5

5

2. REVIEW OF LITERATURE

The available literature on entomopathogenic nematodes (EPNs) in

relation to the investigations reported in the thesis has been reviewed under the following

heads.

2.1 Nematodes associated with insects and their distribution

2.2 Symbiotic relationship of entomopathogenic nematodes with bacteria

2.3 Mass production of entomopathogenic nematodes

2.4 Storage and shelf life of entomopathogenic nematodes

2.5 Efficacy of entomopathogenic nematodes against insect-pests.

2.1 Nematodes associated with insects and their distribution

2.1.1. Distribution of entomopathogenic nematodes in world excluding India

Entomopathogenic nematodes are known since the 17th

century and perhaps the

earlier than this (Nguyen and Smart, 2004), however, extensive studies on

entomopathogenic nematodes were carried out in the 19th

and 20th centuries. These

nematodes have a ubiquitous world-wide distribution. Different species/ genera/ families

of these parasites occur in different habitats/ ecosystems depending on their insect hosts

(Mracek 2008). Heterorhabditids and steinernematids are found in many areas of all

continents, excluding Antarctica. It has been reported that the steinernematids are found

in cooler, temperate regions, whereas heterorhabditids are in warmer, tropical conditions

(Hominick et al. 1996). Hominick et al. (1995) demonstrated that at least some

steinernematids show a distinct habitat preference that may reflect the distribution of

suitable hosts which are adapted for the habitat.

Even though the entopmopathogenic nematodes are ubiquitous, their recovery

from the field is influenced by a number of biotic factors, including host range that is

dependent on the suitability for penetration of different insect hosts by nematodes,

possibility of finding a suitable host in the habitats and by the natural population density

of the nematodes creating epizootics in outbreak sites (Peters 1996). Mracek and Beevas

(2000) emphasized an essential impact of host aggregations on the incidence of

entomopathogenic nematodes.

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6

The first entomopathogenic nematode was described by Steiner as Aplectana

kraussei (now Steinernema kraussei) in 1923 followed by Neoplectana glaseri (Steiner,

1929) from material isolated by Glaser and Fox (1930). It was not until Weiser (1955)

described a European population of Neoplectana carpocapsae from codling moth larvae

and Dutky and Hough (1955) isolated the DD-136 strain of an undescribed

steinernematid from codling moth larvae in eastern North America there after the serious

studies on entomopathogenic nematodes began.

The distribution of entomopathogenic nematodes on a global scale is probably

strongly influenced by climate and chance dispersal events, including those associated

with human activities. The juveniles of heterorhabditids disperse vertically and

horizontally, both actively and passively (Parkman et al. 1994; Timper et al. 1988).

Passively, they may be dispersed by rain, wind, soil, humans or insects (Smart and

Nguyen 1994). Soil texture, vegetation and availability of suitable hosts are amongst the

factors that have been implicated in affecting local distribution patterns. There is growing

evidence of preferences of nematode species for certain habitats. S. affine is found largely

in arable lands and grasslands, and virtually absent in forests (Hominick 2002). More

striking is the association of some species with soil of particular texture, in particular

sand. Heterorhabditis megidis and H. indica are almost exclusively found in sandy soils,

resulting in a mainly coastal distribution (Hara et al. 1991; Amarsinghe et al. 1994;

Griffin et al. 1994, 2000). In Germany, the rate of prevalence of steinernematids was

highest in woodland (50.3%) where S. affinis, S. feltiae, S. intermedium and Steinernema

sp. were the predominant species (Sturhan 1999). Similarly, heterorhabditids were

equally abundant in turf and weedy habitats, but never found in closed-canopy forest

(Stuart and Gaugler 1994).

Of the nematodes associated with insects those belonging to the orders

Mermethida, Aphelenchida, Tylenchida and Rhabditida have been most intensively

studied. However, at present only the Rhabditid genera Heterorhabditis and Steinernema

are widely used for insect control due to their high and rapid infectivity and pathogenicity

and easy manipulation (Mracek 2008).

The heterorhabditids are soil inhabiting and have free living, parasitic and

saprophytic stages. The infective third stage juveniles survive outside the insect and

search for hosts and wait for host to pass by (Mracek 2008). The two genera

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7

Sterinernema and Heterorhabditis contain the most important species of

entomopathgogernic nematodes. Currently, there are 25 species in Steinernema and 11

species in Heterorhabditis. All members of the order Rhabditida are bacteriophagous and

many of them have phoretic associations with insects ((Burnell and Stock 2000)).

Therefore, genera Steinernema and Heterorhabditis are considered here and their

distribution on a global scale is reviewed here under. The available information on

various species of these genera with respect to their distribution, hosts, etc. is summarised

in Table 2.1.

Table 2.1 Distribution of entomopathogenic nematodes in different parts of the

world

Species Habitat Host Country Reference

S. feltiae,

H. bacteriophora

Coastal,

wooded and herbaceous

habitats

- Lebanon Noujeim et al. 2011

H. bacteriophora S. glaseri

S. scarabaei

S. feltiae

- Galleria mellonella

Czech Republic

Hyrsl 2011

H. indica, S. abbasi,

S. cholashanense,

S. feltiae

S. siamkayai

Forest, Agriculture

land

River bank

(weeds) Paddy

Walnut

G. mellonella Nepal Chhetri et al. 2010

S. khoisanae,

Steinernema sp, H. bacteriophora

Fruit orchards Whitegrub and

black vine weevil

(Otiiorhynchus

sulcatus)

South

Africa

Hatting et al. 2009

S. pakistanense, S. asiaticum,

S. abbasi,

S. simkayai, S. feltiae,

H. bacteriophora,

H. indica

- - Pakistan Fayyaz and Javed 2009

S. feltiae, S. affine,

Steinernema sp.,

H. bacteriophora

Cultivated field (vine and

vegetables),

coastal regions, oak forest

- France Emelianoff et al. 2008

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8

Species Habitat Host Country Reference

S. carpocapsae - - Slovenia Laznik et al. 2008 S. anatoliense,

S. carpocapsae,

S. feltiae,

H. bacteriophora

Coniferous,

oak forests,

grasses and

palm

- Stock et al. 2008

Heterorhabditis sp.

Steinernemaspp.

Forest, Pasture,

Coffee,

Vegetables

G. mellonella Kenya Nyasani et al. 2008

S. abassi H. bacteriophora

Agricultural fields

- Pakistan Shahina et al. 2007

S. feltiae,

H. bacteriophora,

H. downesi, H. megidis

Oak and

deciduous

forests, new plantations,

fruit orchards

Melolontha

melolontha

Hungary Toth 2006

S. oregonense

S. riobrave

Oak forest - Arizona Stock and Gress 2006

S. yirgalemense H. bacteriophora

- - Ethiopia Makete et al. 2005

S. carpocapsae

S. arevarium

S. weiseri S. silvaticum

H. bacteriophora

Deciduous,

coniferous

forests, fruit orchards and

crop fields

- Czech

Republic

Mracek et al. 2005

Steinernema n. sp. H. indica

- - Costa Rica Lorio et al. 2005

S. aciari

- - China Qiu et al. 2005

Heterorhabditis sp. Greenhouse Bradysia

agrestis

Korea Kim et al. 2004

S. monticolum Heterorhabditis sp.

Turfgrass Agrotis ipsilon A. segetum

Korea Kang et al. 2004

Steinernema sp.

H. bacteriophora

H. indica

- - Egypt Atwa et al. 2004

S. feltiae

- - Netherland Jgdale et al. 2004

Heterorhabditis sp.

- - Korea Kim et al. 2004

S. hermaphroditum

n sp.

- - Indonesia Stock et al. 2004

S. feltiae

Potato Tecia

solanivora

Colombia

Venezuela

Saenz 2003

S. carpocapsae

S. glaseri

S. longicaudum

Greenhouse Bradysia

agrestis

Korea Kim et al. 2003

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9

Species Habitat Host Country Reference

H. bacteriophora

S. carpocapsae Palm Rhynchophorus

ferrugineus

Japan Iiboshi and Izono 2003

S. carpocapsae H. bacteriophora

Korea Jeon et al. 2003

S. feltiae

S. affine

Steinernema n. sp. H. bacteriophora

- - Turkey Hazir et al. 2003

H. indica

H. baujardi

- - Vietnam Phan et al. 2003

S. carpocapsae

H. bacteriophora

Greenhouse Frankiniella

occidentalis

Korea Kim et al. 2003

S. rarum

S. glaseri

S. carpocapsae

H. bacteriophora

- - USA Shapiro-Ilan et al. 2003

S. arenarium

S. glaseri

S. carpocapsae

Heterorhabditis sp.

