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Immunity of an Alternative Host Can Be Overcome by Higher Densities of Its Parasitoids Palmistichus elaeisis and Trichospilus diatraeae

Immunity of an Alternative Host Can Be Overcome by Higher Densities of Its Parasitoids Palmistichus elaeisis and Trichospilus diatraeae

  • Gilberto Santos Andrade, 
  • José Eduardo Serrão, 
  • José Cola Zanuncio, 
  • Teresinha Vinha Zanuncio, 
  • Germano Leão Demolin Leite, 
  • Ricardo Antonio Polanczyk
PLOS
x
  • Published: October 13, 2010
  • DOI: 10.1371/journal.pone.0013231

Abstract

Interactions of the parasitoids Palmistichus elaeisis Delvare & LaSalle and Trichospilus diatraeae Cherian & Margabandhu (Hymenoptera: Eulophidae) with its alternative host Anticarsia gemmatalis (Hübner) (Lepidoptera: Noctuidae) affect the success or failure of the mass production of these parasitoids for use in integrated pest management programs. The aim of this study was to evaluate changes in the cellular defense and encapsulation ability of A. gemmatalis pupae against P. elaeisis or T. diatraeae in adult parasitoid densities of 1, 3, 5, 7, 9, 11 or 13 parasitoids/pupae. We evaluated the total quantity of circulating hemocytes and the encapsulation rate versus density. Increasing parasitoid density reduced the total number of hemocytes in the hemolymph and the encapsulation rate by parasitized pupae. Furthermore, densities of P. elaeisis above 5 parasitoids/pupae caused higher reduction in total hemocyte numbers. The encapsulation rate fell with increasing parasitoid density. However, parasitic invasion by both species induced generally similar responses. The reduction in defensive capacity of A. gemmatalis is related to the adjustment of the density of these parasitoids to their development in this host. Thus, the role of the density of P. elaeisis or T. diatraeae by pupa is induced suppression of cellular defense and encapsulation of the host, even without them possesses a co-evolutionary history. Furthermore, these findings can predict the success of P. elaeisis and T. diatraeae in the control of insect pests through the use of immunology as a tool for evaluation of natural enemies.

Introduction

Parasitism, and the development of the parasitoid lifestyle, depends on the host being able to meet the nutritional requirements of the parasite and the ability of the parasite to overcome the immune response of the host [1], [2]. Moreover, genetic factors and the host's innate ability to respond to the parasite invasion determine the degree of the host's resistance to parasitism [1], [3][4]. Therefore, parasites rely on their ability to suppress the cellular and humoral defenses of their hosts [5].

Hemocytes are the main defense cells of insects and originate during embryonic development, while maintenance and differentiation of prohemocytes continue the production and circulation of these cells in the hemolymph in adult insects [2]. Membrane receptors in hemocytes recognize invaders and promote subsequent protective cellular reactions including phagocytosis, nodulation or encapsulation [6]. The most common types of insect hemocyte are prohemocytes, plasmatocytes, granulocytes and oenocytoids, although variants can be recognized in different insect species [7, 8–6].

Proteins and peptides also recognize pathogens and then adhere to, and alter, the molecular properties of the invader's cell membrane or cell wall [9]. They then produce proteolytic and toxic molecules that kill the invading parasitoids, bacteria or fungus. Melanin is the final product of these prophenoloxidase cascade reactions, leading to the death of the pathogens [10]. This melanogenesis generates nitrogen and oxygen reactive species that damage the structure of proteins and DNA, and are fatal to pathogens. If invading organisms are to live and develop successfully within a host they must overcome these host defenses [1], [11], [12], [13], [14].

The use of monofilament nylon or micro-injections of Sephadex beads are two methods for estimating the encapsulation of insects in vivo [15], because the capacity to encapsulate abiotic material is related to the ability to encapsulate by non-self recognition [16].The most appropriate method for bioassays in pupae has been the use of implants of nylon monofilament, because microinjections of liquid into pupa with Sephadex microspheres could damage the pupal beg [15].

Parasitoids such as Eulophidae (Hymenoptera) can be used to regulate populations of agricultural insect pests [17], [18]. Effective use of these natural enemies in integrated pest management programs relies on good basic knowledge of the parasitoid's biology [19], [20], because the mass production of parasitoids for release depends on our being able to produce them in quantity within suitable hosts [21].

