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Potential negative effects on biological control by Sancassania polyphyllae (Acari: Acaridae) on an entomopathogenic nematode species
Biological Control 54 (2010) 166–171
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Potential negative effects on biological control by Sancassania polyphyllae (Acari: Acaridae) on an entomopathogenic nematode species Z. Ipek Ekmen a,b, Selcuk Hazir b,*, Ibrahim Cakmak c, Nurdan Ozer a, Mehmet Karagoz c, Harry K. Kaya d a
Hacettepe University, Faculty of Science, Department of Biology, 06800 Beytepe-Ankara, Turkey Adnan Menderes University, Faculty of Arts and Science, Department of Biology, 09010 Aydin, Turkey c Adnan Menderes University, Faculty of Agriculture, Department of Plant Protection, 09010 Aydin, Turkey d Department of Nematology, University of California, One Shields Avenue, Davis, CA 95616, USA b
a r t i c l e
i n f o
Article history: Received 30 December 2009 Accepted 10 May 2010 Available online 13 May 2010 Keywords: Insect-parasitic nematode Nematophagous mite Polyphylla fullo Steinernema feltiae White grub
a b s t r a c t Sancassania polyphyllae (Acari: Acaridae) is associated with larvae of the white grub, Polyphylla fullo (Coleoptera: Scarabaeidae), and will feed on the infective juveniles of entomopathogenic nematodes in the families Steinernematidae and Heterorhabditidae which are important biological control agents of soil insect pests. We conducted laboratory studies to determine the potential negative effects this mite species might have on biological control of soil insect pests. Our objectives in this study were to (1) determine the response of S. polyphyllae adult mites to a nematode-killed insects on agar, (2) evaluate the predation by mites on Steinernema feltiae infective juveniles from nematode-killed insects on agar and in soil, and (3) assess predation efficiency of the mite on the infective juveniles in the soil. On agar, we found (1) significantly more adult female mites near or on a nematode-killed Ceratitis capitata (Diptera: Tephritidae) larva than near or on the freeze-killed larva or a bamboo mimic suggesting that a chemical or an odor from the nematode-killed larva attracted the mites, and (2) 10 mites consumed 96% of infective juveniles that emerged from an insect cadaver. In soil with a nematode-killed insect, the average number of infective juveniles recovered was <30 when mites were present, whereas the average number of infective juveniles recovered was >375 when the mites were absent. When the infective juveniles alone were placed in different depths in relation to the mites in the soil column for 4 and 10 days, S. polyphyllae was not as efficient at finding the infective juveniles when they were separated from each other in the soil lending support to the idea that the mites were cueing in on the cadaver as a food resource. Our data suggest that emerging infective juveniles from an insect cadaver in the soil in the presence of S. polyphyllae can adversely affect biological control because of nematode consumption by the mites. Ó 2010 Elsevier Inc. All rights reserved.
1. Introduction Entomopathogenic nematodes (EPNs) in the families Steinernematidae and Heterorhabditidae live in the soil and are obligate and lethal parasites of insects (Grewal et al., 2005). These nematodes are associated with mutualistic bacteria in the genera, Xenorhabdus and Photorhabdus (Enterobacteriales: Enterobacteriaceae) for Steinernema spp. and Heterorhabditis spp., respectively (Kaya and Gaugler, 1993). The mutualistic bacterial cells are located in the intestine of the infective juveniles which is the only free-living stage outside the insect host. The infective juvenile infects its soil insect host by entering through natural openings (mouth, anus or spiracle), penetrating into the insect’s body cavity and releasing the mutualistic bacterium which reproduces and kills the host within 48 h. The nematodes develop and feed on the bacterial cells * Corresponding author. Fax: +90 256 2135379. E-mail address: [email protected] (S. Hazir). 1049-9644/$ - see front matter Ó 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.biocontrol.2010.05.004
and host tissues, and depending on the species, infective juvenile progeny are normally produced in 7–15 days. These infective juveniles exit the host cadaver and search for new hosts in the soil. EPNs occur naturally in soil in many parts of the world (Hominick, 2002) and are important natural biological control agents. Moreover, they can be effectively applied as infective juveniles against soil insect pests and several species are produced and sold commercially (Koppenhöfer, 2007). However, a limiting factor in the use of these nematodes as biopesticides is the high losses of the infective juveniles occurring within the first few days after application because of abiotic factors (e.g., high temperature, soil type, low moisture, UV light, dehydration) (Smits, 1996; Baur and Kaya, 2001). Once the infective juveniles move into the soil environment away from the harsh abiotic factors, their natural enemies such as nematophagous fungi and invertebrate predators may cause additional mortality (Kaya, 2002). A number of invertebrate predators including tardigrades, turbellarians, collembolans and other insects, mites, predatory nematodes, and oligochaetes
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have been linked to population reduction of many different nematode species (Small, 1988; Hazir et al., 2003). This linkage is weak because of the difficulty in quantifying the effects of these natural enemies against EPNs in the soil habitat (Kaya et al., 1998), and therefore, quantitative data to support the impact of these predators on EPN populations are lacking (Kaya, 2002). Previous studies have demonstrated that different mite groups may be important natural enemies of EPNs. For example, Poinar (1979) reported that mites in the genus Macrocheles fed on Steinernema feltiae (Filipjev) infective juveniles, Ishibashi et al. (1987) found that mites in the genus Eugamasus fed on Steinernema carpocapsae (Weiser) infective juveniles, and Epsky et al. (1988) demonstrated that 12 mite species representing several groups (Mesostigmata, Endeostigmata, Oribatida non-Astigmatina, and Astigmatina) also fed on S. carpocapsae. Karagoz et al. (2007) demonstrated that the female mites of a Sancassania sp. [later identified as Sancassania polyphyllae (Zachvatkin)] consumed most of the infective juveniles of S. feltiae or Heterorhabditis bacteriophora Poinar. In a field study, Wilson and Gaugler (2004) correlated the decline in the infective juveniles to the rise in mites and collembolans but did not observe them feeding on the nematodes or identify the arthropod species. Karagoz et al. (2007) and Cakmak et al. (2010) have gained significant understanding about the interactions between S. polyphyllae and EPNs. Karagoz et al. (2007) reported that the deutonymphal stage of S. polyphyllae was found on living third-instar white grub [Polyphylla fullo (L.) (Coleoptera: Scarabaeidae)] larvae from a strawberry field in Turkey. In a laboratory study, they found the following: (1) two adult females of S. polyphyllae consumed more than 80 of 100 S. feltiae infective juveniles on an agar substrate within 24 h; (2) they fed on more infective juveniles of S. feltiae than of H. bacteriophora (i.e., 10 female S. polyphyllae consumed 99.4 % of 500 S. feltiae infective juveniles and 86% of 500 H. bacteriophora infective juveniles within 48 h); and (3) in 24-well tissue culture plates, S. polyphyllae preyed on more S. feltiae infective juveniles in sandy than in loamy soil. Furthermore, Cakmak et al. (2010) showed that S. polyphyllae successfully completed its development and reproduced when feeding on S. feltiae and H. bacteriophora infective juveniles. Sancassania polyphyllae enters a hypopus stage under unfavorable conditions such as inadequate food supply and attaches to a larval scarab host. This ability may also enhance the mite’s efficacy as a predator of nematodes, especially if its host is killed by EPNs or if the host cadaver is invaded by saprophagous nematodes. Sancassania polyphyllae deutonymphs were found on P. fullo larvae collected from 5–10 cm depth of the soil in a strawberry field. When EPNs are applied, they can kill the larva in situ. Therefore, it is important to understand the spatial relationship of where the nematode-killed larva is in relation to where the infective juveniles emerge and the feeding activity of the mite. We hypothesized that the spatial distribution of the EPN infective juveniles in the presence or absence of nematode-killed insects in relation to the occurrence of S. polyphyllae will affect nematode survival in soil. Accordingly, we conducted laboratory studies to determine the potential negative effects this mite might have on the biological control of soil insect pests. Our objectives in this study were to (1) assess the response of S. polyphyllae adult mites to a nematode-killed insect on agar, (2) evaluate the predation by S. polyphyllae adult mites on S. feltiae infective juveniles from a nematode-killed insect on agar or in a soil column, and (3) assess predation efficiency of the mite on applied infective juveniles in a soil column. The findings obtained from this study will provide useful information on how this mite could affect the survival of the infective juveniles in soil which in turn would have an effect on natural or applied biological control.
