Department of Plant Sciences1, Department of Chemistry and Biochemistry2 and Department of Entomology3, Montana State University, Bozeman, MT 59717, USA
Author for correspondence: Gary A. Strobel. Tel: +1 406 994 5148. Fax: +1 406 994 7600. e-mail: uplgs{at}montana.edu
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Keywords: endophyte, organic volatiles, insects
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Some of these endophytes produce bioactive substances that are involved in the hostmicrobe relationship. While there are many epiphytic micro-organisms associated with plants, the endophytic associations may be more complex since living host tissues are involved. Thus, closer biological associations may have developed between these endophytes in their respective hosts than is the case with epiphytes or soil-related organisms. The result of these associations may be the production, by the endophyte, of bioactive chemicals that are involved in the relationship. This assumption has prompted our search for novel endophytes and an examination of their secondary bioactive products. One remote and extremely biodiverse area of the world in which we focused our search for unique microbeplant relationships is the Manu region of the upper Amazon.
Recently, we described two novel endophytic fungi, Muscodor albus from Cinnamomum zeylanicum from Honduras (Worapong et al., 2001 ), and Muscodor roseus from two monsoonal rainforest trees in Northern Australia (Worapong et al., 2002
). These endophytes produce a mixture of volatile antimicrobials that effectively inhibit and kill a wide spectrum of plant-associated fungi and bacteria (Strobel et al., 2001
). On the other hand, the gases of M. albus did not kill fungi that were related to it, some of which were producers of other lethal gas mixtures (Worapong et al., 2001
, 2002
). Thus, using M. albus as a selection tool, it was possible to isolate still other endophytic fungi that produce volatile compounds of biological interest and importance.
In this report we describe some of the biology and biochemistry of another endophytic Muscodor sp. that was obtained by using the volatiles of M. albus as a selection tool. Representive forest plants, collected in the southern Peruvian Amazon, were examined for the presence of novel Muscodor spp. From this effort we isolated Muscodor vitigenus, which is endophytic on a liana identified as Paullina paullinioides. This unique endophytic fungus produces naphthalene in quantities sufficient to cause modifications in insect behaviour, and this is the subject of this report.
![]() |
METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Quantitative and qualitative analyses of M. vitigenus volatiles.
The gases in the air space above the mycelium of M. vitigenus growing in Petri plates were analysed by trapping the fungal volatiles with a solid-phase micro-extraction syringe and injecting them into a gas chromatograph interfaced to a mass spectrometer, as described by Strobel et al. (2001) . Initial identification of the unknown compounds produced by M. vitigenus was made through library comparison using the NIST database. Control analyses were conducted using Petri plates containing only PDAN; the compounds obtained from these controls were deleted from the analyses done on Petri plates containing PDAN and M. vitigenus.
Synthetic naphthalene (Willert Home Products, St Louis, MO, USA) was used to create an artificial atmosphere mimicking that produced by M. vitigenus for use in preliminary testing of bioactivity. It too was subjected to the GC/MS assay as described above.
Source and handling of insects.
The wheat stem sawfly (Cephus cinctus) was selected as a model for its behavioural responses to naphthalene because of its availability, its economic importance, and the experience that we have in dealing with this insect. It is a major pest in the winter-wheat-growing areas of the Great Plains of North America that cannot be controlled sufficiently with current agricultural management practices (Morrill et al., 2001 ). Overwintering pre-pupae were collected from a wheat field in September after the 2001 harvest in Broadwater County, MT. Stubble was removed from this heavily infested field with a shovel and taken to the laboratory. The uninfested residue was separated from wheat stems containing sawfly larvae. These sawfly-cut stems were stored at 4 °C for at least 100 days to complete diapause. The sawfly-cut stems were then removed from the refrigerator and placed in individual shell vials for 45 weeks to complete metamorphosis. Vials were sealed with a cotton plug that was wetted twice weekly with distilled water. Upon adult emergence, insects were held in darkness, at room temperature, in the shell vial until used.
Collection of volatiles and analytical methods for insect repellency bioassay tests.
