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Department of Plant Pathology, University of Kentucky, 201F Plant Science Building, 1405 Veterans Drive, Lexington, Kentucky 40546
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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The maize anthracnose stalk-rot fungus Colletotrichum graminicola infects its host primarily through wounds in the stalks that are caused by insects. However it also can cause stalk-rot disease without wounding. It is not known how the pathogen enters stalks in the absence of wounds. Studies have suggested that direct invasion through the highly lignified rind tissues is not a viable means of entry. A cytological approach was used to investigate the ability of C. graminicola to penetrate and colonize intact maize stalks. The pathogen had a significant capacity for direct penetration, but this mechanism of infection was much slower and less efficient than penetration through wounds. The fungus breached the lignified rind fibers by passing through small openings in the cell walls via narrow hyphal connections. Epidermal cells and rind fiber cells did not appear to become rotted. Rotting only occurred once the pathogen had penetrated into the pith parenchyma cells. To our surprise the closely related fungus C. sublineolum, which is not normally a pathogen of maize, also was capable of infecting intact maize stalks, although to a lesser degree than C. graminicola. The two species also were observed on intact roots and leaves, and C. sublineolum was incapable of infecting those tissues whereas C. graminicola efficiently colonized both. This suggests the interesting possibility that nonhost resistance to C. sublineolum is conditional and perhaps also tissue-specific.
Key words: corn stalk-rot disease, green fluorescent protein, nonhost resistance
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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). A synergistic relationship has been established between wounding caused by insects such as the European corn borer and the incidence of anthracnose stalk rot and other maize stalk-rot diseases (Bergstrom et al 1983, Keller et al 1986, Gatch and Munkvold 2002a, b). Controlling insect damage by the use of transgenic maize expressing the Bacillus thuringiensis Bt toxin reduces the incidence of some stalk rots (Bergstrom et al 1997, Munkvold et al 1997, Gatch and Munkvold 2002a, b). However anthracnose stalk rot often occurs in the absence of insect wounding and the proportion of stalk rots caused by C. graminicola actually increased in Bt maize (Dodd 1980, Gatch and Munkvold 2002a). We do not know how C. graminicola enters stalk tissues in the absence of wounds. C. graminicola also causes leaf blight and seedling blight of maize, and it has been suggested that the pathogen gets into unwounded stalks via infected leaves or seedling roots (Bergstrom and Nicholson 1999). It also has been proposed that it can penetrate directly through the rind (consisting of the stalk epidermis and subtending rind fiber cells) (Williams and Willis 1963, Bergstrom and Nicholson 1999, Munkvold and Hellmich 1999). However, when attempts were made to produce infections in highly susceptible Jubilee sweet corn plants in the field by placing inoculum behind leaf sheaths in contact with the rind, stalk infections occurred in fewer than 20% of the inoculated plants and these infections usually were associated with stalk borer injuries (White and Humy 1976). Thus the rind appears to be an effective impediment to direct penetration by C. graminicola, perhaps due to the presence there of a layer of highly lignified fiber cells that can act as a physical barrier. Understanding how C. graminicola enters stalks in the absence of wounds will become increasingly important as production of Bt maize continues to expand and direct penetration presumably becomes the predominant means of infection by this important pathogen. To address this question a cytological approach was used to investigate the ability of C. graminicola to penetrate intact Jubilee maize tissues. The results demonstrated that C. graminicola has a significant capacity for penetration of unwounded rind tissues but that this mode of entry is neither as rapid nor as efficient as wound-associated infection.