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2. Materials and methods 2.1. Sources of insects, mites and nematodes The wax moth, Galleria mellonella (L.) (Lepidoptera: Pyralidae), was reared on an artificial medium that included honey, wheat bran, glycerol, soy flour, milk powder, dry yeast and honey bee wax in glass jar at 25 ± 4 °C in the laboratory. S. feltiae (Turkish isolate 09-31) was reared on last instar G. mellonella larvae in the laboratory (Kaya and Stock, 1997). The nematode-killed G. mellonella larvae were placed on a White trap, and the emerging infective juveniles from the cadavers were collected from the water and stored at 10 °C in tissue culture flasks (White, 1927). Infective juveniles were used within 2 weeks for the experiments. Larvae of P. fullo (Coleoptera: Scarabaeidae) were collected from strawberry fields in Aydin, Turkey and brought to the laboratory where the majority of them were stored at 20 °C. Four live P. fullo larvae with the deutonymphal stage of S. polyphyllae were killed, and the larvae were dissected in a petri dish (9 cm diameter) so that the various tissues were exposed to initiate the mite colony. Subsequently, S. polyphyllae was reared in petri dishes containing dissected tissues of previously frozen P. fullo larvae and maintained at room temperature (Karagoz et al., 2007). The larvae of Mediterranean fruit fly (medfly), Ceratitis capitata (Wiedemann) (Diptera: Tephritidae), were obtained from Bornova Plant Protection Research Institute in Izmir, Turkey as needed for the experiments. We used the medfly larvae for our experiments because it is one of the target insects for our research program, pupates in the soil, is susceptible to EPNs, provides a realistic target for EPNs, and produces infective juveniles that can be easily quantified in the soil in contrast to an unnatural host such as G. mellonella which never enters the soil. 2.2. Infection of medfly larvae with nematodes The medfly larvae were infected with S. feltiae in 24-well tissue culture plates. One last instar jumping larva (i.e., that was ready to pupate) was placed on the surface of each well that contained 0.5 g of sterilized, air-dried sand to which 30 infective juveniles in 60 ll distilled water were added. The well plate was covered with Parafilm to prevent the escape of the larvae. The larvae were checked for mortality caused by the infective juveniles daily as indicated by the light brown color of the cadaver. All nematode-killed larvae were transferred individually to White traps and checked three times a day for nematode development as the larval cuticle was transparent and nematode development could be determined. Depending on the experiment, 5- or 6-day-old nematode-killed medfly larvae were used and all experiments were conducted at room temperature (23 ± 1 °C). 2.3. Response of medfly cadaver to mites in a petri dish To assess the response of 10 adult female mites to (1) a 5-dayold nematode-killed medfly larva or (2) a fresh, frozen-killed larva ( 20 °C for 30 min) or (3) a piece of bamboo the same length and width as a medfly larva, the following experiment was conducted. The nematode-killed larva, frozen-killed larva and bamboo were placed near the edge of an experimental arena (100 15 mm plastic petri dish containing 20 ml of 2% water agar), equidistant apart and 3.5 cm from the center of the dish. Then 10 S. polyphyllae adult female mites were released onto the agar surface in the center of the arena, and data on the number of mites at a cadaver or bamboo were taken every 20 min up to 60 min. Ten petri dishes were prepared for the experiment with each plate serving as a replicate, and the experiment was conducted twice.
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2.4. Medfly cadaver, emerging nematodes, and mites in a petri dish For each experimental round, 40 medfly larvae were used at 6days post-nematode infection when the first emergence of infective juveniles from the cadavers was evident. Each nematodekilled medfly larva with emerging infective juveniles was placed in the center of an experimental arena (100 15 mm plastic petri dish containing 20 ml of 2% water agar). Forty petri dishes were prepared for the experiment. Twenty dishes served as the treatment with 10 S. polyphyllae females being released on the agar surface, and 20 dishes served as controls without mites and containing only the medfly larval cadaver. To standardize the experiment, the petri dish with the mites was paired with a control dish with infective juveniles emerging from the larva at about the same time. Three days later, the mites and the larval cadaver were removed from the petri dishes, and the infective juveniles on the agar surface of each dish were rinsed off with distilled water into a 100-ml beaker, and all nematodes in the suspension were counted at 40 magnification with a dissecting microscope. This experiment was conducted twice for a total of 40 petri dishes each for the treatment and control, respectively.
initial set up, whereas in the second experiment, the soil column was destructively examined 10 days after set up. For both experiments, there were 10 tubes (replicates) for each treatment which were kept in the dark, and each experiment was conducted twice for a total of 20 replicates. After 4 or 10 days, the soil in the tubes was handled as in the soil column experiment with the medfly cadaver. For the 4- and 10-day sampling, the experimental design was the same as described above in the soil column experiment with the medfly larval cadaver except that rather than placing a larval cadaver with emerging infective juveniles, 500 S. feltiae infective juveniles in 200 ll of water were pipetted onto the soil surface. In addition, rather than placing 50 adult female mites in the soil column, 10 adult female mites were used. For the 4-day sample, treatment A had the infective juveniles and mites at the soil surface, treatment B had the infective juveniles at 2 cm level of soil and the mites on the top surface, and treatment C had the infective juveniles on the top surface and the mites were at the 2 cm level. Control tubes were prepared for each treatment with 500 infective juveniles but no mites were added. For the 10-day sample, only treatment B and C as described above was prepared.