Sugarcane Mahanarva

Wmbriolata

Brazil Leite et al. 2003

S. carpocapsae

H. bacteriophora

Cassava and

other crops

Cyrtomenus

bergi

Panama Aguilar 2003

H. megidis

S. feltiae

Grains

Maize

Spodoptera

frugiperda Helicoverpa

zea

Mexico Molina–Ochoa et al.

2003a,b,c

S. carpocapsae

- - Japan Iiboshi and Izono, 2003

S. carpocapsae

Heterorhabditis sp.

Coffee Hypothenemus

hampei

- Molina-Acevedo and

Lopez-Nunez 2002, 2003

H. bacteriophora Maize White grubs

Mexico Ruiz-Vega and Aquino-

Bolaños 2002

S. carpocapsae,

S. glaseri

H. bacteriophora

Turfgrass Exomala

orientalis

Korea Choo et al. 2002

S. glaseri H. bacteriophora

Turfgrass Adoretus tenuimaculatus

Korea Lee et al. 2002

S. carpocapsae S. monticolum

H. bacteriophora

Chestnut Parapediasia teterrella

Korea Choo et al. 2001

H. bacteriophora,

H. m

- - USA Mannion et al. 2001

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10

Species Habitat Host Country Reference

S. carpocapsae S. glaseri

S. longicaudam

H. bacteriophora

Vegetables Palpita indica Korea Kim et al. 2001

Heterorhabditis sp.

- - Netherland Wardlow et al. 2001

S. abbasi

- - Taiwan Liao et al. 2001

S. feltiae

Potato Tecia

solanivora

Colombia

Venezuela

Fan et al. 2000

S. carpocapsae Apple Carposina

niponensis

China Yang et al. 2000

Steinernema sp. - -

UK Piggott et al. 2000

S. feltiae

S. carpocapsae

- - Russia Ivanova et al. 2000

Heterorhabditis sp.

Steinernema sp.

- - Azores Rosa et al. 2000

S. krausei S. feltiae

S. affine

S. intermedium S. bicornutum

S. glaseri

- - Czech Republic

Mracek et al. 1999

S. ragum,

S. feltiae, H. bacteriophora

- - Argentina Doucet et al. 1999

S. feltiae

Potato Tecia

solanivora

Colombia

Venezuela

Alvarado et al. 1998

S. glaseri

S. carpocapsae

Turfgrass S. depravata

Parapediasia teterrella

Japan Kinoshita and

Yamanaka 1998

Heterorhbditis sp. - - UK Bennison et al. 1998

H. indica

H. bacteriophora Steinernema sp.

- - Guadeloupe

Islands

Constant et al. 1998

S. feltiae

H. bacteriophora

- - Hungary Lucskai and Mracek

1998

S. bibionis

S. carpocapsae H. bacteriophora

H. indicus

Heterorhabditis spp.Strains HV1,

HV2, HV3, HV4,

HV5, HV6

- - Venezuela Rosales and Saurez

1998

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11

Species Habitat Host Country Reference

H. indicus Steinernema spp.

Crop fields, lawn, turf

grass, sea shore

- Pakistan Shahina et al. 1998

S. kushidai

H. indicus H. megidis

- - Japan Yoshida et al. 1998

S. carpocapsae

S. glaseri

H. bacteriophora S. longicaudum

Turfgrass A. ipsilon

A. segetum

Korea Lee et al. 1997

S. monticolum sp.

nov.

- - Korea Stock et al. 1997

S. carpocapsae Sweet potato Cylas

formicarius Euscepes

postfasciatus

Japan Yamaguchi and

Kawazoe 1997

S. carpocapsae

Banana Cosmopolites

sordidus

Venezuela Rosales and Suarez

1997

Steinernema sp. Potato Premnotrypes

vorax

Chile Garzon et al. 1996

S. carpocapsae

S. glaseri H. bacteriophora

Outhouse Calliphora

lata Muscina

stabulans

Korea Choo et al. 1996

S. carpocapsae,

S. feltiae, S. riobrave,

H. bacteriophora

- - USA Schroeder et al. 1996

S. carpocapsae

- - China Han et al. 1996

S. affinis

S. feltiae S. kraussei

Steinernema spp.

- - Scotland Gwynn and Richardson

1996

H. marelatus n. sp.

- - Oregon Jie and Berry 1996

Steinernema spp. Heterorhabditis

spp.

- - Korea Lee et al. 1996

S. feltiae

S. affinis

- - Belgium Miduturi et al. 1996

S. feltiae Heterorhabditis sp.

- - Spain Pino and Palomo 1996

S. carpocapsae Banana,

Vegetables

Odoiporus

longicollis

China Peng and Han 1996

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12

Species Habitat Host Country Reference

Phyllotreta

striolata

S. carpocapsae Litchi Aristobia

testudo

China Han et al. 1996

H. bacteriophora

- - Cuba Rodriguez et al. 1996

S. carpocapsae Fig Psacothea

hilaris

Japan Tsutsumi and Yamada

1995

S. carpocapsae Litchi Aristobia testudo

China Xu et al. 1995

S. carpocapsae

S. monticolum H. bacteriophora

Forest

Chestnut

Glyphodes

perspectalis

Dichocrocis punctiferalis

Curculio

sikkimensis

Korea Choo et al. 1995

S. carpocapsae

- - China Xu et al. 1995

S. feltiae

S. affinis

H. megidis

- - U.K. Hominick et al. 1995

S. feltiae S. carpocapsae

S. scapterisci

H. bacteriophora

- - Argentina Stock. 1995

S. glaseri

Turfgrass Anomala sp. Japan Yamanaka et al. 1995

S. feltiae Pig sty Musca

domestica

China Xu et al. 1994

S. carpocapsae Palm Sagalassa

valida

Colombia Ortiz-Sarmiento 1994

Heterorhabditis spp.

Steinernema spp.

- - Sri Lanka Amarsinghe et al. 1994

S. affinis

S. feltiae S. intermedia

S. kraussei

S. RFLP type E1 Heterorhabditis sp.

- - Swiss Alps Steiner 1994

S. carpocapsae Apple Carposina

niponensis

China Wang 1993

S. carpocapsae Avenue trees Holcocerus

insularis

China Yang et al. 1993

H. bacteriophora

H. megidis

- - Israel Glazer et al. 1993

Steinernema spp. - - Norway Haukeland 1993

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13

Species Habitat Host Country Reference

S. feltiae

Steinernema spp. H. megidis

Western

Canada

Mracek and Webster

1993

S. carpocapsae

Banana Cosmopolites

sordidus

Brazil Schmitt et al. 1992

S. carpocapsae S. glaseri

H. bacteriophora

Rice

Forest

Chilo suppressalis

Agelastica

coerulea

Korea Choo et al. 1991

Heterorhabditis sp.

- - Peurto Rica Figueroa et al. 1991

Steinernema sp. Heterorhabditis sp.

- - Hawaiian Islands

Hara et al.1991

N. bibionis - - U.K. Hominick and Briscoe

1990

Heterorhabditis sp. S. bibionis

- - U.K. Hominick and Briscoe 1990

S. carpocapsae Strawberry Spodoptera

litura

Japan Gupta et al. 1987

2.1.2. Distribution of entomopathogenic nematodes in India

India is a tropical country having diverse agroclimatic conditions ranging from

the humid, high rainfall north eastern zone to north western semi-arid and arid zones

(Rahaman et al. 2000). The climate is conducive for entomopathogenic nematodes having

no environmental limitation for their commercial exploitation.

In India, the work on entomopathogenic nematodes was first initiated by Rao and

Manjunath (1966) who demonstrated the use of DD-136 strain of S. carpocapsae for the

control of insect-pests of rice, sugarcane and apple. Now there are several reports of

entomopathogenic nematodes parasiting insect pests of rice, maize, groundnut, potato

with a wide host range. The initial work with entomopathogenic nematodes in India was

conducted primarily with exotic species/strains of S. carpocapse, S. glaseri, S. feltiae and

H. bacteriophora. In many cases, these nematodes were found less effective, probably

due to their poor adaptability to the local agro-climatic conditions. India, as in the case

with many other parts of the world, has a rich biodiversity resource because of its varied

geographic, climatic and weather conditions. In India, the search for indigenous strains

have resulted in a number of Indian isolates from different parts of India (Ganguly 2003).