C. sublineolum, which is closely related to C. graminicola, causes stalk rot and leaf blight on sorghum, but this species does not appear to infect maize in the field (Dale 1963, Jamil and Nicholson 1987, LeBeau 1950, Williams and Willis 1963). Thus it was a surprise to discover that C. sublineolum colonized healthy, unwounded Jubilee maize stalk epidermal cells almost as efficiently as C. graminicola and that C. sublineolum also had some ability to penetrate the rind into deeper tissues. This phenomenon was investigated further by comparing the ability of this C. sublineolum strain versus C. graminicola to infect unwounded maize leaves (either seedling or mature) and seedling roots. C. graminicola efficiently infected and colonized all these tissues, but C. sublineolum did not successfully colonize any of them. This suggests the interesting possibility that nonhost resistance of maize to C. sublineolum is conditional and perhaps tissue specific.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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. Protoplasts were transformed with 3–9 µg of plasmid pCT74, which contains an SGFP gene under the control of the Pyrenophora triticirepentis TOX-A promoter, and a hygromycin phosphotransferase gene as a selectable marker (Lorang et al 2001). The plasmid was linearized with EcoR1 before use in transformation. To produce a transformant of C. sublineolum strain CgS11 a protocol developed for Agrobacterium-mediated transformation of falcate spores was used (Flowers and Vaillancourt 2005). Plasmid pBin-GFP-hph, which contains GFP and hygromycin resistance genes under the control of the A. nidulans trpC promoter (O’Connell et al 2004), was used for the transformation. All transformants were single-spored and stored on silica crystals at –80 C.
The two GFP-transformants used in this study also were included in a previous investigation of infection and colonization of wounded maize stalk tissues (Venard and Vaillancourt 2007). Several additional independent GFP transformants were tested in that previous report; they behaved similarly to the representative ones that were chosen for the current analysis. It also was demonstrated in the previous study that the C. graminicola GFP-transformant used here was slightly less pathogenic than the wild-type strain in wounded stalks, whereas the C. sublineolum GFP transformant strain was equivalent to its wild-type progenitor (Venard and Vaillancourt 2007).
Fungal strains were cultured on half-strength oatmeal agar (Difco) at 23 C for 2 wk under continuous light. Falcate spores were collected by adding 10 mL of sterile water and rubbing the surface of the culture gently with a plastic minipestle. The conidial suspension was filtered though sterile glass wool, and the conidia were washed 3x in sterile water. The concentration of conidia was adjusted to 5 x 106 spores per milliliter after the third wash. For leaf and root inoculations, 0.01% Tween-20 was added to the spore suspensions. A total of 10 µL of the spore suspension was applied for each plant inoculation. Controls were inoculated only with water (or water with 0.01% Tween-20, in the leaf and root experiments).
Plant growth and inoculation.— –
The sweet corn hybrid Jubilee was chosen for this study because it is highly susceptible to anthracnose stalk rot and leaf blight; it matures quickly; it does not exceed a manageable mature size for the greenhouse; and it has a historical association with the first epidemics of anthracnose in the United States (Warren et al 1973). Jubilee was used for the field experiments, mentioned above, that tested the ability of C. graminicola to penetrate through the intact rind (White and Humy 1976). Jubilee sweet corn seed was a generous gift of Syngenta (Research Triangle Park, North Carolina), and was obtained from Rogers Seed Co. (Boise Idaho).
For stalk inoculations, plants were grown in the greenhouse as described by Venard and Vaillancourt (2007). Sheath tissue was stripped from the second and third internodes of the plants above the soil line just before anthesis. Plants were placed on their sides, and a drop of spore suspension, or water as a control, was placed in the center of each of the stripped internodes. In some cases only one of the two internodes was inoculated, while in others both were inoculated. In wounded control treatments a small puncture wound 2 mm deep was made through the rind with a dissecting needle and drop of spore suspension was placed on the wound (as described in Venard and Vaillancourt 2007). All drops were covered with detached microfuge tube caps and sealed with parafilm to create a moist chamber, and the plants were left overnight. The tube caps then were removed, and the plants were placed upright. Individual inoculated internodes were collected for observation each day starting at 1 d post-inoculation (dpi) and continuing for up to 9 dpi.
For leaf inoculations 10µL drops of spore suspension were placed in individual wells of microhumidity chambers (Bergstrom and Nicholson 1983). Inoculations were done late in the afternoon. The chambers were clamped onto the leaves, which remained attached to the plants. The chamber wells were covered with adhesive tape, which remained in place overnight. Chambers were removed from the leaves the next morning (14–18 h later) and individual infection sites were excised with a scalpel and examined at 2–14 dpi. Inoculated mature leaves were located at the 2nd internode position of plants just before anthesis, grown as described above. At this stage the leaves were mature but not yet showing signs of senescence. Inoculated seedling leaves were the first, fully expanded true leaves of Jubilee seedlings at the V2 leaf stage in conetainers (one plant per conetainer) containing a mixture of 3 parts Pro Mix BX/2 parts sterilized topsoil.