2.5. Medfly cadaver and mites in a soil column Fifty-ml plastic centrifuge tubes (30 115 mm) (Ratilabo, Karlsruhe, Germany) were used as soil columns and filled with 20 g of moistened (10% w/w) soil (69.4% sand, 30.6 % silt, pH 8.3) collected from a strawberry field in Aydin (see steps A–C below for details). Prior to being moistened, the soil was pasteurized and air-dried for over 2 weeks. The total height of the soil column with 20 g of soil was 9 cm. Three different treatments were employed in this experiment depending on the placement of a 6day-old medfly larval cadaver with emerging infective juveniles and 50 adult female mites in the soil column. The treatments were as follows. A. Twenty g of moistened soil was added into the centrifuge tube and the mites and one medfly larval cadaver with emerging nematodes were placed on the soil surface (referred to as A+). B. Four g of moistened soil was added into the centrifuge tube (2 cm of soil in the tube) and one medfly larval cadaver was placed on the soil surface. The remaining 16 g of the moistened soil was added into the tube and the mites were released to the soil surface resulting in a 7-cm distance between the medfly cadaver and mites (referred to as B+). C. The same protocol as ‘‘B” was followed except that placement of the medfly cadaver and mites were reversed. That is, the mites were placed at the 2 cm level and the medfly larval cadaver was placed on the soil surface (referred to as C+). Twenty tubes were used for each treatment. Control tubes were prepared for each treatment with a medfly larval cadaver with emerging infective juveniles but no mites were added (Referred to as A , B and C , respectively). All experiments were conducted twice and kept in the dark. After 10 days, the soil in each tube was washed and sieved (53-lm mesh) to remove soil particles and obtain the infective juveniles. The different mite stages in the soil were noted but not counted. The collected nematodes were counted at 40 magnification with a dissecting microscope as described for the agar substrate counts. 2.6. Infective juveniles and mites in a soil column Two different experiments were conducted. In the first experiment, the soil column was destructively sampled 4 days after the
2.7. Statistical analysis The data were analyzed using analysis of variance (ANOVA). Means were compared at the P = 0.05 level, and Tukey’s test was used to separate means (SPSS, 2004). The means ± SEM are provided below. 3. Results 3.1. General observations When last instar medfly larvae were individually exposed to 30 infective juveniles in 24-well tissue culture plates, 92% of them were infected by the nematodes, and the first infective juveniles began emerging from the cadavers at 6 days after infection. The total number of infective juveniles emerging from a C. capitata larva in a White trap averaged 2500 ± 510 over a 6-day period. However, in our experiments, less infective juveniles emerged because the duration of the experiments were less than 6 days. In the petri dish agar experiment, the mites fed on intact medfly larval cadavers with nematodes and produced eggs. In the soil column experiments, we recovered adult mites, eggs and larvae from the soil with the infective juveniles only experiment after 4 days, whereas we recovered adults, eggs, and immature mites (larvae, protonymphs and tritonymphs) from the soil with the medfly larval cadaver after 10 days. We did not quantify the number of mites and the various stages recovered because our focus was on counts of the infective juveniles. 3.2. Response of medfly cadaver to mites in a petri dish At 20 min after exposure, the average number of mites at the nematode-killed medfly larva was 1.4 ± 0.4, at the frozen larva was 0.5 ± 0.2, and at the bamboo was 0.1 ± 0.1. At 40 min, the average number of mites at the nematode-killed medfly larva was 2.6 ± 0.5, at the frozen larva was 0.9 ± 0.3, and at the bamboo was 0.1 ± 0.1. Significant differences were seen between the bamboo and the nematode-killed medfly larva at 20 (F = 4.31; df = 2, 27; P < 0.05) and 40 (F = 6.86; df = 2, 27; P < 0.01) min. At 60 min exposure, the average number of mites at the nematode-killed medfly larva was 3.1 ± 0.6, at the frozen larva was 1 ± 0.2, and at the bamboo was 0. Statistical analysis showed that significantly
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10 days, the numbers of infective juveniles recovered were significantly less in soil columns with the mites than without the mites (P < 0.001, Fig. 1). In all cases, the placement of the mites or the larval cadaver did not affect the results. That is, the average number of recovered infective juveniles was <30 infective juveniles when mites were present, whereas the average number of recovered infective juveniles was >375 infective juveniles when the mites were absent. In treatment A (F = 15.04; df = 1, 38; P < 0.001) when the larval cadaver and mites were both placed on the soil surface, an average of <1 infective juvenile was recovered, whereas >340 infective juveniles were recovered from the soil column without mites. In treatment B (F = 22.98; df = 1, 42; P < 0.001) and C (F = 84.87; df = 1, 33; P < 0.001) when the larval cadaver and mites were separated from each other in the soil column, significantly fewer infective juveniles were recovered from the soil column without mites. When the three experimental groups with mites plus cadavers with nematodes (i.e., A+, B+ and C+) were compared, significant differences were observed only between cadaver and mites placed on the surface (A+) and cadaver placed at 2 cm level and the mites placed on the soil surface (B+) (F = 5.19; df = 2, 63; P < 0.05).
more mites were found on the nematode-killed medfly larva than at the freeze-killed medfly larva or at the piece of bamboo (F = 9.87; df = 2, 27; P < 0.001) at 60 min. There was no significant difference between the frozen larva and the bamboo. 3.3. Medfly cadaver, emerging nematodes, and mites in a petri dish On the agar media, significant differences were observed in the numbers of nematodes recovered between larval cadaver with mites (33 ± 8 infective juveniles) and larval cadaver without mites (857 ± 100 infective juveniles) (F = 68.03; df = 1, 82; P < 0.001) after 3 days. Based on what was recovered from the control agar plate, 10 adult female mites consumed 96% of the emerged infective juveniles from the C. capitata cadaver. We observed that mites were around and near the cadaver and attacked the emerging infective juveniles. 3.4. Medfly cadaver and mites in a soil column In all treatments (i.e., A–C), when the medfly larva with nematodes was placed into the soil column with mites or no mites for
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Fig. 1. Mean number ± standard error of consumed Steinernema feltiae infective juveniles that emerged from a Ceratitis capitata larva by Sancassania polyphyllae in a soil column. Treatment A = larval cadaver on the soil surface (solid bars) with A+ being mites placed at the surface and A with no mites. Treatment B = larval cadaver at 2 cm from the bottom (open bars) with B+ being the mites placed on the soil surface and B with no mites. Treatment C = larval cadaver on the soil surface (patterned with black and white squares) with C+ being mites placed at 2 cm from the bottom and C with no mites. Different capital letters (X and Y) above the bars indicate significant differences among the treatments with mites. Different lowercase letters (a and b) above the bars indicate significant differences between each treatment and their control.
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Treatments Fig. 2. Mean number ± standard error of consumed Steinernema feltiae infective juveniles by Sancassania polyphyllae in a soil column after 4 days incubation. Treatment A = infective juveniles placed on the soil surface (solid bars) with A+ being mites placed at the surface and A with no mites. Treatment B = infective juveniles placed at 2 cm from the bottom(open bars) with B+ being the mites placed on the soil surface and B with no mites. Treatment C = infective juveniles placed on the soil surface (patterned with black and white squares) with C+ being mites placed at 2 cm from the bottom and C with no mites. Different capital letters (X, Y and Z) and different lowercase letters (a and b) above the bars indicate significant differences between the treatments and their control.
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3.5. Infective juveniles and mites in a soil column When the infective juveniles alone were placed into the soil column with mites for 4 days, our results showed that the recovery of infective juveniles from soil differed depending on where the infective juveniles were placed in relation to the mites (Fig. 2). For example, in treatment A+ when the infective juveniles and mites were placed on the surface of the soil column together, significantly fewer infective juveniles were recovered from the column that had mites present than the control group that had no mites (F = 87.02; df = 1, 26; P < 0.001) (Fig. 2). Similarly, in treatment B+ when the infective juveniles were placed at 2 cm from the bottom of the soil column and the mites were on the soil surface, significantly fewer infective juveniles were recovered compared to the control group that had no mites (F = 7.52; df = 1, 28; P < 0.05). However, in treatment C+ when the placement of mites and infective juveniles were reversed, no significant difference was observed between mites present and the control group (F = 0.41; df = 1, 26; P > 0.05).When all three experimental groups in which mites and infective juveniles were together (i.e., A+, B+, C+) in the soil column were compared, statistically significant differences were seen among all groups (indicated as X, Y, and Z above the bars) (F = 37.90; df = 2, 57; P < 0.001) (Fig. 2). The same experimental design was prepared with the mites and nematodes incubated for 10 days in the soil column as above, but it only had treatments B and C. The average numbers of infective juveniles were 168 and 158 for B+ and C+ at the end of the 10day incubation period, respectively, and no significant difference was observed between B+ and C+ groups (F = 0.52; df = 1, 18; P > 0.05). However, significantly fewer infective juveniles were recovered from B+ compared with B (F = 24.29; df = 1, 18; P < 0.001 and from C+ compared with C (F = 31.53; df = 1, 18; P < 0.001 (Fig. 3).