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14

Among the indigenous nematode isolates, two have been described as new

species, H. indica (Poinar et al. 1992) from Tamil Nadu and S. thermophilum Ganguly

and Singh (Ganguly and Singh 2000, 2003) from New Delhi. Other species identified as

indigenous isolates include S. carpocapsae (Hussaini et al. 2001), S. bicornutum

(Hussaini et al. 2001), S. riobrave (Ganguly et al. 2002), S. feltiae and H. bacteriophora

(Sivakumar et al. 1989). Hussaini et al. (2001) also identified some of the native

populations of Steinernema by restriction fragment length polymorphism (RFLP)

analysis and analysis of the PCR amplified ITS-rDNA region using 17 restriction

enzymes. In addition, surveys have revealed natural occurrence of several species/strains

of Steinernema and Heterorhabditis in Andaman and Nicobar islands (Prasad et al.

2001), Gujarat (Vyas 2003), Kerala (Banu et al. 1998), New Delhi (Ganguly and Singh

2000) and Tamil Nadu (Bhaskaran et al. 1994). The detailed list showing occurrence and

distribution of entomopathogenic nematodes in India is given in Table 2.2.

Table 2.2 Distribution of entomopathogenic nematodes in India

Species Habitat Host State Reference

H. indica (Meerut

strain)

- - Uttar Pradesh Prasad et al.

2012

H. indica (Meerut

strain)

- - Uttar Pradesh Pal and Prasad

2012

S. meghalayensis sp. - - Meghalaya Ganguly et al.

2011

S. carpocapsae,

H. indica

- - Meghalaya Gitanjalidevi

2011

S. thermophilum, S. riobrave,

S. harryi,

S. meghalayensis

- G. mellonella New Delhi, Gujarat,

Tamil Nadu,

Meghalaya

Kumar and Ganguly 2011

H. indica, S. thermophilum,

S. glaseri

- - Meghalaya Yadav and Lalramliana

2011

Steinernema sp.,

Heterorhabditis sp., Neosteinernema

- - Uttar Pradesh Khan and Haque

2011

H. indica,

S. thermophilum,

S. glaseri

Dry land, wet

land, jhum

land and forests

- Meghalaya Lalramliana and

Yadav 2010

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15

Species Habitat Host State Reference

S. carpocapsae - - Gujarat Shinde et al.

2010

S. siamkayai Coastal

ecosystem

- Puducherry Adiroubane et al.

2010

H. indica,

S. carpocapsae

- - Kerala,

Karnataka

Shakeela and

Hussaini 2009

S. abbasi, H. indica

- - Haryana Sunanda 2009

H. indica (Meerut

strain)

Deserted

honey comb

G. mellonella Andhra

Pradesh

Sankar et al.

2009

H. bacteriophora Apple orchards and

forest

- Himachal Pradesh

Chandel et al. 2009

S. carpocapsae - - Jammu and

Kashmir

Gupta et al. 2008

S. masoodi - - Uttar Pradesh

Khan et al. 2007

H. bacteriophora,

S. feltiae,

Steinernema sp.

Apple

orchards and

forest

- Himachal

Pradesh

Singh and Gupta

2006

H. indicus Grape garden

Scelodonta

strigicollis

Karnataka Prabhuraj et al.

2006

S. carpocapsae,

H. indica

Agriculture

soil

- Karnataka Hussaini et al.

2004

S. abbasi,

S. tami,

S. carpocapsae,

H. indica, H. bacteriophora

Agriculture

soil

- Karnataka

Hussaini et al.

2004

S. bicornutum

H. indica

Potato

Maize

Crucifers

Tobacco

Brinjal (egg

plant)

Banana

Phthorimaea

operculellaChilo

zonellus Swinhoe

Plutella

xylostella S. litura

Lasioderma

serricorne Odoiporus

longicollis

Karnataka Hussaini 2003

Heterorhabditis sp.,

Steinernema sp.

- - Gujarat Vyas 2003

Heterorhabditis sp., Steinernema sp.

- - Gujarat Vyas et al. 2002

Steinernema sp. Mango - Tamil Nadu Ambika and

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16

Species Habitat Host State Reference

orchard, millet

field

Sivakumar 2002

S. carpocapsae Tobacco

nursery

S. litura Karnataka Sitaramaiah et al.

2003

Heterorhabditis sp., - - Rajasthan Rajkumar et al.

2001

Heterorhabditis sp. Cultivated area, forest,

scrub land and

coastal sandy region

- South Andamans

Prasad et al. 2001

S. thermophilum - - New Delhi Ganguly and

Singh 2000

H. indica,

S. riobrave, S. feltiae,

S. carpocapsae

S. Bicornutum

- - Karnataka Hussaini et al.

2000

Steinernema spp. and Heterorhabditis sp.

- - - Kaushal et al. 2000

S. bicornutum,

S. carpocapsae,

H. indica

- - Karnataka Hussaini et al.

2000

H. indica - - Kerala Banu et al. 1998

H. indicus, Steinernema sp.

- - Tamil Nadu Josephrajkumar and Sivakumar

1997

S. glaseri,

S. feltiae, Steinernema sp.,

H. bacteriophora,

Heterorhabditis sp.

- - Rajasthan Bareth et al.

1997

S. feltiae - P. brassicae, Alphitobius

diaperinus,

Oryzaephilus mercator

- Mathur et al. 1994

S. carpocapsae,

H. bacteriophora,

Heterorhabditis sp.

Groundnut Amsacta

albistriga

- Bhaskaran et al.

1994

S. feltiae. Potato A. ipsilon A. segetum

- Singh 1993

H. bacteriophora - - Tamil Nadu Poinar et al.

1992

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17

Species Habitat Host State Reference

S. feltiae - - Andhra

Pradesh

Singh et al.1992

N. bibionis,

Neoplectana sp. and H.

bacteriophora

Apple orchard

and forest soil

- Himachal

Pradesh

Singh 1990

H. bacteriophora - - Tamil Nadu Sivakumar et al.1989

S. feltiae Tobacco S. litura Karnataka Narayanan and

Gopalakrishnan

1987

S. carpocapsae Vegetables Anomala sp. - Rajeswari et al. 1984

S. carpocapsae Maize Rice

C. zonellus Tryporyza

incertulas

Karnataka Rao et al. 1971

S. carpocapsae Maize C. zonellus - Mathur et al.

1966

2.2 Symbiotic relationship of entomopathogenic nematodes with bacteria

Knowledge of the nematode-bacterial symbiosis is essential to understand the

pathogenicity of the complex for target insects and is important for successful mass

production (Grewal et al. 2004). The nematode-bacterial interaction is symbiotic and

each partner can be cultured separately, but when combined they present a high degree of

specificity (Grewal et al. 2004). Sudhaus and Schulte (1998) has suggested that

Heterorhabditis and Steinernema most probably evolved from necromenic nematodes

which developeded a symbiotic association with an entomopathogenic bacterium. Such a

symbiosis specialized for parasiting animals has not been described so far for any other

group of nematodes (Burnell and Stock, 2000). Thomas and Poinar (1979) reported that

symbionts associated with Steinernema are placed in the genus Xenorhabdus, whereas the

bioluminescent symbionts associated with Heterorhabditis are placed in the genus

Photorhabus (Boemare et al. 1993).

According to Bird and Akhurst (1983), the Xenorhabdus occurs in a special

intestinal vesicle of Steinernema infective juveniles. The Photorahabus are mainly

located in the anterior part of the intestine in Heterorhabditis (Boemare et al. 1996).

Symbiont bacteria of both genera are motile, gram negative and belong to the

Enterobacteriaceae (Burnell and Stock 2000). Both genera are negative for nitrate

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18

reductase and Xenorhabdus are negative for catalase (Grewal et al. 2004). Hazir et al.

(2003) also reported that Photorhabdus spp. are luminescent and catalase positive,

whereas Xenorhabdus spp. have no luminescence and are catalase negative. The colonies

of Xenorhabdus are smooth and somewhat granular in appearance, whereas

Photorhabdus colonies are smooth and mucoid in appearance with irregular margins.

Initially they are pale yellow but change to deep yellow and usually red with age

(Thomas and Poinar 1979). The infective stage of nematode carry bacteria within the

intestinal lumen (Poinar and Leuteregger 1968).

The relationship of steinernematids and heterorhabditids with Xenorhabdus spp. is

one of the classical mutualism. The nematodes provide entry to the bacterium and destroy

its inducible antibacterial system and the bacteria optimize nematode reproduction by

providing nutrients and inhibiting many contaminating microorganisms (Poinar and

Thomas 1967). Hu and Webster (2000) reported that an antibiotic 3, 5-dihydroxy-4-

isopropylstibene is produced by P. luminescens C9 which helps in minimizing the

competition from other microorganisms and prevents the putrification of the nematode

infected insect cadaver.