To determine the rate of naphthalene release from M. vitigenus cultures, collections of volatile compounds were made from circular (0·785 cm2) plugs of the fungus growing on PDAN. The volatile collection apparatus used was that described by Heath & Manukian (1992) . Specifically, agar plugs containing the 14-day-old fungus culture were placed in Pyrex glass volatile collection chambers (1·5 cm diameter x 8 cm long) fitted at one end with an air diffuser inlet cap and sealed at both ends with no. 7 ChemThread inlets [0·25 inch (0·64 cm) i.d.] using rubber O-rings (Analytical Research Systems). A volatile collector trap [0·25 inch o.d.x3 inch long (0·64 cmx7·62 cm) glass tube] containing 30 mg SuperQ adsorbent (Alltech Associates) was inserted into one end of the volatile collection chamber. Purified air was delivered at a rate of 0·3 l min-1 over the fungal plugs, and the flow and pressure were maintained by both a compressed air source and a vacuum pump. Volatiles were collected for 3 h. Fungal plugs from four colonies on individual Petri plates containing PDAN were used and the volatile aerations were replicated three times for each colony.
Volatiles were eluted from the SuperQ in each volatile collection trap with 225 µl hexane, and 7 ng decane was added as an internal standard. Volatiles were analysed by coupled GC/MS. The GC was an Agilent Technologies 6890 instrument fitted with a 30 m DB-1MS capillary column (0·25 mm i.d. with a 0·25 µm film thickness; J & W Scientific). The temperature programme increased from 50 °C to 280 °C at 10 °C min1. The MS instrument used was an Agilent Technologies 5973. Authentic naphthalene was used as a standard.
Insect repellency bioassay tests.
Insect repellency bioassay tests were done with a Y-tube olfactometer (Fig. 1). The olfactometer design incorporated two 38 cm long glass tubes upwind of the Y-tube for the stimulus and control odour sources. Each tube was made of Corning 22 mm o.d. glass with a 24/25 inner ground-glass joint, which connected it to either of the upper arms of the Y-tube. A charcoal-purified and humidified airstream was connected to these tubes by a Teflon-lined, threaded 24-410 cap coupled to a 0·64 cm Swagelock union that delivered air through 0·64 cm o.d. Teflon tubing. The airflow was set at 0·8 l min-1 using a flowmeter. The lower arm of the Y-tube was a 30 cm length of 28 mm o.d. Corning glass tubing that branched at 20 cm from the tip to form the upper arms. The interior angle of the upper arms of the Y was 120°, and the arms branched for 4 cm before becoming parallel for the final 10 cm of each arm.
|
Y-tube olfactometry with authentic naphthalene as odour source.
Naphthalene (in acetone) was delivered to a circular (6 mm diameter) piece of filter paper (created using a paper hole punch) and this, along with a wheat plant (natural host of the insect) served as odour sources in the Y-tube (cf. Fig. 1). Naphthalene (10 µl) in acetone was used in two insect behaviour tests: (A) 0·05 g naphthalene (ml acetone)-1, and (B) a 10-fold dilution of (A). The release rate of naphthalene from test system (A) was approximately 12 ng h-1. The rate of release of naphthalene on filter paper was determined using the volatile collection methodology described above. Thirty successful trials were performed for each of the two solutions. The control consisted of a wheat plant and 10 µl acetone placed on a piece of filter paper (also 6 mm diameter). Wheat (Triticum aestivum L., McNeal) was grown in an insect-free growth chamber (15 h light:9 h dark; 22 °C day/20 °C night). Plants at growth stage 30 (Zadoks et al., 1974
) were carefully pulled from the soil and the roots were wrapped with wet cotton prior to use.
Y-tube olfactometry with M. vitigenus as odour source.
In all trials, a wheat plant and a circular (0·785 cm2) fungal plug of M. vitigenus served as odour sources. The control consisted of a wheat plant and a circular (0·785 cm2) plug of uninoculated PDAN. The stage and handling of wheat plants were as described above.
![]() |
RESULTS AND DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
M. vitigenus compounds as insect repellents
M. vitigenus, in limited insect bioassay tests, demonstrated insect repellency. In controlled, replicated experiments, both authentic naphthalene and M. vitigenus were used as odour sources in Y-tube olfactometry tests. The responses of C. cinctus, a major local crop pest, were measured with plugs of the naphthalene-producing fungus as compared to comparable amounts of authentic naphthalene. In 30 successful trials using naphthalene test solution A (naphthalene release rate 12 ng h-1), 24 out of 30 insects responded by moving away from the naphthalene source (Table 1). In a second set of 30 successful trials using naphthalene test solution B, 21 out of 30 insects were repelled by the naphthalene source. Finally, 60 successful trials were performed using 0·785 cm2 fungal plugs (23 weeks old) of M. vitigenus as the odour source. Out of the 60 insects, 49 moved away from the fungus and its volatiles. The results of all three sets of tests were significantly different from the respective controls (Table 1
). Finally, many of the insects tested became more disoriented than usual in the presence of both commercial naphthalene and the M. vitigenus volatiles.
