676
Journal of Food Protection, Vol. 74, No. 4, 2011, Pages 676–680
doi:10.4315/0362-028X.JFP-10-347
Copyright G, International Association for Food Protection
Research Note
Infection and Ultrastructure of Conidia and Pycnidia of
Stenocarpella maydis in Maize
ZHENGJUN XIA,1*{ HONGYAN WU,1{
1Key
AND
PREMILA N. ACHAR2*
Laboratory of Mollisols Agroecology, the Northeast Institute of Geography and Agroecology, Chinese Academy of Sciences, Harbin, People’s Republic
of China; and 2Department of Biology and Physics, Kennesaw State University, Kennesaw, Georgia 30144, USA
MS 10-347: Received 23 August 2010/Accepted 8 December 2010
ABSTRACT
Stenocarpella maydis is the most prevalent ear rot pathogen of maize (Zea mays) in South Africa, the United States, and other
countries. Infection and ultrastructure of propagules of S. maydis in maize were observed by light, scanning, and transmission
electron microscopy. Two-celled conidia of S. maydis were found in the tissues of husk and kernels. Mycelia colonized inter- and
intracellularly in the host tissues. Pycnidia were found abundantly inside the seed tissues of susceptible cultivars; within a single
seed, pycnidia propagated preferentially in embryonic tissues. A pycnidium is composed of morphologically different resting spores
mingled with some degraded organelles of the host cell. In this study, various enzymatic activities led to cell wall degradation,
lacunae in endosperm tissues, and disrupted organelles in susceptible cultivars. In contrast, callose deposition surrounding fungal
hyphae was clearly visible in resistant cultivars. Heavy infection was detected by maceration, even though there was no apparent
symptom on the seed coat. The saprophytic nature and structurally different forms of propagules could contribute to a long-term
survival of this pathogen in the field and during grain storage. Furthermore, S. maydis might pose a threat of diplodiatoxin
intoxication to human and domestic animals when infected maize seeds are consumed.
Ear rot comprises major fungal diseases affecting the
successful production of maize, since it directly influences
yield, grade, and price of the final product (1, 12, 20).
Stenocarpella maydis (Berk.) Sutton, formerly Diplodia
maydis, is the most prevalent ear rot pathogen of maize (Zea
mays L.) in South Africa, the United States, and some other
countries (1, 3, 15). Furthermore, the fungus is reported to
produce diplodiatoxin, which causes diplodiosis in cattle
and other animals (25, 33). Another morphologically related
species, Stenocarpella macrospora, with a markedly bigger,
two-celled conidial spore, is more prevalent in humid
subtropical and tropical zones; it produces symptoms of dry
ear rot, stalk rot, and even leaf striping (23, 35).
Koehler (22) described the ramification of S. maydis
infection in maize. The modes of colonization and infection
of S. maydis was also described (3, 4, 8). Bensch and van
Staden (4) observed that appressoria formed at the hyphal
tips 72 h after inoculation, and penetration by a hypha
resulted in the inter- and intracellular colonization in maize.
Specifically, preferential colonization of the embryo in the
kernel by S. maydis was reported (3). Molecular detection
methods of the pathogen in seeds or in the field were
developed (2, 11, 37). Chitinase might be involved in the
interaction between host and pathogen (24). More recently,
* Authors for correspondence. Tel: z86-451-87501708; Fax: z86-45186603736; E-mail: [email protected] (Xia). Tel: 770-499-3524;
Fax: 770-423-6625; E-mail: [email protected] (Achar).
{ These authors contributed equally to this work.
Naumann and Wicklow (32) demonstrated that susceptibility to modification of a fungal-targeted plant chitinase
differs among inbred lines, suggesting that the LH82 line
allele is a specific genetic determinant, contributing to
resistance to the ear rot caused by S. maydis, whereas the
B73 line allele might contribute to susceptibility. Many
efforts via biological control (7), fungicides (10), and plant
resistance (13, 32) have been made in an attempt to control
the infection. The objective of this study was to observe the
in situ ultrastructure of propagules of S. maydis in maize
tissues to characterize the infective mechanisms of this
pathogen.