4. Discussion
Number of infective juveniles
We demonstrated that the spatial distribution of EPN infective juveniles in the presence or absence of nematode-killed insects in relation to the occurrence of S. polyphyllae affected nematode survival in soil. Accordingly, we showed that S. polyphyllae fed more on infective juveniles emerging from a cadaver in soil than on infective juveniles in the absence of a cadaver (e.g., when infective juveniles are applied as a biopesticide). Thus, our hypothesis, in general, was correct in that the spatial distribution of the infective juveniles
was affected depending upon whether S. polyphyllae was present or absent or what the source of the infective juveniles was. In the initial experiment, we found that significantly more female adult mites of S. polyphyllae were found associated with a nematode-killed medfly larva than to a fresh, freeze-killed larva or a piece of bamboo. The data suggested that a chemical(s) or odor(s) from the nematode-killed larva was produced and served as an attractant for the mites. In the second and third experiments, our data showed that the female mites consumed most of the infective juveniles emerging from a medfly cadaver on an agar substrate and in soil. In the soil column that had nematode-killed medfly larvae with emerging infective juveniles, our data suggest that the 50 female mites and their immature progeny [that is, the protonymphs and tritonymphs are known to feed on the infective juveniles of S. feltiae in the laboratory (see Cakmak et al., 2010)] found the cadaver and probably congregated around it to consume the emerging nematodes over a 10-day period. In fact, the mites were efficient predators of the infective juveniles regardless of whether the cadaver was placed on the top or bottom of the soil assay chamber. These findings suggest that S. polyphyllae may have a negative impact on the biological control of soil insect pests using cadaver containing nematodes (see Shapiro and Lewis, 1999) where S. polyphyllae or other nematophagous mites occur. For example, the application of cadavers with infective juveniles shows considerable promise because the emerging nematodes of some species are more infectious than with infective juveniles applied with water (Shapiro and Lewis, 1999; Shapiro-Ilan et al., 2003). However, if S. polyphyllae is present, it could be an efficient predator of the infective juveniles as they emerged from the cadaver in the soil environment. In addition, the mites may feed on the cadaver before the infective juveniles emerge rendering the cadaver unsuitable for nematode development. Because the mites are small and can easily move through the small soil pores, they may actually cue in on the cadaver and consume the infective juveniles as they emerge from the cadaver as was observed in our agarbased experiment. Thus, if these mites are abundant and near a nematode-killed insect, they may negate the advantages of using a cadaver application. In the fourth experiment, we found that when the free-living infective juveniles alone were placed with 10 mites over a 4-day period, the mites were not as efficient at finding the nematodes compared with the larval cadaver treatments. On the other hand, placing the mites and the free-living infective juveniles in proximity to each other did have a significant negative effect on the numbers of infective juveniles. If the infective juveniles and mites were
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Treatments Fig. 3. Mean number ± standard error of consumed Steinernema feltiae infective juveniles by Sancassania polyphyllae in a soil column after 10 days incubation. Treatment B = infective juveniles placed at 2 cm from the bottom (solid bars) with B+ being the mites placed on the soil surface and B with no mites. Treatment C = infective juveniles placed on the soil surface (open bars) with C+ being mites placed at 2 cm from the bottom and C with no mites. Same capital letters (X) above the bars indicate nonsignificant differences between the treatments with mites. Different lowercase letters (a and b) above the bars indicate significant differences between the treatments and their control.
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spatially separated, the predation rate of the infective juveniles by the mites was reduced or non-existent. These data suggest that the mites were not as efficient at finding the infective juveniles when they were separated from each other in the soil lending support to the idea that the mites were cueing in on the cadaver as a food resource. Although our laboratory study suggests that the presence of S. polyphyllae or a mite species with similar feeding habits has the potential for having a negative effect on biological control by EPN species of soil insect pests, there may be situations where its effect on biological control may be mitigated. From the literature, we know that mites in the genus Sancassania (=Caloglyphus) are biologically diverse group in the family Acaridae and that some species are associated with various arthropods including Coleoptera, Hymenoptera, Orthoptera, Myriapoda and terrestrial Crustacea (Klimov et al., 2004). Sancassania species feed on insect cadavers and microorganisms associated with the cadavers (Krantz, 1978; Krantz and Walter, 2009) as well as on nematodes (Sell, 1988; Karagoz et al., 2007). Our study was done in the absence of other nematode species, and the presence of other nematode species may mitigate the effects of S. polyphyllae. For example, Sell (1988) showed that a Sancassania sp. fed on a plant-parasitic nematode species, and this type of non-selective feeding by S. polyphyllae may allow survival of EPNs. In addition, Ekmen et al. (2010) showed that S. polyphyllae had a preference for tissues of Polyphylla fullo over that of live S. feltiae or H. bacteriophora infective juveniles. If there are a number of insect cadavers in the soil, S. polyphyllae will feed on the cadavers, but this may result in a significant population increase of S. polyphyllae. Once the insect cadavers are consumed, the mites will need alternative food sources which could potentially reduce EPN numbers if they are in the same area. In conclusion, our study strongly suggests that chemicals associated with insect cadavers influence S. polyphyllae behavior. Therefore, they may detect C. capitata cadavers easily in the soil column and consume the infective juveniles as they emerge from the cadavers. On the other hand, the mites had difficulty in finding and preying on the infective juveniles in the absence of a cadaver, especially when the mites and infective juveniles were separated spatially. Although we have gained a greater understanding of S. polyphyllae mites as prey of S. feltiae infective juveniles, our knowledge about the impact of this mite on nematode populations under field conditions remains unknown. Our future studies will involve field studies where the mite naturally occurs and assess if their presence will have significant effect on biological control of soil insect pests. Acknowledgments We thank Baris Gulcu, Derya Asici and Cem Demirtas for their assistance in the experiments. References Baur, M.E., Kaya, H.K., 2001. Persistence of entomopathogenic nematodes. In: Baur, M.E., Fuxa, J.R. (Eds.), Southern Cooperative Series Bulletin, p. 26,.
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Cakmak, I., Ekmen, Z.I., Karagoz, M., Hazir, S., Kaya, H.K., 2010. Development and reproduction of Sancassania polyphyllae (Acari: Acaridae) feeding on entomopathogenic nematodes and tissues of insect larvae. Pedobiologia 53, 235–240. Ekmen, Z.I., Cakmak, I., Karagoz, M., Hazir, S., Ozer, N., Kaya, H.K., 2010. Food preference of Sancassania polyphyllae (Acari: Acaridae): living entomopathogenic nematodes or insect tissues? Biocontrol Science and Technology 20, 553–566. Epsky, N.D., Walter, D.E., Capinera, J.L., 1988. Potential role of nematophagous microarthropods as biotic mortality factors of entomogenous nematodes (Rhabditida: Steinernematidae and Heterorhabditidae). Journal of Economic Entomology 81, 821–825. Grewal, P.S., Ehlers, R.U., Shapiro-Ilan, D.I., 2005. Nematodes as Biocontrol Agents. CABI Publishing, Wallingford, UK. Hazir, S., Kaya, H.K., Stock, S.P., Keskin, N., 2003. Entomopathogenic nematodes (Steinernematidae and Heterorhabditidae) for biological control of soil pests. Turkish Journal of Biology 27, 181–202. Hominick, W.M., 2002. Biogeography. In: Gaugler, R. (Ed.), Entomopathogenic Nematology. CABI Publishing, Wallingford, UK, pp. 115–143. Ishibashi, N., Young, F.Z., Nakashima, M., Agabeyru, C., Haraguchi, N., 1987. Effects of application of DD-136 on silkworm, Bombyx mori, predatory insect, Agriosphodorus dohrni, parasitoid, Trichomalus apanteloctenus, soil mites, and other non-target soil arthropods, with brief notes on feeding behavior and predatory pressure of soil mites, tardigrates, and predatory nematodes on DD136 nematodes. In: Ishibashi, N. (Ed.), Recent Advances in Biological Control of Insect Pests by Entomogenous Nematodes in Japan. Ministry of Education, Culture and Science, Japan, pp. 158–164. Grant No. 59860005 (in