BioMed Research International

BioMed Research International / 2014 / Article

Research Article | Open Access

Volume 2014 |Article ID 378372 |

Bengyella Louis, Sayanika Devi Waikhom, Pranab Roy, Pardeep Kumar Bhardwaj, Chandradev K. Sharma, Mohendro Wakambam Singh, Narayan Chandra Talukdar, "Host-Range Dynamics of Cochliobolus lunatus: From a Biocontrol Agent to a Severe Environmental Threat", BioMed Research International, vol. 2014, Article ID 378372, 9 pages, 2014.

Host-Range Dynamics of Cochliobolus lunatus: From a Biocontrol Agent to a Severe Environmental Threat

Academic Editor: Fengjie Sun
Received22 Feb 2014
Revised28 Apr 2014
Accepted13 May 2014
Published02 Jun 2014


We undertook an investigation to advance understanding of the host-range dynamics and biocontrol implications of Cochliobolus lunatus in the past decade. Potato (Solanum tuberosum L) farms were routinely surveyed for brown-to-black leaf spot disease caused by C. lunatus. A biphasic gene data set was assembled and databases were mined for reported hosts of C. lunatus in the last decade. The placement of five virulent strains of C. lunatus causing foliar necrosis of potato was studied with microscopic and phylogenetic tools. Analysis of morphology showed intraspecific variations in stromatic tissues among the virulent strains causing foliar necrosis of potato. A maximum likelihood inference based on GPDH locus separated C. lunatus strains into subclusters and revealed the emergence of unclustered strains. The evolving nutritional requirement of C. lunatus in the last decade is exhibited by the invasion of vertebrates, invertebrates, dicots, and monocots. Our results contribute towards a better understanding of the host-range dynamics of C. lunatus and provide useful implications on the threat posed to the environment when C. lunatus is used as a mycoherbicide.

1. Introduction

Race specific Cochliobolus species have caused plant disease disaster such as the southern leaf corn blight epidemic of 1970s in the United States of America [1], northern leaf corn blight (Exserohilum turcicum) and corn head smut (Sporisorium reilianum) in northern China in the 1990s [2, 3], and the Great Bengal rice famine of India in 1940s [4, 5]. In the Great Bengal rice famine, more than 2 million people starved to death due to reduction in rice yield of about 40 to 90% [5]. Cochliobolus species often cause diseases to several plant families including Alliaceae, Anacardiaceae, Araceae, Euphorbiaceae, Fabaceae, Malvaceae, Rutaceae, Zingiberaceae and Solanaceae [6].

Cochliobolus lunatus [7] and related species are extensively used as mycoherbicides for controlling weeds in paddies [813]. The host-range of C. lunatus includes plant species, namely, Cynodon sp., Oryza sp., Pennisetum sp., Saccharum sp., Sorghum sp., Triticum sp., and Zea sp. [14]. Geographically, C. lunatus was suggested to be located mainly in Australia, Brazil, Guinea, India, Cameroon, Columbia, Ecuador, Fiji, Gambia, Guadalcanal, Malaysia, Nigeria, Pakistan, Papua New Guinea, Sierra Leone, Sri Lanka, Sudan, Tanzania, Thailand, and USA [14] but not in Europe ( The proposed geographical circumscription and putative hosts of C. lunatus have not been updated.

C. lunatus has emerged in the last decade as a virulent and destructive pathogen [15, 16]. Remarkably, C. lunatus successfully thrives on important crops such as rice (Oryza sativa L.), wheat (Triticum aestivum), cassava (Manihot esculenta), sorghum (Sorghum bicolor), Hymenaches species, strawberry (Fragaria × ananassa), Amaranthus species, and potato [1620]. Decades after Sivanesan’s [14] pioneering study, is C. lunatus solely endemic to the outlined geographical locations? If no, has C. lunatus gained hosts and new geographical zones in the last decade? The aims of this study were (1) to determine the interrelatedness of 5 virulent strains of C. lunatus causing foliar necrosis of potato using morphological descriptors coupled with phylogenetic tools and (2) to establish the current host-range diversity of C. lunatus in the last decade.

2. Materials and Methods

2.1. Study Area and Sampling

Routine survey was performed in potato plantations of Burdwan District (23°14′N, 87°51′E, altitude 150 m, 102.1 km from Kolkata), West Bengal, India, during the winter months of December to March of 2010, 2011, and 2012. Mainly potato cv. Kufri Jyoti is farmed in Burdwan District. The area receives an average annual rainfall of 1173–1442 mm and temperature of 10–20°C during potato farming season. Potato plants showing brown-to-black leaf spot disease previously described [20] were used. Brown-to-black leaf spots were excised and treated with 2% NaClO solution for 2 min and rinsed in sterile water with three changes. The leaf pieces were aseptically plated on V8 agar medium (HiMedia, Mumbai, India) and incubated at 25°C in dark. Developed colonies after 7 days were morphologically identified based on standard monograph taxonomic keys [7].