MATERIALS AND METHODS
Plant materials and inoculation. Two resistant maize
cultivars, RS5206 and PAN6264, and two susceptible ones,
SNK2266 and PAN6140, were used throughout this study. Seeds
were sown in plastic pots filled with compost. Potted seeds were
maintained in an environmentally controlled greenhouse at 30uC,
with 100% humidity. Each pot contained two plants. In addition,
naturally infected maize seeds of cultivar SNK2266 were collected
in fields in KwaZulu-Natal, South Africa.
U2 strains of S. maydis, isolated previously (37), were
incubated separately in potato dextrose broth at 25uC on a shaker.
The conidial suspension was adjusted to <105 spores per ml.
Cultivars were inoculated 15 days after silking. One milliliter of
spore suspension of isolates was injected into the tip of the ears
with a syringe. Each treatment contained six plants; experiments
were performed three times. Two weeks after inoculation, healthy
and diseased tissues were sampled every week.
J. Food Prot., Vol. 74, No. 4
PROPAGULES OF S. MAYDIS IN MAIZE
677
FIGURE 1. Light microscopy micrographs of mycelia and infected embryos.
(A) Infected embryos. (B) Healthy embryos.
(C) Mycelium of Stenocarpella maydis from
infected seed tissues after maceration. The
arrow indicates mycelium with thickened
cell wall.
Scanning electron microscopy. Samples were fixed in 3.0%
glutaraldehyde for 8 h or overnight, washed twice with 0.05 M
sodium cacodylate buffer, and dehydrated with an ethanol serial
dilution (50, 70, 80, and 95% ethanol for 15 min each time, and
then by 100% ethanol [three times] for 10 min each time). Ethanol
absolute (dried by copper sulfate) was used for the last change (19).
Samples were critical point dried with a CPD 750 (Bio-Rad,
Hercules, CA), mounted onto brass stubs, and gold coated with a
Polaron SC500 sputter coater (VG Microtech, Sussex, UK). Hard
kernels were mounted onto brass stubs and gold coated without
any pretreatment. Samples were viewed with a scanning electron
microscope (model 500, Philips, Amsterdam, The Netherlands) at
10 kV (19, 31).
Transmission electron microscopy. Husks and kernels were
cut into sections 2 to 3 by 2 to 3 mm2, and then fixed in 3%
glutaraldehyde overnight. After two washings, samples were
postfixed in aqueous osmium tetroxide, washed in distilled water,
and dehydrated with a graded ethanol series (50% once, 70% once,
95% twice, and 100% twice for 30 min each time). Samples were
pretreated with propylene oxide for 10 min and then subsequently
infiltrated with serial resin (Spurr resin–propylene at 1:1 ratio for
12 h; 3:1 for 12 h; and finally, fresh, pure Spurr resin, overnight).
Samples were embedded in silicone molds with fresh resin for 48 h
at 60 to 70uC (19, 31).
For ultramicrotomy, thin sections (80 nm) were cut with glass
knives. Ultrathin sections with metallic colors (gold to silver) were
picked up on copper grids, stained in uranyl acetate for 20 min, and
washed twice in a distilled water jet stream. Thereafter, grids were
dried and submerged in lead citrate for 10 min. Grids were then
washed in a sodium hydroxide solution, then in distilled water, and
then viewed with a JEM-1010 transmission electron microscope
(Jeol, Tokyo, Japan) at 60 kV (19, 31).
Maceration. All seeds were examined visually for infection
on the seed coat before maceration. Ten seeds from a cultivar were
placed in a 250-ml flask containing 50 ml of maceration solution
(10% [wt/vol] sodium hydroxide and 0.5 g/liter tryptophan blue
[cotton blue]) for 48 h at 25uC. Over 200 seeds were macerated for
each cultivar. After alkaline treatment, the seeds were agitated
gently in a water bath (65uC) for 5 min. Seeds were transferred to a
beaker containing 10 ml of lactophenol. Beakers were placed in a
water bath (65uC) or heated gently over a low flame until embryos
were cleared (27, 34). This enabled complete detachment of the
embryo from the seed coat. Presence and morphology of S. maydis
in seed coats and embryos were examined with light microscopy.