Palmistichus elaeisis Delvare & LaSalle 1993 (Hymenoptera: Eulophidae) parasitize the pupae of Bombycidae, Noctuidae, Arctiidae and Tenebrionidae [22]. Trichospilus diatraeae Cherian & Margabandhu 1942 (Hymenoptera: Eulophidae) parasitize the pupae of Crambidae and Noctuidae [23], [24]. Both parasitoids have been studied with a view to using them to control pests of agricultural crops and forests [22], [24][28]. On the other hand, the reproductive success of these parasitoids has been dependent on varying numbers of them according to host species [22], [23], [24]. One possible explanation is that the lack of co-evolutionary relationship between alternative hosts and these parasitoids hinder the suppression of host immune defense, causing failures in the mass production or even possible failures in the use of these natural enemies to control insects pests.

Understanding the host immune response allows us to predict the success of individual parasitoid species in alternative hosts. The objective of this study was to evaluate the cellular defense of Anticarsia gemmatalis pupae (Hübner, 1818) (Lepidoptera: Noctuidae) and their ability to encapsulate invaders when exposed to different densities of the parasitoids P. elaeisis and T. diatraeae, so as to choose a suitable alternative host for mass rearing of these natural enemies.

Materials and Methods

Host

Eggs of A. gemmatalis were placed in 1100 mL plastic vials and after hatching the larvae were fed on an artificial diet [29] at 25±1°C, 70±10% relative humidity, and 14 h photophase. At the end of fifth instar, larvae were transferred to 1100 mL pots, 1/5 filled with sand previously sterilized at 150°C for two hours, to allow pupation. The pupae were transferred to wooden boxes (30×30×30 cm) supplied with nutrient solution (10.5 g of honey, 60 g of sucrose, 1.05 g of nipagin and 1.05 g of ascorbic acid diluted in 1.05 L of distilled water) embedded in a cotton ball for feeding of the hatched adults. Eggs were collected on white paper sheets placed inside the wooden boxes and transferred to vials provided with the artificial diet.

Parasitoids

Palmistichus elaeisis and T. diatraeae were obtained from the Laboratory of Biological Control of Insects at the Universidade Federal de Viçosa, Minas Gerais and kept at 25±1°C, 70±10% relative humidity, and f 14 h photophase. Six females of P. elaeisis, 72 h after emergence and eight newly emerged T. diatraeae females were each presented with one pupa of A. gemmatalis for 24 h in glass tubes (14×2.2 cm) containing drops of honey for the parasitoids to feed on. The number and age of parasitoid females chosen were those with the best parasitism rates [24, unpublished data].

Hemocyte count in hosts

Pupae of A. gemmatalis (229.72±5.12 mg and 24 h old) were exposed to 1, 3, 5, 7, 9, 11 or 13 adult parasitoids to test the effect of parasitoid densities on the host immune response. The pupae were exposed to 72 h old, mated females of P. elaeisis [26] or newly emerged mated T. diatraeae females. At these ages the parasitoids have mature eggs suitable for oviposition (data not shown). After this period, the pupae were rinsed with 1% sodium hypochlorite for five seconds and then with distilled water. Four microliters samples of pupal hemolymph were collected with micropipettes from a small incision in the thorax and transferred to 20 µL of buffer (98 mM NaOH, 186 mM NaCl, 17 mM Na2 EDTA and 41 mM citric acid, pH 4.5) to prevent the hemocytes aggregation [30].

The 4 µL haemolymph samples were stained with Giemsa and the total numbers of hemocytes, granulocytes, plasmatocytes and other hemocytes types were counted [10], [6][30] using a hemocytometer (Neubauer) with a 40 x objective lens.

Encapsulation Rate

Nylon filaments (2×0.2 mm) were sterilized with 1% sodium hypochlorite, washed with distilled water and implanted into pupae of A. gemmatalis [29]. These pupae were individually placed in glass tubes (14.0×2.2 cm) and exposed to 1, 3, 5, 7, 9, 11 or 13 mated females of P. elaeisis or T. diatraeae for 24 h. Pupae not exposed to parasitoids were used as a control. The filament implants remained inserted into the pupae for 48 h and were then removed, mounted on slides and observed under a light microscope [15].