2.2. Host-Range Diversity

The genomic DNA was isolated from fungal isolates grown in potato dextrose broth (PDB) (HiMedia, Mumbai, India). Approximately 100 mg of mycelia mat was disrupted in the presence of TRI-reagent (Sigma, St. Louis, MO, USA) using mortar and pestle containing 2 mg/mL proteinase K (Merck, Bangalore, India) following the manufacturer instructions. The quality and quantity of the DNA were determined using a 1% agarose gel electrophoresis and a nanodrop spectrophotometer (BioSpec-nano, Shimadzu, Japan), respectively. For molecular identification, the partial sequence of 5.8S rDNA, complete internal transcribed spacer 2 region (ITS2), and partial 28S rDNA region were amplified as previously described [23]. To distinguish the strains, we designed specific primers (forward: 5′-cgatatgcggcatatgca-3′; reverse: 5′-acctacgcattgcggaa-3′) for glyceraldehyde-3-phosphate dehydrogenase (GPDH) gene using C. lunatus (GenBank accession number Gb X58718) sequence. Amplification of GPDH was performed as follows. The PCR mix contained 11 ng genomic DNA, 5 μL Green GoTaq reaction buffer (Promega, Madison, WI, USA), 0.2 mM each of deoxyribonucleoside triphosphate (dNTP), 0.2 μM of each primer, and 1.1 U of GoTaq DNA polymerase in a total reaction volume of 25 μL in triplicates (PCR conditions: 5 min at 95°C, 35 cycles of 1 min at 94°C, 1 min annealing at 53°C, 2 min for extension at 72°C, and a final 5 min extension at 72°C). The quality of the amplicon was checked by performing agarose gel electrophoresis. The PCR products were purified and sequenced. Sequences were assigned to molecular species based on 98–100% sequence similarity threshold in the GenBank with the following accession numbers: JX512810, JX512809, JX907827, JX477595, and JX907828, respectively, for rDNA. GPDH sequences have been submitted in DNA Data Bank of Japan (DDBJ) as accessions AB859034, AB859035, AB859036, AB859037, and AB859038, respectively.

Using GenBank BLAST search tool, a studied set of rDNA sequences deposited in the last decade was collected based on the information associated with the sequences such as GC content, length (>250 bp), and geographic origin of host. Importantly, records with 100% sequence similarity from the same host and geographical coordinates were removed. Unique sequence sets were screened using ElimDupes (available at Sequence alignment was performed using Muscle program [24]. Best substitution model parameters were determined based on Akaike information criterion, corrected (AICc) and Bayesian information criterion (BIC). The evolutionary history was inferred using the maximum likelihood (ML) method, and rooting was performed automatically by saving the generated ML tree in standard Newick format and all the analysis were performed in MEGA 6.06 (updated v. 6140226) software [25]. The strength of the internal branches of the ML tree was statistically tested by performing 1000 bootstrap replications.

3. Results and Discussion

3.1. Identification of C. lunatus Strains Causing Foliar Necrosis of Potato

Basically, most Cochliobolus species have curved conidia, a broad rounded apex cell, a distinct swollen central cell, a tapering to narrowly round base cell, and 4-5 distinct septa. The five strains of Cochliobolus causing brown-to-black leaf spot disease of potato produced varied colonies and conidia (Figure 1) similar to previous studies [6, 7]. The isolates visibly produced different growth patterns (Figure 1). In one isolate Btl26IBSD (DDBJ accession AB859034), brown to whitish mycelium, reddish brown medium, and canoe five-celled conidia without stromatic tissues were observed (Figure 1(a)). In C. lunatus, the stromata are oval or ellipsoidal, 10 to 40 μm in diameter, and located beneath the ascomata. Another isolate Btl27IBSD (DDBJ accession AB859035) produced greyish-brown mycelium and cylindrical clavated fived-celled conidia void of stromatic tissue (Figure 1(b)). Isolate Btl28IBSD (DDBJ accession AB859036) profusely produced yellowish pigmented five-celled conidia, with stromatic tissue, variable shapes, and end at one cell with a thin hilium (Figure 1(c)). Isolates Btl29IBSD (DDBJ accession AB859037) and Btl30IBSD (DDBJ accession AB859038) produced greyish-brown cottony mycelium (Figures 1(d) and 1(e)). Noteworthy, isolate Btl30IBSD profusely produced dark pigments, and with each cell of the conidia bearing a distinctive oval stromata of different sizes. The exact role of stromata in pathogenicity is not known. The stromata are enclosed by a ring of melanin-like pigment, may play a role in preventing desiccation of the conidia, conserved the gene-pool, and ensure survival under adverse conditions. As shown (Figure 1), morphological characters revealed significant intraspecific variations.

