Field Manual of Techniques in Invertebrate Pathology || Resistance to insect pathogens and...

Chapter IX-1

Resistance to insect pathogens and strategiesto manage resistance: An update

A. M. Shelton, P. Wang and J.-Z. ZhaoDepartment of EntomologyCornell UniversityNew York StateAgricultural Experiment StationGeneva, NY 14456, USA

R. T. RoushUC Statewide IPM ProgramDepartment of EntomologyUniversity of CaliforniaDavis, CA 95616, USA

1 Introduction

Resistance to traditional synthetic pesticides hasbecome one of the major driving forces alteringthe development of integrated pest management(IPM) programs worldwide. Early definitionsof resistance focused on the “development ofstrains capable of surviving a dose lethal to amajority of individuals in a normal population”(cited by ffrench-Constant and Roush, 1990).Sawicki (1987) proposed a definition that “resis-tance (is) a genetic change in response toselection by toxicants that may impair controlin the field.” Others have suggested that geneticchanges in behavior, rather than physiology,may effect resistance and these also needto be taken into account (Lockwood et al.,1984; Sparks et al., 1989; but see also Roushand Daly, 1990, for a more skeptical view).In the most recently published survey, therewere over 500 species of arthropods that haddeveloped strains resistant to one or more of thefive principal classes of insecticides (Georghiouand Lagunes-Tejeda, 1991). Interestingly, thesurvey did not include insect pathogens asone of the principal classes, although it listed

the Japanese beetle, Popilla japonica, and theoriental beetle, Anomala (Exomala) orientalis,as having developed resistance to Paenibacillus(Bacillus) popilliae. Because of resistance tothese other classes of insecticides as well asconcern about some of the deleterious effects oftraditional insecticides, there has been increasedinterest in using pathogens for insect control.Will their increased use result in a higherincidence of resistance to these control tacticsas well?

Insect diseases caused by fungi (includingmicrosporidia), viruses, nematodes, protozoa andbacteria are relatively common and widespreadin nature. They can be very important naturalcontrol factors. Widespread high-level prevalenceof naturally occurring diseases (epizootics)frequently reduces pest populations belowdamaging levels (Roberts et al., 1991). Morethan 100 years ago, entomologists proposed usinginsect pathogens for pest control and until recentlymany entomologists believed that resistance topathogens would be unlikely. This unfoundedoptimism rested on the belief that becausepathogens were natural, had short persistence, andhad coexisted with insects over eons, resistance

793L.A. Lacey and H.K. Kaya (eds.), Field Manual of Techniques in Invertebrate Pathology, 793–811.© 2007 Springer.

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would not occur (Roush, 1998). As in many otherinstances, insects have proven us wrong. Indeed,the long evolutionary associations between insectsand pathogens suggest that where genes for resis-tance exist, their frequencies prior to widespreadcommercial use of the pathogens may be higherthan was the case for chemical insecticides (anexample will be given below). While some insectsare or never were susceptible to certain pathogens(and therefore would not be considered resistantin our sense), others have already developedresistance in the laboratory and/or field. Withincreasing use of pathogens and/or their products,it is likely that we will see increasing cases ofresistance. The Resistant Arthropods Database(http://whalonlab.msu.edu/rpmnews/) should bea valuable repository for such information andreaders are encouraged to enter information into it.

2 Documented cases of resistance

A Bacteria

1 Overview

Bacteria which are applied for insect pestmanagement are primarily in the generaBacillus and Paenibacillus. The milky diseasebacteria (Paenibacillus (=Bacillus) popilliae andP. lentimorbus) are primarily used againstthe Japanese beetle, while subspecies of B.thuringiensis (Bt) aizawai and kurstaki aretargeted for Lepidoptera, Bt israelensis (Bti)for mosquitoes and Bt tenebrionis for Coloradopotato beetle, a pest which has developedresistance to most other alternative insecti-cides. By far the most widely used of thesestrains is Bt subsp. kurstaki (Btk) which hasbeen applied for more than four decades tocontrol lepidopteran pests of horticultural crops,ornamentals and forest systems. Especially forBtk and Bt tenebrionis (Btt) and in contrast to theother microbial pesticides we will discuss, it isnot the living organism that is most important incontrol, but bacterial products, the �-endotoxins,that are the main source of mortality (Höfteand Whiteley, 1989). This is well illustrated bythe increasingly dominant use of toxins derivedfrom Bt in transgenic crops (Chapter VIII-1) and,to a lesser extent, in recombinant bacteria such

as Pseudomonas for crop sprays. Although themost extensive use of Btk sprays is probablyfor control of various forest and shade treespests such as gypsy moth, spruce budworm,tent caterpillars and the like, perhaps the mostintensive use has been against lepidopterouspests of cruciferous crops and especially thediamondback moth , Plutella xylostella, the mostimportant insect pest of crucifers worldwide(Talekar and Shelton, 1993).

2 Cases of resistance

The diamondback moth has a long history ofdeveloping resistance to insecticides used againstit. Since it evolved resistance to the first modernsynthetic insecticide (DDT) and has developedresistance to most other available insecticideswithin less than a decade after their wide-spreaduse, it was not surprising that P. xylostellawas the first insect to be detected with resis-tance to Btk in the field. Different strains of Bthave different �-endotoxins (Höfte and Whiteley,1989), and only some of the toxins are activeagainst any particular insect. Extensive use ofsprays of Btk to control P. xylostella has ledto several cases of resistance to Cry1A andrelated Bt toxins in Australia (Ahmad et al.,1998), Florida (Shelton et al., 1993), Hawaii(Tabashnik et al., 1990), Japan (Hama et al.,1992), the Philippines (Ferré et al., 1991),Central America (Perez and Shelton, 1997), andChina (Zhao et al., 1993). Resistance in onefield-collected population was high enough thatinsects could survive on plants engineered toexpress the Cry1A(c) toxin at high levels (Metzet al., 1995). A colony established from field-collected insects from Hawaii (Liu et al., 1996)displayed about 20-fold resistance to Cry1Ctoxins contained in Bt aizawai (Bta), and apopulation collected in South Carolina with aninitial 30-fold resistance level has further beenselected on Cry1C-producing plants to > 100-fold resistance to the Cry1C protein (Cao et al.,1999). In addition to P. xylostella, Bt-resistantpopulations of the cabbage looper, Trichoplusiani, have been recently identified in commercialgreenhouses in Canada, where Bt had been exten-sively applied (Janmaat and Myers, 2003). Inthese greenhouses, T. ni populations showed alevel of resistance to the Btk formulation, Dipel,

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as high as 160-fold. The resistance in these T. nilarvae was primarily to the toxin Cry1Ac andthe resistant individuals can survive on Cry1Actransgenic broccoli plants (Kain et al., 2004).

