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Isolation, structure determination, and phytotoxicity of unusual dioxopiperazines from the phytopathogenic fungus Phoma lingam
Phytochemistry 58 (2001) 905–909 www.elsevier.com/locate/phytochem
Isolation, structure determination, and phytotoxicity of unusual dioxopiperazines from the phytopathogenic fungus Phoma lingam M. Soledade C. Pedras*, Corinne J. Biesenthal Department of Chemistry, University of Saskatchewan, 110 Science Place, Saskatoon SK S7N 5C9, Canada Received 24 May 2001; received in revised form 16 July 2001
Abstract The isolation, chemical structure elucidation, and bioactivity of polanrazines B–F, five new dioxopiperazines produced by isolates of the blackleg fungus [Phoma lingam, perfect stage Leptosphaeria maculans (Desm.) Ces. et de Not.] originating from Poland, are reported. Polanrazines C and E showed moderate but selective toxicity, causing necrotic and chlorotic lesions (1–3 mm diameter) on brown mustard leaves. # 2001 Elsevier Science Ltd. All rights reserved. Keywords: Blackleg disease; Crucifer; Dioxopiperazines; Leptosphaeria maculans; Phoma lingam
1. Introduction Blackleg of cruciferous plants is a devastating fungal disease caused by the phytopathogen Leptosphaeria maculans (Desm.) Ces. et de Not., asexual stage Phoma lingam (Tode ex Fr.) Desm. Virulent isolates of the fungus have caused enormous yield losses in rapeseed (Brassica napus L. and B. rapa L.) and canola (B. napus L. and B. rapa L.) in Canada and worldwide (Gugel and Petrie, 1992). To develop appropriate control practices that prevent blackleg disease outbreaks and a rapid breakdown of plant resistance, it is essential to characterize and differentiate the various fungal isolate types. Initially, two different groups of P. lingam (originally called ‘‘strains’’) were characterized in Canada on the basis of virulence, host range, and cultural tests: a highly virulent type (V) causing leaf spots and severe stem cankers on rapeseed and cabbage (B. oleracea L.) and a weakly virulent type (W) causing only superficial leaf and stem lesions on rapeseed and cabbage (McGee and Petrie, 1978). Subsequently, due to the broad variability of fungal isolates causing blackleg, additional subgroups within each of the V and W groups were proposed (Williams and Fitt, 1999; Howlett et al., 2001). Thus far, isolates belonging to the V group, also * Corresponding author. Tel.: +1-306-966-4772; fax: +1-306-9664730. E-mail address: [email protected] (M.S.C. Pedras).
known as A group or aggressive, appear to be responsible for the largest yield losses in Canada, UK, France, Germany, and Australia, whereas in Poland the W isolates appear to cause significant crop damage as well (Williams and Fitt, 1999). Current evidence indicates that isolates of the blackleg fungus occurring in Canada and previously characterized as W isolates, also known as avirulent, B group, or nonaggressive, are likely P. wasabiae Yokogi, and not P. lingam (Pedras et al., 1995, 1996; Taylor et al., 1995; Pedras, 1998; Reddy et al., 1998); however, since no formal reclassification has occurred yet, these isolates will be hereon referred to as W group. Moreover, we have also reported on isolates Laird 2 and Mayfair 2, thus far two unique W Canadian isolates classified as P. lingam, whose metabolite profile is rather different from those of either V or W isolates (Pedras et al., 1998, 1999). Recent studies of isolates of the species P. lingam collected in different countries suggested that there are at least three distinct groups which may constitute three different Phoma species (Pedras and Biesenthal, 2000). Chemical analysis of the secondary metabolites of such isolates indicated that isolates of P. lingam originating from Poland were clearly different from Canadian, French, German, or Australian isolates (Pedras and Biesenthal, 1998, 2000). To characterize and clarify the relationship among diverse blackleg isolates, we have analyzed fungal isolates from Poland and here wish to report their secondary metabolites, compounds 1–7.
0031-9422/01/$ - see front matter # 2001 Elsevier Science Ltd. All rights reserved. PII: S0031-9422(01)00348-X
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Dioxopiperazines 3–7 represent new metabolites which we named polanrazines B, C, D, E and F. Discussion of the implications of our findings and suggestions for isolate grouping are also provided.
2. Results and discussion Initially, eight Polish isolates (IBCN 19–26) of P. lingam were grown in liquid cultures, the culture broths extracted with EtOAc, and the extracts analyzed by TLC, HPLC, and 1H NMR spectroscopic analyses. Both the HPLC chromatograms and 1H NMR spectra of each extract were similar and suggested the presence of aromatic compounds not available in our library containing P. lingam metabolites isolated to date (Pedras, 1998). Furthermore, bioassay of those extracts (as reported in the Experimental) on leaves of canola, brown mustard (B. juncea), and white mustard (Sinapis alba) suggested that these extracts were more phytotoxic to brown mustard than to canola or white mustard. Subsequently, Polish isolate IBCN 19 was selected to grow on a larger scale for metabolite isolation and chemical structure determination. The EtOAc extract of the culture broth was fractionated by FCC followed by multiple fractionation by prep. TLC, as described in the experimental, to yield phomapyrone A (1) and compounds 2–7. The structures of phomapyrone A (1) (Pedras et al., 1994) and l-valyl-l-tryptophan anhydride (2) (Pedras et al., 1998) were readily established by comparison of their chromatographic and spectroscopic data with that of authentic samples available in our compound library (Pedras and Biesenthal, 2000). Phomapyrone (1) was previously isolated from W Canadian isolates (Pedras et al. 1994), isolates Mayfair 2 and Laird 2 (Pedras et al., 1998, 1999) whereas 2 was only isolated from isolates Mayfair 2 and Laird 2 (Pedras et al., 1998, 1999).