Taxonomic circumscription of Cochliobolus has undergone countless modifications in the last five decades caused by overlapping morphological characters [6, 7, 14, 15]. Furthermore, generic concepts delimiting Bipolaris, Cochliobolus, and Pseudocochliobolus are confused [6, 7]. Thus, ITS region of the ribosomal RNA operon was used to accurately determine the taxonomic placement of the fungi. Based on rDNA locus, we confirmed that the five fungi causing brown-to-black leaf spot disease of potato (Figures 2(a) and 2(b)) were C. lunatus. In the ML tree, the five strains of C. lunatus causing brown-to-black leaf spot disease of potato clustered (Figure 2(c), (I)), closely related to other GenBank type isolates (Figure 2(c), (II)) and distant from other Cochliobolus species. The nucleotide frequencies were A = 25.00%, T/U = 25.00%, C = 25.00%, and G = 25.00%. The transition-transversion bias estimated by K2 + I substitution model [21] was 2.41. The overall rate of heterogeneity between taxa was 0.01. As expected, low level single nucleotide polymorphism (SNP, 5.4%) was observed out of a total of 1188 sites at the rDNA locus. The five strains causing foliar necrosis of potato were weakly supported with bootstrap values ≤61%. As previously reported, rDNA locus do not often provide ample resolution that can allow differentiation of cryptic taxa such as Cochliobolus [6, 15].

The low bootstrap support (≤61%) generated in rDNA ML tree (Figure 2, (I)) made it difficult to determine whether the five strains of C. lunatus causing brown-to-black spot disease of potato in Burdwan Destrict were identical. It could be that all the strains originated from a common source but colonized in different places following dispersion. This is because C. lunatus abundantly produced conidia that can easily be disseminated by air to distant places. To check if the five isolates were identical or not, we used glyceraldehyde 3-phosphate dehydrogenase (GPDH) locus which had been shown to be effective in resolving Cochliobolus species in phylogenetic inference [6, 15]. Partial GPDH locus (Figure 3(a)) was sequenced, as this is one of the house-keeping genes, taken as reference in yeast and fungal systems. Based on sequence alignment for GPDH locus, a total of 340 SNPs out of 708 sites and 325 sites without polymorphism (45.9%) were found. Based on TN93 + G + I substitution model [22], the rate of base transition–transversion was 4.96 and the nucleotide frequencies were A = 23.73%, T/U = 18.55%, C = 33.52%, and G = 24.20% and the overall heterogeneity among taxa was 0.316. The ML tree based on GPDH locus discriminated the five strains of C. lunatus causing foliar necrosis of potato with strong bootstrap support ≥81% (Figure 3(b), (IV)). The overall mean evolutionary distance of 0.03 was observed between the five strains causing foliar necrosis of potato (Figure 3(b), (IV)) relative to other C. lunatus type isolates (Figure 3(b), (I), (II), and (III)).

Importantly, because the five strains of C. lunatus clustered based on GPDH locus (Figures 2(b) and 3(b)), this indicated they were closely related as also revealed on the basis of morphological descriptors (Figure 1). Additionally, bootstrap values were <100% for internal branches within the subcluster I, Figure 3(b). This indicated that the five strains which caused foliar necrosis of potato were different. Although the five C. lunatus strains might have adapted in potato for their nutritional requirements in the same geoclimatic zone, it was not possible to determine their origin. Importantly, it is shown that pathogenic fungi are capable of adapting to the genetic background of their host, thus forming new physiological and virulent races [26]. This is generally a slow progressive process determined mainly by the degree of the pathogen-host specific interactions [27]. Collectively, because of some phenotypical variations such as colonies growth pattern, presence or absence of stromatic tissues, colours of conidia and colonies (Figure 1), and strong bootstrap support (>81%) for clustered and unclustered strains (Figure 3(b), (I), (II), and (III)), C. lunatus strains have evolved divergently.

3.2. Host-Range Diversity

Herein, the term host-range diversity described the group of different hosts on which C. lunatus successfully thrived on such as monocots, dicots, invertebrates, and vertebrates. The known hosts of C. lunatus presented by Sivanesan [14] in 1987 are plant species, namely, Cynodon sp., Oryza sp., Pennisetum sp., Saccharum sp., Sorghum sp., Triticum sp., and Zea sp. There was no up-to-date account on the new host gained by C. lunatus since Sivanesan [14] account. By exploring the public repositories, we found that C. lunatus have gained hosts within host groups such as monocots, dicots, vertebrates, and invertebrates in the last decade (Table 1). New hosts gained in the last decade are Homo sapiens, Musa acuminata, Jatropha curcas, Echinochloa sp., Arecales sp., Cyperaceae sp., Panicum sp., Setaria italic, Solanum tuberosum L., Glycine max L., Nelumbo nucifera, Eugenia jambolana, Actinidia deliciosa, Actinidia sp., Trachymyrmex septentrionalis, and Cyphomyrmex wheeleri (Table 1), geographically distributed across Africa, Asia, North America, South America, and Europe. It is worth noting that Europe was not included in Sivanesan [14] report by 1987 ( Other C. lunatus new hosts reported [19, 2840] in the last decade without nucleotide sequence information are depicted (Table 2).