Prior to the discovery of field resistance, theability of an insect species to evolve resistance toBt in the laboratory was demonstrated with theIndianmeal moth (Plodia interpunctella, a pestof stored grain) in the mid-1980s (McGaughey,1985). In these tests, populations eventuallydeveloped resistance levels of 140-fold. In total,laboratory populations of at least 10 speciesof Lepidoptera, 2 species of Coleoptera and 4species of Diptera have been selected for resis-tance and 9 of the 16 have evolved resistancegreater than 10-fold (Tabashnik, 1994; Ferré andVan Rie, 2002). The most recent example ofresistance to Bt was in Culex pipiens from NewYork. A strain collected from Syracuse displayed33-fold resistance to a preparation of Bt israe-lensis (Paul et al., 2005). This is by far thehighest level of resistance reported to date andmay portent future problems in mosquito control.

3 Resistance mechanism and genetics

The pathogenesis of Bt toxicity in insectsinvolves multiple steps: (a) dissolution ofthe parasporal crystals in the insect midgutlumen; (b) proteolytic cleavage of the protoxinby midgut digestive proteinases, resulting inactivation of the protoxin to an active toxin;(c) passage of the toxin through the midgutperitrophic membrane to reach the midgutepithelium; and (d) binding of the toxin tothe midgut brush border receptors, followedby insertion of the toxin into the midgut cellmembrane or by activation of a cellular signalingpathway, leading to cell lysis and insect death(Schnepf et al., 1998; Bravo et al., 2004; Zhanget al., 2005). Alteration of any of these stepsmay affect the toxicity of Bt toxins in insectsand can be potentially involved in Bt resis-tance. Therefore, various mechanisms have beensuggested in Bt-resistant insects (Ferré and VanRie, 2002; Griffitts and Aroian, 2005).

Solubilization of Cry protein crystals in themidgut is a factor determining the toxicityin insects (Aronson et al., 1991). Therefore,reduced solubilization could be a potentialmechanism for Bt resistance in insects (Schnepf

et al., 1998). Midgut digestive proteinases arecritically involved in both activation of theprotoxins into their active forms and inacti-vation (degradation) of the activated toxins inthe midgut (Oppert et al., 1994, 1997; Forcadaet al., 1996; Shao et al., 1998; Oppert, 1999).Excessive degradation of Bt toxin by the midgutproteinases could potentially contribute to lowtoxicity of Bt toxins in insensitive or Bt-resistantinsect hosts (Forcada et al., 1996; Shao et al.,1998). Similarly, insufficient activation of Bttoxins by midgut serine proteinases can also bea mechanism for Bt resistance (Oppert et al.,1997). Alteration of midgut trypsin activities hasbeen observed in Bt-resistant strains of P. inter-punctella and O. nubilalis (Oppert et al., 1997;Li et al., 2004).

The target site of Bt toxins is the insect midgutepithelium. Active Bt toxins must penetratethrough the midgut peritrophic membrane (PM)to reach the target site. Therefore, the PM can bean important factor for the toxicity of Bt in insects(Rees et al., 2002). Recently, trapping of Bt toxinCry1Ac in the PM was observed in a Cry1Ac-resistant strain of Bombyx mori (Hayakawa et al.,2004). These observations suggest that the PMmay be involved in Bt resistance. Upon contactwith the midgut epithelium, the Bt toxin bindsto the midgut brush border. The specific bindingof Bt toxins to the midgut receptors is a criticalevent for the toxicity of Bt toxins. Studies oninteractions of Bt toxins with several Lepidopteraspecies demonstrated that reduced binding of Bttoxins to the midgut brush border membranescould result in reduced toxin activities in insects(Lee et al., 1999) and is a primary mechanism forBt resistance (Ferré et al., 1991; Tabashnik et al.,1994; Tang et al., 1996). Currently, identifiedmidgut proteins that may serve as the receptor fora Bt toxin include midgut cadherin-like proteins,aminopeptidases N (APNs) and membrane-bound alkaline phosphatase (Sangadala et al.,1994; Knight et al., 1994; Gill et al., 1995;Valaitis et al., 1995; Vadlamudi et al., 1995;Francis and Bullar, 1997; Keeton and Bullar,1997; Gahan et al., 2001; Jenkins and Dean,2001; Jurat-Fuentes and Adang, 2004; Herreroet al., 2005). More recently, midgut glycol-ipids have also been reported to serve asreceptors for Bt toxins (Griffitts et al., 2005).In B. mori midgut, a 252 kD protein has been

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recently identified as a novel Bt toxin bindingprotein (Hossain et al., 2004). Resistance to Crytoxins in several Lepidoptera species has beenattributed to mutations or lack of expressionof a gene coding for the Bt receptor in themidgut (Gahan et al., 2001; Morin et al., 2003;Xu et al., 2005; Herrero et al., 2005; Jurat-Fuentes and Adang, 2004). Recessive mutationsin a cadherin gene are associated with resis-tance to Cry1A in at least three species,i.e., Heliothis virescens, Pectinophora gossyp-iella, and Helicoverpa armigera (Gahan et al.,2001; Morin et al., 2003; Xu et al., 2005).

In addition, other mechanisms involved in Btresistance have been reported to include aggre-gation of Bt toxin proteins by the midgut esterase(Gunning et al., 2005), elevated melanizationactivity of larval hemolymph and the midgut(Rahman et al., 2004; Ma et al., 2005) andpossibly increased antioxidation activities in Bt-resistant insects (Candas et al., 2003). Clearly,mechanisms for Bt resistance in insects may bemultifaceted.