The remaining constituents appeared to be structurally related to dioxopiperazine 2, as their 1H and 13C NMR spectra showed the expected signals of a 2,5dioxopiperazine ring containing 3-methyleneindolyl and isopropyl substituents (Table 1). Compared to l-valyll-tryptophan anhydride (2), the least polar of the dioxopiperazines (3) showed two methyl singlets instead of the methine protons at C20 and C50 . That these methyl groups were likely attached to sulfur atoms was suggested by their chemical shifts, and confirmed by
HRMS (for C18H23N3O2S2). Because no trivial name was assigned to l-valyl-l-tryptophan anhydride (2) we propose that it be named polanrazine A and that compound 3 be named polanrazine B. Similarly, the structures of 4–7 were assigned from analysis of NMR (Table 1) and HRMS data. The regiochemistry of the substituents at C20 and C50 was assigned based on HMBC correlations of the methyl protons to C50 , in compounds 4 and 6, or to C20 , in compound 5. Compound 6 was readily distinguished from 4 based on the chemical shifts of the methyl group (H 3.18, C 51.5 in 6 vs. H 0.81, C 9.7 in 4). The unusually low proton and carbon chemical shifts of the SMe group of 4 might be due to a shielding effect of the indole ring (Lambert et al., 1998). The relative configurations were established by NOE experiments between the methylene protons at C10 and methyne proton at C80 . The absolute configuration of compounds 3–7 was not established; however in previous work we established by total synthesis that the stereogenic centers of 2 (20 and 50 ) had S,S configuration, and are now suggesting a similar assignment for the corresponding stereogenic centers of 3–7. Similar to trivial names of 2 and 3, we propose that 4–7 be named polanrazines C–E. Dioxopiperazines are common secondary metabolites produced by a variety of microorganisms of both marine and terrestrial origin; in fact V isolates of P. lingam produce a variety of epithiodioxopiperazines including the phytotoxins sirodesmins (Pedras, 1998). On the other hand, dioxopiperazines such as polanrazines C-E having OH/OR groups at the a-carbon of the amino acid residues appear to be rather unusual (e.g. Nozawa et al., 1989; King et al., 1992).
The phytotoxicity assays of polanrazines A–E (2–7) on plants resistant and susceptible to P. lingam indicated that both 4 and 6 had moderate but selective toxicity, causing necrotic and chlorotic lesions (1–3 mm diameter) on brown mustard leaves (5 104 M), whereas no damage was observed on canola or white mustard leaves. Lower concentration of these metabolites (2–7) caused no macroscopic effects. Furthermore,
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blackleg disease, was shown to be more susceptible to these ‘‘W Polish’’ isolates, there is a possibility that such isolates will cause new blackleg epidemics. Thus, to prevent additional disease outbreaks, systematic screening of blackleg isolates must be carried out in areas where brown mustard is widely grown.
preliminary inoculations of canola and brown mustard with spores of isolate IBCN 19 have shown that brown mustard is more susceptible to Polish isolates than canola, whereas the W isolates from Canada cause low to no disease symptoms on either species (Gugel and Petrie, 1992). The present studies indicate that blackleg W isolates originating from Poland produce metabolites similar to those obtained from Canadian isolates Laird 2 and Mayfair 2 (Pedras et al., 1998, 1999). It may be that Mayfair 2 and Laird 2 isolates have not originated in Canada but got to North America through seed brought in from European countries (Pedras and Biesenthal, 2000). Further studies to establish the secondary metabolite profiles of blackleg fungal isolates will be of great assistance in the chemotaxonomic classification of the ‘‘complex species’’ presently known as P. lingam/ L. maculans. Polanrazines A–E will be useful chemotaxonomic markers for grouping new isolates in the appropriate ‘‘W group’’ as they do not appear to be produced by other blackleg isolate types. Nonetheless, this statement does not imply that the virulence of the Polish fungal isolates is related with the production of polanrazines A–E, such a conclusion will require additional investigations. Moreover, the metabolite profile obtained for Polish isolates suggest that this group is distinct from the W Canadian group (P. wasabiae). Because brown mustard, a crop traditionally resistant to
3. Experimental 3.1. General All chemicals were purchased from Sigma-Aldrich Canada Ltd., Oakville, ON. All solvents were HPLC grade and used as such, except for CH2Cl2 and CHCl3 which were redistilled. Preparative (prep.) TLC: (Merck, Kieselgel 60 F254), 20 20 cm 0.25 mm; analytical TLC (Merck, Kieselgel 60 F254, aluminum sheets) 5 2 cm 0.2 mm; compounds were visualized by exposure to UV light and by dipping the plates in a 5% aq (w/v) phosphomolybdic acid solution containing a trace of ceric sulfate and 4% (v/v) H2SO4, followed by heating at 200 C. Flash column chromatography (FCC): silica gel Merck, grade 60, mesh size 230–400, 60 A˚. High pressure liquid chromatography (HPLC) analysis was carried out with a high performance liquid chromatograph equipped with quaternary pump, automatic injector, and diode-array detector (wavelength