|
It remains unknown whether M. vitigenus can grow successfully in nature in plants other than P. paullinioides. However, the fungus has proven to be uneven in its production of naphthalene, with a strong dependency upon the presence of starch as an energy source and precursor to naphthalene. On occasion, individual mycelial transfers to a new Petri plate containing PDAN will fail to produce naphthalene (B. H. Daisy & G. A. Strobel, unpublished). It is a common observation that plant-associated fungi will sometimes halt production of interesting active products when cultured in artificial media (Pinkerton & Strobel, 1976 ). This could result from the absence of critical activator compounds that are present in the host (Pinkerton & Strobel, 1976
). We have not yet been able to determine the optimum conditions for the maximum production of naphthalene by this fungus.
The presence of naphthalene in M. vitigenus is important in understanding the biology of the fungus and P. paullinioides as well as their evolutionary relationship. A number of endophytes have been isolated that make biologically active products that have selective activity against potential threats to the host plant, especially in the Muscodor group (Strobel et al., 2001 ). M. vitigenus is quite different from the other Muscodor isolates in that it produces a distinct set of volatile compounds that are less lethal to micro-organisms but much more inhibitory of arthropods. It appears that these gas-producing fungi are not restricted to just one area of the world: they have been recovered from Australia, Honduras, and now Peru. It remains a mystery why these nearly genetically identical fungi with similar functions can exert such biologically diverse phenomena in such a diversity of places. The discovery of this fungus again adds support to the growing necessity to conserve these areas of the world harbouring yet untold numbers of endophytic micro-organisms.
![]() |
ACKNOWLEDGEMENTS |
---|
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Bacon, C. W. & White, J. F., Jr (2000). Microbial Endophytes. New York: Marcel Dekker.
Bolton, D. M. & Eaton, L. G. (1968). In MERCK Index, 8th edn, p. 713. Edited by P. G. Stecher, M. Windholz & D. S. Leahy. Rahway, NJ: Merck.
Chen, J., Henderson, G., Grimm, C. C., Lloyd, S. W. & Laine, R. A. (1998). Termites fumigate their nests with naphthalene. Nature 392, 558-559.
Daisy, B. H., Strobel, G. A., Ezra, D., Castillo, U., Baird, G. & Hess, W. M. (2002). Muscodor vitigenus, sp. nov., an endophyte from Paullinia. Mycotaxon 81 (in press).
Heath, R. R. & Manukian, A. (1992). Development and evaluation of systems to collect volatile semiochemicals from insects and plants using a charcoal-infused medium for air purification. J Chem Ecol 18, 1209-1226.
Morrill, W. L., Weaver, D. K. & Johnson, G. D. (2001). Trap strip and field border modification for management of the wheat stem sawfly (Hymenoptera: Cephidae). J Entomol Sci 36, 34-45.
Pinkerton, F. & Strobel, G. A. (1976). Serinol as an activator of toxin production in attenuated cultures of H. sacchari. Proc Natl Acad Sci USA 73, 4007-4011.[Abstract]
Sokal, R. R. & Rohlf, F. (1981). The Principles and Practice of Statistics in Biological Research. New York: W. H. Freeman.
Stone, J. K., Bacon, C. W. & White, J. F. (2000). An overview of endophytic microbes: endophytism defined. In Microbial Endophytes , pp. 3-29. Edited by C. W. Bacon & J. F. White. New York:Marcel Dekker.
Strobel, G. A., Dirske, E., Sears, J. & Markworth, C. (2001). Volatile antimicrobials from Muscodor albus, a novel endophytic fungus. Microbiology 147, 2943-2950.
Worapong, J., Strobel, G. A., Daisy, B. H., Castillo, U., Baird, G. & Hess, W. M. (2001). Muscodor albus anam. gen. et sp. nov., an endophyte from Cinnamomum zeylanicum. Mycotaxon 79, 67-79.
Worapong, J., Strobel, G. A., Daisy, B. H., Castillo, U., Baird, G. & Hess, W. M. (2002). Muscodor roseus sp. nov., an endophyte from Grevillea pteridifolia. Mycotaxon 81, 463-475.
Zadoks, J. C., Chang, T. T. & Konsak, C. F. (1974). A decimal code for the growth stages of cereals. Weed Res 14, 15-21.
Received 17 May 2002;
revised 19 July 2002;
accepted 22 August 2002.