In Lepidoptera the most common type of resis-tance to Cry1A toxins is known as “Mode 1”resistance, which is characterized by a high levelof resistance to one or more Cry1A toxins,recessive inheritance, reduced binding of one ormore Cry1A toxins to the midgut brush bordermembrane and little or no cross-resistance toCry1C toxin (Tabashnik et al., 1998). “Mode 1”resistance results from an alteration of the midgutbinding sites, which leads to reduced binding ofCry1A toxins to the brush border membranes inhomozygous resistant individuals, but has littleor no effect on the binding in heterozygousindividuals (Tabashnik, 1994; Ferré and Van Rie,2002). Specific binding of a Cry toxin to themidgut brush border membrane is an essentialevent for the toxicity of Cry toxins in insectsand critically determines the toxicity (Schnepfet al., 1998). Domain II, particularly the loopregions of domain II, in the Cry toxins areinvolved in the binding of the toxins to themidgut epithelium. Therefore, individual toxinswithin the same class (e.g., Cry1A with its threesubclasses of Cry1Aa, Cry1Ab and Cry1Ac),which are grouped by their protein sequencesimilarity, likely share the same target site,while toxins in different classes (e.g., Cry1A vs.Cry1C) may have a different target site in the

insect (Crickmore et al., 1998; Ferré and VanRie, 2002). Comparisons of resistance amongP. xylostella, H. virescens, P. gossypiella andPlodia interpunctella show that at least one strainof each of these species has a ‘mode 1’ type ofresistance. On the other hand, at least one strainof each of these three species does not fit themode 1 pattern. Such differences between andwithin species may be important as more sophis-ticated management practices are developed.

In populations of P. xylostella that haveevolved resistance to Cry 1A toxins, the primarymechanism of resistance generally appears to bea reduced binding of the toxin to the epitheliallayer of the brush border membrane of themidgut (Tabashnik et al., 1997). Formulated Btkproducts also contain the HD-1 spore. Testswith a population of P. xylostella collected fromFlorida, which had developed > 1500-fold resis-tance in the field, showed high levels of resis-tance to the HD-1 spore as well as all Cry1Aprotoxins (Tang et al., 1996). When the HD-1spore was combined with a toxin, it synergizedtoxicity to some toxins but not those to whichthe insects had developed resistance.

This same Florida population was used tostudy the inheritance, stability and fitness costof resistance to Btk (Tang et al., 1997). Aswith most other strains (Tabashnik et al., 1998),genetic analysis of F1 and backcross larvaeindicated that resistance was an incompletelyrecessive, autosomal trait probably controlled bya single allele. The instability of resistance hasfigured prominently in resistance managementdebates, although fitness costs seem unlikely tohave a major impact on resistance evolutionunless they are very high for individuals carryingjust one resistance allele (i.e., the heterozygotes)(Roush, 1997). Reports to date indicate that thereis considerable variation in resistance stabilityand fitness costs for Bt among populations ofdiamondback moth, with resistance in somepopulations appearing to have high fitness costsat least to resistant homozygotes (Tabashniket al., 1994), and at least until stabilized bycontinued laboratory selection (Tabashnik et al.,1995). However, the major portion of resis-tance (probably the major gene) in the Floridapopulation has generally seemed to be stable.From the initial > 1�500-fold resistance, resis-tance fell to about 300-fold within 3 generations


in the absence of selection. Unlike previous casesof resistance to Bt (Tabashnik et al., 1994), resis-tance in this Florida colony remained stable atabout 300-fold (Tang et al., 1997), and could berestored to > 1�000-fold after only a single appli-cation of Bt. Further, resistance did not percep-tibly decline in populations with a 50% initialresistance allele frequency.

T. ni is the second insect species that has evolvedresistance to Bt under agricultural conditions, incommercial vegetable greenhouses. Inheritanceanalysis of the resistance to Btk in the greenhouse-evolved T. ni showed a polygenic inheritancepattern and the inheritance was autosomal andincompletely recessive (Janmaat et al., 2004).Further analysis of the greenhouse-evolved resis-tance to the major Cry toxin in Btk, Cry1Ac,showed that the resistance was controlled bya single genetic locus. Similarly, the resistancegene to Cry1Ac was autosomal and incompletelyrecessive (Kain et al., 2004). The resistance ofT. ni to Bt is associated with a fitness cost andthe magnitude of fitness cost negatively correlateswith the suitability of the host plants on which T.ni feeds (Janmaat and Myers, 2005).

The frequency of resistance alleles for Bt hasbeen studied in populations that had not yet beenexposed to significant commercial use of Crytoxins. Gould et al. (1997) estimated that this“initial frequency” of resistance alleles for Cry1A is about 10−3 for H. virescens. Althoughthis estimate may be high (Roush and Shelton,1997; Roush, 1998), it does seem likely on thebasis of successful selection experiments andstudies with genetic markers (cited in Gouldet al., 1997) that resistance alleles for Cry1A maybe on the order of 10−4, which is much higherthan generally supposed for chemical insecti-cides (less than 10−6). Perhaps, resistance to Bttoxins in H. virescens is relatively more commonbecause this species is often exposed to Bt innature, a speculation that may apply to otherspecies and other naturally occurring agents.

B Viruses

1 Overview

Baculoviruses have been the dominantentomopathogenic virus group used in insect pestmanagement. A list of viruses “successfully”

used for insect pest management was suppliedby Roberts et al. (1991), although they notedthat the degree of success is difficult to establishfrom the literature. In a recent review, Moscardi(1999) provides an assessment of the applicationof baculoviruses for control of Lepidoptera anddiscusses how they are used in various cropsthroughout the world.

Viruses have not been cost effective comparedto synthetic insecticides. Their narrow host rangeand relatively slow rate at which they kill insectsplace them at a disadvantage in many cropswith low damage thresholds, but they have beenused successfully in some field crops with higherthresholds. Genetically engineered viruses mayovercome some of the limitations of naturally-occurring viruses. If this happens and virusesbecome more widely and intensively used, resis-tance to them may become problematic. Giventhe history of resistance to insecticides that acthormonally or on the nervous system, includingthe sodium channel (Roush and Tabashnik, 1990;Denholm et al., 1998), resistance may evolvespecifically to some of the genes being intro-duced to the viruses (Roush, 1999).