Japanese with English summary). Karagoz, M., Gulcu, B., Cakmak, I., Kaya, H.K., Hazir, S., 2007. Predation of entomopathogenic nematodes by Sancassania sp. (Acari: Acaridae). Experimental and Applied Acarology 43, 85–95. Kaya, H.K., 2002. Natural enemies and other antagonists. In: Gaugler, R. (Ed.), Entomopathogenic Nematology. CABI Publishing, Wallingford, UK, pp. 169–203. Kaya, H.K., Gaugler, R., 1993. Entomopathogenic nematodes. Annual Review of Entomology 38, 181–206. Kaya, H.K., Stock, S.P., 1997. Techniques in insect nematology. In: Lacey, L.A. (Ed.), Manual of Techniques in Insect Pathology. Academic Press, London, pp. 281– 324. Kaya, H.K., Koppenhöfer, A.M., Johnson, M., 1998. Natural enemies of entomopathogenic nematodes. Japanese Journal of Nematology 28, 13–21. Klimov, P.B., Lekveishvili, M., Dowling, A.P.G., OConnor, B.M., 2004. Multivariate analysis of morphological variation in two cryptic species of Sancassania (Acari: Acaridae) from Costa Rica. Annals of the Entomological Society of America 97, 322–345. Koppenhöfer, A.M., 2007. Nematodes. In: Lacey, L.A., Kaya, H.K. (Eds.), Field Manual of Techniques in Invertebrate Pathology. Springer Publishing, Dordrecht, The Netherlands, pp. 249–264. Krantz, G.W., 1978. A Manual of Acarology, second ed. Oregon State University Book Stores, Inc., Corvallis, OR. Krantz, G.W., Walter, D.E., 2009. A Manual of Acarology, third ed. Texas Tech University Press, Lubbock, TX. Poinar Jr., G.O., 1979. Nematodes for Biological Control of Insects. CRC Press, Boca Raton, FL. Sell, P., 1988. Caloglyphus sp. (Acarina: Acaridae) an effective nematophagous mite on root-knot nematode (Meloidogyne spp.). Nematologica 34, 246–248. Shapiro, D.I., Lewis, E.E., 1999. Infectivity of entomopathogenic nematodes from cadavers vs. aqueous applications. Environmental Entomology 28, 907–911. Shapiro-Ilan, D.I., Lewis, E.E., Son, Y.S., 2003. Superior efficacy observed in entomopathogenic nematodes applied in infected-host cadavers compared with application in aqueous suspension. Journal of Invertebrate Pathology 83, 270–272. Small, R.W., 1988. Invertebrate predators. In: Poinar Jr., G.O., Jansson, H.B. (Eds.), Diseases of Nematodes, vol. 2. CRC Press, Boca Raton, FL, pp. 73–92. Smits, H.P., 1996. Post-application persistence of entomopathogenic nematodes. Biocontrol Science and Technology 6, 379–387. SPSS, 2004. SPSS v.13.0 for Windows, SPSS Inc., Chicago, IL, USA. White, G.F., 1927. A method for obtaining infective nematode larvae from culture. Science 66, 302–303. Wilson, M., Gaugler, R., 2004. Factors limiting short-term persistence of entomopathogenic nematodes. Journal of Applied Entomology 128, 250–253.
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Biological Control journal homepage: www.elsevier.com/locate/ybcon
Potential negative effects on biological control by Sancassania polyphyllae (Acari: Acaridae) on an entomopathogenic nematode species Z. Ipek Ekmen a,b, Selcuk Hazir b,*, Ibrahim Cakmak c, Nurdan Ozer a, Mehmet Karagoz c, Harry K. Kaya d a
Hacettepe University, Faculty of Science, Department of Biology, 06800 Beytepe-Ankara, Turkey Adnan Menderes University, Faculty of Arts and Science, Department of Biology, 09010 Aydin, Turkey c Adnan Menderes University, Faculty of Agriculture, Department of Plant Protection, 09010 Aydin, Turkey d Department of Nematology, University of California, One Shields Avenue, Davis, CA 95616, USA b
a r t i c l e
i n f o
Article history: Received 30 December 2009 Accepted 10 May 2010 Available online 13 May 2010 Keywords: Insect-parasitic nematode Nematophagous mite Polyphylla fullo Steinernema feltiae White grub
a b s t r a c t Sancassania polyphyllae (Acari: Acaridae) is associated with larvae of the white grub, Polyphylla fullo (Coleoptera: Scarabaeidae), and will feed on the infective juveniles of entomopathogenic nematodes in the families Steinernematidae and Heterorhabditidae which are important biological control agents of soil insect pests. We conducted laboratory studies to determine the potential negative effects this mite species might have on biological control of soil insect pests. Our objectives in this study were to (1) determine the response of S. polyphyllae adult mites to a nematode-killed insects on agar, (2) evaluate the predation by mites on Steinernema feltiae infective juveniles from nematode-killed insects on agar and in soil, and (3) assess predation efficiency of the mite on the infective juveniles in the soil. On agar, we found (1) significantly more adult female mites near or on a nematode-killed Ceratitis capitata (Diptera: Tephritidae) larva than near or on the freeze-killed larva or a bamboo mimic suggesting that a chemical or an odor from the nematode-killed larva attracted the mites, and (2) 10 mites consumed 96% of infective juveniles that emerged from an insect cadaver. In soil with a nematode-killed insect, the average number of infective juveniles recovered was <30 when mites were present, whereas the average number of infective juveniles recovered was >375 when the mites were absent. When the infective juveniles alone were placed in different depths in relation to the mites in the soil column for 4 and 10 days, S. polyphyllae was not as efficient at finding the infective juveniles when they were separated from each other in the soil lending support to the idea that the mites were cueing in on the cadaver as a food resource. Our data suggest that emerging infective juveniles from an insect cadaver in the soil in the presence of S. polyphyllae can adversely affect biological control because of nematode consumption by the mites. Ó 2010 Elsevier Inc. All rights reserved.
1. Introduction Entomopathogenic nematodes (EPNs) in the families Steinernematidae and Heterorhabditidae live in the soil and are obligate and lethal parasites of insects (Grewal et al., 2005). These nematodes are associated with mutualistic bacteria in the genera, Xenorhabdus and Photorhabdus (Enterobacteriales: Enterobacteriaceae) for Steinernema spp. and Heterorhabditis spp., respectively (Kaya and Gaugler, 1993). The mutualistic bacterial cells are located in the intestine of the infective juveniles which is the only free-living stage outside the insect host. The infective juvenile infects its soil insect host by entering through natural openings (mouth, anus or spiracle), penetrating into the insect’s body cavity and releasing the mutualistic bacterium which reproduces and kills the host within 48 h. The nematodes develop and feed on the bacterial cells * Corresponding author. Fax: +90 256 2135379. E-mail address: [email protected] (S. Hazir). 1049-9644/$ - see front matter Ó 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.biocontrol.2010.05.004
and host tissues, and depending on the species, infective juvenile progeny are normally produced in 7–15 days. These infective juveniles exit the host cadaver and search for new hosts in the soil. EPNs occur naturally in soil in many parts of the world (Hominick, 2002) and are important natural biological control agents. Moreover, they can be effectively applied as infective juveniles against soil insect pests and several species are produced and sold commercially (Koppenhöfer, 2007). However, a limiting factor in the use of these nematodes as biopesticides is the high losses of the infective juveniles occurring within the first few days after application because of abiotic factors (e.g., high temperature, soil type, low moisture, UV light, dehydration) (Smits, 1996; Baur and Kaya, 2001). Once the infective juveniles move into the soil environment away from the harsh abiotic factors, their natural enemies such as nematophagous fungi and invertebrate predators may cause additional mortality (Kaya, 2002). A number of invertebrate predators including tardigrades, turbellarians, collembolans and other insects, mites, predatory nematodes, and oligochaetes
