Metabolism of the host-selective toxins destruxin B and homodestruxin B: Probing a plant disease resistance trait

Article abstract of DOI:10.1021/ol991042z

(matrix presented). Metabolism of the host-selective toxins destruxin B (1) and homodestruxin B (2) by plants resistant and susceptible to Alternaria blackspot (caused by the fungal pathogen Alternaria brassicae (Berk.) Sacc.) was established using synthetic radiolabeled compounds. The toxins are transformed into the less phytotoxic hydroxydestruxins 3 and 4. The rate of metabolic transformation was correlated with the plant's disease resistance, i.e., significantly faster rates were observed for plants resistant to the pathogen. Efficient syntheses of 1,2,3, and 4 are described.

Full text of DOI:10.1021/ol991042z

ORGANIC
LETTERS
1999
Vol. 1, No. 10
1655-1658
Metabolism of the Host-Selective Toxins
Destruxin B and Homodestruxin B:
Probing a Plant Disease Resistance
Trait
M. Soledade C. Pedras,* Irina. L. Zaharia, Yuanzhu Gai, Kevin C. Smith, and
,†
Dale E. Ward*
Department of Chemistry, UniVersity of Saskatchewan, 110 Science Place, Saskatoon,
Saskatchewan S7N 5C9, Canada
pedras@sask.usask.ca
Received September 13, 1999
ABSTRACT
Metabolism of the host-selective toxins destruxin B (1) and homodestruxin B (2) by plants resistant and susceptible to Alternaria blackspot
(caused by the fungal pathogen Alternaria brassicae (Berk.) Sacc.) was established using synthetic radiolabeled compounds. The toxins are
transformed into the less phytotoxic hydroxydestruxins 3 and 4. The rate of metabolic transformation was correlated with the plant’s disease
resistance, i.e., significantly faster rates were observed for plants resistant to the pathogen. Efficient syntheses of 1, 2, 3, and 4 are described.
Despite the continuing achievements of plant biotechnology,
engineering plants with disease resistance traits remains a
significant challenge. Recent strategies to control fungal
diseases using biotechnology have resulted in a race to obtain
plant resistance genes and transfer them to susceptible, but
agronomically valuable, crops.1 Different approaches have
been used for cloning such genes, a particularly difficult task
in the absence of a gene product. Nonetheless, host-selective
toxins (HSTs) might be one of the most attractive targets to
determine specific disease resistance traits in plants and to
guide the cloning of the corresponding disease resistance
gene(s).2,3 Subsequent plant genetic engineering with such
gene(s) can be carried out to obtain plants with the particular
resistance trait, similar to examples of herbicide, fungicide,
and insect resistance.4 HSTs are secondary metabolites
produced by plant pathogens which facilitate colonization
of host plants by the producing microorganism but do not
significantly affect non host plants. Host plants may be
resistant to a particular pathogen if they produce one or more
specific enzymes that metabolize and detoxify the pathogen’s
HST(s). Consequently, HST-detoxifying enzymes can confer
disease resistance traits to plants and are desirable gene
products useful for cloning disease resistance genes. Perhaps
not surprisingly, this potential application of HSTs has been
demonstrated in very few instances.3a
† e-mail: dale.ward@usask.ca.
As part of a research program aimed at understanding
disease resistance mechanisms, we have been evaluating the
role of HSTs produced by fungal pathogens of important
crops.5 Because destruxin B and homodestruxin B are HSTs
(1) For recent reviews, see, inter alia: (a) Staskawicz, B. J.; et al. Science
1995, 268, 661. (b) Cornelissen, B. J. C.; Melchers, L. S. Plant Physiol.
1993, 101, 709. (c) Lamb, C. J.; et al. In Biotechnology in Agriculture;
You, C. B., Chen, Z., Ding, Y., Eds.; Kluwer Academic Publishers:
Dordrecht, The Netherlands; 1993; p 45.