Images of the nylon implants were made with a photographic camera, Canon PowerShot™ A640, and processed using the computer program RemoteCapture Task ™ with the following settings: white balance (day light); exposure compensation (+1); flash exposure level (zero); metering mode (evaluative); ISO speed (auto); AE Mode (Program AE). The spectral signature of the implants was measured using ImageJ software,National Institutes of Health, USA [31]. The mean absorbance value was adopted as a measure of the rate of encapsulation with values from zero to 255. The mean absorbance of the samples was adjusted by subtracting it from 255, owing to the fact that the computer program indicates the highest encapsulation rate as zero and the lowest one as 255 [31], [32]. The arbitrary values of the implants were adjusted by discounting the background [31].

Statistical analysis

The comparison between parasitoids were conducted using the nonparametric Wilcoxon test (p≤0.05). The hemocyte values at different densities of each parasitoid were compared using regression analysis with computer program SigmaPlot 10.0 (p≤0.05) with 15 repetitions. The dimensionless values for the rate of encapsulation were compared by regression analysis with 20 repetitions (p≤0.05).

Results

Increasing the density of adult P. elaesis reduced the total number of hemocytes in the hemolymph of A. gemmatalis pupae with 117.13×104 and 45.85×104 cells mL−1, at the lowest and highest densities, respectively (F = 68.6945, P≤0.01) (Figure 1). Parasitism by T. diatraeae also reduced the total number of hemocytes in the hemolymph of A. gemmatalis with 116.3×104 cells mL−1 and 72.8×104 cells mL−1 at lowest and highest parasitoid densities, respectively (F = 49.2905, P = 0.0004). Furthermore, the reduction in total circulating hemocytes was greater under parasitism by P. elaeisis in the densities of 5, 7, 9, 11 and 13 (p≤0.05) (Figure 1).

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Figure 1. Hemocyte (mean ± se) of Anticarsia gemmatalis pupae parasitized by densities of Palmistichus elaeisis or Trichospilus diatraeae.

doi:10.1371/journal.pone.0013231.g001

The number of circulating granulocytes in A. gemmatalis pupae decreased linearly as a function of increasing density of P. elaeisis, from 63.35×104 to 11.32×104 cells mL−1 (F = 25.2121, P≤0.01) and similarly for T. diatraeae, from 62.77×104 to 27.65×104 cells mL−1 (F = 55.1502, P≤0.01) from the lowest to the highest densities of parasitoids, respectively (Figure 2). Palmistichus elaeisis showed a greater ability to reduce circulating granulocytes in the hemolymph of A. gemmatalis at densities higher than 5 parasitoids/pupae than T. diatraeae (p≤0.05).

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Figure 2. Total granulocyte (mean ± se) of Anticarsia gemmatalis pupae parasitized by densities of Palmistichus elaeisis or Trichospilus diatraeae.

doi:10.1371/journal.pone.0013231.g002

The number of circulating plasmatocytes was lower with increasing densities of P. elaeisis (F = 63.3011, P≤0.01) and T. diatraeae (F = 19.6201, P≤0.01) (Figure 3). Palmistichus elaeisis showed greater ability to reduce circulating plasmatocytes in the hemolymph of A. gemmatalis at densities of 5, 9 and 13 parasitoids/pupae and a similar reduction at densities below 5 females/pupae. Moreover, T. diatraeae showed a linear reduction in the number of circulating plasmatocytes in the hemolymph of A. gemmatalis with increasing parasite density (Figure 3).

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Figure 3. Total plasmatocytes (mean ± se) of Anticarsia gemmatalis pupaeparasitized by densities of Palmistichus elaeisis or Trichospilus diatraeae.

doi:10.1371/journal.pone.0013231.g003

There is an irregular pattern in the reduction of other types of circulating hemocytes in the hemolymph of A. gemmatalis with increases in parasitoid density (Figure 4). The number of these cells was higher when parasitized by P. elaeisis than T. diatraeae at densities of 5 and 7 parasitoids/pupae, respectively.