Both Xenorhabdus and Photorhabdus occur in two phenotypic forms. Phase I

cells are larger than phase II cells and produce significantly greater amounts of

exoenzymes, toxins, antibiotics than phase II forms. The phase I cells are normally the

cells carried by the infectives (Burnell and Stock 2000). When symbiont bacteria are

released by the nematode into the insect haemolymph, the bacterial cells begin to grow

and the death of insect ensues, either from toxaemia or septicemia, depending on the

sensitivity of the insect and the symbiont strain (Frost et al. 1997). Some strains of

Xenorhabdus and Photorhabdus are highly virulent and an injection of less than 10 cells

of the bacterium into the haemocoel may be sufficient to kill a susceptible insect such as

G. mellonella or Manduca sexta (Poinar and Thomas 1967).

Nagesh et al. (2002) isolated Xenorhabdus and Photorhabdus species from both

surface sterilized infective juveniles of the indigenous isolates of Steinernema and

Heterorhabditis spp., and the haemolymph of G. mellonella larval cadavers infected with

the nematodes. Further, Raman and Bhatnagar (2002) purified the insecticidal toxin

complex produced by the bacterium P. luminescens sub sp. akhurstiiwhich was active

against the larvae of Spodoptera litura and G. mellonella. Similarly, Mahar et al. (2000)

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also recorded high mortality in the larvae of G. mellonella due to P. luminescens isolated

from H. bacteriophora. Lengyel et al. (2005) described four new species viz. X.

budapestensis from S. biocornutum, X. ehlersii from S. serratum, X. innexi from S.

scapterisci and X. szentirmaii from S. rarum.

There are no reports of the isolation of Xenorhabdus and Photorhabdus from soil

and it has been generally assumed that these bacteria cannot exist in soil environment in

the absence of their nematode associates (Burnell and Stock 2000).

There exists a close relationship between the taxonomy of the symbiont species

and their nematode host. Each nematode species is specifically associated with one

symbiont species, although a symbiont species may be associated with more than one

nematode species (Akhurst and Boemare 1990). Grewal et al. (2004) reported that

Xenorhabdus bovienii is associated with four species of Steinernema and X. poinarii with

two species of Steinernema. However, P. luminiscens and P. temperate are both

associated with H. bacteriophora. Kaya and Gaugler (1993) reported that, the nematode

relies upon the bacterium for killing the insect host, creating a suitable environment for

its development by producing antibiotics that suppress competing secondary

microorganisms, breakdown the host tissues into usable nutrients, and serve as food

source. The bacterium requires the nematode for protection from external environment,

penetration into hosts haemocoel, and possibly inhibition of the hosts antibacterial

proteins.

Mohan et al. (2003) tested P. luminiscens isolated from entomopathogenic

nematode, H. indica against Pieris brassicae. They sprayed bacterial formulation

containing 1.8 x 106 CFU/ ml uniformly on the foliage of ornamental nasturtium heavily

infested with larvae of P. brassicae and recorded 100 per cent mortality within 24 hours.

There are reports of successful management of mango mealy bug, Drosicha mangiferae

by using P. luminescens isolated from H. indica (Mohan et al. 2004). Mahar et al. (2004)

reported lethal effects of cell and cell free filtrates of X. nematophila taken from S.

carpocapsae against diamondback moth. It was observed that cells can penetrate into

insects even in the absence of the nematode vector. Cell-free solution containing

metabolites was found equi-effective as bacterial cell suspension.

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Chongachitmate (2005) reported association of symbiotic bacteria Xenorhabdus

sp. with S. siamkayai isolated from haemolymph of Helicoverpa armigera. The bacteria

yielded perfect colonies on NBTA media after 18 hours of infection. Direct application of

either cell solutions or cell-free filterates from X. nematophila has provided good control

of S. exigua, Plutella xylostella, Otiorhynchus sulcatus and nymphs of Schistocera

gregaria. The toxicity of cell suspensions and cell free filtrates persisted for upto 5

months in soil (Mahar et al. 2008). Gerritsen et al. (2005) listed oral toxicity of excretion

products of 52 Photorhabdus and Xenorhabdus strains on Frankliniella occidentalis and

Thrips tabaci and only 6 P. temperate isolates from North America were toxic against

these thrips. There was 90 per cent mortality of thrips after 7 days of feeding from P.

temperate supernatant. Thrips were also killed after sucking from leaves covered with the

toxins.

In France, molecular characterization of isolated entomopathogenic nematodes

depicted three different species of Steinernema, one species of Heterorhabditis, and H.

bacteriophora. The Steinernema species were identified as S. feltiae and S. affine and an

undescribed species. Xenorhabdus symbionts were identified as X. bovienii for both S.

feltiae and S. affine. The Xenorhabdus symbionts from Steinernema species was

identified as X. kozodoii. The bacterial symbionts of H. bacteriophora were identified as

P. luminiscens ssp. kayaii and P. luminescens ssp. laumondii (Emelanoff et al. 2008).

Tsai et al. (2008) isolated a symbiotic bacterium of entomopathogenic nematode S.

abbasi from Taiwan which was determined to be a species of Xenorhabdus. This species

was found similar to X. indica of S. abbasi Oman isolate as based on sequence analysis of

16S r DNA.

2.3 Mass production of entomopathogenic nematodes

For a biocontrol agent to be successful it should be amenable for production on

large scale the ready availability of the organism in required quantity and at competitive

cost makes them acceptable among entrepreneurs and farmers (Rabindra and Hussaini

2003). Mass production of entomopathogenic nematodes has evolved from the first large

scale in vitro solid media production by Glaser et al. (1940), to the in vivo production by

Dutky et al. (1964), to the three dimensional solid media in vitro process (Bedding 1981;

1984) and to the in vitro liquid fermentation production method (Friedman 1990).

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Currently, commercial nematodes are produced monoxenically using the solid media

process developed by Bedding (1981; 1984) or the liquid-fermentation method. The

solid-media process has successfully produced pathogenic steinernematids and

heterorhabditids.

During the past few years, a distinct cottage industry has emerged that produces

entomopathogenic nematodes mostly in vivo for the home lawn and garden markets. The

in vivo process requires minimal expertise and capital investments and may be an

important future sector in nematode commercialization for specific niche markets

(Koppenhofer and Kaya 2001).

Dutky et al. (1964) used larvae of the greater wax moth, G. mellonella to multiply

the DD-136 strain of Neoaplectana carpocapsae in vivo and obtained up to 2,00,000

infective juveniles per larva. House et al. (1965) devised a dog food based medium to

produce the DD- 136 strain of N. carpocapsae on a commercial scale. This method was

later refined by Hara et al. (1981) who stressed monoxenicity, and produced 125 million

nematodes/week from 100 dog food agar Petri dishes at a cost of $ 0.28 per million.

Milstead and Poinar (1978) mass multiplied H. bacteriophora on larvae of G. mellonella

and produced higher yield of up to 3,50,000 infective juveniles per host.

Bedding (1981) soaked shredded plastic foam in pork kidney-beef fat homogenate

and placed in 500 ml flasks. Several species of neoaplectanid and heterorhabditid

nematodes were reared successfully with this method with an average yield of 38 million

N. carpocapsae juveniles per flask, at a cost of less than $ 0.02 per million. As an

improvement to the previous method, Bedding (1984) coated shredded polyether

polyurethane sponge with a homogenate of chiken offal (Steinernematids) or chiken offal

+ 10 per cent beef fats (Heterorhabditds), sterilizing the medium in large autoclavable

bags and adding the appropriate bacterium and nematode. He was able to produce about

50, 000 million IJs of N. bibionis in a week at a cost of less than 1 C/ million.

Deseo et al. (1990) used G. mellonella for multiplication of entomopathogenic

nematodes and reported problems in its mass production. They harvested 550 million

juveniles of Heterorhabdistis spp. from about 1 kg larvae of G. mellonella. Flander et al.

(1996) reported that nematode yield in general is proportional to host size, however

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according to Boff et al.(2000) in vivo production yields are dependent on nematode

doses. In Kashmir, H. bacteriophora and S. carpocapsae were cultured on silk worm,

Bombyx mori. The nematodes did not emerge out of host when applied on 5th

instar

larvae, while on 3rd

instar larvae of silkworm, an average of 2,750 and 48,703 juveniles

of H. bacteriophora and S. carpocapsae were recovered, respectively (Zaki 2000).