Although there are reports of insects becomingincreasingly “resistant” to baculovirus infectionsas they age, this is not really resistance in thesense we use, but rather simply changes indevelopmental susceptibility. Still, some of thesecan be rather dramatic, such as the 34,000-folddifference from the first to the fifth instar ofMamestra brassicae (Evans, 1981). True resis-tance (as we have defined it), however, has beendeveloped through laboratory selections. In anearly summary of studies, Roberts et al. (1991)noted that of six insect species selected overseveral generations for resistance, four becameless susceptible to the virus against which theywere challenged although resistance levels were< 10-fold in two of the species and resistancewas unstable in at least one of them. Includedin this early work were high levels of resistance(140-fold) in the potato tubermoth, Phthorimaeaoperculella, when it was challenged by agranulovirus for only 6 generations (Briese andMende, 1983). Other important agricultural peststhat have developed resistance to viruses through

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laboratory selection are the cabbage looper,Trichoplusia ni (Milks and Theilmann, 2000), theIndianmeal moth, Plodia interpunctella (Bootsand Begon, 1993) and the fall armyworm,Spodoptera frugiperda (Fuxa et al., 1988). Alsoworth noting is the important work on virusresistance in the silkworm, Bombyx mori. Resis-tance of the silkworm to a cypovirus (formerlycytoplasmic polyhedrosis virus) has been clearlydemonstrated and the impact on the silk industryof this virus can be substantial (for an overviewof the early work on virus resistance in thesilkworm, see Watanabe, 1971).

Still, as noted by Fuxa (1993), therewere no documented cases of resistancethrough the early 1990s in the field to avirus employed as a microbial control agent.However, this situation changed with Anticarsiagemmatalis, a pest of soybeans. Populationscollected from Brazil were assayed with theA. gemmatalis multiple-embedded nucleopoly-hedrovirus (AgMNPV) obtained in Brazil in1979. Although the Brazilian populations did notdiffer in susceptibility (based on non-overlap ofthe LC50 values), there was a positive corre-lation with the number of years that field siteshad been sprayed with the virus (Abot et al.,1995). In a follow-up study, Abot et al. (1996)collected colonies from two sites in Brazil andone site in the USA. One of the Brazilian colonieswas initiated from a site that had been treatedfor four seasons with AgMNPV. When thecolonies were challenged for 3–4 generations, theBrazilian populations developed resistance ratiosof > 1�000 while the USA population increasedonly 5-fold. A more recent example is resis-tance to a product containing the codling moth(Cydia pomonella) granulovirus (CpGV). Whenthis product was used over several years, somepopulations in Germany and France developedresistance (Fritsch et al., 2005; Eberle and Jehle,2006; Sauphanor et al., 2006). These examplesmay predict things to come.

3 Resistance mechanism and genetics

The genetics of resistance to viruses has beenstudied in a few species. Naturally occurringresistance in Heliothis subflexa to Baculovirusheliothis appeared to be controlled by a singlemajor gene, as revealed in crosses with the

cotton pest, H. virescens (Ignoffo et al., 1985).In Spodoptera frugiperda when challengedwith a single NPV, Reichelderfer and Benton(1974) reported that “resistance was due toa single gene or genes lacking dominance.”The opposite appears to be the case withC. pomonella since resistance appears to behighly dominant (Sauphanor et al., 2006). In astudy of resistance in B. mori to viral infec-tions, Watanabe (1986) noted that completeprotection against the development of epizooticscan be ensured by rearing silkworm varietieswhich are homozygous for a recessive non-susceptibility gene. In the case of A. gemmatalis(Abot et al., 1996), the large differences inresistance evolution between the populationsfrom Brazil and the USA could have been dueto the occurrence of different genes or genefrequencies resulting from different exposurelevels since AgMNPV does not naturally occur inthe USA. In a recent study in which a laboratorypopulation of A. gemmatalis was challengedby AgMNPV, the population readily regainedsusceptibility to the virus. There were indica-tions that resistant individuals were less fit sincethey produced fewer eggs and had a lowerhatch and longer development times (Fuxa andRichter, 1998). Further tests have indicated thatoptical brighteners, when combined into diet withAgMNPV, enhanced mortality of AgMNPV-resistant insects. Although the mechanismremains unclear (Morales et al., 2001), studieshave shown optical brighteners disrupt theperitrophic membrane (Wang and Granados,2000, Okuno et al., 2003).

Studies with Drosophila have elucidated resis-tance to the sigma rhabdovirus (Wyers et al.,1995). In this case it appears that the ref(2)P gene of Drosophila melanogaster inter-feres with viral replication. This gene is highlyvariable and the different alleles are consideredpermissive or restrictive for viral replication.Similar studies with agricultural or medicalpests are needed for a clearer understanding ofhow resistance to viruses can evolve and bemanaged.

A morphological approach to understandingthe potential mechanisms of resistance wasundertaken in a study of Indianmeal moth.Larvae were inoculated with a granulovirus,dissected over time, and tissues examined with


an electron microscope (Begon et al., 1993). Fourpossible resistance mechanisms were suggested:the peritrophic membrane, which formed abarrier; the basal lamina of the hemocoel, whichimpeded the secondary spread of the infection;hemocytes, which consumed viral particles; andthe midgut cells, which passed viral particlesinto the midgut lumen in the later stages ofinfection.