For seedling root inoculations Jubilee seeds were surface-sterilized by soaking in 10% bleach solution for 10 min, followed by three rinses in sterile milli-Q water. Seeds were rolled into several layers of water-saturated germination paper (Anchor Paper Co., St Paul, Minnesota), forming a loose tube. The tube was placed upright in a beaker of sterile tap water so that the seeds were not in direct contact with the water but were kept moist by the wicking action of the paper, and incubated in the dark at 25 C. Water was replenished as needed to maintain constant saturation of the germination paper. After 2 d primary radicles had emerged from the seeds, and by 5 d these were 1–1.5 inches long. The primary roots were inoculated with 10µL drops of spore suspension just behind the root elongation zone. The roots were left exposed about an hour to let the spores attach, and then the inoculated roots were wrapped again in the saturated germination paper and kept moist in the dark until observation. Individual seedlings were removed from the paper, and infected roots were observed under the microscope at 1, 2 and 3 dpi.
Microscopy.— –
Development of fungi in aerial plant tissues was monitored with a Leica TCS NT confocal microscope (Leica Microsystems Inc., Exton Pennsylvania). Infected leaf pieces were observed without sectioning or staining. Infected stalk tissue sections were cut by hand with a razor blade and observed without further treatment. In both cases GFP was excited at 488 nm. Plant cell walls and chloroplasts autofluoresce at this excitation wavelength, and this property was used to view plant tissues. However at lower magnifications autofluorescence of the plant cell walls was usually not intense enough to be captured in the micrographs. Root tissues were monitored under a Zeiss Axioscop compound microscope equipped with epifluorescence. Longitudinal sections of infected roots were produced by hand with a razor blade. Transverse sections of infected roots were obtained by embedding roots in 5% agarose in PBS buffer and sectioning with a Vibratome (using a protocol adapted from Koltai and Bird 2000). Inoculated roots were cut into 2–3 mm long pieces and immediately fixed in fresh FAA solution at 4 C at least 24 h. The fixed samples were rinsed in PBS buffer and embedded. The agarose was kept at 45 C, just above its solidification temperature, to avoid damaging the samples. When they solidified the blocks of agarose were divided so that each section contained an individual root sample. These smaller blocks were glued for sectioning on a Vibratome, according to the instructions provided by the manufacturer (Vibratome®1000 Plus Sectioning System, The Vibratome Co., St Louis, Missouri). Sections 50–100µm long were cut in a distilled water bath. The agarose usually detached from the root tissues at this point, and the released sections were mounted for viewing in a drop of distilled water under a cover glass.
Statistical analysis.— – Lesion lengths were measured as the maximum length of visible discoloration on the surfaces of uncut, inoculated stalks. Colonization of epidermal cells and of tissues below the epidermal cells was recorded as a percentage, consisting of the number of plants in which these events were seen as a proportion of the total number of plants that were observed. Each experiment included two or three replications of each treatment, and each experiment was repeated at least twice. Data were analyzed with the Mixed procedure that is part of the SAS statistical analysis software package (SAS Institute Inc., 1997). For this procedure, time elapsed postinoculation, fungal strain and treatment (inoculation of the second versus the third internode and inoculation of one versus both internodes on the same plant) were treated as fixed effects, while plants were treated as random effects. The effects of different strains, different treatments and the time elapsed post-inoculation, on lesion length and on the degree of colonization of epidermal cells and deeper tissues, were analyzed.
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RESULTS |
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). No significant effect of internode position (second versus third above the soil line) or of the number of internodes inoculated per plant on lesion length (P = 0.7457) were observed and so the data from different internodes were pooled for all subsequent analyses. Inoculation of stalks with C. graminicola in the absence of wounding did not produce infections as rapidly or as efficiently as wound inoculation. Thus, of the 38 C. graminicola inoculations of unwounded stalk tissues that were observed, only 29 of them (76%) actually became infected at some point during the 9 d course of the experiments, even though the fungus germinated and produced abundant Appressoria on all of the inoculated stalks within 24 h. In contrast all 25 control stalks that were inoculated after wounding became colonized by the fungus within 48 h, a result that was consistent with a previous, more extensive study of infection of wounded stalks (Venard and Vaillancourt 2007).