Husaini et al. (2000) carried out mass multiplication of Steinernema species (SSL 2)

PDBC EN 1321 using four combinations of dog biscuit media in comparison with

Wout’s media. After a culture time of 30 days and an initial inoculum of 500 IJs per 250

ml, maximum yield of 30.58 x 105 IJs was recovered from Wout’s medium. Cost of

production was also minimum in Wout’s medium. Gaugler et al. (2002) developed a

LOTEK system for in vivo mass production of nematodes. They observed high efficacy

in LOTEK system with reduced labour and space requirements relative to the

conventional white trap method. Lewis et al. (2002) collected IJs of H. bacteriophora and

S. carpocapsae in water using standard white trap method and from cadavers through

natural emergence into sand and studied their rates of development. They observed that

when IJs were allowed to emerge from cadavers directly into sand and then allowed to

infect new hosts, they developed into adults at a faster rate than IJs that were collected

with white traps.

Shapiro-Ilan et al. (2002) determined effects of inoculation method, nematode

concentration and host density on in vivo production of S. carpocapsae and H.

bacteriophora in G. mellonella and Tenebrio molitor. It was observed that host

immersion was about 4 times more efficient in time than pipetting inoculum on to the

hosts. Nematode infection increased with nematode concentration and decreased with

host density per unit area. Prabhuraj et al. (2003) tested different hosts for mass

production of entomopathogenic nematodes and recorded highest nematode yield of

1191032.6 and 8807110.9 IJs of S. glaseri and Heterorhabditis sp., respectively, in B.

mori per rupee. However, G. mellonella was cheapest host costing only Rs. 0.026 to

produce single larva, whereas B. mori costed Rs. 0.092.

In order to standardize the optimum inoculum level of H. indicus and S. glaseri

for getting highest level of IJs by in vivo method, Subramanian (2003) used 9 host insects

at room temperature following the filter paper Petri dish exposure method. He observed

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that G. mellonella, H. armigera and S. litura produced a higher number of IJs than the

other hosts. In Pakistan, S. pakistanense, S. asiaticum, S. feltiae and H. indica were mass

produced successfully using chicken offal method by Tabassum and Shahina (2004).

Singh and Gupta (2006) mass produced Himachal isolates of H. bacteriophora

and S. carpocapsae on larvae of Corcyra cephalonica and G. mellonella and on artificial

medium consisting of chicken offal homogenate. In the larvae of C. cephalonica, the

production of H. bacteriophora infectives was 3.2 and 9.5 times higher than that of S.

feltiae and Steinernema sp., respectively. The G. mellonella larvae supported 4.5, 3.0 and

2.5 times more production of H. bacteriophora, S. feltiae and Steinernema sp.,

respectively, than rice moth larvae on artificial medium, H. bacteriophora produced 54.3

per cent more infectives than S. feltiae.

Investigations on mass production of S. masoodi, S. seemae, S. carpocapsae, S.

glaseri and S. thermophilum were conducted at Kanpur by Ali et al. (2008). G. mellonella

was found to be the most suitable host for the production of S. seemae, which yielded

higher infective juveniles than S. carpocapsae. H. armigera was the next best suitable

alternative host, which produced maximum infective juveniles in case of S. seemae

followed by S. masoodi, S. glaseri and S. thermophilum. Rice moth was the least suitable

host. Gupta et al. (2008) used S. litura for in vivo culturing of local isolates of S.

carpocapsae at Jammu. Fifth instar larvae of S. litura @160 IJs/ larva produced

maximum IJs, whereas minimum IJs (0.97 x 105) were obtained from 3

rd instar larvae @

10 IJs/ larva. Bareth and Bhatnagar (2010) standardized an inoculation dose of 150 IJs of

S. glaseri per 5 larvae of G. mellonella which produced maximum yield of up to 21847

IJs/ larva at 250C.

Rishi and Prasad (2012) observed that maximum infective juveniles of Meerut

strain of H. indica are produced from the infective larvae of G. mellonella (50,000-

2,00,000), followed by H. armigera (50,000- 1,50,000), whereas minimum infective

juveniles were produced from infected larvae of P. xylostella (100-2,000).

2.4 Storage and shelf life of entomopathogenic nematodes

After production, the entomopathogenic nematodes often need to be stored for

several weeks. Regardless of how they are formulated, their quality declines with time.

Maximum survival and stability of their infectivity is a goal for long term storage.

General range of storage temperature for steinernematids is 5-100C and 10-15

0C for

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heterorhabditids (Hasan et al. 2009). Steinernematids, especially S. carpocapsae can be

maintained for up to 5 months at room temperature or up to 12 months under

refrigeration (Georgis 1990; Georgis and Hague 1991). Most heterorhabditids whether

formulated or unformulated do not have a long shelf life and suffer high mortalities in

storage (Kaya and Gaugler 1993). Nematode metabolism is temperature driven and warm

temperature (20-300C) increases metabolic activities and reduces nematode viability

(Georgis 1990).

According to Dutky et al. (1964) steinernematids infectives can be stored for up to

5 years without loss of infectivity at 70C in an aerated aqueous suspension. Howell

(1979) reported high survival when juveniles were held on moist filter paper at 30C. An

alternate approach to aqueous storage may be storage in paraffin oil, which holds up to 15

times much oxygen as water (Bedding 1976). According to the method of Bedding

(1984), steinernematids can be stored by placing infective stages on to clean, crumbed

polyether-polyurethane sponge @ 250 million N. bibionis/ 100g of dry sponge and

maintained at 1-20C in aerated polyethylene tubes. Similarly, Greogis et al. (1990) also

reported that entomopathogenic nematodes can be stored by placing IJs in polyether-

polyurethane sponge, alginate, clay, activated charcoal, gel forming polyarylamide,

vermiculite and peat.

The heterorhabditids infectives are best stored in culture flasks, above 120C, but

relatively poorly after extraction (Bedding 1981). However, 2-4 months storage at 4-100C

is considered good for heterorhabditid nematodes (Woodring and Kaya 1988).

Friedman (1990) immobilized active nematodes by maintaining in aqueous

suspension at low temperature (5-150C) to prevent depletion of their lipid and glycogen

reserves. It was observed that steinernematids can be stored for 6-12 months at 4-100C

without much loss of activity, whereas for heterorhabditids 2-4 months of storage at 4-

100C is considered good. According to Pezowiez et al. (1997) the best method of storage

of IJs was the use of gel capsule followed by formalin in 0.1 per cent microporous

synthetic sponge and garden pest. Connick et al. (1993) reported that storage temperature

had greatest effect on recovery of S. carpocapsae juveniles. Nematode recovery after

storage at 210C decreased to zero after 3-6 weeks. Storage of samples at 4

0C and with

high moisture content (19.9-23.1 %) greatly improved nematode viability.

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Hussaini et al. (2000) found liquid paraffin as most suitable for storage of

Steinernema and H. indica isolates at 80C. At 15

0C, distilled water was better than

glycerine and liquid paraffin for all isolates. The viability of IJs of S. bicornutum and H.

indica was more in distilled water than 0.1 per cent formalin at 80C. Strauch et al. (2000)

achieved maximum survival of H. indica at 150C, however, survival of H. bacteriophora

was best at 7.50C and least at 25

0C. High salt concentration, low pH (4-6) and ascorbic

acid had a positive effect on survival of H. indica. For storage at controlled conditions,

aerated water is superior and addition of preservatives will increase survival. The best

storage combinations as reported by Karunakar et al. (2001) are 7.50C, 250 IJs per ml and

120 days, respectively, for S. feltiae; 7.50C, 250 IJs per ml and90 days for S. glaseri and

100C, 250 IJs per ml and 90 days for H. indica.

Hussaini et al. (2003) studied the effect of some optical brighteners (OBS) and

PABA as UV protectant for entomopathogenic nematodes. Addition of OBS and PABA

enhanced the tolerance of IJs of all the indigenous isolates at PDBC Bangalore to UV.

The protective effect was density dependent and irradiation became lethal as the period of

exposure increased. There was no adverse effect of optical brighteners on the

entomopathogenic nematodes. Hussaini et al. (2004) tested the effect of pH on survival of

indigenous isolates of S. capocapse PDBC EN 11 and H. indica PDBC EN 13.3.

Maximum survival for H. indica was obtained at pH 5 and for S. carpocapsae at pH 7. In

general S. carpocapsae tolerated a broader range of pH (4-9) than H. indica. Chaubey et

al. (2004) reported that Steinernema and Heterorhabditis can be stored at 4-100C for 6-12

months without any loss of activity, respectively. In Israel, Chen and Glazer (2005)

developed a novel method for long term storage of IJs of S. feltiae IS-6 at 23 ± 0.30C.