(2) Johal, G. S.; Briggs, S. P. Science 1992, 258, 985.
(3) For recent reviews, see: (a) Walton, J. D. Plant Cell 1996, 8, 1723.
(b) Walton, J. D.; Panaccione, D. G. Annu. ReV. Phytopathol. 1993, 31,
275 and references therein.
(4) Yoneyama, K.; Anzai, H. In Biotechnology in Plant Disease Control;
Wiley-Lyss, 1993; p 115.
10.1021/ol991042z CCC: $18.00 © 1999 American Chemical Society
Published on Web 10/20/1999

produced by one of the most damaging plant pathogenic
fungi (Alternaria brassicae (Berk.) Sacc.) of economically
important oilseed crops (e.g., rapeseed and canola, Brassica
napus, B. rapa),6 we became interested in investigating the
metabolism of destruxins by plants. Here we wish to report
the metabolism and detoxification of destruxin B (1) and
homodestruxin B (2) by plants resistant and susceptible to
the fungal pathogen, as well as the structure elucidation and
chemical synthesis of the metabolic products and their
phytotoxic activity. In addition, a new methodology for
synthesis of 14C-radiolabeled destruxins is summarized.
mate step. Thus, deprotection and coupling of 5a9 and 69
gave the pentadepsipeptide 7a in good yield. Without
purification of the intermediates, [14C-2]-â-alanine7 (8) was
converted into the Boc derivative and coupled with 7a, and
the resulting hexadepsipeptide was subjected to cyclization9
to give 17 in 30% overall yield. Radiolabeled 2 was similarly
prepared from 5b.10
The metabolism of 1 and 2 by leaves of plants resistant
(Sinapis alba cultivar Ochre) and susceptible (B. napus
cultivar Westar) to A. brassicae was investigated. Thus,
solutions of radiolabeled toxins were administered to peti-
olated leaves; after incubation in a growth chamber for
various periods, the leaves were individually extracted and
the extracts analyzed by LSC11 (ca. 70-100% recovery of
radioactive material) and by HPLC (equipped with a radio-
detector) (Table 1).12 HPLC analysis13 indicated that both
toxins were almost completely metabolized by S. alba within
24 h; destruxin B (tR ) 19.3 min) was biotransformed to a
metabolite with tR ) 10.4 min, here on called metabolite A,
and homodestruxin B (tR ) 21.5 min) was biotransformed
to a metabolite with tR ) 12.8 min, here on called metabolite
B. B. napus did not appear to metabolize any of the
destruxins to a detectable level within the same time period;
about 10% transformation occurred in 96 h.
The metabolism of destruxin B (1) and homodestruxin B
(2) by plant tissues was investigated using radiolabeled
compounds7 prepared as shown in Scheme 1.8 The previously
A close inspection of chromatograms shown in Figure 1
indicated that the transformation of 2 occurred within 24 h
for cultivar Ochre. Similar chromatograms were obtained for
1 incubated with Ochre. The data shown in Table 1 indicated
that the peaks observed in chromatograms corresponding to
metabolites A (3) and B (4) represented ca. 90% of the
toxins. That is, both destruxin B (1) and homodestruxin B
(2) were biotransformed by leaves of S. alba to metabolites
significantly more polar than each of the toxins within 24 h.
Scheme 1
reported9 route for synthesis of destruxin B was modified to
allow introduction of a radiolabeled residue in the penulti-
(5) (a) Pedras, M. S. C. Recent Res. DeVel. Agric. Food Chem., 1998, 2,
513. (b) Pedras, M. S. C.; Biesenthal, C. J. Can. J. Micro. 1998, 44, 547.
(c) Ward, D. E.; Va´zquez, A.; Pedras, M. S. C. J. Org. Chem. 1999, 64,
1657.