C Fungi

1 Overview

Virtually all insect orders are susceptible tosome fungal disease. Worldwide there are morethan 700 species of entomopathogenic fungi inapproximately 100 genera, but only 10 specieshave been or are currently being developed forinsect control (Hajek and St. Leger, 1994). As inthe case of viruses, fungi can be introduced intoa population in several ways. Control strategiesthat have been successful include permanentintroduction and establishment, augmentativereleases, and environmental manipulation orconservation (Hajek and St. Leger, 1994).However, the majority of control programs thatrely on fungi as the principal managementtool use fungi essentially as a frequentlyapplied biological insecticide. Such mycoin-secticides have resulted in successful controlof pests in both greenhouse and field situa-tions. There is particular interest in usingfungi to control whiteflies which have becomeincreasingly problematic in recent years (Fariaand Wraight, 2001), especially in cases whereresistance to other insecticides has evolved.In China, one species of fungi, Beauveriabassiana, has been used on over 1,000,000 hafor control of the pine caterpillar (primarilyDendrolimus punctatus) and 300,000 ha of cornfor control of Ostrinia furnacalis (Roberts et al.,1991). In the USA, commercial formulationsof this fungus are registered for control ofwhiteflies, grasshoppers, thrips and aphids onseveral important crops. Entomopathogenic fungiwith enhanced efficacies may contribute in asignificant and sustainable manner to controlvector-borne diseases such as malaria, dengueand filariasis (Scholte et al., 2004).


Cases of true resistance are unknown, althoughthis may be due to the relatively limited use offungi to date or a lack of follow-up studies. Aswith bacteria and viruses, there is considerableintraspecific variability of response to fungiwhich results in differences in pathogenicity.Likewise, there is simultaneous variability in thehost. Successful laboratory selection for resis-tance does not appear to have taken place, andthere are no documented cases of resistance inthe field. However, there are instances in the fieldwhich indicate that real resistance could occurdue to genetic changes in the host. For example,aphid clones susceptible and resistant to Pandoraneoaphidis coexist in Australia (Milner, 1982,1985) and this would be due to genetic variationin host susceptibility. The increased use of fungi,coupled with improved follow-up, may detectresistance in some of the 26 locations wherethere has been permanent introduction of one of19 different host-pathogen systems (Roberts andHajek, 1992). In the case of fungi used to controlmosquito-borne diseases, no cases of resistancehave been observed but such cases may arisein the future because effective control programsrequire repeated rather than single applicationsduring mosquito breeding periods (Scholte et al.,2004).

3 Potential resistance mechanisms and genetics

Pathogenesis is a complex process involvingmany steps. A number of potential mutations ininsects along this pathway may lead to changesin susceptibility. For example, the cuticle servesas multi-layered barrier to fungi. The failure of afungus to penetrate this barrier could be causedby the presence of inhibitory compounds in thecuticle, the lack of factors needed for recog-nition on the cuticle, or lack of proper nutrientsin the cuticle. Once the fungus has entered theinsect’s body, the host may be killed by somecombination of mechanical damage caused bythe fungus, depletion of the host’s nutrients ortoxins produced by the fungus. Again, changesin the insect could alter the insect-pathogen inter-action at any one of these steps and result inresistance.

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D Microsporidia

1 Overview

Microsporidia are now placed with the fungi.They are transmitted orally by ingestion ofspores and some are also transmitted transovar-ially via the eggs or by parasitoids (Robertset al., 1991). Infection by microsporidiausually results only in somewhat reduced pestpopulation densities through lowered fecundityand other such sublethal effects, so theirplace in pest management has been limited.However, two species that might have potentialuse as microbial insecticides are Paranosema(=Nosema) locustae and Varimorpha necatrix.The former is the only microsporidia regis-tered by the EPA and has been used in a baitfor grasshopper control in situations such asrangeland where the economic threshold is high.Due to their limitations in mass production andefficacy, Roberts et al. (1991) suggested that itmight be more realistic to consider microsporidiaas the “microbial counterparts to parasitoids andpredators rather than as candidates for devel-opment as microbial insecticides.”


A careful search of the literature has found nocases of resistance to microsporidia.

3 Potential resistance mechanisms and genetics

Like fungi, infection by microsporidia appearsto be a complex process of interactions betweenthe pathogen and its host. Too little is knownabout this process, and much less is knownabout genetic plasticity to provide insight intothe development of resistance to particularmicrosporidia.

E Nematodes

1 Overview

Although nearly 40 nematode families areassociated with insects, species in three families

(Mermithidae, Steinernematidae and Heterorhab-ditidae) are the predominant nematodes used forinsect control. Steinernematidae and Heterorhab-ditidae are characterized by their mutualisticrelationship with bacterial pathogens in thegenera Xenorhabdus and Photorhabdus, respec-tively (see Chapter IV-5), whereas mermithidsare more like insect parasitoids. In the caseof steinernematids and heterorhabditids, theinfective nematodes enter host insects and releasethe bacterium that multiplies, causes septicemia,and kills the insect within 48 hours. The primarysuccess of these two nematode families asbiological control agents has been in sites wherethe nematodes have been sheltered from environ-mental extremes. The introduction of nematodesinto plant stems or soil environments where theycan then use volatiles from the insect to locatetheir hosts has proven to be successful for suchpests as the black vine weevil, citrus weevil, molecrickets and Japanese beetle. For mermithidstheir most effective use has been against aquaticinsects, especially mosquito larvae.


As with other pathogens, host susceptibilityvaries with different nematode species andstrains. Although not true resistance, hostsusceptibility to nematodes may also changewith age of the host. In the case ofSimulium vittatum infection by Steinernema(=Neoaplectana) carpocapsae, the basis for‘resistance’ was the physical exclusion of thecomparatively large nematodes during the earlyinstars of the host (Gaugler and Molloy, 1981).Because many soil insects have co-evolvedwith steinernematids and heterorhabditids, theyare often ‘resistant’ to these nematodes. Forexample, scarab grubs have sieve plates overtheir spiracles and elaterid larvae have an oralfilter that prevents nematodes from enteringand infecting the larvae (see Chapter IV-5).In a series of studies with nematodes againstJapanese beetle larvae, the infection process bythe nematodes and the defensive mechanisms bythe beetle have been studied in detail. In thecase of Steinernema glaseri and Heterorhabditisbacteriophora, each used different strategies forkilling the larvae but caused similar mortality.S. glaseri tolerated the gut fluid and avoided


the host immune system, while H. bacteriophorahad poor tolerance to the host gut fluid butovercame this by releasing its bacteria duringor soon after penetrating its host (Wang et al.,1995). In a later study Wang and Gaugler (1999)found evidence of a surface coat protein fromS. glaseri that suppresses the immune responseof beetle larvae. In the case of the beetle’sdefensive mechanisms, it displayed aggressivebehaviors such as brushing with the legs andrubbing with its raster to remove attackingnematodes or simply make evasive behaviors inthe presence of nematodes (Gaugler et al., 1994).While these studies do not document cases ofresistance due to selection pressure over time,they do suggest potential mechanisms of resis-tance. However, the mosquito, Culex quinquefas-ciatus, developed bona fide behavioral resistanceto the mermithid, Romanomermis culicivorax,under laboratory selection after 300 genera-tions (Petersen, 1978) and Anopheles quadri-maculatus developed resistance to Diximermispeterseni after 4 years of exposure (Woodwardand Fukuda, 1977). No cases of resistance havebeen reported in the field.