AccessionsHost Host groupGeographic originDate of report

EU828350Allelopathic rice (leaf)Monocots
China (Fujian)16-Jul-2008
GQ179977Musa acuminata China27-Jun-2009
GQ328852Zea sp. (seed)USA (Peoria)25-Jun-2009
JF798505Jatropha curcas Mexico16-Feb-2012
JX256435Oryza sp. (leaf)Thailand10-Sep-2012
HQ248192Arecales (Oil palm, leaf)Colombia31-Oct-2010
AF163082Oryza sp.China (Hong Kong)27-Jul-2000
GQ328851Zea sp. (seed)US (Peoria)05-Aug-2009
FJ040177Oryza sp. (grains)China (Zhejiang)20-Sep-2008
EF189917Echinochloa sp. (leaf)China (Zhejiang)22-Jan-2007
JN207244Cyperaceae sp. (Sedges, leaf)Venezuela (Northwest)22-Jun-2012
JX256436Echinochloa sp. (leaf)Thailand10-Sep-2012
JX256432Panicum sp.Thailand10-Sep-2012
JX256444Panicum sp.Solomon Island10-Sep-2012
HQ130484Panicum virgatum (switchgrass)USA (Tennessee)29-Aug-2012
JN943425Echinochloa sp.Japan (Kochi)21-Dec-2011
JN943426Setaria italica (leaf)Japan (Kagoshima)21-Dec-2011
JN943424Setaria italica (leaf)Japan21-Dec-2011

JX512810*S. tuberosum L. (leaf)Dicots
India (Burdwan)20-Aug-2012
JX512809*S. tuberosum L. (leaf)India (Burdwan)20-Aug-2012
JX907827*S. tuberosum L. (leaf)India (Burdwan)09-Sep-2012
JQ936200Glycine max L. (leaf)Brazil16-Apr-2012
JX477595*S. tuberosum L. (leaf)India (Burdwan)12-Aug-2012
JX907828*S. tuberosum L. (leaf)India (Burdwan)09-Sep-2012
JQ701798Nelumbo nucifera (leaf)China (Jiangxi)01-Jul-2012
JQ765410Ipomoea carnea (leaf)India03-Jul-2012
KC937052S. tuberosum L. (leaf)India12-Aug-2013
KF031026Eugenia jambolana India11-May-13
JX256445Actinidia deliciosa Solomon Island10-Sep-2012

JF819163Actiniaria sp.Invertebrates
China (Yushan)19-Apr-2011
JQ717321CoralesChina (Guangdong)13-Aug-012
HQ608077Trachymyrmex septentrionalis USA (Texas)15-Nov-2011
HQ608020Cyphomyrmex wheeleri Brazil15-Nov-2011
JQ388928Marine sponge Panama01-Jun-2012
HQ607975Cyphomyrmex wheeleri USA (Texas)15-Nov-2011

JX256429Human lungs biopsyVertebrateUSA10-Sep-2012
HE861835Human nasal nostrilsSpain23-Jul-2013
KC288118Human subcutaneous tissueBrazil21-Nov-2012

HQ174562UnknownChina (Shandong)22-Feb-2011
JN943422UnknownUnited kingdom17-Apr-2012
FJ792584Medicinal plantsChina (Jiangsu)30-Mar-2009

JX077054Wetland sedimentSoilChina (Zhejiang)17-Jul-2012

Accessions corresponding to isolates causing brown-to-black leaf spot disease of potato reported in this study.

Host originGeographic originYear of reportReference

Dioscorea sp.Nigeria2005Amusa et al. [28]
Chrysalidocarpus lutescens New Zealand2006Braithwaite et al. [29]
Saccharum officinarum Japan2008Nishi et al. [30]
Passiflora edulis f. flavicarpa Deg.Philippines2009Marvin and Naomi [31]
Pennisetum typhoides PakistanunknownAzhar et al. [32]
Fragaria × ananassa Dutch (Strawberry)India2010Verma et al. [19]
Grewia optiva India2011Cvetomir [33]
Basella rubra India2011Pandey et al. [34]
Mimusops elengi LinnIndia2011Selima et al. [35]
Amaranthus spinosus India2011 Sharma et al. [36]
Vicia faba Egypt2012Saleem et al. [37]
Allium sativum L.India2013Ghangaonkar [38]
Lake water (Fishes)India2013Pratibha et al. [39]
Clerodendrum indicum India2013 Mukherjee et al. [40]

This study seeks to advance insights on the host-range diversity allowing the dynamic movement of C. lunatus observed in the last decade (Tables 1 and 2). From Table 1, it is understood that C. lunatus exploit two kingdoms, notably plant and animal, switching among monocots, dicots, invertebrates, and vertebrates. The paradigm-shift from a plant colonizer sensu stricto to a vertebrate and invertebrate invader (Tables 1 and 2) indicates that C. lunatus have acquired special strategies to switch hosts. The question arises as to why C. lunatus display extensive host-range diversity in a given biota.

