International Journal of Zoology

Volume 2012 (2012), Article ID 303589, 12 pages

doi:10.1155/2012/303589

Review Article

Pathogens Associated with Sugarcane Borers, Diatraea spp. (Lepidoptera: Crambidae): A Review

Víctor M. Hernández-Velázquez,¹ Laura P. Lina-García,¹ Verónica Obregón-Barboza,¹ Adriana G. Trejo-Loyo,² and Guadalupe Peña-Chora²

¹Centro de Investigación en Biotecnología, Universidad Autónoma del Estado de Morelos, Avenida Universidad No. 1001, Colonia Chamilpa, 62210 Cuernavaca, MOR, Mexico

²Centro de Investigaciones Biológicas, Universidad Autónoma del Estado de Morelos, Avenida Universidad No. 1001, Colonia Chamilpa, 62210 Cuernavaca, MOR, Mexico

Received 14 June 2012; Accepted 29 August 2012

Academic Editor: Thomas Iliffe

Copyright © 2012 Víctor M. Hernández-Velázquez 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.

Abstract.

The objective of this paper was to analyze information related to entomopathogenic-associated Diatraea spp. Gaining a better understanding of the effects of these microorganisms will help in the development of successful microbial control strategies against stem borers that attack sugarcane plants.

1. Introduction

The Diatraea spp. (Lepidoptera: Crambidae) complex is only found in the American continent, and it is the most important group of stem borers that principally attack maize and sugarcane, as well as other gramineous crops, including rice, sorghum, and forage grasses [1]. The sugarcane borer (SCB) D. saccharalis Fab. is the most economically important pest in South America [2, 3], whereas the neotropical corn stalk borer (NCB) D. lineolata Walk is primarily found in Central America [4], D. magnifactella Dyar and D. considerata Heinrich are found in Mexico [1], and the southwestern corn borer D. grandiosella Dyar is found in the United States [5].

Unfortunately, commercially available insecticides are not efficient for the control of Diatraea spp. for a variety of reasons, mainly because of the continuous presence of the host plants in fields throughout the year, the concomitant presence of mature and immature forms of the insect, and the cryptic feeding habits of the insect [6]. An alternative strategy is integrated pest management with biological control as the first defense, which includes the use of parasitoids and entomopathogens.

In this paper, we present a perspective on the attempts to control Diatraea spp. using pathogens in the Americas. We also discuss the status of recent attempts to use pathogens in the field.

2. Fungi

One promising field for research is the use of entomopathogenic fungi as biological control agents of insect pests in sugarcane plants. Approximately 80% of the etiological agents involved in insect diseases are fungi, which encompass 90 genera and more than 700 species [7]. A number of fungi (Hypocreales: Clavicipitaceae) including B. bassiana, B. brongniartii, M. anisopliae, P. fumosoroseus, Hirsutella sp., Cylindrocarpon sp., and Nomuraea rileyi have been isolated from Diatraea spp. in the Americas from Argentina to the USA (Table 1). Under certain climatic conditions, B. bassiana has been reported to cause natural epizootics on D. grandiosella [8].

Table 1: Entomopathogens fungi from Diatraea spp.

Host species
Entomopathogen
Country
Reference

D. grandiosella 

D. crambidoides
B. bassiana (1–8%)
Mississippi, United States
Inglis et al., 2000 [5]

D. grandiosella
B. bassiana (epizootics)
Texas High Plains, United States
Knutson and Gilstrap, 1990 [8]

B. bassiana

D. saccharalis
M. anisopliae
Venezuela
Zambrano et al., 2002 [9]

P. fumosoroseus

D. saccharalis
M. anisopliae
PE, Brazil
Alves, et al. 2002 [10]

D. saccharalis
B. bassiana
Pinar del Río, La Habana, Matanzas, Villa Clara, Cienfuegos and Camagüey, Cuba
Estrada et al., 2004 [11]

D. saccharalis
B. bassiana 

M. anisopliae 

Nomuraea rileyi 

Iseria sp.
Tucumán, Argentina
Yasen de Romero et al., 2008 [12]

D. saccharalis
M. anisopliae 

(Ma2 and Ma3)
Mexico
Angel-Sahagún et al., 2005 [13]

B. bassiana

ARSEF: 1489
Pernambuco

1832
Brazil

1834
Brazil

2629
Brazil

3020
Pernambuco

3858
Brazil

D. saccharalis
5500
Pernambuco,
Humber et al., 2009 [14]

5502
Brazil

Brazil

Santa Fe,

Argentina

Santa Fe,

Argentina

D. saccharalis
B. brigniartii 

ARSEF: 1830
Sao Paulo, Brazil
Humber et al., 2009 [14]

D. saccharalis
Cylindrocarpon sp.

ARSEF: 8043

8044
Colima, Mexico
Humber et al., 2009 [14]

M. anisopliae
Colima, Mexico

ARSEF: 3290
Colima, Mexico

D. saccharalis
3291
Colima, Mexico
Humber et al., 2009 [14]

3292
Colima, Mexico

3298
Colima, Mexico

M. anisolpliae

D. magnifactella
B. bassiana
Mexico
Hernández and Velázquez

2004 [15]

Hirsutella sp.