RESULTS AND DISCUSSION
Severely infected maize seeds, albeit no apparent
symptoms on the seed coat. Although molecular detection
of S. maydis has been developed (2, 11, 37), seed maceration (27, 34) still is a quick and reliable approach to
observe the natural appearance of pathogens in seeds. After
maceration, embryos can be detached easily from seed
coats. The infected embryos appeared brown to dark brown
because of the abundant presence of mycelia and pycnidia
(Fig. 1A), in contrast with the clear, transparent, healthy
embryos (Fig. 1B). Some resting structures with thick cell
walls were also observed among normal mycelia (Fig. 1C).
For three batches (more than 200 seeds for each batch) of
naturally infected seeds of susceptible cultivar SNK2266,
the average infection rate by visual examination of
symptoms was 39.6 ¡ 3.3 (standard error), in comparison
with an infection rate of 45.1 ¡ 4.3 (standard error)
revealed by light microscopy after maceration. There was
a significant correlation (r ~ 0.84, P , 0.01) between
infection rates by two parallel examination methods.
However, the discrepancy resulted from severe infections
(detected with light microscopy after maceration for some
seeds), without any apparent symptoms on the seed coat.
Morphological observation of propagules of S.
maydis. In resistant cultivars RS5206 and PAN6264, only
light lesions and infection appeared on the husk and kernel
after inoculation. In comparison, in susceptible cultivars,
symptoms on the surface of the husk appeared on day 2 or 3
after inoculation with conidial suspension of the U2 strain of
S. maydis, indicative of a quick, successful penetration, with
colonization of husks by the fungus. The kernels of
susceptible cultivars were also infected severely by heavily
ramified inter- and intracellular hyphae in the endosperm
tissues (Fig. 2A and 2B). Furthermore, typical two-celled
conidia were found individually in cells of endosperm tissue
(Fig. 2C) 4 to 6 weeks after inoculation. These two-celled
spores resembled morphologically those produced on
synthetic media, as reported by Murphy et al. (30), except
for a large amount of vacuoles. Apart from lytic function,
vacuoles serve in storing ions and amino acids for protein
synthesis (17). In addition, vacuoles might also be very
678
XIA ET AL.
J. Food Prot., Vol. 74, No. 4
FIGURE 3. Electron microscopy micrographs demonstrating the
host–pathogen interaction. (A) Transmission electron microscopy
micrograph showing degradation of the host cell wall (arrowhead)
in susceptible cultivar SNK2266. (B) Scanning electron microscopy micrograph showing disrupted whole endosperm cell in
SNK2266. (C) Transmission electron microscopy micrograph
demonstrating degraded organelles of a host cell (arrows) in
susceptible cultivar SNK2266. H, hyphae. (D) Transmission
electron microscopy micrograph showing callose-like deposition
(arrowhead) in resistant cultivar PAN 6264. H, hyphae.
FIGURE 2. In situ ultrastructure of mycelia and two-celled
conidia. (A) Micrograph showing the penetration and colonization
of kernels by Stenocarpella maydis in susceptible cultivar
SNK2266. H, colonized hyphae. (B) Transmission electron
microscopy micrograph showing infection in endosperm tissues
in kernels of susceptible maize cultivar SNK2266. IH, intercellular
hyphae; IRH, intracellular hyphae. (C) Transmission electron
microscopy micrograph of in situ ultrastructure of two-celled
conidial spores in embryonic tissue of maize. V, vacuole.
important for fungi to synthesize enzymes needed for a
successful colonization, and to moderate the osmotic
pressure from plant if the host plant cells contain large
quantity of vacuoles (9, 17).
Overall, infection progressed in the order of husk,
endosperm, and then kernel, as revealed by both visual
examination of symptoms and microscopic observation of
mycelia under artificial inoculation conditions in this study,
which was inconsistent with previous reports that under
natural conditions, infection occurred in the order of the
base of the ear, the sclerenchyma–placentae–embryos, and
then the whole kernel (6, 8, 13).
