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Strong bottom-up and weak top-down effects in soil: nematode-parasitized insects and nematode-trapping fungi
Soil Biology & Biochemistry 37 (2005) 1011–1021 www.elsevier.com/locate/soilbio
Strong bottom-up and weak top-down effects in soil: nematode-parasitized insects and nematode-trapping fungi B.A. Jaffeea,*, D.R. Strongb a Department of Nematology, University of California at Davis, 1 Shields Avenue, Davis, CA 95616, USA Department of Evolution and Ecology, University of California at Davis, 1 Shields Avenue, Davis, CA 95616, USA
b
Received 25 March 2004; received in revised form 4 May 2004; accepted 7 May 2004
Abstract The soils of the Bodega Marine Reserve (BMR, Sonoma County, California) contain many nematode-trapping fungi and many ghost moth larvae parasitized by entomopathogenic nematodes. The current study determined whether these nematode-parasitized moth larvae, which can produce very large numbers of nematodes, enhanced the population densities of nematode-trapping fungi and whether the fungi trapped substantial numbers of nematodes emerging and dispersing from moths. Wax moths were used in place of ghost moths because the former are easier to obtain. When nematode-parasitized moth larvae were added to laboratory microcosms containing BMR field soil, the population densities of four nematode-trapping fungi increased substantially. The greatest increase in population density was by Arthrobotrys oligospora, which uses adhesive networks to capture nematodes. A. oligospora population density increased about 10 times when the added moth larvae were parasitized by the nematode Heterorhabditis marelatus and about 100 times when added moth larvae were parasitized by the nematode Steinernema glaseri. Other trapping fungi endemic to the soil and enhanced by nematode-parasitized moth larvae included Myzocytium glutinosporum, Drechslerella brochopaga, and Gamsylella gephyropaga, which produce adhesive spores, constricting rings, and adhesive branches, respectively. The data suggest that the previously documented abundance and diversity of nematode-trapping fungi in BMR soil can be explained, at least in part, by nematode-parasitized insects, although that inference requires further studies with ghost moths. The strong bottom-up enhancement of nematode-trapping fungi was not matched by a strong top-down suppression of nematodes, i.e. the fungi trapped fewer than 30% of dispersing nematodes. q 2005 Elsevier Ltd. All rights reserved. Keywords: Arthrobotrys oligospora; Drechslerella brochopaga; Entomopathogenic nematode; Gamsylella gephyropaga; Heterorhabditis marelatus; Myzocytium glutinosporum; Nematode-trapping fungi; Soil food web; Steinernema glaseri
1. Introduction On the coastal headlands of the Bodega Marine Reserve (BMR, Sonoma County, California), nematode-trapping fungi are diverse and often abundant. The soil contains at least 12 species, and overall population densities average 50 propagules/g of soil but can exceed 600 propagules/g of soil. The most common and abundant nematode-trapping fungus at BMR is Arthrobotrys oligospora (Jaffee et al., 1996). Why this soil contains so many nematode-trapping fungi is unclear, but the ghost moth larva and its nematode * Corresponding author. Tel.: C1 530 752 0862; fax: C1 530 752 5809. E-mail address: [email protected] (B.A. Jaffee). 0038-0717/$ - see front matter q 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.soilbio.2004.05.026
parasite may be important. At BMR, ghost moth larvae consume bush lupine roots (Strong et al., 1995). In some years, the larvae kill entire stands of this dominant shrub, but in other years they are suppressed by the entomopathogenic nematode, Heterorhabditis marelatus (Strong et al., 1996, 1999; Preisser, 2003). Like other entomopathogenic nematodes, H. marelatus is an obligate parasite or pathogen of insects (Liu and Berry, 1996; Stock, 1997). Infective juveniles of H. marelatus invade ghost moth larvae in soil or those that have penetrated lupine roots, and once within the insect body cavity, the nematodes release a symbiotic bacterium which kills the insect. The nematodes, which consume both the multiplying bacteria and the degrading insect, then increase in number. Each parasitized moth larva can produce few or many nematodes depending on larval
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size; a large larva can produce more than 400,000 juvenile nematodes (Strong et al., 1996, 1999). When the resource is depleted, the newly-produced juvenile nematodes disperse from the insect cadaver into the soil, where they search for a new host. By potentially producing and releasing many nematodes in a small volume of soil, nematode-parasitized ghost moth larvae may help explain the abundance of nematode-trapping fungi at BMR (Strong, 1999). While data about how the nematode and ghost moth affect trapping fungi are nonexistent, data about how trapping fungi affect H. marelatus do exist but are somewhat conflicting. The more common trapping fungi at BMR capture and consume H. marelatus in vitro (Koppenho¨fer et al., 1996), but their distribution in the field was not correlated with lupine death, as it should be if the fungi suppressed entomopathogenic nematodes and thereby protected ghost moth larvae (Jaffee et al., 1996). In contrast, soil microcosm experiments indicated that the more common species apparently trapped substantial numbers of juvenile H. marelatus as they moved through soil to insect larvae (Koppenho¨fer et al., 1996, 1997). None of these studies measured trapping as the nematodes emerged and dispersed from the nematode-parasitized insect. The current study focuses on the nematode-parasitized insect larva and the surrounding soil. In this soil, which we will not call the ‘larvosphere’, we predicted that emerging nematodes would greatly enhance the population densities of trapping fungi and that the fungi in turn would trap many emerging nematodes. In other words, we predicted strong bottom-up effects of emerging nematodes on fungi and strong top-down effects of trapping fungi on emerging nematodes. Experiments testing these predictions are described in this paper.
Steinernema glaseri, was also used but was not isolated from BMR soil and has been maintained for over 10 years by repeated rearing in wax moth larvae. The infective juvenile of S. glaseri is about twice as long and wide as the infective juvenile of H. marelatus (Poinar, 1978; Liu and Berry, 1996). 2.2. Soil BMR is on the Pacific Coast of California, 80 km north of San Francisco. Soil was collected from a location at BMR called Mussel Point Tip (Jaffee et al., 1996) on 10 June and 18 September 2003. Soil was collected 5–20 cm deep adjacent to lupines. It was sieved (2 mm openings), stored at 10 8C, and used within 30 days (all trials of experiments 1 and 2 and trial 3 of experiment 3) or 68 days (trials 1 and 2 of experiment 3). Mussel Point Tip soil is a coarse sandy loam (Miller, 1972) with a pH in water of 4.9 and with an organic matter content of 4.9% (Jaffee et al., 1996). In the batch of soil collected on 10 June 2003, total nitrogen and carbon contents were 0.19 and 1.83%, respectively, with 4.5 ppm NH4 and 5.9 ppm NO3. A water release curve was determined because we suspected that water potential could greatly affect the results and because a water release curve provides valuable information on the proportion of soil pores that are filled with water or drained. For experiment 3, natural enemies of nematodes were removed for one treatment by heating the soil as follows. A 1-l beaker containing soil was placed in a 75 8C oven. Once the center of the soil mass increased to 70 8C, the oven temperature was reduced to 70 8C. After 2 h at 70 8C, the soil was removed and cooled to room temperature. 2.3. Nematode-parasitized insects
2. Methods 2.1. Nematode isolates One isolate of H. marelatus (referred to here as H. marelatus-k) was obtained from H. K. Kaya. H. marelatus-k was isolated from BMR soil and has been maintained for over 7 years by repeated rearing in Galleria mellonella (wax moth) larvae (Kaya and Stock, 1997). A newer isolate, H. marelatus-b, was used to determine whether repeated rearing of H. marelatus affected how nematophagous fungi responded to the nematode. H. marelatus-b was obtained in July 2003, by baiting BMR soil with wax moth larvae (Kaya and Stock, 1997) and incubating those larvae that died and turned orange in White traps (White, 1927; Strong et al., 1996; Kaya and Stock, 1997); in contrast to H. marelatus-k, H. marelatus-b was reared on wax moth larvae only once (trials 1 and 2 of experiment 3) or twice (trial 3 of experiment 3) after its isolation from the field. The entomopathogenic nematode,
Although ghost moths rather than wax moths are the major hosts of H. marelatus at BMR, wax moth larvae were used because ghost moth larvae are only available several months each year (Strong et al., 1996). In contrast, healthy and uniform wax moth larvae are easily obtained year round. Healthy, last-instar wax moth larvae (about 2 cm long; 220–260 mg fresh mass per larva; purchased from Rainbow Mealworms, Inc., Compton, CA) were placed in a 10-cm diameter Petri dish (10 larvae per dish); the dish contained a 10-cm diameter filter paper which had received 200–500 H. marelatus or S. glaseri in a 1-ml suspension. After 4 days at 20–22 8C, larvae that were dead and showing characteristics of nematode parasitism were added to soil: those exposed to H. marelatus were orange (Strong et al., 1996), whereas those exposed to S. glaseri were grey (Kaya and Stock, 1997). In this paper, ‘nematodeparasitized insects’ refers to moth larvae that have been killed by entomopathogenic nematodes and that contain large numbers of entomopathogenic nematodes.