D. grandiosella
Nosema sp. isolate 167
Marshal County, (Mississippi, USA)
Inglis et al. 2000 [5]

D. grandiosella
Nosema sp. isolates 295, 504
Oktibbeha County, (Mississippi, USA)
Inglis et al. 2000 [5]

D. grandiosella
Nosema sp. isolates 181, 513, 522
Washington County, (Mississippi, USA)
Inglis et al. 2000 [5]

D. saccharalis
Nosema sp.
Colombia
Lastra and Gómez 2000

[16]

D. saccharalis
Granulovirus (DsGV)
Southern United States
Pavan et al. (1983) [17]

D. saccharalis
Densovirus (DsDNV)
Guadeloupe
Meynadier et al. (1977) [18]

D. magnifactella
B. thuringiensis
Mexico
Fonseca-González et al. (2011)

[19]

The life cycle of entomopathogenic fungi in the arthropod hosts is initiated with the germination of conidia that contacts the host integument and produces a germ tube that penetrates the host through a combination of physical pressure and enzymatic degradation of the cuticle. The fungus initially colonizes the host through a yeast phase. Host death usually results from a combination of nutrient depletion, invasion of organs, and the action of fungal toxins. Hyphae usually emerge from the cadaver. The mummified corpse of the insect remains in the environment for several weeks and, in the case of stem borer, it keeps remains protected inside the stem. Therefore, it is more common to detect and isolate fungi compared to other pathogens that destroy the host, such as Bacillus thuringiensis (Bt), viruses, or nematodes.

Entomopathogenic fungi are widely distributed in all regions of the world; these species have wide genetic variation among the different isolates. Pathogenicity and virulence to different species, as well as enzymatic and DNA characteristics, vary among different isolates [14, 20, 21]. Therefore, it is important to evaluate as many geographic isolates as possible from different areas to select the most suitable isolate based on its virulence and growth at high temperatures. Several research groups have verified the pathogenicity and virulence of Hypocreales fungi, such as M. anisopliae and B. bassiana (Table 2), which have become important biocontrol agents used for the microbial control of Diatraea spp. It is possible that a limited selection of available isolates of B. bassiana could identify highly virulent strains from each of the different Diatraea species.

Table 2: Entomopathogenic-fungi bioassays on Diatraea spp.

Host species
Pathogen
Bioassay
Result
Reference

D. saccharalis
B. bassiana
Immersing third instar larvae into a suspension of 10⁸ conidia/mL
TL₅₀ (conidia isolates Bb1 and 5): 4.3 days.

Mortality dry mycelium preparations:

21.3% (Bb1)

82.5% (Bb5) at 7 days after inoculation
Arcas et al., 1999 [26]

D. saccharalis
B. bassiana

Mycotrol strain GHA
Sprayed first, second and third instar larvae with 1 mL
LD₅₀ in spores/mm²

1st instar: 72.1

2nd instar: 384.3

3rd instar: 777.0

Mean days of survival

1st instar: 4.6

2nd instar: 4.8

3rd instar: 6.4
Legaspi et al., 2000 [27]

D. grandiosella
B. bassiana

Eight isolates
Dipped in conidial suspension (dosage not specified)
Mortality from 10 to 21%
Inglis et al., 2000 [5]

D. saccharalis
B. bassiana ATCC 20872 from Solenopsis invicta
Third instar larvae sprayed with suspension of 10⁸ conida/mL
TL₅₀ 3.02 to 4.10 days
Marques et al., 2000 [28]

D. sacccharalis
B. bassiana ATCC 20872 from Solenopsis invicta
Third instar larvae sprayed with 3 mL of yeast-like cells or conidia
LC₅₀ 5.6 × 10⁶ yeast/mL and

4.8 × 10⁶ conidia/mL

Alves et al., 2002 [29]

D. saccharalis
B. bassiana

IBCB from Brazil
1 cm larvae sprayed with 1 mL
CL₅₀ 1.58 × 10⁷ conidia/mL at 6 days after inoculation
Wenzel et al., 2006 [30]

Field evaluations have been performed using M. anisopliae and B. bassiana against several insect pests of sugarcane, including D. saccharalis in Brazil. Application of M. anisopliae at a rate of 1 × 10¹³ spores per hectare caused 58% mortality of D. saccharalis, and B. bassiana at 3.7 × 10⁸ spores per milliliter reduced D. saccharalis damage by 45% (Alves et al., 1984 and 1985, cited in Legaspi et al. [22]). For effectiveness in the field, it is important to consider the contact between the spores and the host, formulation, and the virulence of the pathogens.