Enzymatic activities during host–pathogen interaction of S. maydis. Cell wall degradation surrounding the
penetrating hyphae was clearly visible, implying enzymatic
involvement (Fig. 3A) in embryonic tissues of susceptible
cultivars. Cell wall degradation resulted further in lacunae
on the surface of endosperm cells and disruption of cell
organelles in these tissues (Fig. 3B and 3C). However,
callose-like deposition surrounding fungal hyphae was
observed clearly in embryonic tissues of resistant, but not
susceptible, cultivars (Fig. 3C). Callose was reported to act
as a barrier against fungal colonization (6). Murphy et al.
(29) described callose as a mucus layer that protects hyphae
against dehydration. Plants defend against fungal attack (28)
by physical strengthening of the cell wall via lignification
(36), suberization and callose deposition (6), accumulation
of hydroxyproline-rich glycoprotein (26), synthesis of
J. Food Prot., Vol. 74, No. 4
PROPAGULES OF S. MAYDIS IN MAIZE
679
FIGURE 4. Morphological observation of
pycnidia of Stenocarpella maydis. Pycnidia
in synthetic media (A) and in kernels of
maize (B). (C) Pycnidial spores were
morphologically different in embryos of
susceptible maize cultivar SNK2266.
Arrowheads, degraded organelles of host
cell.
antimicrobial compounds such as phytoalexins, and production of a variety of pathogenesis-related proteins (24).
More recently, it was reported that S. maydis could secrete a
special protein, Stm-cmp, to modify specifically the ChitAF protein alloform, which is encoded by a known allele of
the chiA gene in the strongly susceptible B73 line (32).
Further investigation is needed as to whether the callose
depositions observed in this study are directly associated
with the interaction between Stm-cmp and targeted plant
chitinase (32). More recently, Naumann and Wicklow (32)
demonstrated that the susceptibility to modification of a
fungal-targeted plant chitinase differs among inbred lines,
suggesting that the LH82 and B73 lines’ alleles may be
specific genetic determinants for resistance or susceptibility.
Preferential infection of embryonic tissues and
composition of pycnidia. In resistant cultivars RS5206
and PAN6264, the endosperm was slightly affected by the
pathogen, and no severe damage occurred after artificial
inoculation. However, in the susceptible cultivar, endosperms were severely damaged at later stages (Fig. 3B and
3C). S. maydis favored colonization of the embryonic
tissues by formation of pycnidia once maize kernels were
infected (Fig. 4 A and 4B) (5). In pycnidia, spores appeared
different morphologically, and degraded organelles of host
cells mingled often (Fig. 4C). As colonization progressed,
pycnidia formed in the kernels, especially in the embryos.
The preferential colonization of embryonic tissue by S.
maydis could be ascribed to its richness in nitrogen and
other nutrients, rather than to those nutrients of the
endosperm. Although starch is the primary component of
the endosperm, the embryo also contains significant
amounts. In a previous report, Flett and Wehner (16)
indicated that S. maydis in maize needs additional nutrition
to enhance its sporulation. Guo et al. (18) observed that the
antifungal, ribosome-inactivating protein is located mainly
in the endosperm, whereas zeamatin is located mainly in the
embryo, which uniquely protects kernels from pathogen
attack. The presence of abundant mycelia and pycnidia in
the embryonic tissue could imply that S. maydis might be
insensitive to zeamatin.
The morphologically different propagules and resting
structures of S. maydis in maize along with its saprophytic
nature (14) are likely associated with long-term survival of
the pathogen in the field. This could explain the difficulty in
controlling this disease exclusively by tillage or crop
rotation (12, 15).
Diplodiatoxin might be secreted at the tip of hyphae
during infection and colonization in a way similar to that
of the Fusarium toxin produced in wheat (21). The infected
seeds could pose a long-term threat as food or forage
for humans and domestic animals by diplodiatoxin, a
toxin known for its carcinogenic and neurotoxic effects (25,
33).
680
XIA ET AL.