Table 1 1 H and 13C NMR chemical shifts (ppm) and 1H multiplicities (J in Hz) for compounds 3–7 (solvent) C/H #
2 3 3a 4 5 6 7 7a 10 10 20 30 50 60 80 90 /100 100 /90 110 SMe 120 SMe 110 OMe OH NH NH NH
Polanrazine C 4 (CD3OD)
Polanrazine B 3 (CD2Cl2)
Polanrazine D 5 (CD3OD)
Polanrazine E 6 (CD3CN)
Polanrazine F 7 (CD3OD)
H
C
H
C
H
dC
H
C
H
C
7.20 s – – 7.39 d, 8.0 7.16 dd, 7.5, 7.5 7.21 dd, 7.5, 7.5 7.64 d, 7.8 – 3.28 d, 14.5 3.78 d, 14.5 – – – – 2.22 m 0.14 d, 7 0.92 d, 7 2.23 s 2.44 s – – 5.71 br s 6.09 br s 8.33 br s
125.4 108.3 128.3 111.5 120.3 122.5 119.1 136.4 36.1 36.1 67.4 166.1 69.2 166.7 33.1 16.0 15.0 14.5 14.7 – – – – –
7.15 s – – 7.29 d, 8 7.01 dd, 8, 8 7.07 dd, 7, 7 7.76 d, 8 – 3.13 d, 14 3.74 d, 14 – – – – 2.23 m 1.0 d, 7 1.04 d, 7 0.81 s – – – – – –
125.5 108.0 128.5 111.0 119.0 121.3 119.2 136.8 35.5 35.5 83.2 168.5 72.8 169.2 37.0 17.5 15.5 9.7 – – – – – –
7.11 s – – 7.27 d, 8 7.00 dd, 8, 8 7.05 dd, 7, 7 7.71 d, 8 – 3.17 d, 14 3.72 d, 14 – – – – 2.28 m 0.25 d, 7 0.62 d, 7 2.28 s – – – – – –
125.5 108.1 128.4 111.0 119.0 121.2 119.1 136.7 35.2 35.2 69.4 168.4 84.1 169.0 35.2 15.1 13.2 12.1 – – – – – –
7.10 s – – 7.33 d, 7.5 7.08 dd. 7 7.05 dd, 7 7.65 d, 7.5 – 3.14 d, 14 3.57 d, 14 – – – – 1.97 m 0.04 d, 7 0.62 d, 7 – – 3.18 s 4.75 br s 6.31 br s 7.21 br s 9.17 br s
125.9 108.1 128.6 111.7 119.5 121.9, 119.5 136.8 36.1 36.1 83.3 168.9 89.9 165.6 35.7 15.6 14.1 – – 51.5 – – – –
7.09 s – – 7.28 d, 7.5 6.99 dd. 8 7.05 dd, 7.5 7.70 d, 8 – 3.13 d, 14 3.66 d, 14 – – – – 1.82 m 0.15 d, 7 0.64 d, 7 – – – – – – –
125.2 107.6 128.4 110.9 118.9 121.2, 119.1 136.8 35.2 35.2 83.3 169.4 84.1 168.7 35.1 15.0 13.3 – – – – – – –
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range 190–600 nm), degasser, and a Hypersil ODS column (5 mm particle size silica, 4.6 i.d. 200 mm), equipped with a low-dispersion column-inlet filter. Two different elution methods were employed; method A: linear gradient from 75% H2O–25% CH3CN to 100% CH3CN, at a flow rate 1.0 ml/min over 35 min; method B: isocratic elution 80% H2O–20% CH3CN for 10 min, followed by gradient elutions, 80% H2O–20% CH3CN to 60% H2O–40% CH3CN for 10 min, and 60% H2O– 40% CH3CN to 25% H2O–75% CH3CN for 10 min, at a flow rate 1.0 ml/min. NMR spectra were recorded on a Bruker AMX 300 or 500 spectrometer; for 1H (300 or 500 MHz), values were referenced to CD3OD (CD2HOD at 3.31 ppm), CD2Cl2 (CDHCl2 at 5.32 ppm), or CD3CN (CHD2CN at 1.94 ppm), and for 13C (75.5 or 125.8 MHz) referenced to CD3OD (49.15 ppm), CD2Cl2 (54.00 ppm), or CD3CN (118.69 ppm). The list of coupling constants (J, reported to the nearest 0.5 Hz) corresponds to the order of the multiplicity assignment. Mass spectra (MS) [high resolution (HR), electron impact (EI), or fast atom bombardment (FAB)] were obtained on a VG 70 SE mass spectrometer, employing a solids probe. Specific rotations, []D were determined at ambient temperature on a Perkin-Elmer 141 polarimeter using a 1 ml, 10 cm path length cell; the units are 101 deg cm2 g1 and the concentrations are reported in g/100 ml. Infrared spectra were recorded on a Bio-Rad FTS-40 spectrometer. Spectra were measured by the diffuse reflectance method on samples dispersed in KBr; only diagnostic peaks are reported.
employing a 1:1 MeOH–H20 solution. Plants were incubated in a growth chamber as reported above (16 h light/8 h dark, at 24 2 C); leaves were observed over a 1-week period and the diameter of each lesion was measured after 7 days. 3.3. Isolation of compounds 3–7 The broth and mycelium were separated by gravity filtration, the broth (3 l) was concentrated (400 ml) and extracted with EtOAc (220 mg). Flash column chromatography (FCC) (eluted with CH2Cl2–MeOH, 95:5, 250 ml; 90:10, 250 ml of the EtOAc extract yielded the following fractions: F1 containing compounds 1 and 3, F2 containing compounds 2–4, F3–F4 containing compounds 2 and 5, F6–F9 containing compounds 6 and 7. Prep. TLC separation (CH2Cl2–MeOH, 97:3, double elution) of F1 (20 mg) yielded phomapyrone A (1, 4 mg) and polanrazine B (3), which was combined with a prep. TLC fraction eluted from F2 and further separated to yield chromatographically homogeneous material (3, 3.5 mg). Prep. TLC separation (CH2Cl2–MeOH, 96:4, double elution) of F2 (30 mg) yielded polanrazine B (3) which was combined with a prep. TLC band obtained from F1, and a mixture (20 mg) containing polanrazines C and D (4, 5). Separation of this mixture using multiple prep. TLC (CH2Cl2–MeOH, 96:4, multiple development) yielded chromatographically homogeneous materials 4 (6.5 mg) and 5 (2.5 mg). Finally prep. TLC separation (CH2Cl2–MeOH, 96:4, double elution) of F6-9 (6 mg) yielded polanrazine E (6, 1 mg) and F (7, 1.5 mg).