The nematode suspension was converted into calcium alginate granules which were

sealed into plastic boxes along with a piece of sponge soaked in distilled water. After

storage for 6 months, the survival rate of nematodes in the granules in the plastic boxes

was 95.9 ± 4.3 per cent. Katti et al. (2006) studied the survival and infectivity of

Rhabditis sp. and S. thermophilum at room temperature for varying periods on G.

mellonella and C. cephalonica. There was 100 per cent survival of both species of

entomopathogenic nematodes on the two hosts after storage for up to 50 days and

thereafter, the survival of Rhabditis sp. declined to zero and that of S. thermophilum to 10

per cent after 150 days of storage.

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Hussaini et al. (2005) compared survival and infectivity of IJs of S. tami, S.

abbasi, S. carpocapsae, H. indica and H. bacteriophora stored in distilled water. S. tami

and S. carpocapsae survived up to 40 days without mortality and loss in infectivity and S.

abbasi with 2 per cent mortality. Heterorhabditis spp. survived up to 20 days without

mortality and declined to 60 and 20 per cent after 40 and 60 days of storage, respectively.

Meena (2006) studied the storage life of entomopathogenic nematodes in

glycerine (0.5-1.5 %), liquid paraffin and triton X 100 at different temperatures (5-350C)

up to 60 days. There was less mortality of entomopathogenic nematodes in glycerineas

compared to liquid paraffin and triton X 100. S. carpocapsae had a better storage life in

the media as compared to H. bacteriophora. Gulcu and Hazir (2012) evaluated tetra pack

containers as an alternative to tissue flasks for nematode storage. They observed that tetra

pack containers were an excellent alternative to tissue culture flasks for storage of H.

bacteriophora and will more than likely be useful for other entomopathogenic nematode

species.

2.5 Efficacy of entomopathogenic nematodes against insects-pests.

Entomopathogenic nematodes have been applied successfully against soil

inhabiting insects as soil application as well as above-ground insects as foliar spray in

cryptic habitats (Arthers et al. 2004; Shapiroi-Ilan et al. 2006). The foraging strategies of

entomopathogenic nematodes vary from ambush to cruise (Lewis et al. 1992). Of the

commercially available entomopathogenic nematode species, S. carpocapsae and S.

scapterisci are the most extreme ambushers and may nictate for hours at a time

(Campbell and Gaugler 1993). Ambushing nematode species are usually associated with

highly mobile, surface-dwelling hosts. Cruising nematodes never nictate and spend most

of the IJ stage moving through the soil. Commercially available cruise foraging species

include the Heterorhabditis spp. and S. glaseri (Lewis 2002). These species are usually

effective against relatively sedentary hosts present in the soil (Griffin et al. 2008).

Sometimes, in soil, they have proven superior to chemicals in controlling the target insect

(Gaugler 1981). In soil they have advantages over pesticides and microbial pathogens by

virtue of their attraction and mobility towards an insect. Entomopathogenic nematodes

possess many attributes such as wide host spectrum, active host seeking, long term

efficacy, easy application, compatibility with most chemicals and are environmentally

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safe. However, the pathogenicity, host searching behaviour and survivability of different

nematode species are varied making them suitable in biological control programmes

(Kaya 1990).

The first attempt to control insects with nematodes was made in 1930’s by using

N. glaseri against the Japanese beetle, Popillia japonica (Glaser 1932). Dutky (1959)

obtained 60 per cent mortality of prepupae of codling moth, Cydia pomonella in the

crevices with the application of N. carpocapsae on apple trees. The nematode persisted

from one generation of C. pomonellato the next by penetrating the cracks in the apple

trees to infect the cocoons and the nematode application was superior to the chemical

insecticides. According to Ustimenko-Bakumovskaya and Ishevskij (1979), there was

higher mortality of codling moth on branches during the fall (80-100 %) as compared to

summer (40-75 %) with entomopathogenic nematodes. Kaya et al. (1984) also obtained

90 per cent mortality of overwintering codling moth in winter applications compared to

32 per cent mortality in summer application of N. carpocapsae.

Application of N. carpocapsae in tobacco wet from rains resulted in 80-85 per

cent reduction in larval population of tobacco budworm, Heliothis virescens, within 3-4

days (Chamberlin and Dutky 1958). In another study, the application of N. carpocapsae

to corn ears though yielded high mortality of corn earwom H. zea, but could not prevent

the crop damage (Thanda and Reiner 1962). Experiments conducted on corn, infested

with fall armyworms, S. frugiperda, resulted in 39 per cent larval reduction @ 400

nematodes/plant compared to insecticide check with 74 per cent larval reduction (Landa-

Zabal et al. 1973).

Number of field experiments have been conducted with N. carpocapsae against

foliage feeding lepidopterous insects. There was 73-77 per cent mortality of P. rapae

with nematode application compared to 82-84 per cent mortality with the application of

insecticides (Welch and Briand 1961). Fox and Jaques (1966) also showed that nematode

application was least effective compared to other microbial and chemical insecticides.

However, Pezowicz (1983) reported 100 per cent mortality of B. brassicae, P. brassicae

and Barathra brassicae when treated with 10, 50 and 100 larvae of N.

carpocapsae/insect, respectively. Morris (1985) obtained 95-100 per cent mortality of

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larvae and pupae of P. rapae and P. xylostella with N. carpocapsae @ 250 nematodes/

insect. H. bacteriophora resulted in 50-90 per cent mortality of P. xylostella.

Higher relative humidity conditions give better control with the nematodes than

low relative humidity. Jaques (1967) reported that application of nematode to control

apple defoliators was not successful because of rapid desiccation of nematodes on the

leaves. In citrus, high humidity under the canopy of trees provides ideal conditions

suitable for nematode activity. Srivastava (1978) reported 63.33 per cent mortality of

third instar and 67.67 per cent of fourth and fifth instar larvae of citrus butterfly, Papilio

demoleus with the application of N. carpocapsae.

Soil application of N. carpocapsae reduced cabbage maggot, Delia brassicae

damage but was less effective as compared to chemicals (Welch and Briand 1961).

However, Georgis and Poinar (1983a,b) found that H. bacteriophora was more effective

in controlling cabbage maggot than N. carpocapsae.

Kain et al. (1982) conducted an experiment using nematodes cultured with

associated bacterium and observed 66 per cent reduction in grass grubs. Wright et al.

(1988) also obtained 84 per cent control of Japanese beetle in potted yews with N. glaseri

applied @ 385 nematodes/cm2. The application of H. heliothidis @ 192 nematodes/cm

2

resulted in more than 90 per cent control of the grubs. In another test, Villani and Wright

(1988) reported more than 60 per cent control of Japanese beetle with H. heliothidis when

used @ 310 nematodes/cm2. Against another important species of whitegrub, Rhizotrogus

majalis, H. heliothidis and N. glaseri produced 46-59 per cent mortality of grubs at an

application rate of 388 nematodes/ cm2. In a similar test, Villani and Wright (1988)

obtained 94 and 60 per cent control of grubs with H. heliothidis under laboratory and

field conditions, respectively.

Toba et al. (1983) obtained significant control of wireworms and Colorado potato

beetle with N. carpocapsae. In laboratory conditions, Wright et al. (1987) recorded 80-

90 per cent mortality of grubs of Colorado potato beetle through N. carpocapsae. There

was 88.4-100 per cent reduction in beetle emergence when N. carpocapsae was applied

@ 93-115 nematodes/cm2 under field conditions. H. heliothidis also produced good

results against grubs of Colorado potato beetle both under laboratory and field conditions.

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An adequate control of cutworms, A. segetum has also been reported in Lettuce

with the application of N. bibionis (Theurissen and Fransen, 1984; Lossbroek and

Theunissen, 1985). Their results are comparable with endosulfan. There was 70-86.69 per

cent mortality of A. segetum larvae with the application of S. carpocapsae @ 105-10

6

nematodes/ ring of 5 tomato plants. The number of damaged tomato plants was reduced

from 0-20 per cent. According to Georgis et al. (1989) there was 70-100 per cent control

of black cutworm, A. ipsilon in laboratory containers with anhydrobiotic infectives of N.

carpocapsae.

The use of N. carpocapsae was not successful against lepidopteran pupae (Lewis

and Raun 1978). Kaya and Hara (1980) also reported that pupal stages of soil pupating

lepidopterans under laboratory conditions were less susceptible than pre-pupal stages.