(6) Saharan, G. S. In Breeding Oilseed Brassicas; Labana, K. S., Banga,
S. S., Banga, S. K., Eds.; Springer-Verlag: Berlin, 1993; p 181. Canola
refers to varieties of rapeseed containing very low amounts of erucic acid
and glucosinolates.
Figure 1. HPLC chromatograms (radiodetection) of extracts of
leaves of Sinapis alba, cultivar Ochre (resistant to Alternaria
blackspot), and Brassica napus, cultivar Westar (susceptible to
Alternaria blackspot), incubated with 14C-homodestruxin B (2) up
to 96 h. For HPLC conditions, see ref 13.
(7) [14C-2]-â-Alanine (250 µCi, 55.5 × 107 dpm) was diluted with cold
material: 6 mg was used for 1; 10 mg was used for 2. Specific activities:
destruxin B, 1.28 × 107 dpm/mg; homodestruxin B, 7.94 × 106 dpm/mg.
(8) All new compounds gave satisfactory spectroscopic data.
(9) Ward, D. E.; Lazny, R.; Pedras, M. S. C. Tetrahedron Lett. 1997,
38, 339.
1656
Org. Lett., Vol. 1, No. 10, 1999

Table 1. Metabolism of Host-Selective Toxins (HST) Destruxin B (1) and Homodestruxin B (2) by Leaves of Sinapis alba, Cultivar
Ochre (resistant to A. brassicae), and Brassica napus, Cultivar Westar (susceptible to A. brassicae)
HST conversion (%)
per leafb determined by HPLCc
% of total radioactivity
recovered per leafb (LSC)
HST
(radioactivitya
added per leafb)
incubation
period
Ochre
Westar
Ochre
Westar
destruxin B (1)
(1.44 × 105)
0 h
22 ( 8
no conversion
97 ( 2
97 ( 4
24 h
48 h
72 h
0 h
87 ( 25
complete conversion
complete conversion
43 ( 2
no conversion
3 ( 4
6 ( 3
no conversion
96 ( 4
82 ( 7
93 ( 3
92 ( 1
81 ( 21
96 ( 3
95 ( 2
93 ( 3
homodestruxin B
(2) (1.22 × 105)
24 h
48 h
72 h
97 ( 2
90 ( 20
complete conversion
no conversion
4 ( 3
76 ( 11
77 ( 21
70 ( 8
92 ( 1
91 ( 1
93 ( 5
9 ( 6
a Units are in disintegration per minute (dpm). b Results are averages of experiments conducted in triplicate. c HPLC analysis as described in footnote 13;
areas of peaks determined in counts and converted to dpm utilizing a calibration curve. The total HPLC peak areas were within 10-20% of the LSC total
counting.
To isolate the metabolites, scale-up experiments were carried
out with nonlabeled destruxins utilizing ca. 300 leaves per
toxin (1 mL per leaf, 2 × 10-5 M in 2% aqueous CH3CN,
v/v). Following an incubation period of 48 h, the leaves were
extracted with EtOAc and the extracts fractionated, while
monitoring the presence of metabolites A and B by HPLC.
Multiple chromatographic fractionations yielded ca. 2 mg
of each metabolite.14 HR-EIMS analysis of metabolite A
(C30H51N5O8) indicated an additional oxygen compared to
singlets at δH 1.33, δC 31.4 and δH 1.30, δC 29.5 and an
additional quaternary carbon at δC 70.0. Thus, the spectro-
scopic data suggested that metabolite A contained a hydroxyl
group located either at the valyl or at the 2-hydroxy-3-
methylpentanoyl residue. Further proton-decoupling experi-
ments allowed the unambiguous assignment of structure 3
to metabolite A. Similarly, the spectroscopic data of me-
tabolite B obtained from homodestruxin B indicated it to
have structure 4. Although structures of at least 30 destruxins
are known,16 neither compound 3 or 4 appears to have been
previously reported. As expected, parallel experiments carried
out with leaves of B. napus and S. alba incubated under
similar conditions but without destruxins did not allow
detection of metabolites 3 or 4.
