34, orthorhombic, P212121, a 5.18407(15), b 8.7085(3), c 29.2276(8) Å, V 1319.50(7) Å3 Z = 4, Dcalc = 1.190 Mg/m3, μ 0.598 mm−1, F(000) 520, crystal size 0.32 × 0.07 × 0.02 mm3, θ range for data collection 5.30−75.78°, Reflns coll. 7685, Obs. reflns 2466, Rint 0.0396, GoF 1.031, R1 [I > 2σ(I)] 0.0350, wR2 (all data) 0.0858, Flack 0.02(16), largest diff. peak and hole 0.167 and −0.189 e·Å−3.

■

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jnatprod.7b00616. 1D and 2D NMR of 1, 14, and 15 in CDCl3; images of voucher specimens for comparison (PDF) Crystallographic data for cedranediol (CIF) Crystallographic data for 14 (CIF) 408

DOI: 10.1021/acs.jnatprod.7b00616 J. Nat. Prod. 2018, 81, 405−409

Journal of Natural Products

■

Note

(23) Métra, P. L.; Sutherland, M. D. Tetrahedron Lett. 1983, 24, 1749−1752. (24) Baggaley, K. H.; Erdtman, H.; Norin, T. Tetrahedron 1968, 24, 3399−3405. (25) Ihara, M.; Makita, K.; Takasu, K. J. Org. Chem. 1999, 64, 1259− 1264. (26) Ghisalberti, E. L.; Jefferies, P. R.; Skelton, B. W.; White, A. H.; Williams, R. S. F. Tetrahedron 1989, 45, 6297−6308. (27) Adams, R. P. Identification of Essential Oil Components by Gas Chromatography/Mass Spectrometry; Allured Publishing Corporation: Carol Stream, IL, 2007. (28) Linstrom, P. J.; Mallard, W. NIST Chemistry WebBook; NIST Standard Reference Database No. 69., 2001. (29) Sheldrick, G. SHELXS-2014 and SHELXL-2014: Program for Xray Crystal Structure Determination; Göttingen University: Göttingen, Germany, 2014. (30) Sheldrick, G. M. Acta Crystallogr., Sect. A: Found. Crystallogr. 2008, 64, 112−122. (31) Sheldrick, G. M. Acta Crystallogr., Sect. C: Struct. Chem. 2015, 71, 3−8. (32) Barbour, L. J. J. Supramol. Chem. 2001, 1, 189−191. (33) Dolomanov, O. V.; Bourhis, L. J.; Gildea, R. J.; Howard, J. A. K.; Puschmann, H. J. Appl. Crystallogr. 2009, 42, 339−341. (34) Hooft, R. W. W.; Straver, L. H.; Spek, A. L. J. Appl. Crystallogr. 2008, 41, 96−103.

AUTHOR INFORMATION

Corresponding Author

*Tel: +612 6773 2402. E-mail: [email protected]. ORCID

Christopher J. Sumby: 0000-0002-9713-9599 Ben W. Greatrex: 0000-0002-0356-4966 Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors thank Associate Professor Rohan Davis (Griffith University) for his helpful assistance in supplying the Ph.D. thesis of E. Barnes. The authors acknowledge the assistance of the N.C.W. Beadle Herbarium and Mrs Alison Downing (Macquarie University) for providing the voucher specimen from Liu et al.15 The authors thank Associate Professor Brendan F. Abrahams, The University of Melbourne for collecting diffraction data for compound 14.

■

REFERENCES

(1) Chinnock, R. J. Eremophila and Allied Genera: A Monograph of the Plant Family Myoporaceae; Rosenberg Pub Pty Limited: Kenthurst, NSW, 2007; pp 216−220. (2) Dastlik, K. A.; Forster, P. G.; Ghisalberti, E. L.; Jefferies, P. R. Phytochemistry 1989, 28, 1425−1426. (3) Singab, A. N.; Youssef, F. S.; Ashour, M. L.; Wink, M. J. Pharm. Pharmacol. 2013, 65, 1239−1279. (4) Richmond, G. S.; Ghisalberti, E. L. Econ. Bot. 1994, 48, 35−59. (5) Ghisalberti, E. L. Phytochemistry 1993, 35, 7−33. (6) Barnes, E. C.; Carroll, A. R.; Davis, R. A. J. Nat. Prod. 2011, 74, 1888−1893. (7) Ghisalberti, E.; Jefferies, P.; Skelton, B.; White, A.; Williams, R. Tetrahedron 1989, 45, 6297−6308. (8) Wagner, H.; Hörhammer, L.; Höer, R.; Murakami, T.; Farkas, L. Tetrahedron Lett. 1969, 10, 3411−3414. (9) Chang, S.-L.; Chiang, Y.-M.; Chang, C. L.-T.; Yeh, H.-H.; Shyur, L.-F.; Kuo, Y.-H.; Wu, T.-K.; Yang, W.-C. J. Ethnopharmacol. 2007, 112, 232−236. (10) Beattie, K. D.; Waterman, P. G.; Forster, P. I.; Thompson, D. R.; Leach, D. N. Phytochemistry 2011, 72, 400−408. (11) Massy-Westropp, R.; Reynolds, G. Aust. J. Chem. 1966, 19, 303− 307. (12) Barnes, E. C. Chemical Diversity of Eremophila Species and Screening Library Generation. Ph.D. Thesis, Griffith University, Nathan, QLD, 2012. (13) Barnes, E. C.; Choomuenwai, V.; Andrews, K. T.; Quinn, R. J.; Davis, R. A. Org. Biomol. Chem. 2012, 10, 4015−4023. (14) Zhao, G.; He, M.; Li, H.; Duan, S.; Yuan, Z.; Xie, X.; She, X. Chem. Commun. 2015, 51, 17321−17323. (15) Liu, Q.; Harrington, D.; Kohen, J. L.; Vemulpad, S.; Jamie, J. F. Phytochemistry 2006, 67, 1256−1261. (16) Ndi, C. P.; Semple, S. J.; Griesser, H. J.; Pyke, S. M.; Barton, M. D. J. Nat. Prod. 2007, 70, 1439−1443. (17) Ndi, C. P.; Semple, S. J.; Griesser, H. J.; Pyke, S. M.; Barton, M. D. Phytochemistry 2007, 68, 2684−2690. (18) Council of Heads of Australasian Herbaria. Australia’s Virtual Herbarium, 2017. http://avh.chah.org.au (accessed 03/05/2017). (19) Sadgrove, N. J.; Collins, T. L.; Legendre, S. V.-M.; Klepp, J.; Jones, G. L.; Greatrex, B. W. Nat. Prod. Commun. 2016, 11, 1211− 1214. (20) Sadgrove, N. J.; Jones, G. L.; Greatrex, B. W. J. Ethnopharmacol. 2014, 154, 758−766. (21) Birch, A.; Massy-Westropp, R.; Wright, S. Aust. J. Chem. 1953, 6, 385−390. (22) Matsuo, K.; Arase, T.; Ishida, S.; Sakaguchi, Y. Heterocycles 1996, 43 (6), 1287−1300. 409

DOI: 10.1021/acs.jnatprod.7b00616 J. Nat. Prod. 2018, 81, 405−409