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2.4. Experimental arenas The arena was based on the original ‘White trap’ (White, 1927) and the ‘modified White trap’ (Kaya and Stock, 1997). The original trap was designed to collect and quantify nematode parasites of mammals as they emerge and disperse from nematode-infested feces, and the modified trap was designed to collect and quantify entomopathogenic nematodes as they emerge and disperse from nematode-parasitized insects. Both traps use a smaller dish that sits in a larger, water-filled dish. The smaller dish contains the feces or insect-parasitized nematodes, and the nematodes are collected in the surrounding water (hereafter called the ‘moat’). To assemble an arena, 40 g soil (dry weight equivalent) at 14% water content (g water per 100 g soil) was placed in the bottom of a small plastic Petri dish (8.7-cm diameter and 1.5 cm deep). The soil was tamped; the height and bulk density of the soil column were about 5 mm and 1.3 g/cm3. Soil water content was adjusted as indicated for each experiment, and one or no nematode-parasitized wax moth larva was placed on the surface of the soil in the center of the dish. The small dish, which was uncovered, was placed in a large plastic Petri dish (15 cm diameter and 2.5 cm deep). Sufficient distilled water (45 ml) was added to the large dish so that the water covered the bottom of the large dish and rose part way up the sides of the small dish. A rubber band (76!1.6 mm) placed beneath the smaller dish prevented it from sliding to the side of the larger dish and becoming filled with water. As in the White trap, the moat was a key feature of our experimental arenas. To reach the safety of the moat, juveniles emerging and dispersing from insect cadavers had to traverse at least 5 cm of soil that contained nematodetrapping fungi and other natural enemies (unless the soil had been heat-treated). Nematodes in the moats were identified and counted, as described for each experiment. The soil in the inner dish was used to quantify fungus population density, as described for experiment 1 below. Nematodes and other organisms in the soil were not quantified. Arenas were assembled and grouped by replicate, sealed in plastic bags (one replicate per bag) with moist paper towels, and placed in a 20 8C incubator without light. After 7–10 days, all replicates were examined for weed seedlings, which were removed to prevent nematodes from moving up the weed stems rather than moving through the soil. Other details are described for each experiment. 2.5. Experiment 1 Experiment 1 examined the numerical response of nematode-trapping fungi to nematode-parasitized insects in BMR soil (i.e. the bottom-up effect of parasitized insects on trapping fungi); the nematode was H. marelatus-k. Soil water content was selected as a variable because we
Fig. 1. Water release curve of soil from Mussel Point Tip, the Bodega Marine Reserve.
suspected that the most abundant trapping fungus in BMR soil, A. oligospora (Jaffee et al., 1996), might be more active at high soil water potentials. Soil water content was adjusted to 14, 20, 26, 32, or 38% by adding the appropriate volume of distilled water to the soil surface; 14% was considered a moderate water content and 38% was considered wet; at 14%, the larger soil pores were drained and smaller pores were water-filled, whereas at 38%, all but the largest soil pores were water-filled (Fig. 1). Experiment 1 was ended 35 days after the moth larvae were exposed to H. marelatus (31 days after the arenas were assembled and larvae were placed on the soil), and the following were quantified: nematodes in the moat, A. oligospora sporulation on the soil surface, the population density of nematode-trapping fungi in the soil, and soil water content. To count nematodes, the water and nematodes in the moat were placed in a 1-l beaker, the volume was increased to 800 ml, the suspension was mixed, 1 ml was placed in each of two small dishes, and the nematodes were counted with the aid of a dissecting microscope. To quantify the nematode-trapping fungi in the soil, the soil was removed from each arena, placed in a plastic bag, and mixed with a spatula, and 10 g (dry wt equivalent) was placed in a sterile 125-ml flask. The volume of each flask was increased to 50 ml with sterile distilled water, and the suspension was dispersed on a wrist action shaker for 9 min. A 10-fold dilution series with three dilutions was then prepared, and 0.1 ml of each dilution was placed on each of five quarter-strength corn meal agar plates. About 1000 bait nematodes (healthy S. glaseri) were added to each plate. After 7, 14, and 21 days, each plate was examined for nematode-trapping fungi, and a computer program was used to determine the most probable number (Klee, 1993; Jaffee, 2003). The remaining soil from each arena was used to determine water content; soil mass was compared before and after drying.
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Experiment 1 was performed twice (trials 1 and 2). There were five replications for each treatment in each trial. 2.6. Experiment 2 Experiment 2 was identical to experiment 1 except that the wax moths were parasitized by S. glaseri rather than H. marelatus-k. We decided to examine the effect of S. glaseri because H. marelatus-parasitized insects did not stimulate extensive A. oligospora sporulation on the soil surface as we had expected based on preliminary experiments. Experiment 2 was also performed twice (trials 1 and 2). There were five replications for each treatment in each trial. 2.7. Experiment 3 Experiment 3 had three objectives. One was to quantify how trapping fungi and other natural enemies reduced dispersal of nematodes from parasitized insects. A second objective was to directly compare the effects of S. glaseriand H. marelatus-parasitized wax moth larvae on the fungi. Although experiments 1 and 2 indicated that nematodetrapping fungi responded more strongly to S. glaseri- than to H. marelatus-parasitized wax moth larvae, these experiments were independent, and differences may have reflected other uncontrolled effects. A third objective was to compare the effects of two H. marelatus isolates; one isolate had been cultured in the laboratory for many years and the other had not. Arenas contained heat-treated and nonheat-treated soil, which was packed at 14% water content and then increased to 32% as described for experiment 1. Each arena received one wax moth larva parasitized with either H. marelatus-k, H. marelatus-b, or S. glaseri. Other procedures were as described in experiments 1 and 2, except that experiment 3 was ended after 45 rather than 35 days and nematodes were counted four times rather than once. We extended experiment 3 because H. marelatus emerged from the wax moth larvae about 10 days after S. glaseri in experiments 1 vs 2 (data not shown), and we suspected that the trapping fungi might require more time to respond to emerging H. marelatus. We removed nematodes from the moat on four dates to prevent them from being parasitized and degraded by A. oligospora and Myzocytium glutinosporum. Nematodes in the moat were collected and counted on day 18, 25, 32, and 45 (moat water was replaced each time except the last). Observations in experiments 1 and 2 suggested that moth larvae degraded faster when parasitized by S. glaseri than H. marelatus. In experiment 3, degradation of each moth larva was subjectively assessed from 0 (no degradation) to 100% (complete degradation). Experiment 3 was performed three times (trials 1–3). There were six replications for each treatment in each trial.