1
destruxin B (C30H51N5O7). H and 13C NMR spectroscopic
data15 of metabolite A (3) were very similar to those of
destruxin B except for the presence of two distinct methyl
(10) Prepared in 65% overall yield from Cbz-MeAla-NHNH-Boc, Cbz-
MeIle-OH, and Cbz-Ile-OH using PyBrop/DIEA for coupling.9
(11) Liquid scintillation counting was performed on a Beckman LS-6500
connected to a Wyse WY-370 data system, and results are quench corrected
and reported in dpm.
8.5 Hz, 1H), 2.07-1.91 (m, 4H), 1.87 (dd, 15, 3 Hz, 1H), 1.43 (ddd, J )
13.5, 7.5, 3.5 Hz, 1H), 1.33 (s, 3H), 1.31 (d, J ) 5 Hz, 3H), 1.30 (s, 3H),
1.26 (m, 1H), 0.93 (d, J ) 6.5 Hz, 3H), 0.89 (d, J ) 6.5 Hz, 3H), 0.87 (d,
J ) 7 Hz, 3H), 0.86 (t, J ) 7 Hz, 3H); 13C NMR (75.5 MHz, CDCl3) δ
173.8 (s), 173.6 (s), 171.3 (s), 171.2 (s), 170.0 (s), 169.9 (s), 71.1 (d), 70.0
(s), 61.0 (d), 58.3 (d), 55.7 (d), 53.9 (d), 46.7 (t), 43.2 (t), 37.7 (d), 34.8 (t),
33.4 (t), 31.4 (q), 31.1 (q), 29.5 (q), 29.2 (t), 28.3 (q), 27.5 (d), 24.6 (t),
24.3 (t), 20.2 (q), 19.9 (q), 15.6 (q), 15.4 (q), 11.6 (q); EIMS m/z (%) 609
(M+, 10), 552 (28), 86 (60), 70 (100); HREIMS m/z calcd for C30H51N5O8
(609.3738), found 609.3735; FTIR (cm-1) 3382, 3300, 2966, 1729, 1668,
1631, 1516, 1442, 1182. Spectroscopic data of metabolite B (4): HPLC tR
) 12.2 min; [R]24D ) -196 (c 0.14, MeOH); 1H NMR (300 MHz, CDCl3)
δ 8.23 (d, J ) 8.5 Hz, 1H), 7.17 (d, J ) 9 Hz, 1H), 5.18 (br q, J ) 6.5 Hz,
1H), 5.17 (dd, J ) 8.5, 3 Hz, 1H), 5.02 (d, J ) 11 Hz, 1H), 4.87 (dd, J )
9, 6.5 Hz, 1H), 4.66 (d, J ) 7 Hz, 1H), 4.05 (m, 1H), 3.90 (br t, J ) 9 Hz,
1H), 3.61 (br q, J ) 9 Hz, 1H), 3.20 (s, 3H), 3.08 (dddd, J ) 13.5, 11.5,
2, 2 Hz, 1H), 2.72 (s, 3H), 2.68 (ddd, J ) 18.5, 11, 2 Hz, 1H), 2.58 (ddd,
J ) 18.5, 5, 1.5 Hz, 1H), 2.46 (m, 1H), 2.11 (dd, 15, 8.5 Hz, 1H), 2.07-
1.91 (m, 5H), 1.87 (dd, 15, 3 Hz, 1H), 1.50-1.37 (m, 2H), 1.32 (s, 3H),
1.30 (d, J ) 5.5 Hz, 3H), 1.29 (s, 3H), 1.28 (m, 1H), 0.93 (m, 1H), 0.91
(m, 3H), 0.86 (t, J ) 7.5 Hz, 3H), 0.85 (d, J ) 6.5 Hz, 6H); 13C NMR
(75.5 MHz, CDCl3) δ 173.8 (s), 173.6 (s), 171.3 (s, 2×), 169.9 (s, 2×),
71.1 (d), 70.1 (s), 61.0 (d), 57.0 (d), 55.7 (d), 53.9 (d), 46.7 (t), 43.2 (t),
37.7 (d), 34.8 (t), 33.8 (d), 33.4 (t), 31.4 (q), 31.2 (q), 29.5 (q), 29.2 (t),
28.3 (q), 26.0 (t), 24.7 (t), 24.3 (t), 16.5 (q), 15.6 (q), 15.4 (q), 11.6 (q),
11.3 (q); EIMS m/z (%) 623 (M+, 12), 566 (28), 524 (18), 467 (10), 196
(14), 100 (100); HREIMS m/z calcd for C31H53N5O8 (623.3894), found
623.3891; FTIR (cm-1) 3385, 3297, 2961, 1730, 1672, 1631, 1514, 1441,
1230, 1182.