Although studies have shed light on specific aspects of C. lunatus pathogenicity such as induce-virulence variations on maize crop [41], virulence differentiation on maize crop [42], secretome weaponries on potato crop [43], and heat-dependent virulence on Lolium spp. [44], the nutritional evolution of C. lunatus is unresolved. Intriguingly, host shifting dynamics is not well understood and it has been argued that (1) close proximity to host is prerequisite for pathogens to jump from a natural host to a new host [45], (2) the future host must act as the substrate [15, 45], and (3) compatible factors promoting infection must be present [45, 46]. Importantly, most host-switching pathogens self-protect themselves by producing high level pigment such as melanin to deal with the host defense [42, 47, 48]. Additionally, C. lunatus profusely produced melanized colonizing hyphae during invasion in potato [43] and non-host-specific toxin such as methyl-5-[hydroxymethylfuran-2-carboxylate] in maize [49], to suppress the host defense.

Nonetheless, the above-mentioned factors seem more likely to be limited in explaining how C. lunatus gain access to hosts and not how C. lunatus spreads in a given biota and prevails as an environmental hazard. C. lunatus had extensively been used as mycoherbicide formulations in the past decade [813]. Remarkably, Zhang et al. [12] fused the protoplast of Helminthosporium gramineum and C. lunatus, to generate a strain with high potential to produce conidia, phytotoxin ophiobolin, and improved potential to control rice weed. Introduction of genetically manipulated strains and unmodified strains of C. lunatus could have hazardous implications to the environment. This is because, in some cases, C. lunatus failed to control the targeted weeds but caused severe damages in economically important crops in the same biota. For instance, C. lunatus isolated from barnyardgrass and used as mycoherbicide failed to control competitive weeds in rice fields but severely damaged bean varieties [8]. Nevertheless, the effectiveness of a biocontrol in the fields depends on the environmental conditions of a given biota, especially humidity and temperature [10]. C. lunatus exhibits a temperature-dependent virulence [43, 44] and its introduction into the environment without a precise prediction of the geoclimatic conditions, that is, humidity and temperature, can prove harmful; consequently, it disequilibrates the interaction dynamics of the organisms dwelling in the same biota.

Owing to the divergence in evolution (Figure 1, (I) and (II)) and the emergence of unclustered strains (Figure 2, (I) and (II)), it is clear that strains of C. lunatus have coevolved with their different hosts translated by their different placement in ML tree inference and speciation in their nutritional requirements (Tables 1 and 2). In keeping with the results of the evolutionary disparity, the global control of C. lunatus diseases would require tremendous exertion. This is because in an intermixed network of host-groups, C. lunatus strains from different hosts, genetically distant ((IV) versus (I), (II), and (III), Figure 2(b)) and found in the same geographic zone, would readily invade putative hosts regardless of their temporal host-groups. For instance in the last three years in India, C. lunatus have invaded strawberry [19], Mimusops elengi [37], Amaranthus spinosus [36], Grewia optiva [33], Clerodendrum indicum [40], and potato [20, 48]. Noteworthy, these hosts were spatially and temporally distant. With this illustration, it is clear that the host-pathogen proximity hypothesis and host relatedness hypothesis, where a given pathogen switches to new species closely related to the original host, might all apply for C. lunatus.

4. Conclusions

From an evolutionary viewpoint, the variations observed in C. lunatus colony, conidia size, conidia colour, conidia texture, and the presence or absence of stromata should be regarded as prominent acquired adaptational traits. These characteristic traits were not consistent between the five strains causing foliar necrosis of potato but provided indicators for generic circumscription. Phenotypic intraspecific variations can obscure placement of Cochliobolus species and make correlation to phylogeny difficult. As shown, C. lunatus have considerable ecological and economic importance being a highly successful colonizer in monocots, dicots, vertebrates, and invertebrates. On this basis, the purpose as a biocontrol agent is overshadowed by its virulent and indiscriminate destructive potential in the ecosystem. For this reason, we suggest that the use of C. lunatus as mycoherbicide should be stopped.

Conflict of Interests

The authors declare there is no conflict of interests.


This research was jointly supported by the Academy of Sciences for Developing World (TWAS), Trieste, Italy, and the Department of Biotechnology, Government of India (Program no. 3240223450).