3. Microsporidia

Microsporidia (Eukaryota: Fungi) is the most ubiquitous group among insect populations [23, 24]. Microsporidia are tiny unicellular organisms (from 2 to 40 μm in diameter), that are opportunistic and obligate intracellular parasites and attack different groups of invertebrate and vertebrate animals. Microsporidia generally produce chronic diseases and reduce the physiological and reproductive ability of their hosts. Many species of microsporidia infect arthropods, especially insects such as Lepidoptera and Coleoptera [23–25].

In general, these microorganisms live as parasites in cells of the midgut epithelium, where they complete their development, and the cycle starts when the infective states of microsporidia (spores) arrive at the digestive tube and colonize this region all the way to the excretory system. The spores germinate because of the acid intestinal pH, the microorganisms penetrate the midgut cells, and the intestinal activity is paralyzed between 14 to 21 days later because the insect cannot assimilate nutrients. Nosema locustae infects the adipocytes of fat body, which interferes with the adequate function of the insect’s intermediary metabolism and competes with the insect for energetic reserves. Microsporidia produce effects that depend on the species and concentration; however, they generally produce weakness and eventually lead to death [31, 32].

Only some species in this group have the possibility to be potentially relevant for natural or classical control. There are many studies worldwide on the pathogenic effects of the microsporidia N. pyrausta (Payllot) and Vairimorpha necatrix (Kramer) on borer organisms such as Ostrinia nubilalis (Hϋbner) (European corn borer), Lymantria dispar (Gypsy moth), and the grasshopper [5, 32]. In this group, only the microsporidium N. locustae has been registered as a microbial insecticide for the control of grasshoppers in grasslands [33].

In many areas of the USA and Europe, Nosema is the main agent used for the control of grasshoppers; however, in Latin America, few studies have been performed with Nosema [33]. For example, Inglis et al. [5] reported six isolates of Nosema on larvae in the winter diapause stage from D. grandiosella Dyer collected during 1998 in corn stems from three locations in Mississippi, USA. However, the frequency of infection in the field was very low (1, 3, and 15% in the counties of Marshal, Oktibbeha, and Washington, resp.), and no isolates were found in D. crambidoides (Grote) (Table 3).

Table 3: Entomopathogenic-microsporidia bioassays on Diatraea spp.

Host specie
Pathogen
Bioassay
Results
Reference

D. grandiosella
Nosema sp isolate 167
Surface of diet (squares), concentration not mentioned. Larvae and pupae
Mortality of 1–15%
Inglis et al., 2000 [5]

D. grandiosella
Nosema sp isolate 295, 504
Surface of diet (squares), concentration not mentioned. Larvae and pupae
Mortality of 1–15%
Inglis et al., 2000 [5]

D. grandiosella
Nosema sp isolate 181, 513, 522
Surface of diet (squares), concentration not mentioned. Larvae and pupae
Mortality of 1–15%
Inglis et al., 2000 [5]

D. saccharalis
Vairimorpha necatrix
1st instar larvae, Surface of diet
Microsporidiosis LC₅₀ = 48.4 spores/mm²

Gut damage LC₅₀ = 8941 spores/mm²
Fuxa, 1981 [32]

D. grandiosella
Nosema sp isolate 506
5-day-old larvae and pupae, leaf pieces ~5 cm² with 10 l inoculum (10¹ to 10⁷ spores)
Median infective dose of 2 × 10³ spores per larva, pupae small, decreased egg production
Inglis et al., 2003 [34]

When the mortality produced by the Nosema isolates was assessed in the laboratory using larvae that had stayed in environmental-like natural winter diapause conditions, variations in mortality between 0 and 55% were observed in larvae, and variations between 7 and 29% were observed in pupae; homogenization of the dead larvae revealed a large amount of Nosema spores. However, in the surviving adults, a large number of larvae were positive for the Nosema spores when they were analyzed under light microscopy using staining techniques and electron microscopy.

In these experiments, the larvae were reared in the winter diapause stage, and the temperature for Nosema was not optimal for the development of infection; in the field during the winter, the prediapause larvae migrate to the base of the corn stem just below the surface of the soil [5], and it is possible that Nosema can produce infection in natural conditions as the temperature inside the cane is higher than the external temperature.