However, adults emerging in soil infested with N. carpocapsae were infested by the

nematodes as they worked their way through the soil to the surface (Kaya and Grieve

1982).

The carpenter worm, Prionoxytus robiniae are completely suppressed in

commercial fig orchards by N. carpocapsae (Lindegren and Barnette 1982). Similarly,

this nematode killed 85-90 per cent larvae of the Zeuzera pyrina,a pest of fruit trees in

Italy (Deseo 1982; Foschi and Deseo, 1983).

The large scale use of Neoaplectana spp. has been developed to control wood

borers in the family Sessidae. More than 90 per cent mortality of borer, Synanthedon

tipuliformis has been obtained on currants with the application of N. bibionis (Bedding

and Miller 1981; Miller and Bedding 1982). In another test, Deseo and Miller (1985)

observed 74-94 per cent infection of larvae of apple clearing moth, S. myopaeformis with

the application of N. bibionis @ 1.30 x 106 nematodes/ tree. N. carpocapsae produced

76-92 per cent mortality of clearing moth larvae.

Bong and Sikorowski (1983) applied N. carpocapsae in early June on maize crop

and reported 88 per cent mortality of H. zea. Although the larval mortality was high, but

the economic damage to ears was not prevented. In India, there was 100 per cent

mortality of H. armigera with Steinernema sp. under laboratory conditions (Vela et

al.1998).

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According to Narayanan and Gopalakrishnan (1987) pupae of S. litura were less

susceptible than pre pupa and adults of S. feltiae. They obtained complete mortality of

prepupa, pupa and adults of S. litura @ 10000, 1000 and 100 nematode/ml. Jackson and

Brooks (1989) compared susceptibility of third instar larvae of Diabrotica virgifera

virgifera to Agriotos, A11, Breton and Mexican strains of S. feltiae. Larvae were most

susceptible to Mexican strain and least susceptible to A11 strain. Further experiments

with Mexican strain of S. carpocapsae against Diabrotica virgifera virgifera showed that

nematode should be applied when second and third instar root worm larvae are

predominant in the field (Jackson and Brooks 1995). There was considerable reduction in

root damage and adult emergence when S. carpocapsae and H. bacteriophora were

applied @ 2,00,000 nematodes/plant against Diabrotica virgifera virgifera in vegetable

crops (Jackson 1996).

Against H. armigera, the heterorhabditids and steinernematids produced more

than 80 per cent and 95 per cent mortality and the LD50 values were calculated to be 50

IJs and 55 IJs/insect, respectively. The youngest instars were the most susceptible to

nematode infection (Glazer and Navon 1990). Glazer and Wysoki (1990) achieved 80 per

cent mortality of giant looper, Boarmia selenaria (Schiffermuller) in Petri dish bioassay,

with a minimum of 20,000 IJs of S. carpocapsae strain ‘A11’. The LD50 was 4,250 IJs/

larva. Similar results were obtained with S. carpocapsae, stain ‘Mexican’, Steinernema

sp., strain ‘CR’; H. bacteriophora, strain ‘HP88’ and Heterorhabditis sp., strain ‘IS’. The

first and second instars were the most susceptible stages to nematode infection.

Field studies on the use of entomopathogenic nematodes for the control of

Diaprepes abbreviates on citrus was conducted by Schroeder (1990). There was 50 per

cent reduction in adult emergence after treatment with infective juveniles of S.

carpocapsae @ 100 or 500 IJs/cm2 or H. bacteriophora (HP 88 and Florida strains) @

100 IJs/cm2. Choo et al. (1991) tested S. carpocapsae and H. bacteriophora against rice

borer, Chilo suppressalis and concluded that both these nematodes hold potential to

control this pest.

Ishibashi and Choi (1991) obtained good control of A. segetum by mixed

application of entomopathogenic nematode, S. carpocapsae and fungivorous nematode,

Aphelenchus avenae. Buhler and Gibb (1994) observed significantly higher mortality of

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A. ipsilon in plots treated with entomopathogenic nematodes as compared to untreated

plots after one day of treatment. S. carpocapsae was slightly better than S. glaseri and

gave little higher control of cut worms. Shanthi and Sivakumar (1991) assessed the

virulence of S. glaseri (Strain NC-34) and S. carpocapsae (Strain DD-136) in laboratory

against third instar grubs of Holotrichia consanguinea using filter paper method. Strain

NC 34 of S. glaseri was found more virulent. In Japanese beetle, Popillia japonica, H.

bacteriophora and S. carpocapsae gave 60 and 51 per cent control of larvae in a field test

in turf grass plots after 34 days, respectively (Klein and Georgis 1992).

Hara et al. (1993) tested 20 strains of steinernematids and heterorhabditids against

Liriomyza trifolii. The larval mortality ranged from 48-98 per cent. Against plum

curculio, Conotrachelus nenuphar on apple, S. carpocapsae was most effective under

laboratory conditions and a dose of 50 IJs/larva produced 71.8 per cent mortality (Olthof

and Hagley 1993). Sosa et al. (1993) applied S. carpocapsae, H. bacteriophora and S.

glaseri separately to the diet of sugarcane borer, Diatraea saccharalis @ 5,000

nematodes/ml. There was complete mortality with S. carpocapsae and H. bacteriophora,

whereas S. glaseri produced only 30 per cent mortality of third and fourth instar larvae of

D. saccharalis.

In storage application of H. bacteriophora HP88 either as aqueous suspension or

G. mellonella cadavers infected with HP88 proved highly effective against sweet potato

weevil (Lecrore 1994). In Germany, Sulistyanto and Ehlers (1996) evaluated liquid

culture of H. megidis and H. bacteriophora @ 0.5 and 1.5 million dauer

juveniles/m2against Aphodius contaminatus and Phyllopertha horticola on the golf

course. There was 40 and 62 per cent reduction of A. contaminates, whereas in P.

horticola the reduction was 70 and 83 per cent with H. megidis and H. bacteriophora,

respectively.

Bareth et al. (1997) tested seven species of entomopathogenic nematodes viz S.

feltiae, Steinernema sp., H. bacteriophora, Heterorhabditis sp. and two unidentified

strains isolated from local population of A dimidiata. And H. consanguineain Jaipur

against final instar larvae of A. ipsilon and A. flammatra and wireworms. The LT50 values

ranged from 3.43-7.10 days depending upon the species of entomopathogenic nematodes,

host insect and method of exposure. Depending upon the size of host, 450-45,000 IJs/

insect were collected and wireworms were more susceptible to nematodes as compared to

cutworms.

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In India, Shinde and Singh (2000) tested eight nematode species/strains against

diamondback moth. They found that H. bactertiophora was most virulent showing

minimum LD50 (9.16 IJs/larva, LT50 (43.26 hrs) and Lex T50 (3.24 hrs). The average

population potential was 271.4 IJs/mg host body weight and, it was maximum among all

the tested species. Hussaini et al. (2000) conducted sand column assay of S. bicornutum

(Isolate PDBC 3.1, PDBC 3.2 and PDBC 2.1), S. carpocapsae (PDBC 66.1, PDBC 13.1

and PDBC 6.11) and H. indica (PDBC 6.71 and PDBC 13.3) against larvae and pupae of

A. segetum and A. ipsilon. There was 100 per cent mortality of larvae, pupae of A.

segetum with S. bicornutum (PDBC 3.2) and H. indica (PDBC 13.3), respectively. H.

iondica (PDBC 6.71) and S. carpocapsae (PDBC 66.1) gave complete kill of larvae and

pupae of A. ipsilon, respectively. In further studies, using filter paper and sand column

assay for six indigenous Steinernema isolates, there was cent per cent mortality of both A.

ipsilon and A. sgetum (Hussaini et al. 2001).

Hussaini et al. (2003) tested the efficacy of different formulations of S.

carpocapsae, S. abbasi and H. indica against A. ipsilon. Maximum mortality of A. ipsilon

(60-80 %) after 96 hours of inoculation was recorded with alginate formulation in soil

assay. The mortality of A. ipsilon ranged from 33-47 per cent in filter paper assay

method. With Indian isolate of S. riobrave, there was significant reduction in cutworm

population and plant damage in potato crop after six days of treatment. In

entomopathogenic nematode treatment, the plant damage averaged 10.92 per cent as

compared to 28.29 per cent in control (Mathasoliya et al. 2004). Shapiro-Ilan (2005)

compared the potential of newly discovered H. mexicana (MX4) with other

entomopathogenic nematodes against a number of insect-pests. They observed higher

mortality in A. ipsilon with S. carpocapsae as compared to H. mexicana. In Spain, S.

carpocapsae @ 2,00,000 DJs/m2 was as effective as cypermethrin at any of the

concentrations against A. segetum, however, there was no effect of nematode

concentration (Lopez Robles and Hague 2003).