  1. A. J. Ullstrup, “The impact of the southern corn leaf blight epidemics of 1970-71,” Annual of Review of Phytopathology, vol. 10, pp. 37–50, 1972. View at: Google Scholar
  2. J. Chen, H. H. Yan, Z. G. Gao, C. S. Xue, and J. H. Zhuang, “Identification techniques for physiological differentiation of Curvularia lunata in maize,” ACTA Phytopathologica Sinica, vol. 33, no. 2, pp. 121–125, 2003. View at: Google Scholar
  3. H. H. Yan, J. Chen, Z. G. Gao, S. C. Xia, and R. Q. Zhang, “The heredity and variation of the interaction between Curvularia lunata and plant,” Journal of Maize Science, vol. 13, no. 2, pp. 119–120, 2005. View at: Google Scholar
  4. S. Y. Padmanabhan, “The great Bengal famine,” Annual Review of Phytopathology, vol. 11, pp. 11–26, 1973. View at: Google Scholar
  5. R. P. Scheffer, The Nature of Disease in Plants, Cambridge University Press, 1997.
  6. D. S. Manamgoda, L. Cai, A. H. Bahkali, E. Chukeatirote, and K. D. Hyde, “Cochliobolus: an overview and current status of species,” Fungal Diversity, vol. 51, pp. 3–42, 2011. View at: Publisher Site | Google Scholar
  7. R. R. Nelson and F. A. Haasis, “Cochliobolus lunatus,” Mycologia, vol. 56, no. 2, p. 316, 1964. View at: Google Scholar
  8. P. S. Bisen, “Production of toxin metabolites by Curvularia lunata and its role in leaf spot disease of bean,” Acta Botanica Indica, vol. 11, no. 3, pp. 235–238, 1983. View at: Google Scholar
  9. H. Tsukamoto, M. Tsuda, M. Gohbara, and T. Fujimori, “Effect of water management on mycoherbicidal activity of Exserohilum monoceras against Echinochloa oryzicola,” Annals of the Phytopathological Society of Japan, vol. 64, no. 6, pp. 526–531, 1998. View at: Google Scholar
  10. S. W. Huang, L. Q. Yu, G. F. Duan, and K. Luo, “Study on barnyard grass (Echinochloa crus-galli) control by Helminthosporium gramineum and Exserohilum monoceras,” Acta Agronomica Academiae Scientiarum Hungaricae, vol. 35, no. 1, pp. 66–72, 2005. View at: Google Scholar
  11. M. R. S. Motlagh, “Evaluation of Curvularia lunata as an biological control agent in major weeds of rice paddies,” Life Science Journal, vol. 8, no. 2, pp. 81–91, 2011. View at: Google Scholar
  12. Z. B. Zhang, N. R. Burgos, J. P. Zhang, and L. Q. Yu, “Biological control agent for rice weeds from protoplast fusion between Curvularia lunata and Helminthosporium gramineum,” Weed Science, vol. 55, no. 6, pp. 599–605, 2007. View at: Publisher Site | Google Scholar
  13. G. Jyothi, K. R. N. Reddy, R. N. Reddy, and A. R. Podile, “Exploration of suitable solid media for mass multiplication of Cochliobolus lunatus and Alternaria alternata used as mycoherbicide for weed management (barnyard grass) in rice,” Journal of Experimental Biolology and Agricultural Sciences, vol. 1, no. 4, pp. 281–284, 2013. View at: Google Scholar
  14. A. Sivanesan, Graminicolous Species of Bipolaris, Curvularia, Drechslera, Exerohilum and Their Telemorphs, CAB International, 1987.
  15. M. L. Berbee, M. Pirseyedi, and S. Hubbard, “Cochliobolus phylogenetics and the origin of known, highly virulent pathogens, inferred from ITS and glyceraldehyde-3-phosphate dehydrogenase gene sequences,” Mycologia, vol. 91, no. 6, pp. 964–977, 1999. View at: Google Scholar
  16. I. Ahmad, S. Iram, and J. Cullum, “Genetic variability and aggressiveness in Curvularia lunata associated with rice-wheat cropping areas of Pakistan,” Pakistan Journal of Botany, vol. 38, no. 2, pp. 475–485, 2006. View at: Google Scholar
  17. W. Msikita, J. S. Yaninek, M. Ahounou, H. Baimey, and R. Fagbemissi, “First report of Curvularia lunata associated with stem disease of cassava,” Plant Disease, vol. 81, no. 1, p. 112, 1997. View at: Publisher Site | Google Scholar
  18. E. E. John and K. P. Louis, “Seed mycoflora for grain mold from natural infection in sorghum germplasm grown at Isabela, Puerto Rico and their association with kernel weight and germination,” Plant Pathology Journal, vol. 5, no. 1, pp. 106–112, 2006. View at: Google Scholar