2.8. Statistical analyses After transformation (log for numerical counts and arcsine for percentages), data were subjected to an analysis of variance (SAS for Windows, version 8.02; SAS Institute, Inc., Cary, NC) to determine whether treatment affected (P!0.05) fungus population densities in all three experiments and nematode population densities in experiment 3. In experiments 1 and 2, soil water content was considered a categorical variable and a continuous variable in separate analyses. In experiment 3, partial Spearman correlation analysis was used to determine whether nematode population density and fungus population density were related; a separate analysis was done for each combination of nematode and fungus and with data from both heat-treated and nonheat-treated soil or with data only from nonheattreated soil. The data for detection frequency were not statistically analyzed because, in our view, there were too few replications to support statistical inferences. Note that each trial produced only one value per fungus for frequency of detection (a percentage or proportion of replicates that were positive in that trial). Thus, there were only three values for detection frequency for each fungus in experiment 3.
3. Results 3.1. Experiment 1 (with H. marelatus-k) Of the nematode-trapping fungi detected (Table 1), A. oligospora and M. glutinosporum were most numerous. A. oligospora was detected in most arenas, even when wax moth larvae were absent and was detected in all arenas except one when H. marelatus-parasitized wax moth larvae were present (Fig. 2). A. oligospora population density was 12 times greater in arenas with wax moths but was unaffected by soil water content (Fig. 3A). Averaged across trials and soil water content, A. oligospora population Table 1 Fungi detected in this study Fungusa
Infective structuresb
Arthrobotrys oligospora Myzocytium glutinosporum A. eudermata A. musiformis A. paucispora Gamsylella gephyropaga Drechslerella brochopaga
Adhesive networks Adhesive sporesc Adhesive networks Adhesive networks Adhesive networks Adhesive branches Constricting rings
a
These generic designations follow Scholler et al. (1999). Once the infective structure has firmly contacted the nematode, all of these fungi directly penetrate the nematode cuticle and then grow through and consume the nematode body. c M. glutinosporum produces zoospores that usually encyst near the parasitized nematode. Each encysted spore forms a bud which adheres to the surface of passing nematodes (Barron, 1976). b
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Fig. 2. Detection frequency of nematophagous fungi in arenas of experiments 1 and 2. Each arena contained nonheat-treated soil and no wax moth larva or a wax moth larva parasitized with Heterorhabditis marelatus isolate k (experiment 1) or Steinernema glaseri (experiment 2). Wax moth larvae were exposed to entomopathogenic nematodes on day 0 and placed in arenas on day 4; experiments 1 and 2 were ended on day 35. Detection frequency indicates the proportion of 10 replicates (trials 1 and 2 combined) with the indicated fungus. Open bars and striped bars indicate data from arenas without and with nematodeparasitized larvae, respectively.
density was 207G39 propagules/g soil (meanG1 standard error) or 17G2 propagules/g soil in arenas with or without H. marelatus-parasitized wax moth larvae. In contrast to A. oligospora, M. glutinosporum was not detected unless H. marelatus-parasitized wax moth larvae were present (Fig. 2); and even in the presence of H. marelatus-parasitized wax moth larvae, M. glutinosporum was detected in only 13 of 48 total arenas and never when soil water was at the driest level tested (Fig. 3B). When detected, M. glutinosporum was abundant: averaged across trials and soil water contents, and including arenas both with and without M. glutinosporum (nZ48), M. glutinosporum population density was 582G245 or 0G0 propagules/g soil in arenas with or without
H. marelatus-parasitized wax moth larvae. Including only those arenas in which M. glutinosporum was detected (nZ13), the mean population density of M. glutinosporum was 2148G768 propagules/g soil. Detection of fungi that form adhesive branches (Gamsylella gephyropaga) or constricting rings (Drechslerella brochopaga) increased in the presence of H. marelatusparasitized moth larvae (Fig. 2), but population densities (averaged over all water levels) remained less than 20 propagules/g soil (data not shown). Detection of fungi other than A. oligospora that form adhesive networks (A. eudermata, A. musiformis, and A. paucispora) was either unaffected or possibly suppressed by wax moth larvae (Fig. 2), and population densities of these fungi (averaged
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Fig. 3. Population densities of Arthrobotrys oligospora and Myzocytium glutinosporum in nonheat-treated soil as affected by the presence or absence of nematode-parasitized insects and soil water content. Experiment 1 (A and B) was done with Heterhorhabditis marelatus isolate k, and experiment 2 (C and D) was done with Steinernema glaseri. Solid bars and open bars indicate data from arenas without and with nematode-parasitized larvae, respectively. Values are the meansC1 standard error of 10 replicates (trials 1 and 2 combined).
over all water levels) was less than 2 propagules/g soil (data not shown). The total population density of nematode trapping fungi was 812 or 23 propagules/g soil in arenas with or without H. marelatus-parasitized larvae. In the former case, M. glutinosporum made up 72% and A. oligospora made up 25% of the total. We frequently observed A. oligospora conidiophores and conidia, which are distinctive (Cooke and Godfrey, 1964), on the soil surface and the insect cadaver. To quantify this sporulation, the soil surface and cadaver were examined with reflected light and a dissecting microscope (35–70!magnification), and sporulation was rated on a scale from 0 to 3 (0, no A. oligospora conidiophores; 1, fewer than 10 A. oligospora conidiophores per arena; 2, 10–75 A. oligospora conidiophores per arena; 3, O75 A. oligospora conidiophores per arena). In the presence of H. marelatus-parasitized wax moth larvae, the percentage of arenas rated 0, 1, 2, or 3 was 54, 35, 10, and 0%, respectively. Conidiophores like those produced by A. eudermata, G. gephyropaga, or D. brochopaga, were infrequently observed. None of these structures was observed in arenas without nematodeparasitized insects. Large numbers of H. marelatus (143,000G7000) dispersed through soil and into the moats of arenas receiving H. marelatus-parasitized wax moths, and this number was
unaffected by soil water content or trial; no H. marelatus were found in moats from arenas lacking wax moth larvae. Soil water content was less at the end of the experiment than at the start. Averaged over both trials, soil water content decreased from 14.0 to 11.2%, 20.0 to 16.2%, 26.0 to 22.6%, 32.0 to 27.6%, and 38.0 to 33.7%. One arena in trial 2 was discarded because the soil had become saturated; apparently water from the moat splashed into the soil when the arena was moved. 3.2. Experiment 2 (with S. glaseri) Data for experiment 2, which used S. glaseri- rather than H. marelatus-parasitized wax moth larvae, were similar to those of experiment 1 (Figs. 2 and 3), except that addition of S. glaseri-parasitized wax moth larvae in experiment 2 caused a much greater increase in the population density of A. oligospora than did addition of H. marelatus-parasitized larvae in experiment 1 (Fig. 3A vs C); the increase in A. oligospora was unaffected by soil water content or trial. Averaged across trials and soil water contents, A. oligospora population density was 5095G556 or 23G3 propagules/g soil in arenas with or without S. glaseri-parasitized wax moth larvae. Note that some estimates of A. oligospora population density and variance in experiment 2 may be low because some arenas with S. glaseri-parasitized wax moth larvae generated dilution plates that were all positive for
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A. oligospora. To use the most probable number program for these data, we assumed that an additional dilution would have generated all negative plates. Of the 10 arenas for each water level (five for each of two trials), the number requiring this assumption were zero at 14% water, one at 20% water, two at 26% water, three at 32% water, and two at 38% water. Arenas without S. glaseri-parasitized wax moth larvae did not require this assumption. As with H. marelatus-parasitized larvae in experiment 1, M. glutinosporum increased from nondetectable numbers to large numbers in the presence of S. glaseri-parasitized larvae (Figs. 2 and 3D). Averaged across trials and soil water content, and including arenas both with and without M. glutinosporum (nZ49), M. glutinosporum population density was 0G0 propagules/g soil in the absence of wax moth larvae and 187G101 propagules/g soil in the presence of S. glaseri-parasitized wax moth larvae. Including only those arenas in which M. glutinosporum was detected (nZ14), the mean population density of M. glutinosporum was 655G329 propagules/g soil. The total population density of nematode trapping fungi was 5370 or 29 propagules/g soil in arenas with or without S. glaseri-parasitized larvae. In the latter case and in contrast to experiment 1, M. glutinosporum made up only 3% and A. oligospora made up 94% of the total. In the absence of S. glaseri-parasitized wax moth larvae, no A. oligospora sporulation was observed on the surface of the soil. In the presence of S. glaseri-parasitized wax moth larvae, the percentage of arenas rated 0, 1, 2, or 3 for A. oligospora sporulation on the soil surface was 14, 19, 12, and 4%, respectively. When sporulation was heavy, A. oligospora conidiophores occurred on the cadaver and covered the surface of soil, even 5 cm from the cadaver. Large numbers of S. glaseri (24,980G1537) dispersed through soil and into the moat of arenas receiving S. glaseriparasitized wax moths, and this number was unaffected by soil water content or trial; no S. glaseri were found in moats from arenas lacking wax moth larvae. Soil water content was less at the end of the experiment than at the start, but the difference was much less that in experiment 1. Averaged over both trials, soil water content decreased from 14.0 to 13.7%, 20.0 to 19.5%, 26.0 to 24.9%, 32.0 to 31.3%, and 38.0 to 36.7%. The incubator contained more arenas during experiment 1 than experiment 2, and this may explain the greater water loss in experiment 1. One arena in trial 2 of experiment 2 was discarded because the soil had become saturated. 3.3. Experiment 3 In contrast to experiments 1 and 2, all arenas in experiment 3 received nematode-parasitized moth larvae. In nonheat-treated soil, A. oligospora was detected in most replicates but other fungi were not (Fig. 4). Detection seemed unaffected by nematode species or nematode isolate, except that D. brochopaga may have been detected
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Fig. 4. Detection frequency of nematophagous fungi in nonheat-treated soil as affected by entomopathogenic nematode in experiment 3. Arenas received wax moth larvae parasitized by Heterhorhabditis marelatus isolate b, Heterhorhabditis marelatus isolate k, or Steinernema glaseri. Detection frequency indicates the proportion of replicates of 18 total replicates (trials 1–3 combined) with the indicated fungus.