Phoofolo et al. [35] observed a similar behavior, and they reported that the O. pyrausta infection by Nosema is chronic. Although no immediate mortality is produced, the longevity and fecundity of the adults are reduced. Likewise, Phoofolo mentioned the possibility of a response to other factors of mortality that regulate the population dynamics of borers, such as low temperature, the host plant, and crowding, which have chronic effects on Nosema infection. Therefore, although the pest did not die directly because of microsporidia infection, the population size was eventually reduced.

This effect has been previously observed by Fuxa [32], when he evaluated the susceptibility of larvae in the first and third instars of six species of Lepidoptera, including D. saccharalis, to the microsporidium Vairimorpha necatrix. He described two routes of infection that resulted in mortality; one resulted from the chronic effects produced by the exposure of larvae to low doses of spores, which led to lethal septicemia (microsporidiosis) just before pupating, and the other route was due to the intake of a large number of spores, which apparently damaged the gut because of the introduction of a large number of spore polar filaments. In this study, Fuxa [32] concluded that this pathogen could be promising for the remaining five species, although not for D. saccharalis, as he had observed direct mortality by damage to the gut and indirect mortality by septicemia and because a high concentration of spores was necessary.

Inglis et al. [34] reported that one strain of Nosema (506), which had been previously isolated [5], can infect other species of insects in the Crambidae family, including O. nubilalis and D. crambidoides (Grote), but not other species of Noctuidae. However, they also observed that the infection can be transmitted transovarially, although at a very low frequency (a difference with other Nosema species that can be transmitted frequently in the vertical rout, such as the genus Ostrinia).

Solter et al. [36] researched the vertical and horizontal transmission of seven species of microsporidia, including two strains of Nosema sp. (isolated from their natural hosts in the field D. saecharalis and Eoreuma loftini); although five of these strains were transmitted at a low percentage horizontally and vertically, two Nosema strains did not behave similarly.

In assessing the infectivity percentage under laboratory conditions when applying high concentrations of spores, they observed very low mortality in the larvae of D. sacharalis and E. loftini (2 and 4%, resp.), although the same strains produced high mortality at low concentrations in the other species studied, including O. nubilalis. They explained the low mortality, even at high doses, as a function of low horizontal and vertical transmission because few infective or abnormal spores were produced.

In this case, it is important to clarify that they could obtain live infected larvae without mortality; however, they only measured mortality and did not measure other parameters, such as fecundity or larvae hatching in the next generation. As mentioned by Fuxa [32] and Inglis et al. [5], infection does not always lead to mortality, and occasionally the effects are observed long term and can be inferred from a reduction in fecundity and susceptibility to other stress situations.

Lastra and Gómez [16] implemented a system to produce natural enemies of D. saccharalis in CENICAÑA (Clombian Sugarcane Research Center, Colombia) and began their colony with larvae collected in the field. Their most significant result was the detection of one “protozoa,” possibly Nosema, identified as the causal agent for the diminution in larvae production. They observed refractile spores in the fluids of the malpighi tubes or hemolymph using phase contrast microscopy in healthy adults. Macroscopically, the diseased larvae were dwarfed and white.

In addition, different pathogenic microorganisms have been isolated from Diatraea sp. in experimentally transmitted infections in the laboratory; however, there is little information about the natural presence of entomopathogens belonging to the microsporidia group in this species of borer. It is important to perform a systematic search for infected larvae, pupae, or adults of Diatraea, and studies need to include microscopy techniques to complement the mortality bioassays because infections could be asymptomatic. As many microsporidia do not produce insecticide activity quickly and because many species have complex life cycles that involve more than one host, very few attempts have been made to implement microsporidia and use them as agents of biological control.

4. Nematodes

Pathogenic nematodes of the Heterorhabditidae and Steinernematidae families live in the soil where they are parasitic to certain soil-dwelling insects. The free-living third juvenile stage (infective juveniles, IJs) locates a suitable host and penetrates the host in different ways. Once inside the insect, the IJs initiate its development, the nutritional tract becomes functional, and the symbiotic bacterium Xenorhabdus or Photorhabdus is released through the anus and begins to multiply in the hemocoel, which kills the insect by septicemia and creates suitable conditions for the reproduction of the nematode. Nematodes feed on bacteria and the dead tissue of the host, and they pass through several generations until new IJs are produced, which emerge from the cadaver [37]. They have a ubiquitous distribution [37]; therefore, it is not unusual to find them in the soil of sugarcane plantations (Pizano et al., 1985 cited in Khan et al. [38, 39]).

However, there are few reports of nematodes infecting stem borers of the Diatraea genus in natural and experimental situations. In Costa Rica, several entomopathogenic species, including nematodes, have been isolated from D. tabernella [40]; however, the author does not mention the nematode species.