  19. V. S. Verma, R. D. Brahmana, and V. S. Gupta, “First Report of Curvularia lunata causing root rot of strawberry in India,” Plant Disease, vol. 94, no. 4, pp. 477–477, 2010. View at: Google Scholar
  20. B. Louis, P. Roy, S. D. Waikhom, and N. C. Talukdar, “Report of foliar necrosis of potato caused by Cochliobolus lunatus in India,” Africa Journal Biotechnology, vol. 12, no. 8, pp. 833–835, 2013. View at: Google Scholar
  21. M. Kimura, “A simple method for estimating evolutionary rates of base substitutions through comparative studies of nucleotide sequences,” Journal of Molecular Evolution, vol. 16, no. 2, pp. 111–120, 1980. View at: Google Scholar
  22. K. Tamura and M. Nei, “Estimation of the number of nucleotide substitutions in the control region of mitochondrial DNA in humans and chimpanzees,” Molecular Biology and Evolution, vol. 10, no. 3, pp. 512–526, 1993. View at: Google Scholar
  23. T. J. White, T. D. Bruns, S. B. Lee, and J. W. Taylor, “Amplification and sequencing of fungal ribosomal RNA genes for phylogenetics,” in PCR-Protocols and Applications. A Laboratory Manual, N. Innis, D. Gelfand, J. Sninsky, and T. C. White, Eds., pp. 315–322, Academic Press, New York, NY, USA, 1990. View at: Google Scholar
  24. R. C. Edgar, “MUSCLE: multiple sequence alignment with high accuracy and high throughput,” Nucleic Acids Research, vol. 32, no. 5, pp. 1792–1797, 2004. View at: Publisher Site | Google Scholar
  25. K. Tamura, G. Stecher, D. Peterson, A. Filipski, and S. Kumar, “MEGA6: molecular evolutionary genetics analysis version 6. 0,” Molecular Biology and Evolution, vol. 30, no. 1, pp. 2725–2729, 2014. View at: Google Scholar
  26. L. G. Barrett, P. H. Thrall, P. N. Dodds et al., “Diversity and evolution of effector loci in natural populations of the plant pathogen melampsora lini,” Molecular Biology and Evolution, vol. 26, no. 11, pp. 2499–2513, 2009. View at: Publisher Site | Google Scholar
  27. J. W. Kirchner and B. A. Roy, “Evolutionary implications of host-pathogen specificity: fitness consequences of pathogen virulence traits,” Evolutionary Ecology Research, vol. 4, no. 1, pp. 27–48, 2002. View at: Google Scholar
  28. N. A. Amusa, A. A. Adegbite, and M. O. Oladapo, “Investigations into the role of weeds, soil and plant debris in the epidemiology of foliar fungal diseases of yam in western Nigeria,” International Journal of Botany, vol. 1, no. 2, pp. 111–115, 2005. View at: Google Scholar
  29. M. Braithwaite, C. F. Hill, S. Ganev, J. M. Pay, H. G. Pearson, and B. J. R. Alexander, “A survey of sub-tropical nursery plants for fungal diseases in northland,” New Zealand Plant Protection, vol. 59, no. 1, pp. 132–136, 2006. View at: Google Scholar
  30. N. Nishi, T. Muta, M. Nakamura, M. Takemure, and T. Tsukiboshi, “Leaf spot of Saccharum officinarum caused by Curvularia lunata,” Japanese Journal of Phytopathology, vol. 74, no. 1, pp. 118–120, 2008. View at: Google Scholar
  31. A. R. Marvin and G. T. Naomi, “Diseases of passion fruit (Passiflora edulis f. flavicarpa Deg.) in three municipalities of North and South Cotabato, Philippines,” USM R&D Journal, vol. 17, no. 2, pp. 123–129, 2009. View at: Google Scholar
  32. H. Azhar, A. A. Safdar, G. M. Sahi, Q. Abbas, and Imran, “Seed borne fungal pathogens associated with pearl millet (Pennisetum typhoides) and their impact on seed germination,” Pakistan Journal Phytopathology, vol. 21, no. 1, pp. 55–60, 2009. View at: Google Scholar
  33. M. D. Cvetomir, “New records of fungi-like organisms and slime moulds from Europe and Asia,” Mycologia Balcanica, vol. 8, no. 1, pp. 173–175, 2011. View at: Google Scholar
  34. R. K. Pandey, P. K. Gupta, M. Srivastava, S. R. Singh, and G. Robin, “First report of brown leaf spot disease caused by Curvularia lunata infecting Indian spinach or poi (Basella rubra),” Indian Phytopathology, vol. 64, no. 2, p. 207, 2011. View at: Google Scholar
  35. K. Selima, C. Urgar, C. Manoranjan, S. Ojha, and N. C. Chatterjee, “Biochemical defense against die-back disease of a traditional medicinal plant Mimusops elengi linn,” European Journal of Medicinal Plants, vol. 1, no. 4, pp. 40–49, 2011. View at: Google Scholar