(12) Leaves were cut at the base of their petiole and immediately placed
in Eppendorf tubes containing the 14C-toxin (1 mL of 2.0 × 10-5
Μ
dissolved in 2% aqueous CH3CN). After the solution was taken up (this
time ) t0), an additional 0.5 mL of H2O was added to ensure the complete
uptake of compound. Leaves were incubated in a growth chamber under
fluorescent lighting (16-h light (22 °C)/8-h dark (18 °C) cycle) for 12, 24,
48, 72, or 96 h, keeping the petiole immersed in dd H2O.
(13) HPLC analysis was carried out with a high performance Hewlett-
Packard liquid chromatograph equipped with quaternary pump, automatic
injector, and photodiode array detector (wavelength range 190-600 nm)
connected with a Canberra Packard Radiomatic 150TR flow scintillation
analyzer (fitted with a 210 µL high performance flow cell Solarscint),
degasser, and a Hypersil ODS column (5 µm particle size silica, 4.6 i.d. ×
200 mm), equipped with an in-line filter. Mobile phase: 75% H2O-25%
CH3CN to 100% CH3CN, for 35 min, linear gradient, and a flow rate 1.0
mL/min. The 14C counting window was 15-100 keV with an update time
of 6 s.
(14) The EtOAc residue was fractionated by multiple column chroma-
tography (C-18 reversed phase silica gel, J. T. Baker, 40 µm) eluting with
acetonitrile-water, gradient elution 30:70 to 100:0. The fractions were
combined after HPLC analysis and further fractionated by PTLC on silica
gel, 250 µm, developed in acetone-hexane (65:35) to yield chromato-
graphically homogeneous material (TLC and HPLC).
(15) Spectroscopic data of metabolite A (3): HPLC tR ) 10.2 min;
25.5
1
[R]D
) -255 (c 0.10, MeOH); H NMR (300 MHz, CDCl3) δ 8.20 (d,
J ) 8.5 Hz, 1H), 7.18 (d, J ) 9 Hz, 1H), 5.19 (br q, J ) 7 Hz, 1H), 5.17
(dd, J ) 8.5, 3 Hz, 1H), 4.95 (d, J ) 11 Hz, 1H), 4.88 (dd, J ) 9, 6.5 Hz,
1H), 4.66 (d, J ) 7 Hz, 1H), 4.05 (m, 1H), 3.90 (br t, J ) 9 Hz, 1H), 3.61
(br q, J ) 9 Hz, 1H), 3.23 (s, 3H), 3.08 (dddd, J ) 13.5, 11.5, 2, 2 Hz,
1H), 2.72 (s, 3H), 2.68 (ddd, J ) 18.5, 11, 2 Hz, 1H), 2.56 (ddd, J ) 18.5,
5, 2 Hz, 1H), 2.47 (m, 1H), 2.32 (dsept, J ) 11, 6.5 Hz, 1H), 2.11 (dd, 15,
(16) All the destruxins reported so far were isolated from diverse fungal
species; for recent reports see, for example: (a) Krasnoff, S. B.; Gibson,
D. M. J. Nat. Prod. 1996, 59, 485. (b) Cai, P.; Smith, D.; Katz, B.; Pearce,
C.; Venables, D.; Houck, D. J. Nat. Prod. 1998, 61, 290.