Khan et al. [38] reviewed the world bibliography on rice stem borers and found five reports of entomopathogenic nematodes associated with D. saccharalis, two reports of Steinernema (=Neoaplectana) glaseri in Brazil and three reports of Steinernema (=Neoaplectana) carpocapsae in the USA and Guadeloupe. However, in one study, they report that the aim of the research was the use of D. saccharalis to produce S. glaseri in controlled conditions because of its susceptibility to the nematode. Another study by Folegatti et al. [41] used entomopathogenic nematodes and D. saccharalis in laboratory conditions to produce S. carpocapsae in vivo, using larvae of the sugarcane borer as the host.

We believe that pathogenic nematodes have good potential for the biological control of Diatraea species because of their presence in sugarcane-producing regions (as mentioned above) and because the species S. feltiae, S. glaseri, S. rarum, Heterorhabditis heliothis, and H. bacteriophora have demonstrated high infectivity in D. saccharalis [42–45] (Table 4) as well as in E. loftini (some of them) [46, 47] in experimental studies. We consider that performing a systematic search for infected larvae or pupae of Diatraea could be helpful in finding new nematode strains and species adapted to local environmental conditions and pest species that could be used in the future.

Table 4: Entomopathogenic-nematode bioassays on Diatraea spp.

Host specie
Pathogen
Bioassay
Result
Reference

Steinernema feltiae [Neoaplectana carpocapsae]

100% mortality with 5000 nematodes/larvae

Diatraea saccharalis
Heterorhabditis heliothidis
Several concentrations for all species
100% mortality with 5000 nematodes/larvae
Sosa et al., 1993 [42]

S. glaseri

30% mortality with 5000 nematodes/larvae

Diatraea saccharalis
S. feltiae

S. rarum

Heterorhabditis

bacteriophora
500 IJs/5 larvae in Petri dishes with two moistened filter papers disks
Adults and IJs produced. Larval mortality >90%
De Doucet et al., 1999 [43]

Diatraea saccharalis
H. bacteriophora JPM4 isolated from soil in Larvae MG, Brazil
Larvae were exposed to 25 5 IJs/100 L for 48 h in sand-filled Petri dishes
100% larval mortality within 5 days. LT₅₀ 2.1 d and LT₉₅ 4.4 d
Molina et al., 2007 [44]

Diatraea saccharalis
Heterorhabditis bacteriophora
500 mL/L applied to larvae
93% of larvae mortality
Aguila et al., 2008 [45]

Although we could not find any report on field trials, we are aware that there have been mass rearings of H. bacteriophora in Cuba since 1987 to control soil pests, such as D. saccharalis [45].

5. Bacteria

Bacillus thuringiensis (Bt) is a Gram-positive bacterium that has been isolated from several sources, including soil, water, phylloplane, and insect cadavers. Bt produces various insecticidal crystal proteins during the onset of sporulation that are toxic to insects, acari, and nematodes [48]. The steps involved in the mode of action of the proteins after their ingestion are as follows: (a) solubilization of the crystals by the highly alkaline pH of the midgut, (b) activation of the proteins by proteases, (c) binding of the toxins to specific receptors located on the microvilli membrane of the midgut columnar epithelium cells, and (d) the insertion of the toxin into the membrane, which forms a pore and induces cell lysis. There are no reports from the American continent on the isolation of Bt strains from Diatraea spp. larvae cadavers; however, some Bt strains and pure proteins have been evaluated against these pests.

Bohorova et al. [49] evaluated Cry1Aa, Cry1Ab, Cry1Ac, Cry1B, Cry1C, Cry1D, Cry1E, and Cry1F Bt pure proteins against four species of Lepidoptera that are pests of maize, including D. saccharalis. The proteins were diluted in water and added to the diet at doses of 10 and 100 mg/mL of diet, and mortality was recorded after seven days. D. saccharalis was susceptible to Cry1B protein at 10 mg/g, and the LC₅₀ was 113.6 mg/g of meridic diet (Table 5).

Table 5: Entomopathogenic-bacteria bioassays on Diatraea spp.