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Substantial colonization of the epidermal cells of unwounded stalks always preceded invasion of rind fibers and deeper tissues, but deeper colonization did not always occur when epidermal infection was observed, at least within the course of this study. The percentage of deeper colonization did increase for both strains over time (P = 0.0023). Inoculation with C. graminicola resulted in colonization of deeper tissues 60% of the time by 7–9 dpi. Deeper colonization was significantly less frequent for C. sublineolum (P = 0.0062), occurring in only 15% of inoculation sites at 7–9 dpi. It also was delayed, with no deeper colonization appearing earlier than 7 dpi, whereas for C. graminicola colonization of deeper tissues occurred in as few as 2 dpi.
Infection of unwounded maize stalks by C. graminicola and C. sublineolum.— –
Infections of control wounded maize stalks by both strains proceeded as described previously (Venard and Vaillancourt 2007). The process by which epidermal cells of unwounded stalks were infected was similar to published descriptions of infection of leaf epidermal cells (Bergstrom and Nicholson 1999, Mims and Vaillancourt 2002, Politis and Wheeler 1973) and also to our own observations of this process (see below). Appressoria formed on stalk epidermal cell surfaces within 24 h postinoculation (hpi), and the first infection hyphae were visible within 48 h. Appressoria were formed by both strains, with no obvious differences between them in the efficiency of this process. Within 2 d a network of hyphae was visible for both strains within the epidermal cells in some of the stalks (FIG. 1a, g). The hyphae passed from cell to cell via extremely narrow connections (FIG. 1c). Discoloration of the epidermal tissues occurred within 1 or 2 d, and then formation of stromata by both strains was observed (FIG. 1b, i). These stromata produced acervuli and conidia if they were incubated in a moist environment (not shown).
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). Movement from cell to cell again occurred via narrow connections through cell walls that otherwise appeared to remain intact. Once the parenchyma cells were invaded, signs of rotting finally began to appear, including destruction of cell walls and formation of a cavity within the pith (not shown). C. sublineolum entered the tissues beneath the epidermis less frequently, but when it did so it appeared to use a mechanism similar to that of C. graminicola (FIG. 1h)
Infection of leaves by C. graminicola.— –
The ability of C. sublineolum to colonize unwounded stalk epidermal tissues so efficiently was surprising, and this was investigated further by comparing this strain with C. graminicola regarding its ability to colonize other types of unwounded Jubilee maize tissues. A total of 38 seedling leaves were inoculated with C. sublineolum and examined, and the following descriptions represent the consensus of all observations. C. sublineolum formed abundant appressoria on all inoculated leaves, but only three successful penetrations into the tissues resulted (out of many hundreds of appressoria on each leaf), and only one primary infection hypha was observed (FIG. 2a). C. graminicola in contrast entered all of the seedling leaves onto which it was inoculated within 48 hpi and had proliferated and begun to cause significant tissue collapse 24 h later. The fungus was observed to produce numerous swollen primary infection hyphae and progress from cell to cell in a manner similar to that described for epidermal colonization in stalks (i.e. through narrow connections) (FIG. 2b, c). At 3 dpi typical anthracnose lesions were observed on these leaves, consisting of a necrotic lesion surrounded by a yellow halo. In the necrotic center of the lesion C. graminicola produced stromata (FIG. 2d). Colonization of the mesophyl cells continued to expand at the edges of the lesion, and hyphae often were observed associated with the bundle sheath cells (FIG. 2e).
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). It could be seen frequently inside cells surrounding the vascular bundles (FIG. 2h, i). At the margins of the lesions the fungus continued to expand and colonize the mesophyl cells (FIG. 2f).