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Figure 4. Total of other cell types (mean ± se) of Anticarsia gemmatalis pupae parasitized by densities of Palmistichus elaeisis or Trichospilus diatraeae.

doi:10.1371/journal.pone.0013231.g004

The encapsulation rate was demonstrated by hemocyte adherence to the nylon filaments, followed by darkening of the filaments. Increasing parasitoid density decreased the melanization rate from 65.24±4.61 in non-parasitized pupae to 44.44±3.82 and 42.22±5.10 at higher densities of P. elaeisis and T. diatraeae, respectively (Figure 5). At 7 parasitoids/pupa, a higher encapsulation rate occurred in pupae parasitized by P. elaeisis than by T. diatraeae (Figure 5).

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Figure 5. Encapsulation rate pupae of Anticarsia gemmatalis pupae parasitized by densities of Palmistichus elaeisis or Trichospilus diatraeae.

doi:10.1371/journal.pone.0013231.g005

Discussion

Changes in total hemocytes circulating in the haemolymph, and encapsulation rates of parasitized A.gemmatalis pupae are critical steps in the suppression of the host defenses by P. elaeisis and T. diatraeae, allow them to develop immature parasitoids. These features are also likely to be important in the parasitism of pupae of the pest Acrolepiopsis assectella (Zeller, 1839) (Lepidoptera: Yponomeutoidae) [15].

The greater reduction in the number of granulocytes in pupae attacked by five or more P. elaeisis suggests that host defenses may vary according to the parasitoid species involved. However, since we found similar encapsulation rates for both parasitoids, we suggest that there is link between the number of hemocytes and the humoral response [33], [34].

Changes in the total circulating hemocyte numbers in A. gemmatalis pupae parasitized by P. elaeisis and T. diatraeae are similar to those reported in larvae of Putella xylostella L. (Lepidoptera: Plutellidae) parasitized by Cotesia plutellae (Kurdjumov) (Hymenoptera: Braconidae) (Ibrahim and Kim, 2006). The polyphagous larval endoparasitoid Meteorus pulchricornis (Wesmael, 1835) (Hymenoptera: Braconidae) also induces changes in the numbers of hemocytes of its host [35] indicating that this is a common pathway for host immune suppression in parasitoids. The means by which parasitoids alter the number of circulating hemocytes in the hemolymph has been discussed. Braconidae inject venom associated with teratocytes [35], while Eulophidae inject venom during oviposition [36]. The reduction in the number of circulating hemocytes ensures a favorable environment for the development of the parasitoid larvae and prevents the host producing prophenloxidase, oxygen and nitrogen intermediate reactive species, and melanin [37], [38].

Detailed differences in the suppression of the immune response, and changes in the circulating hemocytes of A. gemmatalis by P. elaeisis and T. diatraeae, suggest that these parasitoids may use different strategies to suppress host immunity, such as through the death of hemocytes or modification of their adhesion properties [1, 30, 39. 40, 41]. These differences may result from toxic substances in ovarian fluids, rich in proteins that induce changes in hemocytes, released by the parasitoids during oviposition [42]. Increases in the concentration of fluids injected during oviposition may enhance the physiological changes in the host hemocytes [43], which may explain the decrease in circulating hemocytes and encapsulation rates in parasitized A. gemmatalis pupa.

The similar number of other types of hemocytes circulating in the hemolymph with increasing density of parasitoids may be because these cells play a role in tissue and organ restructuring during the pupal stage, unlike plasmatocytes and granulocytes [44].

The lower melanization of nylon implants in A. gemmatalis pupae with the highest densities of parasitoids suggests an effect of parasitoid densities on the population of circulating hemocytes, which participate in the initial process of pathogen recognition and triggering of humoral defenses. Higher densities allow more parasitoid eggs to develop by lowering the ability of the host to encapsulate them [43]. Oviposition of greater numbers of eggs in the same host can counteract a high mortality rate of immature stages of the parasitoid by the host immune system, and result in greater reproductive success of gregarious parasitoids compared to solitary ones [45]. The reduction of humoral and cellular defenses of parasitized A. gemmatalis pupae indicates the importance of the gregarious habit for the reproductive success of P. elaeisis and T. diatraeae. This is demonstrated by the increase in offspring of Plutella xylostella (L., 1758) (Lepidoptera: Plutellidae) super-parasitised by Oomyzus sokolowskii (Kurdjumov, 1912) (Hymenoptera: Braconidae) [46]. In this case the host immune system is the main factor causing parasitoid mortality in the immature phase, and the degree of encapsulation has strong implications for the reproductive success of its natural enemies [7][10], [37][47]. Moreover, increase in competition for the food resources provided by the host resulting from super-parasitism may reduce the quality of the parasitoids, and shows the importance of the appropriate density of parasitoids per host individual [48], [49].