  36. P. Sharma, N. Singh, and O. P. Verma, “First report of curvularia lunata associated with leaf spot of Amaranthus spinosus,” Asian Journal of Plant Pathology, vol. 5, no. 2, pp. 100–101, 2011. View at: Publisher Site | Google Scholar
  37. A. Saleem, A. H. M. El-Said, T. A. Maghraby, and A. Hamid, “Pathogenicity and pectinase activity of some facultative mycoparasites isolated from Vicia faba diseased leaves in relation to photosynthetic pigments of plant,” Journal Plant Pathology and Microbiology, vol. 3, no. 6, p. 141, 2012. View at: Google Scholar
  38. N. M. Ghangaonkar, “Incidence of mycoflora on garlic (Allium sativum L.) bulbs,” International Research Journal of Biological Sciences, vol. 2, no. 7, pp. 64–66, 2013. View at: Google Scholar
  39. V. Pratibha, S. Shailu, and S. Ranjana, “Seven species of Curvularia isolated from three lakes of Bhopal,” Advances in Life Science and Technology, vol. 8, pp. 13–15, 2013. View at: Google Scholar
  40. A. Mukherjee, A. Bandhyopadhyay, and S. Dutta, “New report of leaf spot disease of clerodendrum indicum caused by Curvularia lunata,” International Journal of Pharma and Bio Sciences, vol. 4, no. 3, pp. B808–B812, 2013. View at: Google Scholar
  41. S. Gao, T. Liu, Y. Li, Q. Wu, K. Fu, and J. Chen, “Understanding resistant germplasm-induced virulence variation through analysis of proteomics and suppression subtractive hybridization in a maize pathogen Curvularia lunata,” Proteomics, vol. 12, no. 23-24, pp. 3524–3535, 2012. View at: Publisher Site | Google Scholar
  42. S. Xu, J. Chen, L. Liu, X. Wang, X. Huang, and Y. Zhai, “Proteomics associated with virulence differentiation of Curvularia lunata in maize in China,” Journal of Integrative Plant Biology, vol. 49, no. 4, pp. 487–496, 2007. View at: Publisher Site | Google Scholar
  43. B. Louis, S. D. Waikhom, P. Roy et al., “Secretome weaponries of Cochliobolus lunatus interacting with potato leaf at different temperature regimes reveal a CL[xxxx]LHM—motif,” BMC Genomics, vol. 15, article 213, 2014. View at: Google Scholar
  44. J. J. Muchovej and H. B. Couch, “Colonization of bent-grass turf by Curvularia lunata after leaf clipping and heat stress,” Plant Disease, vol. 71, pp. 873–875, 1987. View at: Google Scholar
  45. P. Van Baarlen, A. Van Belkum, R. C. Summerbell, P. W. Crous, and B. P. H. J. Thomma, “Molecular mechanisms of pathogenicity: how do pathogenic microorganisms develop cross-kingdom host jumps?” FEMS Microbiology Reviews, vol. 31, no. 3, pp. 239–277, 2007. View at: Publisher Site | Google Scholar
  46. R. M. Kepler, G.-H. Sung, Y. Harada et al., “Host jumping onto close relatives and across kingdoms by Tyrannicordyceps (Clavicipitaceae) gen. nov. and Ustilaginoidea (Clavicipitaceae),” American Journal of Botany, vol. 99, no. 3, pp. 552–561, 2012. View at: Publisher Site | Google Scholar
  47. J. D. Nosanchuk and A. Casadevall, “Impact of melanin on microbial virulence and clinical resistance to antimicrobial compounds,” Antimicrobial Agents and Chemotherapy, vol. 50, no. 11, pp. 3519–3528, 2006. View at: Publisher Site | Google Scholar
  48. B. Louis, S. D. Waikhom, M. S. Wakambam, N. C. Talukdar, and R. Pranab, “Diversity of ascomycetes at the potato interface: new devastating fungal pathogens posing threat to potato farming,” Plant Pathology Journal, vol. 13, no. 1, pp. 18–27, 2014. View at: Google Scholar
  49. J. Goa, T. Liu, and J. Chen, “Insertional mutagenesis and cloning of the gene required for the biosynthesis of the non-host specific toxin in Cochliobolus lunatus that causes maize leaf spot,” Phytopathology, vol. 104, no. 4, pp. 332–339, 2014. View at: Google Scholar

Copyright © 2014 Bengyella Louis et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

More related articles

 PDF Download Citation Citation
 Download other formatsMore
 Order printed copiesOrder

Related articles

Article of the Year Award: Outstanding research contributions of 2020, as selected by our Chief Editors. Read the winning articles.