3 Resistance mechanisms and genetics

As with fungi and microsporidia, the complexinteractions between nematodes and their hostsinvolve multiple steps, any one of which may bealtered by changes in the genome of the host.A prime limitation of nematodes has been theirsusceptibility to environmental factors and hencelack of persistence. Selection and/or developmentof strains that have increased persistence maylead to increased use and perhaps potential resis-tance problems. Insect immunity to nematodesmight result from encapsulation, intracellularmelanization, changes in behavior, or resistanceto the bacterium Xenorhabdus. Perhaps thesefactors could also occur through genetic changesin previously susceptible insects and therebycreate truly resistant populations. In the case ofA. quadrimaculatus, resistance appears to haveoccurred because the resistant host was muchmore active and attempted to remove attachedpreparasites. The resistance mechanism ofC. p. quinquefasciatus is not known.

3 Characteristics of pathogensinfluencing resistance

The most spectacular documented case of resis-tance in the field to any of the pathogens listedabove is with P. xylostella and Bt. We speculatethat this is due not to some inherent charac-teristic of this system, but rather to the heavyuse of this bacterially produced pesticide againstthis specific pest. In this case, use was increasedbecause other control tactics had failed (Sheltonet al., 1993). In the case of field resistance inA. gemmatalis to AgMNPV in Brazil, resistanceoccurred only after several years of wide-spreaduse, but such use may have been due to economicand environmental reasons rather than to thefailure of other tactics.

However, one can speculate that the morecomplex the infection process, the more likelythat the host population may carry rare resistancealleles to combat a critical step in the process. Forexample, when conidia of Metarhizium aniso-pliae attach to a host, they produce appressoriaon the cuticle, infection pegs in the epicuticle,hyphae that penetrate plates in the procuticle andyeast-like hyphal bodies within the hemocoel,which eventually lead to the death of the insect(Hajek and St. Leger, 1994). At various stages,genetic variants could physiologically challengethe fungal invasion.

Perhaps, the factors which would mostinfluence the development of resistance areapplication, use pattern and formulation,especially in terms of persistence of exposureof the control agent. As noted above, in Floridause of Bt increased when synthetic productsfailed, and we estimate that high levels of resis-tance to Btk in P. xylostella occurred withina 4 year period (Shelton et al., 1993). Themore a microbial insecticide is used, the morelikely resistance to it will evolve. In the caseof Bt toxins that are engineered into plants andthereby significantly increase persistence, it isespecially appropriate that resistant managementstrategies be developed and utilized when theplants are first deployed (Chapter VIII-1). Projec-tions indicate that Bt transgenic plants have thepotential to capture one-third of the insecticidemarket (Krattiger, 1997). This will likely lead to

802 Shelton et al.

further selection for Bt resistance to foliar appli-cations as well as when expressed in plants, iffor no other reason than that Bt toxins will findgreatly increased use.

Although plants which have been engineeredto express a microbial toxin effectively make itmore persistent and may increase the likelihoodfor resistance, transgenic plants may also producesuch a consistently high dose as to delay resis-tance more effectively than sprays in the presenceof a “refuge” of untreated individuals (Roush,1994). Companies have produced plants whichexpress Bt toxins and utilized a high dose strategywhich calls for high expression of the toxin(> LC95 of heterozygous RS insects) in combi-nation with a refuge of non-transformed plant(Chapter VIII-1). The benefits of utilizing arefuge have been proved in greenhouse studies(Shelton et al., 1998, Tang et al., 2001). Infield studies with P. xylostella when appliedas a spray, development of resistance to Btkhas been slower with lower rates, but thisalso resulted in increased damage to the crop(Perez et al., 1997b). This indicates the need tobalance rate with efficacy in an overall resis-tance management program. Stability of resis-tance can also be influenced by another aspectof formulation besides application. In the caseof Bt, the two major strains that have been usedagainst P. xylostella are Bta and Btk. While theyshare some of the same toxins, e.g., Cry1A,Bta has an additional toxin, Cry1C, which hasa different receptor site and consequently cancontrol insects that have developed resistanceonly to Btk. However, using a Bta productagainst a population of P. xylostella can maintainor increase resistance to Cry1A, which mightotherwise have declined if a pure Cry 1C productwas used (Tang et al., 1995). In contrast tosprays, however, models suggest that pyramidingtwo toxins into the same cultivar may greatlydelay resistance (Roush, 1994, 1997, 1998),and laboratory selection tests on H. armigera(Zhao et al., 1999) and greenhouse selectionexperiments on P. xylostella (Zhao et al., 2003)have provided direct evidence. Thus, one needsto be concerned not only about the method ofdelivering Bt, whether it be as a spray or incorpo-rated into plants, but also about the concentrationand the composition of the toxins.