Females of the parasitoids P. elaeisis and T. diatraeae reduce the immune response of the alternative host, A. gemmatalis, by reducing the number of circulating hemocytes in the host by increasing the densities of the attacking parasitoids.

Author Contributions

Conceived and designed the experiments: GSA JES JCZ. Performed the experiments: GSA. Analyzed the data: GSA TVZ GLDL. Contributed reagents/materials/analysis tools: GSA JES TVZ RAP. Wrote the paper: GSA JES JCZ.

References

  1. 1. Strand MR, Pech LL (1995) Immunological basis for compatibility in parasitoid host relationships. Annu Rev Entomol 40: 31–56.
  2. 2. Strand MR (2008) The insect cellular immune response. Insect Science 15: 1–14.
  3. 3. Abdel-latief M, Hilker M (2007) Innate immunity: eggs of Manduca sexta are able to respond to parasitism by Trichogramma evanescens. Insect Biochem Molec Biol 38: 136–145.
  4. 4. Yamamoto D, Henderson R, Corley LS, Iwabuchi K (2007) Intrinsic, inter-specific competition between egg, egg-larval, and larval parasitoids of plusiine loopers. Ecol Entomol 32: 221–228.
  5. 5. Bae S, Kim Y (2004) Host physiological changes due to parasitism of a braconid wasp, Cotesia plutellae, on diamondback moth, Plutella xylostella. Comp Biochem Physiol, A 138: 39–44.
  6. 6. Ribeiro C, Brehélin M (2006) Insect haemocytes: What type of cell is that? J Insect Physiol 52: 417–429.
  7. 7. Lavine MD, Strand MR (2002) Insect hemocytes and their role in immunity. Insect Biochem Molec Biol 32: 1295–1309.
  8. 8. Theopold U, Schmidt O, Söderhäll K, Dushay MS (2004) Coagulation in arthropods: defense, wound closure and healing. Trends Immunol 25: 289–294.
  9. 9. Cheng TC, Zhang YL, Liu C, Xu PZ, Gao ZH, et al. (2008) Identification and analysis of toll-related genes in the domesticated silkworm, Bombyx mori. Dev. Comp. Immunol 32: 464–475.
  10. 10. Jiravanichpaisal P, Lee BL, Söderhäll K (2006) Cell-mediated immunity in arthropods: Hematopoiesis, coagulation, melanization and opsonization. Immunobiology 211: 213–236.
  11. 11. Richards EH, Edwards JP (2000) Parasitism of Lacanobia oleracea (Lepidoptera) by the ectoparasitoid, Eulophus pennicornis, is associated with a reduction in host haemolymph phenoloxidase activity. Comp Biochem Physiol B 127: 289–298.
  12. 12. Beckage NE, Gelman DB (2004) Wasp parasitoid disruption of host development: implications for new biologically based strategies for insect control. Annu Rev Entomol 49: 299–330.
  13. 13. Narayanan K (2004) Insect defense: its impact on microbial control of insect pest. Curr Sci 86: 800–814.
  14. 14. Reed DA, Luhring KA, Stafford CA, Hansen AK, Millar JG, et al. (2007) Host defensive response against an egg parasitoid involves cellular encapsulation and melanization. Biol Control 41: 214–222.
  15. 15. Renault S, Petit A, Bénédet F, Bigot S, Bigot Y (2002) Effects of the Diadromus pulchellus ascovirus, DpAV-4, on the hemocytic encapsulation response and capsule melanization of the leek-moth pupa, Acrolepiopsis assectella. J Insect Physiol 48: 297–302.
  16. 16. Rantala MJ, Roff D A (2007) Inbreeding and extreme outbreeding cause sex differences in immune defence and life history traits in Epirrita autumnata. Heredity 98: 329–336.