more frequently with H. marelatus-b than with H. marelatus-k or S. glaseri. Heat-treatment of soil eliminated nematophagous fungi from all arenas of trials 2 and 3. In trial 1, two of 18 arenas with heat-treated soil and H. marelatus-b parasitized wax moth larvae contained A. oligospora (10 propagules/g soil) and one arena with heat-treated soil and S. glaseriparasitized wax moth larvae contained A. oligospora (1199 propagules/g soil) (Fig. 5A). In an analysis of A. oligospora population density that only included data from nonheat-treated soil, the effects of trial and nematode isolate were significant. But because the interaction of trial and isolate was not significant, the data from the three trials were combined to simplify the presentation. All three nematode isolates supported large A. oligospora population densities, but A. oligospora population density was 11–17 times greater with S. glaseriparasitized wax moth larvae than with H. marelatusparasitized wax moth larvae (Fig. 5A). As in experiment 1, M. glutinosporum was the most common fungus after A. oligospora in experiment 3. It was not detected in heat-treated soil, but its overall population density in nonheat-treated soil was 185 propagules/g of soil if all replications were included and O500 propagules/g soil if arenas without M. glutinosporum were excluded (Fig. 5B). In nonheat-treated soil, the total population density of nematode-trapping fungi was 500, 712, or 4811 propagules/g soil when moth larvae were parasitized with H. marelatus-b, H. marelatus-k, or S. glaseri, respectively. The percentages of the total made up of M. glutinosporum or A. oligospora were 13 and 83% with H. marelatus-b, 60 and 38% with H. marelatus-k, and 1 and 98% with S. glaseri.
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Fig. 5. Population densities of Arthrobotrys oligospora and Myzocytium glutinosporum in heat-treated soil (solid bars) and nonheat-treated soil (open bars) as affected by nematode-parasitized insects (experiment 3). Values are the meansC1 standard error of 18 replicates (trials 1–3 combined).
In arenas containing nonheat-treated soil, A. oligospora sporulated on the soil surface in 28, 22, and 28% of the arenas containing H. marelatus-b, H. marelatus-k, and S. glaseri-parasitized wax moth larvae, respectively. But sporulation on the soil surface was seldom heavy and was not a good indicator of population density in soil. For example, 13 of 18 arenas with S. glaseriparasitized larvae exhibited no A. oligospora sporulation but contained 3168G747 A. oligospora propagules/g of soil. The strength of top-down effects of trapping fungi and other natural enemies on nematodes was measured as the fraction of dispersing nematodes that reached the moat in arenas with heat-treated vs nonheat treated soil. We excluded the three arenas of heat-treated soil that contained A. oligospora. H. marelatus-k dispersal was 22% less (P!0.05) and S. glaseri dispersal was 29% less (PZ0.08) in nonheat-treated soil than in heat-treated soil (Fig. 6A and C). In contrast, H. marelatus-b dispersal was greater in nonheat-treated than in heat-treated soil (Fig. 6B). Many more H. marelatus than S. glaseri were recovered from moats (Fig. 6A and B vs C). Most S. glaseri recovered from moats had dispersed there by day 18, but most H. marelatus did not disperse to moats until day 25 or 32.
Fig. 6. Cumulative numbers of entomopathogenic nematodes recovered from arena moats (experiment 3) as affected by heat treatment of soil and species of nematode used to infect the insects. Wax moth larvae were exposed to entomopathogenic nematodes on day 0 and placed in arenas on day 4; experiment 3 was ended on day 45. A (Heterhorhabditis marelatus isolate k), B (H. marelatus isolate b), and C (Steinernema glaseri). Values are the meansG1 standard error of 18 replicates (trials 1–3 combined).
After they emerged from the insect cadaver, S. glaseri but not H. marelatus infective juveniles aggregated and remained on the surface of the cadaver for several days. Steinernema glaseri-parasitized cadavers were more degraded than H. marelatus-parasitized cadavers at the end of experiment 3; the percentage degradation was 64G2% for H. marelatus-parasitized cadavers, regardless of isolate or soil treatment, whereas the percentage degradation for S. glaseri-parasitized cadavers was 78G4% in heat-treated soil and 92G1% in nonheat-treated soil. In nonheat-treated soil, more fungal hyphae (unidentified) surrounded S. glaseriparasitized cadavers than H. marelatus-parasitized cadavers.
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The biomasses of H. marelatus and S. glaseri infective juveniles were calculated using a published formula (Andra´ssy, 1956) and published morphometric data (Liu and Berry, 1996; Poinar, 1978; Stock, 1997). The calculated biomass of a H. marelatus or S. glaseri infective juvenile was 0.32 or 1.47 mg, respectively. If the mean numbers of nematodes in the moats (about 175,000 for H. marelatus and 25,000 for S. glaseri) were used to estimate the number of nematodes produced per wax moth larva, then each larva produced about 56 mg of H. marelatus or about 37 mg of S. glaseri. Nematode and fungus numbers were uncorrelated when the analyses included data from both heat-treated and nonheat-treated soil. When the analyses included data only from nonheat-treated soil, numbers of H. marelatus-k in the moat were positively correlated with the population density of A. oligospora (PZ0.04); other correlations were not significant. Soil water at the end of experiment 3 was unaffected by nematode isolate or the interaction of trial and soil treatment, but was significantly affected by trial and soil treatment. Averaged over soil treatments, soil water, initially at 32.0%, was 31.2, 28.0, and 25.1% at the end of trial 1–3, respectively. Averaged over the three trials, soil water content was 28.6% in nonheat-treated soil and 27.6% in heat-treated soil. One arena in trial 3 of experiment 3 was discarded because the soil had become saturated.