3.2. Fungal cultures and bioassays 3.4. Polanrazine B (3) The eight Polish isolates of P. lingam, IBCN 19-26, were obtained from the IBCN blackleg collection at Agriculture and Agri-Food Canada, Saskatoon. Liquid cultures of these isolates were grown as previously reported (Pedras and Biesenthal, 2000). For isolation of metabolites, isolate IBCN 19 was grown in 250 ml Erlenmeyer flasks containing 100 ml minimal medium (total 4 l) inoculated with fungal spores at 2 109 per flask were incubated on a shaker at 150 rpm, at 24 2 C for 6 days, and worked up as described below. Plants were grown in a growth chamber for 3 weeks, with 16 h light (fluorescent and incandescent, 450–530 mmol s1 m2)/8 h dark, at 24 2 C. Three different species of known resistance to V isolates of Phoma lingam were employed in the bioassays: Brassica napus, cv. Westar (susceptible) and B. juncea, cv. Cutlass (resistant), Sinapis alba, cv. Ochre (resistant). EtOAc extracts (2 mg/ml) or pure compounds (5104, 1104, 2105M) where dissolved in MeOH–H20 (1:1). Leaves were punctured with a needle (6 punctures per leaf) and 10 ml droplets (6 droplets per leaf) were applied on punctured sites. Control leaves were treated similarly
HPLC tR=22.4 min (solvent system B); []D60 (c 0.2 CHCl3); HR–EI–MS m/z (% relative abundance) measured: 377.1230 (377.1232 calc. for C18H23N3O2S2); FTIR max: 3388, 3190, 2926,1665, 1452, 1413, 1075 cm1; lmax (log ") 218 (4.5) nm. 3.5. Polanrazine C (4) HPLC tR=10.6 min (solvent system B); []D 16 (c 0.18 MeOH); HR–EI–MS m/z (% relative abundance) measured: 347.1306 (347.1304 calcd. for C17H21N3O3S1); FTIR vmax: 3335 3220, 2956, 2926, 1673, 1435, 1075 cm1; lmax (log ") 219 (4.3) nm. 3.6. Polanrazine D (5) HPLC tR=11.3 min (solvent system B); []D8.2 (c 0.18 MeOH); HREIMS m/z (% relative abundance) measured: 347.1306 (347.1303 calc. for C17H21N3O3S1. FTIR vmax: 3444, 3347, 2968, 2926, 1677, 1625, 1465, 1427, 1058 cm1; lmax( log ") 219 nm (4.0).
M.S.C. Pedras, C.J. Biesenthal / Phytochemistry 58 (2001) 905–909
3.7. Polanrazine E (6) HPLC tR=9.6 min (solvent system B); []D6 (c 0.07 MeOH); HREIMS m/z (% relative abundance) measured: 331.1531 (331.1532 calc. for C17H21N3O4). FTIR vmax: 3342, 3223, 2962, 2927, 2857, 1682, 1429, 1094 cm1; lmax (log ") 222 (4.3), 282 (3.7) nm. 3.8. Polanrazine F (7) HPLC tR=7.0 min (solvent system B); []D10 (c 0.26 MeOH); HR–EI–MS m/z (% relative abundance) measured: 317.1377 (317.1376 calcd. for C16H19N3O4); FTIR vmax: 3319, 2970, 2931, 1680, 1426, 1086, 1043 cm1; lmax ( log ") 219 (4.2), 281 (3.5) nm.
Acknowledgements We would like to thank G. Se´guin-Swartz and R. Gugel, Agriculture and Agri-Food Canada, Saskatoon Research Station, Saskatchewan, for kindly providing isolates from the International Blackleg of Crucifers Network (IBCN) collection available in Saskatoon, and K. Brown, Department of Chemistry, University of Saskatchewan, for NMR spectral measurements. Financial support from the Natural Sciences and Engineering Research Council (Canada) and the University of Saskatchewan is gratefully acknowledged. References Gugel, R.K., Petrie, G.A., 1992. History, occurrence, impact, and control of blackleg of rapeseed. Canadian Journal of Plant Pathology 14, 36–45. Howlett, B.J., Idnurm, A., Pedras, M.S.C., 2001. Leptosphaeria maculans the causal agent of blackleg disease of Brassicas. Fungal Genetics and Biology 33, 1–14. King, R.R., Lawrence, C.H., Calhoun, L.A., 1992. Chemistry of phytotoxins associated with Streptomyces scabies, the causal organism
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of potato common scab. Journal of Agriculture and Food Chemistry 40, 834–837. Lambert, J.B., Shurvell, H.F., Lightner, D.A., Cooks, R.G., 1998. Organic Structural Spectroscopy. Prentice-Hall, New Jersey. McGee, D.C., Petrie, G.A., 1978. Variability of Leptosphaeria maculans in relation to blackleg of oilseed rape. Phytopathology 68, 625– 630. Nozawa, K., Udagawa, S-I., Nakajima, S., Kawai, K-I., 1989. A dioxopiperazine derivative from Penicillium megasporum. Phytochemistry 28, 929–931. Pedras, M.S.C., 1998. Towards an understanding and control of plant fungal diseases in Brassicaceae. Recent Research Developments in Agricultural and Food Chemistry 2, 513–532. Pedras, M.S.C., Biesenthal, C.J., 1998. Production of the host-selective toxin phomalide by isolates of Leptosphaeria maculans and its correlation with sirodesmin PL production. Canadian Journal of Microbiology 44, 547–553. Pedras, M.S.C., Biesenthal, C.J., 2000. HPLC analyses of cultures of Phoma spp.: differentiation among groups and species through secondary metabolite profiles. Canadian Journal of Microbiology 46, 685–691. Pedras, M.S.C., Erosa-Lo´pez, C.C., Quail, J.W., Taylor, J.L., 1999. Phomalairdenone: a new host-selective phytotoxin from a virulent type of the blackleg fungus Phoma lingam. Bioorganic and Medicinal Chemistry Letters 9, 3291–3294. Pedras, M.S.C., Morales, V.M., Taylor, J.L., 1994. Phomapyrones: three metabolites from the blackleg fungus. Phytochemistry 36, 1315–1318. Pedras, M.S.C., Smith, K.C., Taylor, J.L., 1998. Production of a 2,5dioxopiperazine by a new isolate type of the blackleg fungus Phoma lingam. Phytochemistry 49, 1575–1577. Pedras, M.S.C., Taylor, J.L., Morales, V.M., 1995. Phomaligin A and other yellow pigments in Phoma lingam and P. wasabiae. Phytochemistry 38, 1215–1222. Pedras, M.S.C., Taylor, J.L., Morales, V.M., 1996. The blackleg fungus of rapeseed: how many species? Acta Horticulturae 407, 441– 446. Reddy, P.V., Patel, R., White Jr., J.F., 1998. Phylogenetic and developmental evidence supporting reclassification of cruciferous pathogens Phoma lingam and Phoma wasabiae in Plenodomus. Canadian Journal of Botany 76, 1916–1922. Taylor, J.L., Pedras, M.S.C., Morales, V.M., 1995. Horizontal transfer in the phytopathogenic fungal genus Leptosphaeria and hostrange expansion. Trends in Microbiology 3, 202–206. Williams, R.H., Fitt, B.D., 1999. Differentiating A and B groups of Leptosphaeria maculans, causal agent of stem canker (blackleg) of oilseed rape. Plant Pathology 48, 161–175.