  17. 17. Pennacchio F, Strand MR (2006) Evolution of developmental strategies in parasitic Hymenoptera. Annu Rev Entomol 51: 233–58.
  18. 18. Mendel Z, Protasov A, Blumberg D, Brand D, Saphir N, et al. (2007) Release and recovery of parasitoids of the eucalyptus gall wasp Ophelimus maskelli in Israel. Phytoparasitica 35: 330–332.
  19. 19. Andrade GS, Pratissoli D, Torres JB, Barros R, Dalvi LP, et al. (2009) Parasitismo de ovos de Heliothis virescens por Trichogramma spp. pode ser afetado por cultivares de algodão. Acta Sci Agron 31: 569–573.
  20. 20. Pratissoli D, Bueno AF, Bueno RCOF, Zanuncio JC, Polanczyk RA (2009) Trichogramma acacioi parasitism capacity at different temperatures and factitious hosts. Rev Bras Entomol 53: 151–153.
  21. 21. Pratissoli D, Zanuncio JC, Vianna UR, Andrade JS, Pinon TBM, et al. (2005) Thermal requirements of Trichogramma pretiosum and T. acacioi (Hym.: Trichogrammatidae), parasitoids of the avocado defoliator Nipteria panacea (Lep.: Geometridae), in eggs of two alternative hosts. Braz Arch Biol Technol 48: 523–523.
  22. 22. Zanuncio JC, Pereira FF, Jacques GC, Tavares MT, Serrão JE (2008) Tenebrio molitor Linnaeus (Coleoptera: Tenebrionidae), a new alternative host to rear the pupae parasitoid Palmistichus elaeisis Delvare & LaSalle (Hymenoptera: Eulophidae). Coleopt Bull 62: 64–66.
  23. 23. Paron MR, Berti Filho E (2000) Capacidade reprodutiva de Trichospilus diatraeae (Hymenoptera: Eulophidae) em pupas de diferentes hospedeiros (Lepidoptera). Scientia Agric 57: 355–358.
  24. 24. Pereira FF, Zanuncio TV, Zanuncio JC, Pratissoli D, Tavares MT (2008b) Species of Lepidoptera defoliators of eucalypt as new hosts for the polyphagous parasitoid Palmistichus elaeisis (Hymenoptera: Eulophidae). Braz J Biol 51: 259–262.
  25. 25. Pereira FF, Zanuncio JC, Tavares MT, Pastori PL, Jacques GC, et al. (2008a) New record of Trichospilus diatraeae as a parasitoid of the eucalypt defoliator Thyrinteina arnobia in Brazil. Phytoparasitica 36: 304–306.
  26. 26. Pereira FF, Zanuncio JC, Serrão JE, Oliveira HN, Fávero K, et al. (2009a) Progênie de Palmistichus elaeisis Delvare & LaSalle (Hymenoptera: Eulophidae) parasitando pupas de Bombyx mori L. (Lepidoptera: Bombycidae) de diferentes idades. Neotrop Entomol 38: 660–664.
  27. 27. Pereira FF, Zanuncio JC, Serrão JE, Pastori PL, Ramalho FS (2009b) Reproductive performance of Palmistichus elaeisis (Hymenoptera: Eulophidae) with previously refrigerated pupae of Bombyx mori (Lepidoptera: Bombycidae). Braz J Biol 69: 865–869.
  28. 28. Soares MA, Gutierrez CT, Zanuncio JC, Pedrosa RPP, Lorenzon AS (2009) Superparasitismo de Palmistichus elaeisis (Hymenoptera: Eulophidae) y comportamiento de defensa de dos hospederos. Revista Colombiana de Entomología 35: 62–67.
  29. 29. Greene GL, Leppla NC, Dickerson WA (1976) Velvetbean caterpillar: A rearing procedure and artificial diet. J Econ Entomol 69: 487–488.
  30. 30. Ibrahim AMA, Kim Y (2006) Parasitism by Cotesia plutellae alters the hemocyte population and immunological function of the diamondback moth, Plutella xylostella. J Insect Physiol 52: 943–950.
  31. 31. Souza DJ, van Vlaenderen J, Moret Y, Lenoir A (2008) Immune response affects ant trophallatic behaviour. J Insect Physiol 54: 828–832.