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have been linked to population reduction of many different nematode species (Small, 1988; Hazir et al., 2003). This linkage is weak because of the difficulty in quantifying the effects of these natural enemies against EPNs in the soil habitat (Kaya et al., 1998), and therefore, quantitative data to support the impact of these predators on EPN populations are lacking (Kaya, 2002). Previous studies have demonstrated that different mite groups may be important natural enemies of EPNs. For example, Poinar (1979) reported that mites in the genus Macrocheles fed on Steinernema feltiae (Filipjev) infective juveniles, Ishibashi et al. (1987) found that mites in the genus Eugamasus fed on Steinernema carpocapsae (Weiser) infective juveniles, and Epsky et al. (1988) demonstrated that 12 mite species representing several groups (Mesostigmata, Endeostigmata, Oribatida non-Astigmatina, and Astigmatina) also fed on S. carpocapsae. Karagoz et al. (2007) demonstrated that the female mites of a Sancassania sp. [later identified as Sancassania polyphyllae (Zachvatkin)] consumed most of the infective juveniles of S. feltiae or Heterorhabditis bacteriophora Poinar. In a field study, Wilson and Gaugler (2004) correlated the decline in the infective juveniles to the rise in mites and collembolans but did not observe them feeding on the nematodes or identify the arthropod species. Karagoz et al. (2007) and Cakmak et al. (2010) have gained significant understanding about the interactions between S. polyphyllae and EPNs. Karagoz et al. (2007) reported that the deutonymphal stage of S. polyphyllae was found on living third-instar white grub [Polyphylla fullo (L.) (Coleoptera: Scarabaeidae)] larvae from a strawberry field in Turkey. In a laboratory study, they found the following: (1) two adult females of S. polyphyllae consumed more than 80 of 100 S. feltiae infective juveniles on an agar substrate within 24 h; (2) they fed on more infective juveniles of S. feltiae than of H. bacteriophora (i.e., 10 female S. polyphyllae consumed 99.4 % of 500 S. feltiae infective juveniles and 86% of 500 H. bacteriophora infective juveniles within 48 h); and (3) in 24-well tissue culture plates, S. polyphyllae preyed on more S. feltiae infective juveniles in sandy than in loamy soil. Furthermore, Cakmak et al. (2010) showed that S. polyphyllae successfully completed its development and reproduced when feeding on S. feltiae and H. bacteriophora infective juveniles. Sancassania polyphyllae enters a hypopus stage under unfavorable conditions such as inadequate food supply and attaches to a larval scarab host. This ability may also enhance the mite’s efficacy as a predator of nematodes, especially if its host is killed by EPNs or if the host cadaver is invaded by saprophagous nematodes. Sancassania polyphyllae deutonymphs were found on P. fullo larvae collected from 5–10 cm depth of the soil in a strawberry field. When EPNs are applied, they can kill the larva in situ. Therefore, it is important to understand the spatial relationship of where the nematode-killed larva is in relation to where the infective juveniles emerge and the feeding activity of the mite. We hypothesized that the spatial distribution of the EPN infective juveniles in the presence or absence of nematode-killed insects in relation to the occurrence of S. polyphyllae will affect nematode survival in soil. Accordingly, we conducted laboratory studies to determine the potential negative effects this mite might have on the biological control of soil insect pests. Our objectives in this study were to (1) assess the response of S. polyphyllae adult mites to a nematode-killed insect on agar, (2) evaluate the predation by S. polyphyllae adult mites on S. feltiae infective juveniles from a nematode-killed insect on agar or in a soil column, and (3) assess predation efficiency of the mite on applied infective juveniles in a soil column. The findings obtained from this study will provide useful information on how this mite could affect the survival of the infective juveniles in soil which in turn would have an effect on natural or applied biological control.
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2. Materials and methods 2.1. Sources of insects, mites and nematodes The wax moth, Galleria mellonella (L.) (Lepidoptera: Pyralidae), was reared on an artificial medium that included honey, wheat bran, glycerol, soy flour, milk powder, dry yeast and honey bee wax in glass jar at 25 ± 4 °C in the laboratory. S. feltiae (Turkish isolate 09-31) was reared on last instar G. mellonella larvae in the laboratory (Kaya and Stock, 1997). The nematode-killed G. mellonella larvae were placed on a White trap, and the emerging infective juveniles from the cadavers were collected from the water and stored at 10 °C in tissue culture flasks (White, 1927). Infective juveniles were used within 2 weeks for the experiments. Larvae of P. fullo (Coleoptera: Scarabaeidae) were collected from strawberry fields in Aydin, Turkey and brought to the laboratory where the majority of them were stored at 20 °C. Four live P. fullo larvae with the deutonymphal stage of S. polyphyllae were killed, and the larvae were dissected in a petri dish (9 cm diameter) so that the various tissues were exposed to initiate the mite colony. Subsequently, S. polyphyllae was reared in petri dishes containing dissected tissues of previously frozen P. fullo larvae and maintained at room temperature (Karagoz et al., 2007). The larvae of Mediterranean fruit fly (medfly), Ceratitis capitata (Wiedemann) (Diptera: Tephritidae), were obtained from Bornova Plant Protection Research Institute in Izmir, Turkey as needed for the experiments. We used the medfly larvae for our experiments because it is one of the target insects for our research program, pupates in the soil, is susceptible to EPNs, provides a realistic target for EPNs, and produces infective juveniles that can be easily quantified in the soil in contrast to an unnatural host such as G. mellonella which never enters the soil. 2.2. Infection of medfly larvae with nematodes The medfly larvae were infected with S. feltiae in 24-well tissue culture plates. One last instar jumping larva (i.e., that was ready to pupate) was placed on the surface of each well that contained 0.5 g of sterilized, air-dried sand to which 30 infective juveniles in 60 ll distilled water were added. The well plate was covered with Parafilm to prevent the escape of the larvae. The larvae were checked for mortality caused by the infective juveniles daily as indicated by the light brown color of the cadaver. All nematode-killed larvae were transferred individually to White traps and checked three times a day for nematode development as the larval cuticle was transparent and nematode development could be determined. Depending on the experiment, 5- or 6-day-old nematode-killed medfly larvae were used and all experiments were conducted at room temperature (23 ± 1 °C). 2.3. Response of medfly cadaver to mites in a petri dish To assess the response of 10 adult female mites to (1) a 5-dayold nematode-killed medfly larva or (2) a fresh, frozen-killed larva ( 20 °C for 30 min) or (3) a piece of bamboo the same length and width as a medfly larva, the following experiment was conducted. The nematode-killed larva, frozen-killed larva and bamboo were placed near the edge of an experimental arena (100 15 mm plastic petri dish containing 20 ml of 2% water agar), equidistant apart and 3.5 cm from the center of the dish. Then 10 S. polyphyllae adult female mites were released onto the agar surface in the center of the arena, and data on the number of mites at a cadaver or bamboo were taken every 20 min up to 60 min. Ten petri dishes were prepared for the experiment with each plate serving as a replicate, and the experiment was conducted twice.
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2.4. Medfly cadaver, emerging nematodes, and mites in a petri dish For each experimental round, 40 medfly larvae were used at 6days post-nematode infection when the first emergence of infective juveniles from the cadavers was evident. Each nematodekilled medfly larva with emerging infective juveniles was placed in the center of an experimental arena (100 15 mm plastic petri dish containing 20 ml of 2% water agar). Forty petri dishes were prepared for the experiment. Twenty dishes served as the treatment with 10 S. polyphyllae females being released on the agar surface, and 20 dishes served as controls without mites and containing only the medfly larval cadaver. To standardize the experiment, the petri dish with the mites was paired with a control dish with infective juveniles emerging from the larva at about the same time. Three days later, the mites and the larval cadaver were removed from the petri dishes, and the infective juveniles on the agar surface of each dish were rinsed off with distilled water into a 100-ml beaker, and all nematodes in the suspension were counted at 40 magnification with a dissecting microscope. This experiment was conducted twice for a total of 40 petri dishes each for the treatment and control, respectively.
initial set up, whereas in the second experiment, the soil column was destructively examined 10 days after set up. For both experiments, there were 10 tubes (replicates) for each treatment which were kept in the dark, and each experiment was conducted twice for a total of 20 replicates. After 4 or 10 days, the soil in the tubes was handled as in the soil column experiment with the medfly cadaver. For the 4- and 10-day sampling, the experimental design was the same as described above in the soil column experiment with the medfly larval cadaver except that rather than placing a larval cadaver with emerging infective juveniles, 500 S. feltiae infective juveniles in 200 ll of water were pipetted onto the soil surface. In addition, rather than placing 50 adult female mites in the soil column, 10 adult female mites were used. For the 4-day sample, treatment A had the infective juveniles and mites at the soil surface, treatment B had the infective juveniles at 2 cm level of soil and the mites on the top surface, and treatment C had the infective juveniles on the top surface and the mites were at the 2 cm level. Control tubes were prepared for each treatment with 500 infective juveniles but no mites were added. For the 10-day sample, only treatment B and C as described above was prepared.