4. Discussion As predicted, the population density of several nematode-trapping fungi increased substantially when nematodeparasitized insects were added to BMR soil. The strongest numerical response was by A. oligospora; in the presence of parasitized insects, this fungus frequently ‘bloomed’ in that its population density increased 10–100 times and sometimes exceeded 10,000 propagules/g of soil. In some arenas the bloom was literal: A. oligospora conidiophores and conidia covered the soil surface. Three other fungi (M. glutinosporum, D. brochopaga, and G. gephyropaga) were seldom detected unless nematode-parasitized insects were present. Numbers of D. brochopaga and G. gephyropaga remained relatively small, but M. glutinosporum often increased to numbers as large as those of A. oligospora. The resources underlying this strong numerical response are unclear because the nematode-parasitized insect comprises several potential resources and because A. oligospora in particular is thought to be a ‘facultative’ trapper of nematodes, i.e. one that uses nematodes as a nitrogen source but decomposing organic matter as a carbon and energy source (Cooke, 1963; Barron, 1992). Additional research is needed to determine whether A. oligospora and the other fungi responded to the entomopathogenic nematodes, the dead insect, bacterivorous nematodes stimulated by the dead
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insect, or to some combination of these. We expect that other substrates at BMR (e.g. decomposing lupine leaves, mammal feces, isopod feces, dead arthropods not killed by nematodes, etc.) will also elicit numerical responses, but we doubt that these responses will be as strong as those documented in this study. Some trapping fungi were not enhanced by nematodeparasitized insects, and these were A. eudermata, A. musiformis, and A. paucospora. All three are similar to A. oligospora in that they use adhesive nets to capture nematodes. One possible explanation for this result is that the three species could not increase and were even suppressed because their niches overlap with that of A. oligospora. Koppenho¨fer et al. (1997) provided some evidence that A. oligospora competes with and suppresses A. eudermata and A. paucospora. In contrast to A. oligospora, M. glutinosporum has been infrequently studied. Its morphology and how it infects nematodes have been described (Barron, 1976), but its population biology in soil has not. The current study suggests that even though its zoospores swim poorly and encyst rapidly to form adhesive structures (Barron, 1976), M. glutinosporum prefers high soil water potentials. The dominant fungus in this study, A. oligospora, responded to both species of entomopathogenic nematode but much more to S. glaseri than to H. marelatus. Gross differences in nematode susceptibility cannot explain this result, because both nematodes are highly susceptible to the fungus on agar (Koppenho¨fer et al., 1996). But differences in susceptibility too small to detect on agar might be accentuated in soil. In particular, Heterorhabditis spp. retain their second-stage cuticle longer than Steinernema spp., and the extra cuticle can increase the chances of escaping a fungal trap (Timper and Kaya, 1989; Timper et al., 1991; Timper and Brodie, 1995). Other possible differences underlying the stronger response to S. glaseri than to H. marelatus include nematode biomass produced per insect, the timing of nematode emergence, the behavior of nematodes after emergence, the effect of nematode infection on the insect cadaver, and the effect of the symbiotic bacteria. Differences in nematode biomass, however, cannot explain the stronger effect of S. glaseri because S. glaseri produced less rather than more nematode biomass per insect. S. glaseri did emerge from the cadaver sooner than did H. marelatus, but how this might result in stronger enhancement of the fungi is unclear. Behavior after emergence seems a better explanation for the greater response of fungi to S. glaseri: unlike H. marelatus, S. glaseri aggregated on the cadaver for several days after emerging, and this may have increased exposure to adjacent fungi. Such behavior has been described for S. glaseri (Lewis and Gaugler, 1994) but not for H. marelatus. The effect of the nematode on the cadaver may also be important, because S. glaseri-parasitized cadavers degraded much faster and seemed leakier than H. marelatus-parasitized cadavers. Perhaps materials
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leaking from the cadaver supported saprophytic growth of nematode-trapping fungi. Differences in cadaver degradation could result from how the nematodes emerged or from differences in the species of symbiotic bacteria or their antibiotics (Zhou et al., 2002). To quantify nematode mortality caused by trapping fungi and other natural enemies, we compared numbers of nematodes that emerged and moved from parasitized insects through at least 5 cm of heat-treated (natural enemies removed) or nonheat-treated soil (natural enemies present) and into a moat. Mortality was 22% with H. marelatus-k and 29% with S. glaseri. In the case of H. marelatus-b, dispersal was greater in nonheat-treated than in heat-treated soil, suggesting an artifact or inadequate replication relative to the variance; a previous study reported a large variance in the number of H. marelatus produced per ghost moth or wax moth (Strong et al., 1996). Regardless, the data failed to support the hypothesis that trapping fungi kill substantial proportions of nematodes (O50%) emerging from parasitized insects (Strong, 1999). Given the very strong bottom-up effects of nematodeparasitized insects on trapping fungi, the relatively weak top-down effects described in the previous paragraph are difficult to explain. Perhaps the fungi responded too slowly, i.e. perhaps the numerical response occurred only after many nematodes had already dispersed. Perhaps the rapid emergence and dispersal of many nematodes simply overwhelmed the traps. Perhaps A. oligospora and M. glutinosporum are inefficient at trapping [for more data on the failure of A. oligospora to trap many nematodes in soil, see Jaffee (2002, 2003, 2004) but very efficient at converting captured nematodes into fungal propagules. Finally, these fungi may have used substrates other than or in addition to nematodes. The use of wax moth rather than ghost moth larvae requires discussion. As noted previously, we used wax moths in place of the natural ghost moths because ghost moth larvae are often unavailable. While sharing many similarities, wax moths and ghost moths also differ. For example, wax moths are more susceptible to H. marelatus infection (Strong et al., 1996); this difference, however, is probably unimportant because H. marelatus-parasitized ghost moths are abundant at BMR and because they produce at least as many nematodes as do parasitized wax moths of similar size (Strong et al., 1996). In contrast, moth larva size is probably important. Ghost moth larvae that hatch from eggs and move through soil to roots are smaller than last instar wax moth larvae and don’t become large until after they penetrate roots. It is reasonable to expect that small, nematode-parasitized ghost moth larvae in the soil will produce fewer nematodes and elicit a smaller fungal response than did last instar wax moths. But ghost moth larvae in roots, which can be as large or larger than the wax moths used in this study, are also parasitized by nematodes and are exposed to trapping fungi. This happens because roots penetrated by the larvae often
crack (unpublished observations, D.R. Strong). When the cracking is extensive, the large, nematode-parasitized ghost moth larvae are fully exposed to soil and to nematodetrapping fungi. It follows that data generated with nematode-parasitized wax moths may be similar to data generated from large, nematode-parasitized ghost moths. But because ghost moth larvae in the field range from small to large and because ghost moths may differ from wax moth in other important ways, future research should compare fungal responses to wax moths and ghost moths and should determine the significance of moth size. Future research should also explore the presence and activity of trapping fungi in the penetrated lupine roots for two reasons. First, as insects move through soil, they can superficially acquire the conidia of trapping fungi (Fowler and Garcia, 1989), and we hypothesize that ghost moths in lupine root galleries are frequently contaminated with A. oligosopora. Second, A. oligospora has cellulolytic activity (Barron, 1992; Jaffee, 2004), and the combination of dead lupine wood and nematode-parasitized insects might induce both strong numerical responses and abundant trapping of nematodes. In addition to being relevant to the community ecology of BMR, the current data are also relevant to the general ecology of nematode-trapping fungi and to biological control of plant-parasitic nematodes. From the perspective of fungal ecology, the very large numerical response to nematode-parasitized wax moths is exciting and worth understanding. That trapping fungi can be enhanced by organic amendments to soil is well documented (Stirling, 1991), but to the best of our knowledge no amendment has elicited so large a response as that reported here by nematode-parasitized insects. And from the perspective of biological control, at least some researchers who study natural enemies of plant-parasitic nematodes will want to understand this numerical response, how it relates to nematode mortality, and how it might be used to manage pest nematodes. Two important questions are: ‘Which resources are the fungi using?’ and ‘Will other soils exhibit both strong bottom-up effects on trapping fungi and strong top-down effects on nematodes?’ Acknowledgements We thank Harry Kaya and Francine Farrell for comments on an earlier version of this paper. References Andra´ssy, I., 1956. The determination of volume and weight of nematodes. Acta zoologica (Hungarian academy of science) 2, 1–15, in: Zuckerman, B.B., Brzeski, M.W., Deubert, K.H. (Eds.), English Translation of Selected East European papers in Nematology, 1967. University of Massachusetts, Cranberry Experiment Station, East Wareham, Massachusetts, pp. 73–84. Barron, G.L., 1976. Nematophagous fungi: three new species of Myzocytium. Canadian Journal of Botany 22, 752–762.