Isolation, structure determination, and phytotoxicity of unusual dioxopiperazines from the phytopathogenic fungus Phoma lingam M. Soledade C. Pedras*, Corinne J. Biesenthal Department of Chemistry, University of Saskatchewan, 110 Science Place, Saskatoon SK S7N 5C9, Canada Received 24 May 2001; received in revised form 16 July 2001
Abstract The isolation, chemical structure elucidation, and bioactivity of polanrazines B–F, five new dioxopiperazines produced by isolates of the blackleg fungus [Phoma lingam, perfect stage Leptosphaeria maculans (Desm.) Ces. et de Not.] originating from Poland, are reported. Polanrazines C and E showed moderate but selective toxicity, causing necrotic and chlorotic lesions (1–3 mm diameter) on brown mustard leaves. # 2001 Elsevier Science Ltd. All rights reserved. Keywords: Blackleg disease; Crucifer; Dioxopiperazines; Leptosphaeria maculans; Phoma lingam
1. Introduction Blackleg of cruciferous plants is a devastating fungal disease caused by the phytopathogen Leptosphaeria maculans (Desm.) Ces. et de Not., asexual stage Phoma lingam (Tode ex Fr.) Desm. Virulent isolates of the fungus have caused enormous yield losses in rapeseed (Brassica napus L. and B. rapa L.) and canola (B. napus L. and B. rapa L.) in Canada and worldwide (Gugel and Petrie, 1992). To develop appropriate control practices that prevent blackleg disease outbreaks and a rapid breakdown of plant resistance, it is essential to characterize and differentiate the various fungal isolate types. Initially, two different groups of P. lingam (originally called ‘‘strains’’) were characterized in Canada on the basis of virulence, host range, and cultural tests: a highly virulent type (V) causing leaf spots and severe stem cankers on rapeseed and cabbage (B. oleracea L.) and a weakly virulent type (W) causing only superficial leaf and stem lesions on rapeseed and cabbage (McGee and Petrie, 1978). Subsequently, due to the broad variability of fungal isolates causing blackleg, additional subgroups within each of the V and W groups were proposed (Williams and Fitt, 1999; Howlett et al., 2001). Thus far, isolates belonging to the V group, also * Corresponding author. Tel.: +1-306-966-4772; fax: +1-306-9664730. E-mail address: [email protected] (M.S.C. Pedras).
known as A group or aggressive, appear to be responsible for the largest yield losses in Canada, UK, France, Germany, and Australia, whereas in Poland the W isolates appear to cause significant crop damage as well (Williams and Fitt, 1999). Current evidence indicates that isolates of the blackleg fungus occurring in Canada and previously characterized as W isolates, also known as avirulent, B group, or nonaggressive, are likely P. wasabiae Yokogi, and not P. lingam (Pedras et al., 1995, 1996; Taylor et al., 1995; Pedras, 1998; Reddy et al., 1998); however, since no formal reclassification has occurred yet, these isolates will be hereon referred to as W group. Moreover, we have also reported on isolates Laird 2 and Mayfair 2, thus far two unique W Canadian isolates classified as P. lingam, whose metabolite profile is rather different from those of either V or W isolates (Pedras et al., 1998, 1999). Recent studies of isolates of the species P. lingam collected in different countries suggested that there are at least three distinct groups which may constitute three different Phoma species (Pedras and Biesenthal, 2000). Chemical analysis of the secondary metabolites of such isolates indicated that isolates of P. lingam originating from Poland were clearly different from Canadian, French, German, or Australian isolates (Pedras and Biesenthal, 1998, 2000). To characterize and clarify the relationship among diverse blackleg isolates, we have analyzed fungal isolates from Poland and here wish to report their secondary metabolites, compounds 1–7.
0031-9422/01/$ - see front matter # 2001 Elsevier Science Ltd. All rights reserved. PII: S0031-9422(01)00348-X
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M.S.C. Pedras, C.J. Biesenthal / Phytochemistry 58 (2001) 905–909
Dioxopiperazines 3–7 represent new metabolites which we named polanrazines B, C, D, E and F. Discussion of the implications of our findings and suggestions for isolate grouping are also provided.