Host species
Pathogen
Bioassay
Result
Reference

Diatraea saccharalis
Bacillus thuringiensis
Diet incorporated spores-crystals
LC50 113.6 mg/g diet
Bohorova et al., 1997 [49]

D. saccharalis
B. thuringiensis

LC₅₀ 33.21 g/mL
Rosas-Garcia et al., 2004 [50]

D. saccharalis
B. thuringiensis
Diet incorporated spores-crystals complex
LC₅₀ 13.06 g/L
Gitahy et al., 2007 [51]

D. saccharalis
Photorhabdustemperata
Cells injected

LD₅₀ 16.2 bacterial cells

LT₅₀ 33.8 h
Carneiro et al., 2008 [52]

Diatraea grandiosella
Bacillus thuringiensis
Diet incorporated spores-crystals
LC₅₀ 5.2 mg/g diet
Bohorova et al., 1997 [49]

Diatraea grandiosella
Bacillus thuringiensis

Biotrol BTB 183-25 Nutrilite Products Incorporated, Kansas City, Missouri, USA

Immersing stem seedling in spores suspensions: 1.25 × 10⁸, 6.25 × 10⁷, 2.5 × 10⁷ and 1.25 × 10⁷

Mean mortality on 5-day-old larvae: 32, 31, 25, 15 out of 40.

Control mean mortality 0.01

Mean mortality on 10-day-old larvae: 15, 15, 12 and 6 out of 20.
Sikorowski and Davis, 1970 [53]

Twelve Bt strains were evaluated by Rosas García et al. [50] at a dosages of 50 and 500 μg of total protein and spores per milliliter against 2-day-old D. saccharalis larvae. The strains used were HD1, HD2, HD9, HD29, HD37, HD59, HD133, HD137, and HD559, as well as the GM7, GM10, and GM34 native strains. The strains that killed more than 50% of larvae were selected to obtain the LC₅₀. Strains HD133, HD559, GM7, GM10, and GM34 were toxic; however, GM34 was the most toxic with an LC₅₀ of 33.21 μg/mL (Table 5). PCR analysis was performed to determine the cry1 genes of the toxic strains: HD133 cryAa, cry1Ab, cry1C; HD559 and GM7 cry1Aa, cry1Ab, and cry1B; GM10 cry1Aa, cry1Ab, cry1Ac, and cry1C; GM34 cry1Aa, cry1Ab, and cry1Ac.

Gitahy et al. [51] evaluated a spore-crystal complex in vitro from five native Bt strains (S48, S76, S90, S105, and S135) and HD1, which served as a positive control, on second instar larvae of D. saccharalis and mortality was recorded after 5 days. The strain S76 caused 100% mortality at 72 h, HD1 caused 69% mortality, and 3% of the other native strains caused mortality at 500 μg L⁻¹ of the spore-crystal complex. The LC₅₀ values of the S76 and HD1 strains were determined, and S76 was 11-fold more toxic (13.06 μg/L) than HD1 (143.88 μ/L) (Table 5). The S76 strain carries cryAa, cry1Ab, cryAc, cry2Aa, and cry2Ab genes, which is similar to HD1.

There are other species of bacteria with potential insecticide activity; Carneiro et al. [52] evaluated Photorhabdus temperata, which is a bacterium associated with Heterorhabditis entomopathogenic nematodes. Cells were injected with a volume of 10 μL of phosphate-buffered saline directly into the hemocoel of fourth instar D. saccharalis larvae, and the LD₅₀ was 16.2 bacterial cells with an LT₅₀ of 33.8 h.

Sikorowski and Davis [53] determined the susceptibility of 5- and 10-day-old D. grandiosella larvae with a Bt commercial product (Biotrol BTB 183-25) using 1.25 × 10⁸, 6.25 × 10⁷, 2.5 × 10⁷, and 1.25 × 10⁷ spores per milliliter (Table 5) where two-inch stem seedlings were dipped for 30 min in spore suspensions. One 10-day-old or two 5-day-old larvae were allowed to feed on each stem. In addition, a known number of spores were placed or injected into 5-day-old stem seedlings at  inch sections. Stems dipped in water were used as controls. Mortality was recorded after 48 h. With 10-day-old larvae, five replicates with 20 larvae for each treatment were used; for the 5-day-old larvae, two replicates with 40 larvae per treatment were used. With 5- and 10-day-old larvae at 1.25 × 10⁸ spores per milliliter, a mean of 32 dead larvae out of 40 and 15 dead larvae out of 20 were recorded (Table 5), respectively. Thus, it was concluded that this species is highly susceptible to B. thuringiensis.

6. Viruses

Viruses that infect insects have received great attention as biological control agents because of their specificity on insect populations; they have little or no impact on the environment and are an ecological friendly alternative to chemical pesticides [58]. Entomopathogenic viruses are grouped in 33 genera within fifteen families [59]. However, only a few of these families, such as Baculoviridae, Poxviridae, and Reoviridae, have potential as biological control agents and have been successfully used in microbial control programs. A common characteristic among these entomopathogenic viruses is that the virions (infective unit) are occluded within a crystalline protein matrix to form an occlusion body (OB) [60], which is a unique characteristic of viruses that infect insects. In these entomopathogenic viruses, OBs have independently evolved as a protective mechanism to environmental factors, which gives the viruses a great advantage as biological control agents [61]. Baculoviruses (BVs) and entomopoxviruses (EPVs: subfamily Entomopoxvirinae) have a large double-stranded DNA genome, and cypoviruses (CPVs: family Reoviridae, genera Cypovirus) contain segmented double-stranded RNA viruses.