B.A. Jaffee, D.R. Strong / Soil Biology & Biochemistry 37 (2005) 1011–1021 Barron, G.L., 1992. Lignolytic and cellulolytic fungi as predators and parasites, in: Carroll, G.C., Wicklow, D.T. (Eds.), The Fungal Community: Its Organization and Role in the Ecosystem. Marcel Dekker, New York, pp. 311–326. Cooke, R.C., 1963. The predaceous activity of nematode-trapping fungi added to soil. Annals of Applied Bioogy 51, 295–299. Cooke, R.C., Godfrey, B.E.S., 1964. A key to the nematode-destroying fungi. Transactions of the British Mycological Society 47, 61–74. Fowler, H.G., Garcia, C.R., 1989. Parasite-dependent protocooperation. Naturwissenschaften 76, 26–27. Jaffee, B.A., 2002. Soil cages for studying how organic amendments affect nematode-trapping fungi. Applied Soil Ecology 21, 1–9. Jaffee, B.A., 2003. Correlations between most probable number and activity of nematode-trapping fungi. Phytopathology 93, 1599–1605. Jaffee, B.A., 2004. Wood, nematodes, and the nematode-trapping fungus Arthrobotrys oligospora. Soil Biology & Biochemistry 36, 1171–1178. Jaffee, B.A., Strong, D.R., Muldoon, A.E., 1996. Nematode-trapping fungi of a natural shrubland: tests for food chain involvement. Mycologia 88, 554–564. Kaya, H.K., Stock, S.P., 1997. Techniques in insect nematology, in: Lacey, L. (Ed.), Techniques in Insect Nematology. Academic Press, San Diego, pp. 281–324. Klee, A.J., 1993. A computer program for the determination of most probable number and its confidence limits. Journal of Microbiological Methods 18, 91–98. Koppenho¨fer, A.M., Jaffee, B.A., Muldoon, A.E., Strong, D.R., Kaya, H.K., 1996. Effect of nematode-trapping fungi on an entomopathogenic nematode originating from the same field site in California. Journal of Invertebrate Pathology 68, 246–252. Koppenho¨fer, A.M., Jaffee, B.A., Muldoon, A.E., Strong, D.R., 1997. Suppression of an entomopathogenic nematode by the nematodetrapping fungi Geniculifera paucispora and Monacrosporium eudermatum as affected by the fungus Arthrobotrys oligospora. Mycologia 89, 220–227. Lewis, E.E., Gaugler, R., 1994. Entomopathogenic nematode (Rhabdita: Steinernematidae) sex ratio relates to foraging strategy. Journal of Invertebrate Pathology 64, 238–242. Liu, J., Berry, R.E., 1996. Heterorhabditis marelatus n. sp. (Rhaditida: Heterorhabditidae) from oregon. Journal of Invertebrate Pathology 67, 48–54. Miller, V.C., 1972. Soil Survey of Sonoma County California. USDA Forest Service and Soil Conservation Service, University of California Agricultural Experiment Station.
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Poinar Jr.., G.O., 1978. Generation polymorphism in Neoaplectana glaseri Steiner (Steinernematidae: Nematoda), redescribed from Strigoderma arboricola (Fab.) (Scarabaeidae: Coleoptera) in North Carolina. Nematologica 24, 105–114. Preisser, E.L., 2003. Field evidence for a rapidly cascading underground food web. Ecology 84, 869–874. Scholler, M., Hagedorn, G., Rubner, A., 1999. A reevaluation of predatory Orbiliaceous fungi. II. A new generic concept. Sydowia 51, 89–113. Stirling, G.R., 1991. Biological Control of Plant Parasitic Nematodes. CAB International, Wallingford, UK, p. 282. Stock, S.P., 1997. Heterorhabditis hepialus Stock, Strong and Gardner, a junior synonym of H. marelatus Liu and Berry, 1996 (Rhabditida: Heterorhabditidae) with a rediscription of the species. Nematologica 43, 455–463. Strong, D.R., 1999. Predator control in terrestrial ecosystems: the underground food chain of bush lupine, in: Olff, H., Brown, V.K., Drent, R.H. (Eds.), Herbivores: Between Plants and Predators. Blackwell Science, Oxford, UK, pp. 577–602. Strong, D.R., Maron, J.L., Connors, P.G., Whipple, A., Harrison, S., Jefferies, R.L., 1995. High mortality, fluctuation in numbers, and heavy subterranean insect herbivory in bush lupine, Lupinus arboreus. Oecologia 104, 85–92. Strong, D.R., Kaya, H.K., Whipple, A.V., Child, A.L., Kraig, S., Bondonno, M., Dyer, K., Maron, J.L., 1996. Entomopathogenic nematodes: natural enemies of root-feeding caterpillars on bush lupine. Oecologia 108, 167–173. Strong, D.R., Whipple, A.V., Child, A.L., Dennis, B., 1999. Model selection for a subterranean trophic cascade: root-feeding caterpillars and entomopathogenic nematodes. Ecology 80, 2750–2761. Timper, P., Brodie, B.B., 1995. Interaction of the microbivorous nematode Teratorhabditis dentifera and the nematode-pathogenic fungus Hirsutella rhossiliensis. Biological Control 5, 629–635. Timper, P., Kaya, H.K., 1989. Role of second-stage cuticle of entomogenous nematodes in preventing infection by nematophagous fungi. Journal of Invertebrate Pathology 54, 314–321. Timper, P., Kaya, H.K., Jaffee, B.A., 1991. Survival of entomogenous nematodes in soil infested with the nematode-parasitic fungus Hirsutella rhossiliensis (Deuteromycotina: Hyphomycetes). Biological Control 1, 42–50. White, G.F., 1927. A method for obtaining infective nematode larvae from cultures. Science 66, 302–303. Zhou, X.S., Kaya, H.K., Heungens, K., Goodrich-Blair, H., 2002. Response of ants to a deterrent factor(s) produced by the symbiotic bacteria of entomopathogenic nematodes. Applied and Environmental Microbiology 68, 6202–6209.