2. Results and discussion Initially, eight Polish isolates (IBCN 19–26) of P. lingam were grown in liquid cultures, the culture broths extracted with EtOAc, and the extracts analyzed by TLC, HPLC, and 1H NMR spectroscopic analyses. Both the HPLC chromatograms and 1H NMR spectra of each extract were similar and suggested the presence of aromatic compounds not available in our library containing P. lingam metabolites isolated to date (Pedras, 1998). Furthermore, bioassay of those extracts (as reported in the Experimental) on leaves of canola, brown mustard (B. juncea), and white mustard (Sinapis alba) suggested that these extracts were more phytotoxic to brown mustard than to canola or white mustard. Subsequently, Polish isolate IBCN 19 was selected to grow on a larger scale for metabolite isolation and chemical structure determination. The EtOAc extract of the culture broth was fractionated by FCC followed by multiple fractionation by prep. TLC, as described in the experimental, to yield phomapyrone A (1) and compounds 2–7. The structures of phomapyrone A (1) (Pedras et al., 1994) and l-valyl-l-tryptophan anhydride (2) (Pedras et al., 1998) were readily established by comparison of their chromatographic and spectroscopic data with that of authentic samples available in our compound library (Pedras and Biesenthal, 2000). Phomapyrone (1) was previously isolated from W Canadian isolates (Pedras et al. 1994), isolates Mayfair 2 and Laird 2 (Pedras et al., 1998, 1999) whereas 2 was only isolated from isolates Mayfair 2 and Laird 2 (Pedras et al., 1998, 1999).
The remaining constituents appeared to be structurally related to dioxopiperazine 2, as their 1H and 13C NMR spectra showed the expected signals of a 2,5dioxopiperazine ring containing 3-methyleneindolyl and isopropyl substituents (Table 1). Compared to l-valyll-tryptophan anhydride (2), the least polar of the dioxopiperazines (3) showed two methyl singlets instead of the methine protons at C20 and C50 . That these methyl groups were likely attached to sulfur atoms was suggested by their chemical shifts, and confirmed by
HRMS (for C18H23N3O2S2). Because no trivial name was assigned to l-valyl-l-tryptophan anhydride (2) we propose that it be named polanrazine A and that compound 3 be named polanrazine B. Similarly, the structures of 4–7 were assigned from analysis of NMR (Table 1) and HRMS data. The regiochemistry of the substituents at C20 and C50 was assigned based on HMBC correlations of the methyl protons to C50 , in compounds 4 and 6, or to C20 , in compound 5. Compound 6 was readily distinguished from 4 based on the chemical shifts of the methyl group (H 3.18, C 51.5 in 6 vs. H 0.81, C 9.7 in 4). The unusually low proton and carbon chemical shifts of the SMe group of 4 might be due to a shielding effect of the indole ring (Lambert et al., 1998). The relative configurations were established by NOE experiments between the methylene protons at C10 and methyne proton at C80 . The absolute configuration of compounds 3–7 was not established; however in previous work we established by total synthesis that the stereogenic centers of 2 (20 and 50 ) had S,S configuration, and are now suggesting a similar assignment for the corresponding stereogenic centers of 3–7. Similar to trivial names of 2 and 3, we propose that 4–7 be named polanrazines C–E. Dioxopiperazines are common secondary metabolites produced by a variety of microorganisms of both marine and terrestrial origin; in fact V isolates of P. lingam produce a variety of epithiodioxopiperazines including the phytotoxins sirodesmins (Pedras, 1998). On the other hand, dioxopiperazines such as polanrazines C-E having OH/OR groups at the a-carbon of the amino acid residues appear to be rather unusual (e.g. Nozawa et al., 1989; King et al., 1992).
The phytotoxicity assays of polanrazines A–E (2–7) on plants resistant and susceptible to P. lingam indicated that both 4 and 6 had moderate but selective toxicity, causing necrotic and chlorotic lesions (1–3 mm diameter) on brown mustard leaves (5 104 M), whereas no damage was observed on canola or white mustard leaves. Lower concentration of these metabolites (2–7) caused no macroscopic effects. Furthermore,
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blackleg disease, was shown to be more susceptible to these ‘‘W Polish’’ isolates, there is a possibility that such isolates will cause new blackleg epidemics. Thus, to prevent additional disease outbreaks, systematic screening of blackleg isolates must be carried out in areas where brown mustard is widely grown.