Singh et al. (2001) reported efficacy of H. bacteriophora against whitegrubs after

4 days of exposure. In filter paper impregnation method, the LD50 values for instars I-III

were 110, 326 and 989 IJs/ grub, respectively. However, in soil inoculation method, the

LD50 values were very high as 1875, 5097 and 8942 IJs/ grub, respectively.Jothi and

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Mehta (2002) tested H. indica, H. bacteriophora and S. glaseri against third, fourth and

fifth instar larvae of H. armigera. S. glaseri was more effective causing 63.9 per cent

mortality and third instar larvae were more sensitive with mean mortality of 59 per cent,

followed by fourth instar (56.4 %) and fifth instar (52.9 %).

Chandel et al. (2005) applied H. indica after mixing with FYM during first week

of July in potato at Kufri in Shimla hills. There was 8.13 per cent tuber infestation due to

Brahmina coriacea whitegrubs in treated plots as compared to 11.28 per cent in control

plots. However, under laboratory conditions, they observed 100 per cent mortality of

second instar grubs of B. corecea after 28 days of inoculation. In third instar grubs, 80.76

per cent mortality was recorded. According to Prabhuraj et al. (2006) mortality in H.

armigerawas directly related to increased doses of H. indicus. They also observed that

third instar larve of H. armigera are more susceptible than fourth instar. There was

complete mortality of third instar larvae @ 100 IJs/larva, whereas in fourth instar, 94.7

per cent mortality was noticed.

Singh and Gupta (2006) reported the occurrence of entomopathogenic nematodes

in Himachal Pradesh and tested their pathogenicity against wide range of insects

belonging to four orders. H. bacteriophora was found to be more pathogenic than either

S. feltiae or Sterinernema sp. The whitegrub, B. corecea and greasy cutworm, A. ipsilon

were more susceptible to these entomopathogenic nematodes.In Tamil Nadu, H. indica

and S. glaseri were evaluated agaist H. armigera, S. litura, P. xylostella and

Cnaphalocrosis medinalis. Larvae of P. xylostella were more susceptible to both these

nematodes with LC50 values of 2.01 and 2.53 IJs/ larva, followed by C. medinalis (LC50:

5.46 and 5.17 IJs/larva), S. litura (LC50: 7.32 and 9.04 IJs/larva) and H. armigera

(LC50:9.40 and 10.51 IJs/larva). For pupal stage, the LC50 values of H. indica were 73.29,

86.28, 104.45 and 120.79 IJs/pupa for S. litura, P. xylostella, H. armigera and C.

medinalis, respectively. The LC50 values of S. glaseri for pupal stages of P. xylostella, C.

medinalis, S. litura and H. armigera are reported to be 95.29, 161.67, 108.12 and 122.73

IJs/pupa, respectively (Saravanapriya and Subramanian 2007).

In Kenya, Nyasani et al. (2007) revealed that entomopathogenic nematodes have a

great potential in diamondback moth management. They evaluated five different Kenyen

entomopathogenic nematodes viz S. karii, H. indica, S. waiseri, Heterorhabditis sp and

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Steinernema sp. The ET50 (Exposure time 50) values of tested entomopathogenic

nematodes ranged from 20.27-38.12 hours and it was significantly higher for S. karii than

that of H. indica.

In Kashmir, local isolate of S. carpocapsae produced 100 per cent mortlity of 3-5

instar larvae of S. litura after 96 hours of treatment in laboratory. Fifth instar larvae of S.

litura @ 160 IJs/larva produced maximum IJs (3.29 x 105). The LC50 values varied

between 11.41 and 27.17 IJs/larva in all the instars (Gupta et al. 2008). In Shillong, H.

indica, S. thermophilum and S. glaseri were evaluated @ 10-100 IJs/larva against P.

brassicae. On the basis of LC50 value, S. feltiae was found highly effective (LC50: 30.2

IJ/larva) however, progeny production by larvae of P. brassicae was highest only in case

of H. indica (Lalramliana and Yadav 2009).

In Himachal Pradesh, Chandel et al. (2009) tested the potential of H.

bacteriophora to control A. segetum. The testing was done against 3rd

, 4th

and 5th

instar

larvae @ 10-40 IJs/ cm2 in Petri plates and @ 1000-5000 IJs/ kg soil. In soil, 1000 IJs/kg

were sufficient to initiate infection and produced up to 61.3 per cent insect kill of fifth

instar larvae. There was increase in larval mortality with increase in exposure time and as

the age increased, the mortality decreased. Fetoh et al. (2009) evaluated S. carpocapsae

and H. bacteriophora against A. ipsilon under laboratory and field conditions. Both these

species of entomopathogenic nematodes were found more effective @ 100 IJs as

compared to 25 IJs/larva which clearly indicates that with increase in nematode dose, the

mortality increases.

Sankar et al. (2009) studied the pathogenicity of H. indica against G. mellonella

and tested its compatability with other biopesticides. The combination of Pseudomonas

fluorescence with H. indica was most efficient causing 100 per cent mortality on G.

mellonella after 24 hours of storage. Progeny produced by H. indica on G. mellonella

was maximum (1,40,108 IJs/larva) in the combination treatment with Trichoderma

viridae. Pathogenicity influence of H. indica when exposed with other biopesticides on

host larva proved to be more virulent and compatible.

Divya et al (2010) conducted bioassay experiment to evaluate the pathogenicity of

H. indica against larval stages of H. armigera, S. litura and G. mellonella. All the tested

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stages of larvae of tested insects were highly susceptible to H. indica. However, degree of

susceptibility differed according to instars, dose and periods of exposure. In the dose

response bioassay, second and third instar larvae were found susceptible @ 300 IJs of H.

indica/larva than fourth and fifth instar larvae exposed for 24 hours. Under greenhouse

conditions, per cent larval mortality of H. armigera and S. litura with H. indica @ 25 ml/

plant was significantly more on final instar larvae (62.87 and 56.75 %, respectively) after

60 hours of treatment.

Adiroubane et al (2010) isolated S. siamkayai from Karaikal region of Puducherry

and tested its efficacy against S. litura, P. xylostella, Leucinodes orbonalis, Earais vitella

and C. medinalis. There was increase in susceptibility with an increase in exposure time.

Hyrsl (2011) reported that H. bacteriophora, S. glaseri, S. scarabaei and S.

feltiaeare widely used againstinsect pests of commercial crops. Pathogenicity studies

against G. mellonella revealed that infective stages of entomopathogenic nematodes

killed host within 48 hours, and mortality of host insect was correlated with number of

invaded infective juveniles. The invasion process is very fast, with infective juveniles

entering insect host within a few hours.

Kumar and Ganguly (2011) tested S. thermophilum (New Delhi strain), S.

meghalayensis (Meghalaya strain), S. riobrave (Gujarat strain), S. harryi n. Sp. (Tamil

Nadu strain) against third instar nymphs of solenopsis mealybug (Phenacoccus

solenopsis), adult cotton aphid (Aphis gossypii) and second instar nymphs of cotton

whitefly (Bemisia tabaci@ 50 and 500 IJs/ml in sand well and leaf disc assays. S.

thermophilum caused 83.00 per cent mortality of mealybugs within 72 hours after

inoculation at 50 IJs/ml and 100 per cent within 48 hours at 500 IJs/ml. Against aphid, S.

thermophilum caused 66 and 83% mortality @ 50 and 500 IJs/ml, respectively within 3

days of treatment. All tested Steinernema spp. were ineffective against whitefly at 50

IJs/ml, however, at 50 IJs/ml, S. riobrave produced 66 per cent mortality within 72 hours

after inoculation. S. meghalyensis was the least effective strain.

Prasad et al. (2012) evaluated Meerut- strain of H. indica using filter paper and

soil column assay against different lepidopteran and coleopteran pests. The larvae of P.

xylostella, H. armigera, L. orbonalis and Earais vitella were highly susceptible to H.

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indica causing 73–100 per cent mortality after 48 hours of exposure. The LC50 values

were calculated to be 132, 290, 475 and 810 IJs per larva of P. xylostella, H. armigera, L.

orbonalis and E. vitella, respectively. Against S. litura, Spilosoma obliqua, P. brassicae

and H. consanguinea low susceptibility was recorded.