Strong bottom-up and weak top-down effects in soil: nematode-parasitized insects and nematode-trapping fungi B.A. Jaffeea,*, D.R. Strongb a Department of Nematology, University of California at Davis, 1 Shields Avenue, Davis, CA 95616, USA Department of Evolution and Ecology, University of California at Davis, 1 Shields Avenue, Davis, CA 95616, USA
b
Received 25 March 2004; received in revised form 4 May 2004; accepted 7 May 2004
Abstract The soils of the Bodega Marine Reserve (BMR, Sonoma County, California) contain many nematode-trapping fungi and many ghost moth larvae parasitized by entomopathogenic nematodes. The current study determined whether these nematode-parasitized moth larvae, which can produce very large numbers of nematodes, enhanced the population densities of nematode-trapping fungi and whether the fungi trapped substantial numbers of nematodes emerging and dispersing from moths. Wax moths were used in place of ghost moths because the former are easier to obtain. When nematode-parasitized moth larvae were added to laboratory microcosms containing BMR field soil, the population densities of four nematode-trapping fungi increased substantially. The greatest increase in population density was by Arthrobotrys oligospora, which uses adhesive networks to capture nematodes. A. oligospora population density increased about 10 times when the added moth larvae were parasitized by the nematode Heterorhabditis marelatus and about 100 times when added moth larvae were parasitized by the nematode Steinernema glaseri. Other trapping fungi endemic to the soil and enhanced by nematode-parasitized moth larvae included Myzocytium glutinosporum, Drechslerella brochopaga, and Gamsylella gephyropaga, which produce adhesive spores, constricting rings, and adhesive branches, respectively. The data suggest that the previously documented abundance and diversity of nematode-trapping fungi in BMR soil can be explained, at least in part, by nematode-parasitized insects, although that inference requires further studies with ghost moths. The strong bottom-up enhancement of nematode-trapping fungi was not matched by a strong top-down suppression of nematodes, i.e. the fungi trapped fewer than 30% of dispersing nematodes. q 2005 Elsevier Ltd. All rights reserved. Keywords: Arthrobotrys oligospora; Drechslerella brochopaga; Entomopathogenic nematode; Gamsylella gephyropaga; Heterorhabditis marelatus; Myzocytium glutinosporum; Nematode-trapping fungi; Soil food web; Steinernema glaseri
1. Introduction On the coastal headlands of the Bodega Marine Reserve (BMR, Sonoma County, California), nematode-trapping fungi are diverse and often abundant. The soil contains at least 12 species, and overall population densities average 50 propagules/g of soil but can exceed 600 propagules/g of soil. The most common and abundant nematode-trapping fungus at BMR is Arthrobotrys oligospora (Jaffee et al., 1996). Why this soil contains so many nematode-trapping fungi is unclear, but the ghost moth larva and its nematode * Corresponding author. Tel.: C1 530 752 0862; fax: C1 530 752 5809. E-mail address: [email protected] (B.A. Jaffee). 0038-0717/$ - see front matter q 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.soilbio.2004.05.026
parasite may be important. At BMR, ghost moth larvae consume bush lupine roots (Strong et al., 1995). In some years, the larvae kill entire stands of this dominant shrub, but in other years they are suppressed by the entomopathogenic nematode, Heterorhabditis marelatus (Strong et al., 1996, 1999; Preisser, 2003). Like other entomopathogenic nematodes, H. marelatus is an obligate parasite or pathogen of insects (Liu and Berry, 1996; Stock, 1997). Infective juveniles of H. marelatus invade ghost moth larvae in soil or those that have penetrated lupine roots, and once within the insect body cavity, the nematodes release a symbiotic bacterium which kills the insect. The nematodes, which consume both the multiplying bacteria and the degrading insect, then increase in number. Each parasitized moth larva can produce few or many nematodes depending on larval
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size; a large larva can produce more than 400,000 juvenile nematodes (Strong et al., 1996, 1999). When the resource is depleted, the newly-produced juvenile nematodes disperse from the insect cadaver into the soil, where they search for a new host. By potentially producing and releasing many nematodes in a small volume of soil, nematode-parasitized ghost moth larvae may help explain the abundance of nematode-trapping fungi at BMR (Strong, 1999). While data about how the nematode and ghost moth affect trapping fungi are nonexistent, data about how trapping fungi affect H. marelatus do exist but are somewhat conflicting. The more common trapping fungi at BMR capture and consume H. marelatus in vitro (Koppenho¨fer et al., 1996), but their distribution in the field was not correlated with lupine death, as it should be if the fungi suppressed entomopathogenic nematodes and thereby protected ghost moth larvae (Jaffee et al., 1996). In contrast, soil microcosm experiments indicated that the more common species apparently trapped substantial numbers of juvenile H. marelatus as they moved through soil to insect larvae (Koppenho¨fer et al., 1996, 1997). None of these studies measured trapping as the nematodes emerged and dispersed from the nematode-parasitized insect. The current study focuses on the nematode-parasitized insect larva and the surrounding soil. In this soil, which we will not call the ‘larvosphere’, we predicted that emerging nematodes would greatly enhance the population densities of trapping fungi and that the fungi in turn would trap many emerging nematodes. In other words, we predicted strong bottom-up effects of emerging nematodes on fungi and strong top-down effects of trapping fungi on emerging nematodes. Experiments testing these predictions are described in this paper.
Steinernema glaseri, was also used but was not isolated from BMR soil and has been maintained for over 10 years by repeated rearing in wax moth larvae. The infective juvenile of S. glaseri is about twice as long and wide as the infective juvenile of H. marelatus (Poinar, 1978; Liu and Berry, 1996). 2.2. Soil BMR is on the Pacific Coast of California, 80 km north of San Francisco. Soil was collected from a location at BMR called Mussel Point Tip (Jaffee et al., 1996) on 10 June and 18 September 2003. Soil was collected 5–20 cm deep adjacent to lupines. It was sieved (2 mm openings), stored at 10 8C, and used within 30 days (all trials of experiments 1 and 2 and trial 3 of experiment 3) or 68 days (trials 1 and 2 of experiment 3). Mussel Point Tip soil is a coarse sandy loam (Miller, 1972) with a pH in water of 4.9 and with an organic matter content of 4.9% (Jaffee et al., 1996). In the batch of soil collected on 10 June 2003, total nitrogen and carbon contents were 0.19 and 1.83%, respectively, with 4.5 ppm NH4 and 5.9 ppm NO3. A water release curve was determined because we suspected that water potential could greatly affect the results and because a water release curve provides valuable information on the proportion of soil pores that are filled with water or drained. For experiment 3, natural enemies of nematodes were removed for one treatment by heating the soil as follows. A 1-l beaker containing soil was placed in a 75 8C oven. Once the center of the soil mass increased to 70 8C, the oven temperature was reduced to 70 8C. After 2 h at 70 8C, the soil was removed and cooled to room temperature. 2.3. Nematode-parasitized insects
2. Methods 2.1. Nematode isolates One isolate of H. marelatus (referred to here as H. marelatus-k) was obtained from H. K. Kaya. H. marelatus-k was isolated from BMR soil and has been maintained for over 7 years by repeated rearing in Galleria mellonella (wax moth) larvae (Kaya and Stock, 1997). A newer isolate, H. marelatus-b, was used to determine whether repeated rearing of H. marelatus affected how nematophagous fungi responded to the nematode. H. marelatus-b was obtained in July 2003, by baiting BMR soil with wax moth larvae (Kaya and Stock, 1997) and incubating those larvae that died and turned orange in White traps (White, 1927; Strong et al., 1996; Kaya and Stock, 1997); in contrast to H. marelatus-k, H. marelatus-b was reared on wax moth larvae only once (trials 1 and 2 of experiment 3) or twice (trial 3 of experiment 3) after its isolation from the field. The entomopathogenic nematode,
Although ghost moths rather than wax moths are the major hosts of H. marelatus at BMR, wax moth larvae were used because ghost moth larvae are only available several months each year (Strong et al., 1996). In contrast, healthy and uniform wax moth larvae are easily obtained year round. Healthy, last-instar wax moth larvae (about 2 cm long; 220–260 mg fresh mass per larva; purchased from Rainbow Mealworms, Inc., Compton, CA) were placed in a 10-cm diameter Petri dish (10 larvae per dish); the dish contained a 10-cm diameter filter paper which had received 200–500 H. marelatus or S. glaseri in a 1-ml suspension. After 4 days at 20–22 8C, larvae that were dead and showing characteristics of nematode parasitism were added to soil: those exposed to H. marelatus were orange (Strong et al., 1996), whereas those exposed to S. glaseri were grey (Kaya and Stock, 1997). In this paper, ‘nematodeparasitized insects’ refers to moth larvae that have been killed by entomopathogenic nematodes and that contain large numbers of entomopathogenic nematodes.