preliminary inoculations of canola and brown mustard with spores of isolate IBCN 19 have shown that brown mustard is more susceptible to Polish isolates than canola, whereas the W isolates from Canada cause low to no disease symptoms on either species (Gugel and Petrie, 1992). The present studies indicate that blackleg W isolates originating from Poland produce metabolites similar to those obtained from Canadian isolates Laird 2 and Mayfair 2 (Pedras et al., 1998, 1999). It may be that Mayfair 2 and Laird 2 isolates have not originated in Canada but got to North America through seed brought in from European countries (Pedras and Biesenthal, 2000). Further studies to establish the secondary metabolite profiles of blackleg fungal isolates will be of great assistance in the chemotaxonomic classification of the ‘‘complex species’’ presently known as P. lingam/ L. maculans. Polanrazines A–E will be useful chemotaxonomic markers for grouping new isolates in the appropriate ‘‘W group’’ as they do not appear to be produced by other blackleg isolate types. Nonetheless, this statement does not imply that the virulence of the Polish fungal isolates is related with the production of polanrazines A–E, such a conclusion will require additional investigations. Moreover, the metabolite profile obtained for Polish isolates suggest that this group is distinct from the W Canadian group (P. wasabiae). Because brown mustard, a crop traditionally resistant to
3. Experimental 3.1. General All chemicals were purchased from Sigma-Aldrich Canada Ltd., Oakville, ON. All solvents were HPLC grade and used as such, except for CH2Cl2 and CHCl3 which were redistilled. Preparative (prep.) TLC: (Merck, Kieselgel 60 F254), 20 20 cm 0.25 mm; analytical TLC (Merck, Kieselgel 60 F254, aluminum sheets) 5 2 cm 0.2 mm; compounds were visualized by exposure to UV light and by dipping the plates in a 5% aq (w/v) phosphomolybdic acid solution containing a trace of ceric sulfate and 4% (v/v) H2SO4, followed by heating at 200 C. Flash column chromatography (FCC): silica gel Merck, grade 60, mesh size 230–400, 60 A˚. High pressure liquid chromatography (HPLC) analysis was carried out with a high performance liquid chromatograph equipped with quaternary pump, automatic injector, and diode-array detector (wavelength
Table 1 1 H and 13C NMR chemical shifts (ppm) and 1H multiplicities (J in Hz) for compounds 3–7 (solvent) C/H #
2 3 3a 4 5 6 7 7a 10 10 20 30 50 60 80 90 /100 100 /90 110 SMe 120 SMe 110 OMe OH NH NH NH
Polanrazine C 4 (CD3OD)
Polanrazine B 3 (CD2Cl2)
Polanrazine D 5 (CD3OD)
Polanrazine E 6 (CD3CN)
Polanrazine F 7 (CD3OD)
H
C
H
C
H
dC
H
C
H
C
7.20 s – – 7.39 d, 8.0 7.16 dd, 7.5, 7.5 7.21 dd, 7.5, 7.5 7.64 d, 7.8 – 3.28 d, 14.5 3.78 d, 14.5 – – – – 2.22 m 0.14 d, 7 0.92 d, 7 2.23 s 2.44 s – – 5.71 br s 6.09 br s 8.33 br s
125.4 108.3 128.3 111.5 120.3 122.5 119.1 136.4 36.1 36.1 67.4 166.1 69.2 166.7 33.1 16.0 15.0 14.5 14.7 – – – – –
7.15 s – – 7.29 d, 8 7.01 dd, 8, 8 7.07 dd, 7, 7 7.76 d, 8 – 3.13 d, 14 3.74 d, 14 – – – – 2.23 m 1.0 d, 7 1.04 d, 7 0.81 s – – – – – –
125.5 108.0 128.5 111.0 119.0 121.3 119.2 136.8 35.5 35.5 83.2 168.5 72.8 169.2 37.0 17.5 15.5 9.7 – – – – – –
7.11 s – – 7.27 d, 8 7.00 dd, 8, 8 7.05 dd, 7, 7 7.71 d, 8 – 3.17 d, 14 3.72 d, 14 – – – – 2.28 m 0.25 d, 7 0.62 d, 7 2.28 s – – – – – –
125.5 108.1 128.4 111.0 119.0 121.2 119.1 136.7 35.2 35.2 69.4 168.4 84.1 169.0 35.2 15.1 13.2 12.1 – – – – – –
7.10 s – – 7.33 d, 7.5 7.08 dd. 7 7.05 dd, 7 7.65 d, 7.5 – 3.14 d, 14 3.57 d, 14 – – – – 1.97 m 0.04 d, 7 0.62 d, 7 – – 3.18 s 4.75 br s 6.31 br s 7.21 br s 9.17 br s
125.9 108.1 128.6 111.7 119.5 121.9, 119.5 136.8 36.1 36.1 83.3 168.9 89.9 165.6 35.7 15.6 14.1 – – 51.5 – – – –
7.09 s – – 7.28 d, 7.5 6.99 dd. 8 7.05 dd, 7.5 7.70 d, 8 – 3.13 d, 14 3.66 d, 14 – – – – 1.82 m 0.15 d, 7 0.64 d, 7 – – – – – – –
125.2 107.6 128.4 110.9 118.9 121.2, 119.1 136.8 35.2 35.2 83.3 169.4 84.1 168.7 35.1 15.0 13.3 – – – – – – –
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range 190–600 nm), degasser, and a Hypersil ODS column (5 mm particle size silica, 4.6 i.d. 200 mm), equipped with a low-dispersion column-inlet filter. Two different elution methods were employed; method A: linear gradient from 75% H2O–25% CH3CN to 100% CH3CN, at a flow rate 1.0 ml/min over 35 min; method B: isocratic elution 80% H2O–20% CH3CN for 10 min, followed by gradient elutions, 80% H2O–20% CH3CN to 60% H2O–40% CH3CN for 10 min, and 60% H2O– 40% CH3CN to 25% H2O–75% CH3CN for 10 min, at a flow rate 1.0 ml/min. NMR spectra were recorded on a Bruker AMX 300 or 500 spectrometer; for 1H (300 or 500 MHz), values were referenced to CD3OD (CD2HOD at 3.31 ppm), CD2Cl2 (CDHCl2 at 5.32 ppm), or CD3CN (CHD2CN at 1.94 ppm), and for 13C (75.5 or 125.8 MHz) referenced to CD3OD (49.15 ppm), CD2Cl2 (54.00 ppm), or CD3CN (118.69 ppm). The list of coupling constants (J, reported to the nearest 0.5 Hz) corresponds to the order of the multiplicity assignment. Mass spectra (MS) [high resolution (HR), electron impact (EI), or fast atom bombardment (FAB)] were obtained on a VG 70 SE mass spectrometer, employing a solids probe. Specific rotations, []D were determined at ambient temperature on a Perkin-Elmer 141 polarimeter using a 1 ml, 10 cm path length cell; the units are 101 deg cm2 g1 and the concentrations are reported in g/100 ml. Infrared spectra were recorded on a Bio-Rad FTS-40 spectrometer. Spectra were measured by the diffuse reflectance method on samples dispersed in KBr; only diagnostic peaks are reported.