  32. 32. Rantala MJ, Kortet R (2004) Male dominance and immunocompetence in the field cricket (Gryllus bimaculatus). Behav Ecol 15: 187–191.
  33. 33. Cotter SC, Kruuk LEB, Wilson K (2004) Costs of resistance: genetic correlations and potential trade-offs in an insect immune System. J Evol Biol 17: 421–429.
  34. 34. Hoch G, Solter LF, Schopf A (2004) Hemolymph melanization and alterations in hemocyte numbers in Lymantria dispar larvae following infections with different entomopathogenic microsporidia. Entomol Exp Appl 113: 77–86.
  35. 35. Suzuki M, Tanaka T (2007) Development of Meteorus pulchricornis and regulation of its noctuid host, Pseudaletia separata. J Insect Physiol 53: 1072–1078.
  36. 36. Uckan F, Sinan S, Savasci S, Ergin E (2004) Determination of venom components from the endoparasitoid wasp Pimpla turionellae L. (Hymenoptera: Ichneumonidae). Ann Entomol Soc Am 97: 775–780.
  37. 37. Pech LL, Strand MR (1996) Granular cells are required for encapsulation of foreign targets by insect haemocytes. J Cell Sci 109: 2053–2060.
  38. 38. Carton Y, Poirié M, Nappi AJ (2008) Insect immune resistance to parasitoids. Insect Science 15: 67–87.
  39. 39. Asgari S, Schmidt O, Theopold U (1997) A polydnavirus-encoded protein of an endoparasitoid wasp is an immune suppressor. J Gen Virol 78: 3061–3070.
  40. 40. Shelby KS, Webb BA (1999) Polydnavirus-mediated suppression of insect immunity. J Insect Physiol 45: 507–514.
  41. 41. Amaya KE, Asgari S, Jung R, Hongskula M, Beckage NE (2005) Parasitization of Manduca sexta larvae by the parasitoid wasp Cotesia congregate induces an impaired host immune response. J Insect Physiol 51: 505–512.
  42. 42. Wu ML, Ye GY, Zhu JY, Chen XX, Hu C (2008) Isolation and characterization of an immunosuppressive protein from venom of the pupa-specific endoparasitoid Pteromalus puparum. J Invertebr Pathol 99: 186–191.
  43. 43. Nalini M, Ibrahim AMA, Hwang I, Kim Y (2009) Altered actin polymerization of Plutella xylostella (L.) in response to ovarian calyx components of an endoparasitoid Cotesia plutellae (Kurdjumov). Physiol Entomol 34: 110–118.
  44. 44. Chapman RF (1998) The insects: structure and function, Cambrigde University Press, UK.
  45. 45. Alphen van, Visser ME (1990) Superparasitism as an adaptive strategy for insect parasitoids Annu Rev Entomol 35: 59–79.
  46. 46. Silva-Torres C, Ramos Filho IT, Torres JB, Barros R (2009) Superparasitism and host size effects in Oomyzus sokolowskii, a parasitoid of diamondback moth. Entomol Exp Appl 133: 65–73.
  47. 47. Tena A, Kapranas A, Garcia-Mari F, Luck RF (2008) Host discrimination, superparasitism and infanticide by a gregarious endoparasitoid. Anim Behav 76: 789–799.
  48. 48. Pereira FP, Barros R, Pratissoli D, Parra JRP (2004) Biologia e exigências térmicas de Trichogramma pretiosum Riley e T.exiguum Pinto & Platner (Hymenoptera: Trichogrammatidae) criados em ovos de Plutella xylostella (L.) (Lepidoptera: Plutellidae). Neotrop Entomol 33: 231–236.
  49. 49. Moreira MD, Santos MCF, Beserra EB, Torres JB, Almeida RP (2009) Parasitismo e superparasitismo de Trichogramma pretiosum Riley (Hymenoptera: Trichogrammatidae) em ovos de Sitotroga cerealella (Oliver) (Lepidoptera: Gelechiidae). Neotrop Entomol 38: 237–242.