Org. Lett., Vol. 1, No. 10, 1999
1657

Next, to establish if the hydroxylation of destruxins
represented in fact a metabolic detoxification, it was crucial
to determine the phytotoxicity of hydroxydestruxins 3 and
4 to plants resistant and susceptible to A. brassicae. Total
synthesis of 3 and 4 was necessary because of the lengthy
and difficult separations required to obtain these hydroxy-
destruxins from plant tissue. The required (2R)-2,4-dihy-
droxy-4-methylpentanoic acid residue has been obtained in
very low yield by resolution but was found to be highly prone
to lactonization and racemization.17 To prevent these pro-
cesses, the silyl ether 10 was prepared from the readily
available lactone 918 (Scheme 2).8 Enantioselective hydroxyl-
yield. In analogy with the previously described synthesis of
1,5a coupling of fragments 5a and 13 followed by cyclization
provided 3 in 41% overall yield; similar processing of 5b
and 13 gave 4 in 37% yield. As expected, the chemical
synthesis of both hydroxydestruxins 3 and 4 provided
products identical in all respects with metabolites A (3) and
B (4), thus corroborating the chemical structure assignments.
With both compounds 3 and 4 in hand, leaf puncture and
leaf uptake bioassays were carried out; results of both
bioassays indicated that the hydroxylated compounds were
less phytotoxic than each of the destruxins. In addition, a
bioassay with cell suspension cultures of both B. napus and
S. alba was employed to compare 1 and 3; the toxicity of
both 1 and 3 was evaluated by determining the percentage
of viable cells after exposure to each compound for different
periods. The results of these bioassays were similar to those
observed with the leaf assays and indicated that the percent-
age of viability of cells exposed to destruxin B (1) was about
50% lower than the percentage of viability of cells exposed
to hydroxydestruxin B (3). Moreover, 3 was no more
phytotoxic to the susceptible cultivar Westar than to the
resistant cultivar Ochre. Altogether these bioassay results
indicate that the metabolism of destruxins 1 and 2 to the
respective hydroxylated products are detoxifications occur-
ring at a faster rate in Ochre plants, i.e., plants resistant to
A. brassicae. Most importantly, investigation with two
additional plant cultivars, a disease resistant and a susceptible,
showed similar behavior, indicating that destruxin and
homodestruxin B detoxification correlated with Alternaria
blackspot resistance. Consequently, it is probable that this
biochemical trait is involved in the blackspot resistance
shown by Ochre; if this is true Westar might be expected to
become blackspot resistant if transformed with the Ochre
destruxin B hydroxylating gene.
Scheme 2
Acknowledgment. Financial support from the Natural
Sciences and Engineering Research Council of Canada
(Strategic Project Grant to M.S.C.P. and D.E.W.) and the
University of Saskatchewan is gratefully acknowledged.
ation of 10 using the method of Evans et al.19 provided 12
in good yield. We were unable to hydrolyze 12 without
extensive concomitant γ-lactonization. By contrast, acylation
of 12 with Boc-â-Ala followed by hydrolysis and coupling
with Pro-OBn gave the tridepsipeptide 13 in good overall
OL991042Z
(18) Stevens, C. M.; Tarbell, D. S. J. Org. Chem. 1954, 19, 1996.
(19) Evans, D. A.; Morrissey, M. M.; Dorow, R. L. J. Am. Chem. Soc.
1985, 107, 4346.
(17) Mori, K.; Takigawa, T.; Matsuo, T. Tetrahedron 1979, 35, 933.
1658
Org. Lett., Vol. 1, No. 10, 1999