employing a 1:1 MeOH–H20 solution. Plants were incubated in a growth chamber as reported above (16 h light/8 h dark, at 24 2 C); leaves were observed over a 1-week period and the diameter of each lesion was measured after 7 days. 3.3. Isolation of compounds 3–7 The broth and mycelium were separated by gravity filtration, the broth (3 l) was concentrated (400 ml) and extracted with EtOAc (220 mg). Flash column chromatography (FCC) (eluted with CH2Cl2–MeOH, 95:5, 250 ml; 90:10, 250 ml of the EtOAc extract yielded the following fractions: F1 containing compounds 1 and 3, F2 containing compounds 2–4, F3–F4 containing compounds 2 and 5, F6–F9 containing compounds 6 and 7. Prep. TLC separation (CH2Cl2–MeOH, 97:3, double elution) of F1 (20 mg) yielded phomapyrone A (1, 4 mg) and polanrazine B (3), which was combined with a prep. TLC fraction eluted from F2 and further separated to yield chromatographically homogeneous material (3, 3.5 mg). Prep. TLC separation (CH2Cl2–MeOH, 96:4, double elution) of F2 (30 mg) yielded polanrazine B (3) which was combined with a prep. TLC band obtained from F1, and a mixture (20 mg) containing polanrazines C and D (4, 5). Separation of this mixture using multiple prep. TLC (CH2Cl2–MeOH, 96:4, multiple development) yielded chromatographically homogeneous materials 4 (6.5 mg) and 5 (2.5 mg). Finally prep. TLC separation (CH2Cl2–MeOH, 96:4, double elution) of F6-9 (6 mg) yielded polanrazine E (6, 1 mg) and F (7, 1.5 mg).
3.2. Fungal cultures and bioassays 3.4. Polanrazine B (3) The eight Polish isolates of P. lingam, IBCN 19-26, were obtained from the IBCN blackleg collection at Agriculture and Agri-Food Canada, Saskatoon. Liquid cultures of these isolates were grown as previously reported (Pedras and Biesenthal, 2000). For isolation of metabolites, isolate IBCN 19 was grown in 250 ml Erlenmeyer flasks containing 100 ml minimal medium (total 4 l) inoculated with fungal spores at 2 109 per flask were incubated on a shaker at 150 rpm, at 24 2 C for 6 days, and worked up as described below. Plants were grown in a growth chamber for 3 weeks, with 16 h light (fluorescent and incandescent, 450–530 mmol s1 m2)/8 h dark, at 24 2 C. Three different species of known resistance to V isolates of Phoma lingam were employed in the bioassays: Brassica napus, cv. Westar (susceptible) and B. juncea, cv. Cutlass (resistant), Sinapis alba, cv. Ochre (resistant). EtOAc extracts (2 mg/ml) or pure compounds (5104, 1104, 2105M) where dissolved in MeOH–H20 (1:1). Leaves were punctured with a needle (6 punctures per leaf) and 10 ml droplets (6 droplets per leaf) were applied on punctured sites. Control leaves were treated similarly
HPLC tR=22.4 min (solvent system B); []D60 (c 0.2 CHCl3); HR–EI–MS m/z (% relative abundance) measured: 377.1230 (377.1232 calc. for C18H23N3O2S2); FTIR max: 3388, 3190, 2926,1665, 1452, 1413, 1075 cm1; lmax (log ") 218 (4.5) nm. 3.5. Polanrazine C (4) HPLC tR=10.6 min (solvent system B); []D 16 (c 0.18 MeOH); HR–EI–MS m/z (% relative abundance) measured: 347.1306 (347.1304 calcd. for C17H21N3O3S1); FTIR vmax: 3335 3220, 2956, 2926, 1673, 1435, 1075 cm1; lmax (log ") 219 (4.3) nm. 3.6. Polanrazine D (5) HPLC tR=11.3 min (solvent system B); []D8.2 (c 0.18 MeOH); HREIMS m/z (% relative abundance) measured: 347.1306 (347.1303 calc. for C17H21N3O3S1. FTIR vmax: 3444, 3347, 2968, 2926, 1677, 1625, 1465, 1427, 1058 cm1; lmax( log ") 219 nm (4.0).
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3.7. Polanrazine E (6) HPLC tR=9.6 min (solvent system B); []D6 (c 0.07 MeOH); HREIMS m/z (% relative abundance) measured: 331.1531 (331.1532 calc. for C17H21N3O4). FTIR vmax: 3342, 3223, 2962, 2927, 2857, 1682, 1429, 1094 cm1; lmax (log ") 222 (4.3), 282 (3.7) nm. 3.8. Polanrazine F (7) HPLC tR=7.0 min (solvent system B); []D10 (c 0.26 MeOH); HR–EI–MS m/z (% relative abundance) measured: 317.1377 (317.1376 calcd. for C16H19N3O4); FTIR vmax: 3319, 2970, 2931, 1680, 1426, 1086, 1043 cm1; lmax ( log ") 219 (4.2), 281 (3.5) nm.
Acknowledgements We would like to thank G. Se´guin-Swartz and R. Gugel, Agriculture and Agri-Food Canada, Saskatoon Research Station, Saskatchewan, for kindly providing isolates from the International Blackleg of Crucifers Network (IBCN) collection available in Saskatoon, and K. Brown, Department of Chemistry, University of Saskatchewan, for NMR spectral measurements. Financial support from the Natural Sciences and Engineering Research Council (Canada) and the University of Saskatchewan is gratefully acknowledged. References Gugel, R.K., Petrie, G.A., 1992. History, occurrence, impact, and control of blackleg of rapeseed. Canadian Journal of Plant Pathology 14, 36–45. Howlett, B.J., Idnurm, A., Pedras, M.S.C., 2001. Leptosphaeria maculans the causal agent of blackleg disease of Brassicas. Fungal Genetics and Biology 33, 1–14. King, R.R., Lawrence, C.H., Calhoun, L.A., 1992. Chemistry of phytotoxins associated with Streptomyces scabies, the causal organism
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