2.5. Medfly cadaver and mites in a soil column Fifty-ml plastic centrifuge tubes (30 115 mm) (Ratilabo, Karlsruhe, Germany) were used as soil columns and filled with 20 g of moistened (10% w/w) soil (69.4% sand, 30.6 % silt, pH 8.3) collected from a strawberry field in Aydin (see steps A–C below for details). Prior to being moistened, the soil was pasteurized and air-dried for over 2 weeks. The total height of the soil column with 20 g of soil was 9 cm. Three different treatments were employed in this experiment depending on the placement of a 6day-old medfly larval cadaver with emerging infective juveniles and 50 adult female mites in the soil column. The treatments were as follows. A. Twenty g of moistened soil was added into the centrifuge tube and the mites and one medfly larval cadaver with emerging nematodes were placed on the soil surface (referred to as A+). B. Four g of moistened soil was added into the centrifuge tube (2 cm of soil in the tube) and one medfly larval cadaver was placed on the soil surface. The remaining 16 g of the moistened soil was added into the tube and the mites were released to the soil surface resulting in a 7-cm distance between the medfly cadaver and mites (referred to as B+). C. The same protocol as ‘‘B” was followed except that placement of the medfly cadaver and mites were reversed. That is, the mites were placed at the 2 cm level and the medfly larval cadaver was placed on the soil surface (referred to as C+). Twenty tubes were used for each treatment. Control tubes were prepared for each treatment with a medfly larval cadaver with emerging infective juveniles but no mites were added (Referred to as A , B and C , respectively). All experiments were conducted twice and kept in the dark. After 10 days, the soil in each tube was washed and sieved (53-lm mesh) to remove soil particles and obtain the infective juveniles. The different mite stages in the soil were noted but not counted. The collected nematodes were counted at 40 magnification with a dissecting microscope as described for the agar substrate counts. 2.6. Infective juveniles and mites in a soil column Two different experiments were conducted. In the first experiment, the soil column was destructively sampled 4 days after the
2.7. Statistical analysis The data were analyzed using analysis of variance (ANOVA). Means were compared at the P = 0.05 level, and Tukey’s test was used to separate means (SPSS, 2004). The means ± SEM are provided below. 3. Results 3.1. General observations When last instar medfly larvae were individually exposed to 30 infective juveniles in 24-well tissue culture plates, 92% of them were infected by the nematodes, and the first infective juveniles began emerging from the cadavers at 6 days after infection. The total number of infective juveniles emerging from a C. capitata larva in a White trap averaged 2500 ± 510 over a 6-day period. However, in our experiments, less infective juveniles emerged because the duration of the experiments were less than 6 days. In the petri dish agar experiment, the mites fed on intact medfly larval cadavers with nematodes and produced eggs. In the soil column experiments, we recovered adult mites, eggs and larvae from the soil with the infective juveniles only experiment after 4 days, whereas we recovered adults, eggs, and immature mites (larvae, protonymphs and tritonymphs) from the soil with the medfly larval cadaver after 10 days. We did not quantify the number of mites and the various stages recovered because our focus was on counts of the infective juveniles. 3.2. Response of medfly cadaver to mites in a petri dish At 20 min after exposure, the average number of mites at the nematode-killed medfly larva was 1.4 ± 0.4, at the frozen larva was 0.5 ± 0.2, and at the bamboo was 0.1 ± 0.1. At 40 min, the average number of mites at the nematode-killed medfly larva was 2.6 ± 0.5, at the frozen larva was 0.9 ± 0.3, and at the bamboo was 0.1 ± 0.1. Significant differences were seen between the bamboo and the nematode-killed medfly larva at 20 (F = 4.31; df = 2, 27; P < 0.05) and 40 (F = 6.86; df = 2, 27; P < 0.01) min. At 60 min exposure, the average number of mites at the nematode-killed medfly larva was 3.1 ± 0.6, at the frozen larva was 1 ± 0.2, and at the bamboo was 0. Statistical analysis showed that significantly
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10 days, the numbers of infective juveniles recovered were significantly less in soil columns with the mites than without the mites (P < 0.001, Fig. 1). In all cases, the placement of the mites or the larval cadaver did not affect the results. That is, the average number of recovered infective juveniles was <30 infective juveniles when mites were present, whereas the average number of recovered infective juveniles was >375 infective juveniles when the mites were absent. In treatment A (F = 15.04; df = 1, 38; P < 0.001) when the larval cadaver and mites were both placed on the soil surface, an average of <1 infective juvenile was recovered, whereas >340 infective juveniles were recovered from the soil column without mites. In treatment B (F = 22.98; df = 1, 42; P < 0.001) and C (F = 84.87; df = 1, 33; P < 0.001) when the larval cadaver and mites were separated from each other in the soil column, significantly fewer infective juveniles were recovered from the soil column without mites. When the three experimental groups with mites plus cadavers with nematodes (i.e., A+, B+ and C+) were compared, significant differences were observed only between cadaver and mites placed on the surface (A+) and cadaver placed at 2 cm level and the mites placed on the soil surface (B+) (F = 5.19; df = 2, 63; P < 0.05).
more mites were found on the nematode-killed medfly larva than at the freeze-killed medfly larva or at the piece of bamboo (F = 9.87; df = 2, 27; P < 0.001) at 60 min. There was no significant difference between the frozen larva and the bamboo. 3.3. Medfly cadaver, emerging nematodes, and mites in a petri dish On the agar media, significant differences were observed in the numbers of nematodes recovered between larval cadaver with mites (33 ± 8 infective juveniles) and larval cadaver without mites (857 ± 100 infective juveniles) (F = 68.03; df = 1, 82; P < 0.001) after 3 days. Based on what was recovered from the control agar plate, 10 adult female mites consumed 96% of the emerged infective juveniles from the C. capitata cadaver. We observed that mites were around and near the cadaver and attacked the emerging infective juveniles. 3.4. Medfly cadaver and mites in a soil column In all treatments (i.e., A–C), when the medfly larva with nematodes was placed into the soil column with mites or no mites for
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Fig. 1. Mean number ± standard error of consumed Steinernema feltiae infective juveniles that emerged from a Ceratitis capitata larva by Sancassania polyphyllae in a soil column. Treatment A = larval cadaver on the soil surface (solid bars) with A+ being mites placed at the surface and A with no mites. Treatment B = larval cadaver at 2 cm from the bottom (open bars) with B+ being the mites placed on the soil surface and B with no mites. Treatment C = larval cadaver on the soil surface (patterned with black and white squares) with C+ being mites placed at 2 cm from the bottom and C with no mites. Different capital letters (X and Y) above the bars indicate significant differences among the treatments with mites. Different lowercase letters (a and b) above the bars indicate significant differences between each treatment and their control.
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Treatments Fig. 2. Mean number ± standard error of consumed Steinernema feltiae infective juveniles by Sancassania polyphyllae in a soil column after 4 days incubation. Treatment A = infective juveniles placed on the soil surface (solid bars) with A+ being mites placed at the surface and A with no mites. Treatment B = infective juveniles placed at 2 cm from the bottom(open bars) with B+ being the mites placed on the soil surface and B with no mites. Treatment C = infective juveniles placed on the soil surface (patterned with black and white squares) with C+ being mites placed at 2 cm from the bottom and C with no mites. Different capital letters (X, Y and Z) and different lowercase letters (a and b) above the bars indicate significant differences between the treatments and their control.