4 Detecting resistance

As noted previously, not all insects withina population are equally susceptible to allpathogens and, for those that are similarlysusceptible, there may be large differencesin susceptibility depending on the stage. Itis within this framework that entomologistsmust use techniques that detect true resistance.Tremendous improvements have been made ininsecticide resistance detection methods (ffrench-Constant and Roush, 1990) and our currentunderstanding of biochemistry and ecologicalgenetics of resistance to chemical insecticides(Roush and Tabashnik, 1990; Denholm et al.,1998). Outside of P. xylostella and Bt toxinsand the ref (2)P gene of D. melanogaster whichinterferes with virus replication, however, wehave little knowledge of fundamental aspects thatcan be used to detect resistance of pathogens ortheir products and must rely more on standard-izing methods for monitoring resistance throughdose-response and, more practically, diagnosticdose methods (for a more thorough treatment seeffrench-Constant and Roush, 1990; Halliday andBurnham, 1990; Chapter VIII-1).

Testing of microbials should be done ina fashion similar to how the insect wouldbecome exposed in the field to the pathogen.The leaf-dip bioassay technique has been usedmost frequently to assess P. xylostella resis-tance to Bt (Tabashnik et al., 1990; Sheltonet al., 1993) and the LC50 and the corre-sponding 95% CL have also been the mostfrequent criterion used to compare the suscepti-bility of two or more populations. Field-collectedlarvae can be brought back to the laboratory,but it is usually necessary to rear them forone or more generations so that enough insectsare available for testing. Perez et al. (1997a)compared P. xylostella in leaf-dip bioassays withthe mortality of larvae caused by residues of fieldapplications of Btk and mortality of larvae inthree diagnostic concentrations of a Btk productincorporated in an artificial diet. They concludedthat LC50s of a B. thuringiensis product (Javelin)> 0�6 mg (AI)/liter in leaf dip bioassays can beassociated with low levels of mortality in fieldapplications and that the diagnostic concentrationof 20.5 mg/ml artificial diet can be used as an on-farm assay for monitoring P. xylostella resistance


to Btk. In those cases where toxins of Bt havebeen incorporated into a plant, one can use theplant as an assay substrate and simply put thelarvae on the plant (Chapter VIII-1).

Realistically, the sample sizes needed to reliablydetect resistance across a wide geographic areaare so large that resistance management must beseen as a preventative rather than reactive activity.Even with excellent diagnostic doses, one needs asamplesizeofat least3000individualsper locationto detect resistance phenotypes even when theyare at a frequency of 0.1% (Roush and Miller,1986), a frequencysohigh that failuresdue to resis-tance may occur in just a few more generations(Roush, 1994). Even reliance on a sample of 3000is optimistic, because perfect diagnostic dosesrarely exist given the inconsistencies and varia-tions typical in bioassays (Halliday and Burnham,1990). Even if the diagnostic dose is coupledto the F2 screen as discussed by Caprio et al.(Chapter VIII-1), the labor requirements wouldbe enormous. It was proved that the diagnosticdiet assay was a better F2 screen method todetect alleles in P. xylostella than using transgenicplants expressing a high level of a Bt toxin (Zhaoet al., 2002). The estimated probabilities of falsepositives and false negatives were 33% and 1%,respectively, for detecting Cry1Ac resistance atthe allele frequency of 0.012 using the diagnosticdiet assay. Careful validation of the screeningmethod for each insect-crop system is necessarybefore the F2 screen can be used to detect rare Btresistance alleles in field populations. Given thatresistance can often evolve on very local scales(e.g., Tabashnik et al., 1987; Roush et al., 1990),many locations would have to be monitored toassure the detection of resistance before it becamewidespread.

A DNA-based method was recently developedthat could detect the mutation in genomic DNA ofeach of the three Bt resistant alleles (r1, r2, r3) of acadheringene in P.gossypiella (Morinetal., 2004)

5 Management of resistance

A Characteristics of insects influencingresistance

The proportion of a population that is exposedto a selection agent is among the most important

factors influencing the evolution of resistance,particularly when coupled with long rangedispersal to ensure mating between selected andunselected individuals. In many comparisons ofsimilar or closely related pests (even with thesame species in different habitats), resistanceevolves more quickly where a high proportionof the population is exposed each generation(Tabashnik and Croft, 1985; Roush and Daly,1990), i.e., when there are only small “refuges”of untreated individuals. In the presence ofrefuges, larger scale insect migrations furtherassist the delay of resistance by allowing aninflux of potentially susceptible alleles intoan area where selection pressure is occurring,and thereby help dilute resistant alleles. InBrazil, where one million ha of soybeans weretreated annually with AgMNPV, this representedonly 10% of the total soybean acreage. Fuxa(1993) suggests quite sensibly that one factorpreventing wide-spread development of resis-tance is the migration of adults from soybeanareas not treated with AgMNPV to areas wherethe pathogen has been extensively used.

In contrast, many other aspects of pest biologythat are commonly thought to influence the rateof resistance evolution have much less clear orconsistent influences. A large number of gener-ations per year do not consistently accelerateresistance (Rosenheim and Tabashnik, 1990,1991); some species with few annual genera-tions, such as the Colorado potato beetle (Roushet al., 1990; Zhao et al., 2000), evolve resis-tance very quickly, apparently due to intenseselection within each generation. The resistancegenes themselves can have a strong influenceon the rate of evolution. Resistance will evolvefaster with dominant alleles (i.e., where theheterozygous carriers of one resistance alleleand one susceptible allele survive) (Tabashnikand Croft, 1982; Roush, 1994, 1997, 1998).However, as noted above, fitness costs to resis-tance do not seem to have a significant effecton slowing resistance unless the costs arelarge and affect heterozygotes (Roush, 1997),which seems generally not to be the case(Roush and Daly, 1990). However, an alter-native view about the value of fitness costsin slowing resistance evolution was suggestedin P. gossypiella (Carrieré et al., 2001a, b;Tabashnik et al., 2005).

804 Shelton et al.

B Promote refuges: apply agent less oftenand only on “hot spots”

Given that one of the most obvious and consistentinfluences on the rate at which resistance evolvesis the proportion of the population in “refuges”each generation, it seems clear that a key tacticfor the management of resistance is to providerefuges where possible. The best way to increaserefuges for pests with a history of resistanceevolution would be to reduce the percentageof the pest population that is sprayed withinthe crop.