EPVs have been reported to infecting insects of a variety of orders, such as Coleoptera, Lepidoptera, Orthoptera, and Diptera. Some EPVs have two distinct OBs spheroids and spindles. The spheroids occlude virions, whereas the spindles do not [62]. The most abundant proteinaceous component of spheroids and spindles is proteins called spheroidin and fusolin, respectively [63–66]. EPV fusolin is an enhancing factor (EF) that increases BVs infection and has been characterized as a chitin-binding protein [67]. The fusolin mechanism of action is similar to that of Calcofluor, which facilitates BV infection by disrupting or preventing the formation of the peritrophic membrane [68]. EPVs are pathogenic but are scarcely virulent; infected larvae exhibit extreme longevity and take up to 70 days to die. However, EPVs have potential as biological control agents for pest insects where BVs have not been isolated [61]. Because of the activity of spindles, the EF of EPVs can be used as a synergistic agent to increase BV infectivity or generate genetically modified organisms, such as BVs and transgenic plants.

CPVs have been mainly isolated from Lepidoptera insects. OBs are dissolved in the midgut, and virions only infect epithelial cells; therefore, CPVs are very pathogenic but act slowly and frequently to produce chronic infections [69]. At this time, no commercial bioinsecticides based on CPVs have been developed. However, Caballero and Williams [61] suggest that their greatest potential as biological control agents is through inoculative or augmentative releases.

BVs predominantly infect insects within the Lepidoptera order, which includes important agricultural insect pests [70]. The Baculoviridae family includes two genera: nucleopolyhedrovirus (NPV), which forms large, polyhedric OBs where many enveloped virions are occluded [71], and Granulovirus (GV), which forms small, granular OBs that occlude only one enveloped virion each [72]. BVs are safe for humans and wildlife. Their specificity is usually very narrow and often is species specific. Because of their specificity and other characteristics, such as elevated virulence and pathogenicity, BVs are by far the most studied and extensively used as commercial biopesticides for the control of a variety of insect pests in many countries around the world [60, 61, 73].

Stem borers of the Lepidoptera order attack gramineous crops throughout the world [74, 75]. Diatraea stem borers (DSB) are widely distributed in the Americas and attack a wide variety of host plants, including maize and sugarcane [1]. Prediapause larvae of southwestern corn borers migrate to the base of the stalk of the corn plant below the soil surface to survive the winters [76]; cryptic habits make the chemical control of these insect pests difficult. Degaspari et al. [77] have argued that chemical control of D. saccharalis in Brazil is not economically feasible. To solve this problem, surveys to isolate endemic entomopathogens of stem borer populations in maize and sugarcane crops have been developed.

Inglis et al. [5] developed an exhaustive survey to isolate entomopathogens from the southern corn borer, D. grandiosella, and the southern corn stalk borer, D. crambidoides, and larvae in the diapause stage were collected from crops located in Mississippi and North Carolina. These authors did not observe OBs in any of the collected larvae and concluded that there are no naturally occurring viruses in these Diatraea populations [5]. According to Inglis et al. [5], Pavan and Ribeiro [55] mentioned that natural populations of the SCB in Brazil do not exhibit endemic viral pathogens.

Currently, there are only two records of endemic entomopathogenic viruses isolated from Diatraea spp. larvae. Pavan et al. [17] isolated a GV from the sugarcane borer (SCB), D. saccharalis, from sugarcane crops in the southern United States (Table 6). These authors developed bioassays with D. saccharalis third instar larvae (Table 6). External symptoms of GV-infected larvae were similar to those reported for other lepidopterous larvae, and the symptoms and ultrastructures were determined using electronic microscopy as well as replication of DsGV, which are typical of GVs. The LD₅₀ for the third instar larvae was 42.3 OBs/larva (Table 6) with 14.5 and 123.6 OBs/larva as the lower and upper limits at 95% probability, respectively [16]. These authors stated that this work was the first of a series of publications; however, there have been no additional studies produced to date, such as the molecular and biological characterization, of this GV, DsGV was introduced into Brazil [16], and Moscardi [78] mentioned that this virus is currently being applied at a small scale as an experimental product on sugarcane.

Table 6: Entomopathogenic-virus bioassays on Diatraea spp.