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3.5. Infective juveniles and mites in a soil column When the infective juveniles alone were placed into the soil column with mites for 4 days, our results showed that the recovery of infective juveniles from soil differed depending on where the infective juveniles were placed in relation to the mites (Fig. 2). For example, in treatment A+ when the infective juveniles and mites were placed on the surface of the soil column together, significantly fewer infective juveniles were recovered from the column that had mites present than the control group that had no mites (F = 87.02; df = 1, 26; P < 0.001) (Fig. 2). Similarly, in treatment B+ when the infective juveniles were placed at 2 cm from the bottom of the soil column and the mites were on the soil surface, significantly fewer infective juveniles were recovered compared to the control group that had no mites (F = 7.52; df = 1, 28; P < 0.05). However, in treatment C+ when the placement of mites and infective juveniles were reversed, no significant difference was observed between mites present and the control group (F = 0.41; df = 1, 26; P > 0.05).When all three experimental groups in which mites and infective juveniles were together (i.e., A+, B+, C+) in the soil column were compared, statistically significant differences were seen among all groups (indicated as X, Y, and Z above the bars) (F = 37.90; df = 2, 57; P < 0.001) (Fig. 2). The same experimental design was prepared with the mites and nematodes incubated for 10 days in the soil column as above, but it only had treatments B and C. The average numbers of infective juveniles were 168 and 158 for B+ and C+ at the end of the 10day incubation period, respectively, and no significant difference was observed between B+ and C+ groups (F = 0.52; df = 1, 18; P > 0.05). However, significantly fewer infective juveniles were recovered from B+ compared with B (F = 24.29; df = 1, 18; P < 0.001 and from C+ compared with C (F = 31.53; df = 1, 18; P < 0.001 (Fig. 3).
4. Discussion
Number of infective juveniles
We demonstrated that the spatial distribution of EPN infective juveniles in the presence or absence of nematode-killed insects in relation to the occurrence of S. polyphyllae affected nematode survival in soil. Accordingly, we showed that S. polyphyllae fed more on infective juveniles emerging from a cadaver in soil than on infective juveniles in the absence of a cadaver (e.g., when infective juveniles are applied as a biopesticide). Thus, our hypothesis, in general, was correct in that the spatial distribution of the infective juveniles
was affected depending upon whether S. polyphyllae was present or absent or what the source of the infective juveniles was. In the initial experiment, we found that significantly more female adult mites of S. polyphyllae were found associated with a nematode-killed medfly larva than to a fresh, freeze-killed larva or a piece of bamboo. The data suggested that a chemical(s) or odor(s) from the nematode-killed larva was produced and served as an attractant for the mites. In the second and third experiments, our data showed that the female mites consumed most of the infective juveniles emerging from a medfly cadaver on an agar substrate and in soil. In the soil column that had nematode-killed medfly larvae with emerging infective juveniles, our data suggest that the 50 female mites and their immature progeny [that is, the protonymphs and tritonymphs are known to feed on the infective juveniles of S. feltiae in the laboratory (see Cakmak et al., 2010)] found the cadaver and probably congregated around it to consume the emerging nematodes over a 10-day period. In fact, the mites were efficient predators of the infective juveniles regardless of whether the cadaver was placed on the top or bottom of the soil assay chamber. These findings suggest that S. polyphyllae may have a negative impact on the biological control of soil insect pests using cadaver containing nematodes (see Shapiro and Lewis, 1999) where S. polyphyllae or other nematophagous mites occur. For example, the application of cadavers with infective juveniles shows considerable promise because the emerging nematodes of some species are more infectious than with infective juveniles applied with water (Shapiro and Lewis, 1999; Shapiro-Ilan et al., 2003). However, if S. polyphyllae is present, it could be an efficient predator of the infective juveniles as they emerged from the cadaver in the soil environment. In addition, the mites may feed on the cadaver before the infective juveniles emerge rendering the cadaver unsuitable for nematode development. Because the mites are small and can easily move through the small soil pores, they may actually cue in on the cadaver and consume the infective juveniles as they emerge from the cadaver as was observed in our agarbased experiment. Thus, if these mites are abundant and near a nematode-killed insect, they may negate the advantages of using a cadaver application. In the fourth experiment, we found that when the free-living infective juveniles alone were placed with 10 mites over a 4-day period, the mites were not as efficient at finding the nematodes compared with the larval cadaver treatments. On the other hand, placing the mites and the free-living infective juveniles in proximity to each other did have a significant negative effect on the numbers of infective juveniles. If the infective juveniles and mites were
500 400
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Treatments Fig. 3. Mean number ± standard error of consumed Steinernema feltiae infective juveniles by Sancassania polyphyllae in a soil column after 10 days incubation. Treatment B = infective juveniles placed at 2 cm from the bottom (solid bars) with B+ being the mites placed on the soil surface and B with no mites. Treatment C = infective juveniles placed on the soil surface (open bars) with C+ being mites placed at 2 cm from the bottom and C with no mites. Same capital letters (X) above the bars indicate nonsignificant differences between the treatments with mites. Different lowercase letters (a and b) above the bars indicate significant differences between the treatments and their control.
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spatially separated, the predation rate of the infective juveniles by the mites was reduced or non-existent. These data suggest that the mites were not as efficient at finding the infective juveniles when they were separated from each other in the soil lending support to the idea that the mites were cueing in on the cadaver as a food resource. Although our laboratory study suggests that the presence of S. polyphyllae or a mite species with similar feeding habits has the potential for having a negative effect on biological control by EPN species of soil insect pests, there may be situations where its effect on biological control may be mitigated. From the literature, we know that mites in the genus Sancassania (=Caloglyphus) are biologically diverse group in the family Acaridae and that some species are associated with various arthropods including Coleoptera, Hymenoptera, Orthoptera, Myriapoda and terrestrial Crustacea (Klimov et al., 2004). Sancassania species feed on insect cadavers and microorganisms associated with the cadavers (Krantz, 1978; Krantz and Walter, 2009) as well as on nematodes (Sell, 1988; Karagoz et al., 2007). Our study was done in the absence of other nematode species, and the presence of other nematode species may mitigate the effects of S. polyphyllae. For example, Sell (1988) showed that a Sancassania sp. fed on a plant-parasitic nematode species, and this type of non-selective feeding by S. polyphyllae may allow survival of EPNs. In addition, Ekmen et al. (2010) showed that S. polyphyllae had a preference for tissues of Polyphylla fullo over that of live S. feltiae or H. bacteriophora infective juveniles. If there are a number of insect cadavers in the soil, S. polyphyllae will feed on the cadavers, but this may result in a significant population increase of S. polyphyllae. Once the insect cadavers are consumed, the mites will need alternative food sources which could potentially reduce EPN numbers if they are in the same area. In conclusion, our study strongly suggests that chemicals associated with insect cadavers influence S. polyphyllae behavior. Therefore, they may detect C. capitata cadavers easily in the soil column and consume the infective juveniles as they emerge from the cadavers. On the other hand, the mites had difficulty in finding and preying on the infective juveniles in the absence of a cadaver, especially when the mites and infective juveniles were separated spatially. Although we have gained a greater understanding of S. polyphyllae mites as prey of S. feltiae infective juveniles, our knowledge about the impact of this mite on nematode populations under field conditions remains unknown. Our future studies will involve field studies where the mite naturally occurs and assess if their presence will have significant effect on biological control of soil insect pests. Acknowledgments We thank Baris Gulcu, Derya Asici and Cem Demirtas for their assistance in the experiments. References Baur, M.E., Kaya, H.K., 2001. Persistence of entomopathogenic nematodes. In: Baur, M.E., Fuxa, J.R. (Eds.), Southern Cooperative Series Bulletin, p. 26,
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