While this advice may at first seem imprac-tical, it is completely consistent with the long-term thrust of IPM: reduce the frequency ofsprays or portion of the crop covered, atleast where sprays fail to provide an economicreturn (i.e., pests exceed economic or actionthreshold densities). In particular, there aremany examples in which “spot treatment” isoften sufficient to control pests without treatingthe entire population. For example, Coloradopotato beetles are often concentrated along fieldmargins (Roush and Tingey, 1992). Those areasinternal to the field serve as refuges for insectsthat are not sufficiently dense to cause loss butwhich can dilute resistance.

C Avoid persistent formulations

Closely related to the notion of spraying lessoften is to avoid highly persistent formula-tions (Roush, 1989, 1999). Users want enoughpersistence to control the pest, but excessivepersistence can continue to select for resis-tance long after the pest has been suppressedbelow damaging numbers, just as would calendarspraying. Most biopesticides probably will haveshort persistence, which would tend to avoidresistance, but it may be technically feasible touse them in slow release devices. Such deliverysystems may be counterproductive in the longterm by creating persistent selection.

D Avoid high application rates

Contrary to popular belief, there is no generaladvantage to applying high rates in sprays, eitherin theory or experiments (Tabashnik and Croft,1982; Roush 1989, 1998). There may be an

advantage for transgenic crops (Roush, 1994,1997, 1998) or closed systems like granaries(Roush, 1989) where there can be tight controlof the application rate even if the exposure ispersistent, but sprays will inherently suffer fromless than thorough coverage and decay, such thatthe 95% mortality of heterozygotes needed for a“high dose strategy” (Chapter VIII-1) is simplynot possible (Tabashnik and Croft, 1982; Roush,1994, 1998). Further, excessively good controlcan eliminate prey for natural enemies such thatthey starve even if not killed by the spray itself,which has the potential for inducing a pesticidetreadmill that increases dependence on the spraysand faster resistance. On the other hand, thereis also no clear evidence or theory that a lowerrate of application will in itself significantlyslow resistance, at least not within the range ofrates that will still control the pests, althoughlow rates may be more easily integrated withthe survival and use of predators or parasitoids(Roush, 1989, 1994).

E Avoid pesticide mixtures

Contrary to another popular belief, it is not neces-sarily true that mixtures of pesticides will delayresistance. Experimental studies have failed toconsistently find any advantage to mixtures oversequential or rotational use of the same insecti-cides (Tabashnik, 1989; Immaraju et al., 1990),and theoretical models showed that mixtures willsignificantly delay resistance only when severalconditions are met (Gould, 1986; Roush, 1989,1997, 1998). To be most effective, mixturesmust meet several requirements, especially thatthere must be high mortality from each of theagents when they are sprayed alone. Almost allindividuals resistant and exposed to one pesticidemust be killed by the other for mixtures to behighly effective (Roush, 1989, 1998).

These conditions will probably be rarely metfor sprays of microbial agents. For example,experiments with mixtures of Bt serotypes,applied at concentrations that did not providehigh levels of control when used individually,failed to delay resistance in Indianmeal moth(McGaughey and Johnson, 1992). Not only isthis experiment a good model for the field (wherecontrol with microbials probably rarely exceeds90%), the results are just as would be predicted


from the modeling papers listed above. We havepreviously mentioned problems of dual selectionwith mixtures of Cry 1A and Cry 1C toxins asfound in Bta.

F Use rotations rather than mosaics

Rotating the use of pesticides over an entirearea in a so-called “window strategy” based ona seasonal calendar has proven to be a veryeffective resistance management tactic both interms of adoption and efficacy (Roush, 1989;Forrester et al., 1993). The use of differentcompounds at roughly the same time in neigh-boring fields creates a mosaic of treatmentpatterns and should be avoided, as should veryshort-term rotations within a generation (Roush,1989). The problem for both is that you havesimultaneous selection with several pesticides,resulting in much lower pesticide durability (asillustrated by Roush, 1993).

To make sense of this, consider a case whereselection for resistance is so strong that resis-tance occurs in a single generation. If you havetwo pesticides and use the first pesticide, youstill have the second pesticide for the nextgeneration, giving a total of two generationsof control. If instead, you split the populationin half, roughly half of the population will beresistant to each of the pesticides in the nextgeneration (depending on what assumptions youmake about the dominance of resistance), whichwould most likely result in failure to controlthe population in the field after one generation.Long term rotations, over at least a month inlength and in some cases for 2 years, have provedto be extremely successful for cotton insecti-cides and acaricides in Australia and Zimbabwe(Roush, 1989; Forrester et al., 1993). Recentlywe evaluated the development of resistance inP. xylostella to three insecticides (Bt, spinosad,and indoxacarb) when these products are usedin a rotation strategy compared to when theproducts are used in a mosaic fashion in whichall products are used simultaneously in green-house cage experiments. After 9 generations ofselection, the overall results showed that rotationof each insecticide was better than a mosaic ofall three insecticides for resistance management(Zhao et al., unpublished data).

6 Conclusions

It has been the general experience of entomol-ogists that resistance will evolve to any controlagent that is intensively applied on populationswith small refuges. Even if we exclude Bt asa unique case, resistance will almost certainlyoccur to at least some insect pathogens, as hasalready been found for viruses and, to a lesserextent, nematodes. Determining whether or notresistance has historically been a problem for thepest in question is a simple test of the need for aresistance management strategy.

Although resistance management is oftenperceived as a complex problem, the list ofpotential practical tactics is generally so short thatchoosing which would be useful is not difficultwith a minimal amount of information about pestbiology (Roush, 1989). Most of these tactics arealso complementary and are most effective whenadopted before selection commences. Thus, eventhough resistance management can always beimproved later with additional data, one shouldaim to adopt a resistance management plan at thefirst introduction of the product to keep resistancefrequencies as low as is reasonably practical.

The most common problem in resistancemanagement is not one of assembling the obvioustactics, but of gaining implementation of thestrategy. The history of resistance managementimplies quite strongly that for problematic pests,it is at least in the interests of the pesticide userand the local community to find a path to imple-mentation.

7 Acknowledgments

The authors thank H. Collins, J. Curtis,R. Gaugler, R. Granados, J. Grant, H. Kaya,R. Milner and M. Villani for their help on thischapter.

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