ACKNOWLEDGMENTS
We thank the Department of Microbiology and Electron Microscopy
Unit, University of Kwazulu-Natal, South Africa, and the Department of
Biology and Physics, Kennesaw State University, GA, for infrastructural
and technical assistance. This work was supported by two Chinese grants,
2009ZX08009-013B and KZCX2-EW-303.
REFERENCES
1. Andrew, R. H. 1954. Breeding for stalk-rot resistance in maize.
Euphytica 3:43–48.
2. Barros, E., M. Crampton, G. Marais, and S. Lezar. 2008. A DNAbased method to quantify Stenocarpella maydis in maize. Maydica
53:125–129.
3. Bensch, M. J. 1995. Stenocarpella maydis (Berk.) Sutton colonization of maize ear. J. Phytopathol. 43:597–599.
4. Bensch, M. J., and J. van Staden. 1992. Ultrastructural histopathology
of infection and colonization of maize by Stenocarpella maydis
(~ Diplodia maydis). J. Phytopathol. 136:312–318.
5. Bensch, M. J., J. van Staden, and F. H. J. Rijkenberg. 1992. Time and
site of inoculation of maize for optimum infection of ears by
Stenocarpella maydis. J. Phytopathol. 136:265–269.
6. Bonhoff, A., B. Rieth, J. Golecki, and H. Grisebach. 1987. Race:
cultivar-specific differences in callose deposition in soybean roots
following infection with Phytophthora megasperma f. sp. glycinea.
Planta 172:101–105.
7. Bressan, W., and J. E. F. Figueiredo. 2005. Biological control of
Stenocarpella maydis in maize seed with antagonistic Streptomyces
sp. isolates. J. Phytopathol. 153:623–626.
8. Chambers, K. R. 1988. Effect of time of inoculation on Diplodia
stalk and ear rot inoculation of maize in South Africa. Plant Dis. 72:
736–739.
9. Dodd, J. L. 1980. The role of plant stress in the development of corn
stalk rots. Plant Dis. 64:533–537.
10. Duarte, R. P., F. C. Juliatti, B. V. Lucas, and P. T. de Freitas. 2009.
Performance of different maize genotypes with leaf application of
fungicides for the incidence of fungi causing sour kernel. Biosci. J.
25:112–122.
11. Fessehaie, A., C. C. Block, L. M. Shepherd, and M. K. Misra. 2007.
Quantitative TaqMan real-time PCR assay for Stenocarpella maydis,
the causal agent of Diplodia ear and stalk rot of maize.
Phytopathology 97:S35.
12. Flett, B. C. 1991. Crop plants as host and non-hosts of Stenocarpella
maydis. Phytophylactica 23:237–238.
13. Flett, B. C., and N. W. McLaren. 1994. Optimum disease potential for
evaluating resistance to Stenocarpella maydis ear rot in corn hybrids.
Plant Dis. 78:587–589.
14. Flett, B. C., and N. W. McLaren. 1998. Incidence of ear rot pathogens
under alternating corn tillage practices. Plant Dis. 82:781–784.
15. Flett, B. C., and F. C. Wehner. 1989. The effect of temperature and
relative humidity on Stenocarpella maydis spore survival. Phytophylactica 21:100. (Abstract.)
16. Flett, B. C., and F. C. Wehner. 1991. Incidence of Stenocarpella and
Fusarium cob rots in maize monoculture under different tillage
systems. J. Phytopathol. 133:327–333.
17. Griffin, D. H. 1994. Fungal physiology, 2nd ed. John Wiley and
Sons, Inc., New York.
18. Guo, B. Z., T. E. Cleveland, R. L. Brown, N. W. Widstrom, R. E.
Lynch, and J. S. Russin. 1999. Distribution of antifungal proteins
in maize kernel tissues using immunochemistry. J. Food Prot. 62:
295–299.
J. Food Prot., Vol. 74, No. 4
19. Hall, J. L., and C. Hawes. 1991. Electron microscopy of plant cells,
p. 1–66. Academic Press, New York.