Infection of maize roots by C. graminicola.— –
It has been suggested that C. graminicola might enter stalks via roots, and it is known that the pathogen can cause decay of the roots of highly susceptible seedlings (Warren and Nicholson 1975), but the process of root infection has not been investigated closely. A total of 71 inoculation sites on Jubilee seedling roots were sectioned and examined. The following descriptions represent the consensus of all observations. By 24 hpi roots inoculated with C. graminicola became visibly discolored at the point of infection, while roots inoculated with C. sublineolum appeared no different from the water control (data not shown). At 24 hpi conidia of both Colletotrichum species had germinated on the roots and formed melanized Appressoria (FIG. 3a). Appressoria were observed in 82.3% of the samples for C. graminicola and 60% of the samples for C. sublineolum. The appressoria in both cases appeared to be identical to those formed on aerial plant parts. In some cases as early as 24 hpi C. graminicola had invaded the root epidermal cells (in 13.6% of the samples) and cortical cells (in 31.7% of the samples) (FIG. 3b, c, d). It was necessary to end the experiment at 3 dpi due to limits on the ability to keep the seedlings healthy in the germination papers, and in that time the fungus was never seen to move into the stele where it might be possible for it to initiate systemic movement into the above-ground portions of the plant. In contrast to the stalks, but like the leaves, C. sublineolum did not successfully colonize root tissues; development was arrested at the point of appressorium formation, and structures that might have been papillae usually were observed associated with the arrested appressoria (FIG. 3a). In a total of 30 samples production of recognizable primary infection hyphae in root tissues by C. sublineolum was never observed.
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DISCUSSION |
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) and Magnaporthe oryzae in rice leaves (Kankanala et al 2007). For M. oryzae the evidence suggested that the fungus progressed from cell to cell by penetrating the plasmodesmata (Kankanala et al 2007).
Lesion expansion was somewhat slower for C. sublineolum than C. graminicola, suggesting that quantitative resistance might be operating to slow the growth of this organism. However it should be mentioned that C. sublineolum also grows more slowly than C. graminicola in culture (Vaillancourt and Hanau 1992). The fact that not all inoculations of unwounded stalks resulted in infections over the course of the experiment, while all inoculations of wounded stalks did result in infection, shows that, even in this highly susceptible maize genotype, the intact rind does have some capacity to protect the inner stalk tissues.
White and Humy (1976) reported that only about 20% of the Jubilee sweet corn plants they inoculated behind leaf sheaths in the field developed stalk rot after 4 wk and a majority of those were associated with corn borer damage. They did report a high degree of darkening of the stalk and adjacent leaf epidermis, which could correlate with the epidermal infection and sclerotial formation observed in the current study. However they did not describe this darkening further. The rate of infection of unwounded inner tissues at 7–9 dpi was much higher in the current study (60% ) than the 20% observed by White and Humy (1976). This was in spite of the fact that White’s and Humy’s experiment lasted more than twice as long. It is possible that this discrepancy was due to our plants being environmentally stressed, perhaps by the relatively low light intensities in the greenhouse versus in the field, and therefore they were less able to resist pathogen ingress. It also is possible that there are moderating influences of other organisms living behind intact leaf sheaths in the field that were not duplicated in this study where the rinds were exposed and the leaf sheaths removed. Thus, even though the current study demonstrates the potential for direct invasion of stalks by C. graminicola, this does not necessarily mean that it occurs commonly in the field.
It has been suggested that stalks might become colonized via leaves that are infected with anthracnose leaf blight (Bergstrom and Nicholson 1999). Fibers and sheath cells associated with vascular bundles in both seedling and mature leaves were colonized readily by C. graminicola. Given the direct connection between leaf bundles and those in the sheaths, it seems quite possible that the pathogen could be introduced into the sheaths from the leaves, and perhaps from there into the stalk epidermal cells, which are typically in direct contact with sheath tissues.