Host species
Pathogen
Bioassay conditions
Result
Reference

D. grandiosella
Autographa californica

MNPV (AcMNPV)
4 and 7 day-old larvae were fed strips of corn leaf soaked in different dilutions of virus.
DL₅₀: 1.32 10³ PIB*/larvae (for 4-day-old larvae)

DL₅₀: 1.32 10⁴ PIB/larvae (for 7-day-old larvae).
Davis and Sikorowsy, 1978 [54]

D. saccharalis
Diatraea saccharalis Granulosis Virus

(DsGV)
3rd instar larvae feeding individually on small artificial diet discs treated with 2.7 μL of virus dilutions.
LD₅₀: 42 PIB/larva at 26˚C

LT₅₀: From 29 to 63 for 107 and10² PIB/larvae, respectively.
Lastra and Gómez

et al., 1983 [16]

D. saccharalis
Anticarsia gemmatalis MNPV (AgMNPV)

3rd instar larvae fed with artificial diet discs treated with 5 different doses of PIB through 20 serial passages.
LD₅₀: From 7.9 10⁵ to 5.3 10² for 1 and 20 serial passages, respectively.

Pavan and Ribeiro, 1989 [55]

D. saccharalis
AgMNPV wt

AgMNPV-D10
3rd instar larvae individually fed small artificial diet discs containing 10⁵ PIB of virus and kept individually at 10 different temperatures.
LT₅₀: From 10 to 13 for

39 and 30˚C, respectively

From 9 to 29 for 39 and 22˚C, respectively
Ribeiro and Pavan, 1994 [56]

D. saccharalis
Trichoplusia ni MNPV (TnMNPV) wt

TnMNPV-D11
3rd instar larvae fed individually with small artificial diet discs containing 10⁵ PIB of virus and kept individually at 10 different temperatures.
LT₅₀: From 9 to 16 for

39 and 24˚C, respectively

From 6 to 46 for 39 and 17˚C, respectively
Ribeiro and Pavan, 1994 [56]

D. sacccharalis
AgMNPV
3rd instar larvae fed on formalin free diet disc (3 mm) inoculated with 2.7 μL of the viral solution from 10 clonal isolates of the AgMNPV-Ds20
LD₅₀: From 5.3 × 10³  PIB/larvae (7A genotypic variant) to 8 × 10⁴ PIB/larvae (33B genotypic variant)

Ribeiro et al., 1997 [57]

*PIB: Polyhedral inclusion bodies.

Because of their high pathogenicity combined with a limited host range, Densoviruses (DNVs) have potential as effective insecticides [79]. Meynadier et al. [80] isolated a DNV of D. saccharalis (DsDNV) from the Guadeloupe sugarcane borer (Table 6). Kouassi et al. [81] tested the pathogenicity of DsDNV on its host. These authors observed that the infected larvae exhibited infection symptoms from the fourth day postinfection, such as anorexia and lethargy followed by flaccidity and inhibition of molting and metamorphosis. Larvae became paralyzed and stopped feeding after 7 days. The cumulative mortality of infected larvae increased significantly and reached 60% after 12 days and 100% at 21 days postinfection [81]. Although DNVs have no potential for large scale use as a biological control agent because they have no OBs and are related to vertebrate pathogenic viruses, the genes involved with anorexia and paralysis could be used to produce transgenic BVs or plants.

Most BVs have a limited host range and may be species specific in some cases, although there are several NPVs, such as Autographa californica MNPV, Anagrapha falcifera MNPV, and Anticarsia gemmatalis MNPV, which have broader host ranges within the Lepidoptera order [60]. Because of the scarcity of reports on natural isolates of entomopathogenic viruses from Diatraea spp. populations, cross-infectivity of AgMNPV, TnMNPV, and AgMNPV has been evaluated in D. saccharalis and D. grandiosella larvae (Table 6).

Although only two reports of entomopathogenic viruses isolated from Diatraea stem borers exist in the Americas, entomopathogenic viruses, such as NPVs and GVs, have been reported to occur in Africa and Asia in cereal stem borers within the Crambidae family, including maize and sugarcane borers such as Chilo sp. [82], Chilo sacchariphagus [83, 84], Ch. infuscatellus [84, 85], Ch. partellus [86], and Eldana saccharina [87]. As the Mexican territory is the origin of some Diatraea stem borer species, we hypothesize that a great diversity of pathogenic viruses in Diatraea populations that attack sugarcane crops exists in Mexico.

Acknowledgments

This study was financed in part by the project “Selección de enemigos naturales de barrenadores de la caña de azúcar de Colima y Morelos con potencial como agentes de control biológico (SEP-PROMEP),” of Mexico. The authors also thank Mrs. Ingrid Masher for reviewing the paper and for editorial assistance.

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