20. Heald, F. D., E. M. Wilcox, and V. W. Pool. 1909. The life-history
and parasitism of Diplodia zeae (Schw.) Lev., p. 1–7. Nebraska
Agricultural Experimental Station 22nd Annual Report, 1908.
Nebraska Agricultural Experimental Station, Lincoln.
21. Kang, Z., and H. Buchenauer. 2002. Studies on the infection process
of Fusarium culmorum in wheat spikes: degradation of host cell wall
components and localization of trichothecene toxins in infected
tissue. Eur. J. Plant Pathol. 1087:653–660.
22. Koehler, B. 1942. Natural mode of entrance of fungi into corn ears
and some symptoms that indicate infection. J. Ag. Res. 64:291–307.
23. Latterell, F. M., and A. E. Rossi. 1983. Stenocarpella macrospora
(~ Diplodia macrospora) and S. maydis (~ D. maydis) compared as
pathogens of corn. Plant Dis. 67:725–729.
24. Legrant, M., S. Kauffman, P. Geoffroy, and B. Fritig. 1987.
Biological function of pathogenesis-related proteins: four PR proteins
of tobacco are chitinases. Proc. Natl. Acad. Sci. USA 84:6750–6754.
25. Marasas, W. F. O. 1977. Diplodiosis in cattle, p. 163–165. In T. D.
Wyllie and L. G. Morehouse (ed.), Mycotoxic fungi, mycotoxins,
mycotoxicoses, vol. 3. Marcel Dekker, Inc., New York.
26. Mazau, D., and M. T. Esquerretugay. 1986. Hydroxyproline-rich
glycoprotein accumulation in the cell walls of plants infected by
various pathogens. Physiol. Mol. Plant Pathol. 29:147–157.
27. McGee, D. C. 1988. Maize diseases: a reference for seed
technologists. American Phytopathological Society Press, St. Paul,
MN.
28. Mendgen, K., M. Hahn, and H. Deising. 1996. Morphogenesis and
mechanisms of penetration by plant pathogenic fungi. Annu. Rev.
Phytopathol. 34:367–386.
29. Murphy, J. A., L. L. Campbell, and A. J. Paperlis. 1974.
Morphological observations of Diplodia maydis on synthetic and
natural substrates as revealed by scanning electron microscopy.
J. Appl. Microbiol. 61:232–250.
30. Murphy, J. A., M. R. Thompson, and A. J. Pappelis. 1976.
Ultrastructure and elemental composition of dormant and germinating
Diplodia maydis spores. J. Bacteriol. 127:1465–1471.
31. Naidoo, Y., and G. Naidoo. 1998. Sporobolus virginicus leaf salt
glands: morphology and ultrastructure. S. Afr. J. Bot. 64:198–204.
32. Naumann, T. A., and D. T. Wicklow. 2010. Allozyme-specific
modification of a maize seed Chitinase by a protein secreted
by the fungal pathogen Stenocarpella maydis. Phytopathology 100:
645–654.
33. Rabie, C. J., J. J. du Preez, and J. P. Hayes. 1987. Toxicity of
Diplodia maydis to broilers, ducklings and laying chicken hens.
Poult. Sci. 66:1123–1128.
34. Shetty, H. S., A. K. Khanzada, S. B. Mathur, and P. Neergaard. 1978.
Procedures for detecting seed-borne inoculum of Sclerospora
graminicola in pearl millet (Pennisetum typhoides). Seed Sci.
Technol. 6:935–941.
35. Sutton, B. C., and J. M. Waterston. 1966. Diplodia macrospora. In
CMI descriptions of pathogenic fungi and bacteria, no. 83. CAB
International, Wallingford, UK.
36. Vance, C. P., T. K. Kirk, and R. T. Sherwood. 1980. Lignification
as a mechanism of disease resistance. Annu. Rev. Phytopathol. 18:
259–288.
37. Xia, Z., and P. N. Achar. 2001. Random amplified polymorphic DNA
and polymerase chain reaction markers for the differentiation and
detection of Stenocarpella maydis in maize seeds. J. Phytopathol.
149:35–44.
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.