Entry into the stalk via the roots also has been suggested, and although it is known that C. graminicola can infect roots there appears to have been no further investigation of this question in the literature. The conidia of both Colletotrichum species germinated on roots, formed melanized appressoria that appeared identical to those formed on aerial plant parts. This is different from a report of root infection by Magnaporthe grisea, which also produces melanized appressoria on leaves, but which formed nonmelanized penetration structures on roots (Sesma and Osbourne 2005). C. graminicola was able to invade the root epidermal and cortical cells and grow within them, but within the time constraints of the experiments reported here the fungus never moved into the stele where it might initiate systemic movement, as was described for M. grisea. It is possible that this would have occurred given more time because hyphae were seen close to the stele (FIG. 3d). One batch of 12 inoculated seedlings was planted in soil and grown to postanthesis in the greenhouse, but no sign was observed of stalk-rot development and GFP-expressing fungus could not be recovered from samples of stalk and leaf tissues of the mature plants. However the plants did become severely infested with spider mites and thrips as they senesced, and so that experiment was not repeated. Our results thus are inconclusive on the question of whether seedling roots are a viable entry point for stalk rot.
C. sublineolum is not normally found as a pathogen of maize, even in areas where maize and sorghum commonly are grown together (Dale 1963, Jamil and Nicholson 1987, LeBeau 1950, Williams and Willis 1963). Thus the ability of this fungus to infect maize stalk epidermal cells so efficiently was surprising. These observations suggest that nonhost resistance to C. sublineolum by maize is conditional and perhaps also tissue specific. One possibility is that the lower light conditions under which stalk epidermal cells are produced might decrease their basal levels of resistance. Low light is known to increase susceptibility of leaves to C. graminicola (Hammerschmidt and Nicholson 1977). Because the stalk epidermis develops beneath the sheath it is not exposed to as much light as the leaves and so it may not have the same capacity to mount an effective defense. This will be interesting to investigate further.
C. sublineolum did not germinate on older leaves, even though the same batch of inoculum applied to unwounded stalks germinated and formed appressoria efficiently. Spores of C. graminicola must sense a hydrophobic substrate for germination to be induced (Chaky et al 2001), and so it is possible the older leaves are not sufficiently hydrophobic to induce germination of this species; leaves do become less hydrophobic as they age and as the cuticle is eroded (Lindow and Brandl 2003). In addition it is known that older (nonsenescent) leaves express a greater degree of resistance to C. graminicola than younger leaves (Leonard and Thompson 1976), and so it is possible that the lack of germination was due directly to expression of resistance by the leaves, although it would be surprising to see it expressed even pregermination because resistance to Colletotrichum typically occurs at the stage of initial penetration.
Conditional nonhost resistance is an interesting finding, which is consistent however with previous observations from our lab that suggested that drought- or light-stressed maize stalks were prone to significant rotting by C. sublineolum (Vaillancourt unpublished) and that C. sublineolum can complete its life cycle in wounded stalk tissues (Venard and Vaillancourt 2007). Conditional nonhost resistance might explain the controversy in earlier papers where some sources suggested that maize was susceptible to isolates of Colletotrichum from sorghum, whereas others found strict host specificity (Chowdhury 1936, Dale 1963, Jamil and Nicholson 1987, LeBeau 1950, Williams and Willis 1963, Wheeler et al 1974). Although there is no evidence that cross-infection occurs routinely in the field, this work suggests that the potential exists and this might pose a risk, particularly in areas where both crops are commonly grown and where conditions for maize might be suboptimal so that the plant may be stressed and less able to mount a defense. Evidence has been increasing that host resistance and nonhost resistance are closely related processes (reviewed by Thordal-Christensen 2003). There may be relatively little difference mechanistically between host-specific resistance of maize to C. graminicola and nonhost resistance to C. sublineolum. C. sublineolum clearly has the potential to cause disease in maize, and maize clearly has the potential to be susceptible. Why then is this interaction usually incompatible? Understanding the answer to this question could help us to develop improved methods for control of the anthracnose stalk-rot disease in the field.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Current address: Department of Entomology S–225, Agricultural Science Building North, Lexington, Kentucky 40546. 
1 Corresponding author. E-mail: vaillan{at}uky.edu
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LITERATURE CITED |
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  J. C. Inguagiato, J. A. Murphy, and B. B. Clarke Anthracnose Severity on Annual Bluegrass Influenced by Nitrogen Fertilization, Growth Regulators, and Verticutting Crop Sci., July 1, 2008; 48(4): 1595 - 1607. [Abstract] [Full Text] [PDF]
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