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2.4. Experimental arenas The arena was based on the original ‘White trap’ (White, 1927) and the ‘modified White trap’ (Kaya and Stock, 1997). The original trap was designed to collect and quantify nematode parasites of mammals as they emerge and disperse from nematode-infested feces, and the modified trap was designed to collect and quantify entomopathogenic nematodes as they emerge and disperse from nematode-parasitized insects. Both traps use a smaller dish that sits in a larger, water-filled dish. The smaller dish contains the feces or insect-parasitized nematodes, and the nematodes are collected in the surrounding water (hereafter called the ‘moat’). To assemble an arena, 40 g soil (dry weight equivalent) at 14% water content (g water per 100 g soil) was placed in the bottom of a small plastic Petri dish (8.7-cm diameter and 1.5 cm deep). The soil was tamped; the height and bulk density of the soil column were about 5 mm and 1.3 g/cm3. Soil water content was adjusted as indicated for each experiment, and one or no nematode-parasitized wax moth larva was placed on the surface of the soil in the center of the dish. The small dish, which was uncovered, was placed in a large plastic Petri dish (15 cm diameter and 2.5 cm deep). Sufficient distilled water (45 ml) was added to the large dish so that the water covered the bottom of the large dish and rose part way up the sides of the small dish. A rubber band (76!1.6 mm) placed beneath the smaller dish prevented it from sliding to the side of the larger dish and becoming filled with water. As in the White trap, the moat was a key feature of our experimental arenas. To reach the safety of the moat, juveniles emerging and dispersing from insect cadavers had to traverse at least 5 cm of soil that contained nematodetrapping fungi and other natural enemies (unless the soil had been heat-treated). Nematodes in the moats were identified and counted, as described for each experiment. The soil in the inner dish was used to quantify fungus population density, as described for experiment 1 below. Nematodes and other organisms in the soil were not quantified. Arenas were assembled and grouped by replicate, sealed in plastic bags (one replicate per bag) with moist paper towels, and placed in a 20 8C incubator without light. After 7–10 days, all replicates were examined for weed seedlings, which were removed to prevent nematodes from moving up the weed stems rather than moving through the soil. Other details are described for each experiment. 2.5. Experiment 1 Experiment 1 examined the numerical response of nematode-trapping fungi to nematode-parasitized insects in BMR soil (i.e. the bottom-up effect of parasitized insects on trapping fungi); the nematode was H. marelatus-k. Soil water content was selected as a variable because we
Fig. 1. Water release curve of soil from Mussel Point Tip, the Bodega Marine Reserve.
suspected that the most abundant trapping fungus in BMR soil, A. oligospora (Jaffee et al., 1996), might be more active at high soil water potentials. Soil water content was adjusted to 14, 20, 26, 32, or 38% by adding the appropriate volume of distilled water to the soil surface; 14% was considered a moderate water content and 38% was considered wet; at 14%, the larger soil pores were drained and smaller pores were water-filled, whereas at 38%, all but the largest soil pores were water-filled (Fig. 1). Experiment 1 was ended 35 days after the moth larvae were exposed to H. marelatus (31 days after the arenas were assembled and larvae were placed on the soil), and the following were quantified: nematodes in the moat, A. oligospora sporulation on the soil surface, the population density of nematode-trapping fungi in the soil, and soil water content. To count nematodes, the water and nematodes in the moat were placed in a 1-l beaker, the volume was increased to 800 ml, the suspension was mixed, 1 ml was placed in each of two small dishes, and the nematodes were counted with the aid of a dissecting microscope. To quantify the nematode-trapping fungi in the soil, the soil was removed from each arena, placed in a plastic bag, and mixed with a spatula, and 10 g (dry wt equivalent) was placed in a sterile 125-ml flask. The volume of each flask was increased to 50 ml with sterile distilled water, and the suspension was dispersed on a wrist action shaker for 9 min. A 10-fold dilution series with three dilutions was then prepared, and 0.1 ml of each dilution was placed on each of five quarter-strength corn meal agar plates. About 1000 bait nematodes (healthy S. glaseri) were added to each plate. After 7, 14, and 21 days, each plate was examined for nematode-trapping fungi, and a computer program was used to determine the most probable number (Klee, 1993; Jaffee, 2003). The remaining soil from each arena was used to determine water content; soil mass was compared before and after drying.
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Experiment 1 was performed twice (trials 1 and 2). There were five replications for each treatment in each trial. 2.6. Experiment 2 Experiment 2 was identical to experiment 1 except that the wax moths were parasitized by S. glaseri rather than H. marelatus-k. We decided to examine the effect of S. glaseri because H. marelatus-parasitized insects did not stimulate extensive A. oligospora sporulation on the soil surface as we had expected based on preliminary experiments. Experiment 2 was also performed twice (trials 1 and 2). There were five replications for each treatment in each trial. 2.7. Experiment 3 Experiment 3 had three objectives. One was to quantify how trapping fungi and other natural enemies reduced dispersal of nematodes from parasitized insects. A second objective was to directly compare the effects of S. glaseriand H. marelatus-parasitized wax moth larvae on the fungi. Although experiments 1 and 2 indicated that nematodetrapping fungi responded more strongly to S. glaseri- than to H. marelatus-parasitized wax moth larvae, these experiments were independent, and differences may have reflected other uncontrolled effects. A third objective was to compare the effects of two H. marelatus isolates; one isolate had been cultured in the laboratory for many years and the other had not. Arenas contained heat-treated and nonheat-treated soil, which was packed at 14% water content and then increased to 32% as described for experiment 1. Each arena received one wax moth larva parasitized with either H. marelatus-k, H. marelatus-b, or S. glaseri. Other procedures were as described in experiments 1 and 2, except that experiment 3 was ended after 45 rather than 35 days and nematodes were counted four times rather than once. We extended experiment 3 because H. marelatus emerged from the wax moth larvae about 10 days after S. glaseri in experiments 1 vs 2 (data not shown), and we suspected that the trapping fungi might require more time to respond to emerging H. marelatus. We removed nematodes from the moat on four dates to prevent them from being parasitized and degraded by A. oligospora and Myzocytium glutinosporum. Nematodes in the moat were collected and counted on day 18, 25, 32, and 45 (moat water was replaced each time except the last). Observations in experiments 1 and 2 suggested that moth larvae degraded faster when parasitized by S. glaseri than H. marelatus. In experiment 3, degradation of each moth larva was subjectively assessed from 0 (no degradation) to 100% (complete degradation). Experiment 3 was performed three times (trials 1–3). There were six replications for each treatment in each trial.
2.8. Statistical analyses After transformation (log for numerical counts and arcsine for percentages), data were subjected to an analysis of variance (SAS for Windows, version 8.02; SAS Institute, Inc., Cary, NC) to determine whether treatment affected (P!0.05) fungus population densities in all three experiments and nematode population densities in experiment 3. In experiments 1 and 2, soil water content was considered a categorical variable and a continuous variable in separate analyses. In experiment 3, partial Spearman correlation analysis was used to determine whether nematode population density and fungus population density were related; a separate analysis was done for each combination of nematode and fungus and with data from both heat-treated and nonheat-treated soil or with data only from nonheattreated soil. The data for detection frequency were not statistically analyzed because, in our view, there were too few replications to support statistical inferences. Note that each trial produced only one value per fungus for frequency of detection (a percentage or proportion of replicates that were positive in that trial). Thus, there were only three values for detection frequency for each fungus in experiment 3.
3. Results 3.1. Experiment 1 (with H. marelatus-k) Of the nematode-trapping fungi detected (Table 1), A. oligospora and M. glutinosporum were most numerous. A. oligospora was detected in most arenas, even when wax moth larvae were absent and was detected in all arenas except one when H. marelatus-parasitized wax moth larvae were present (Fig. 2). A. oligospora population density was 12 times greater in arenas with wax moths but was unaffected by soil water content (Fig. 3A). Averaged across trials and soil water content, A. oligospora population Table 1 Fungi detected in this study Fungusa
Infective structuresb
Arthrobotrys oligospora Myzocytium glutinosporum A. eudermata A. musiformis A. paucispora Gamsylella gephyropaga Drechslerella brochopaga
Adhesive networks Adhesive sporesc Adhesive networks Adhesive networks Adhesive networks Adhesive branches Constricting rings
a
These generic designations follow Scholler et al. (1999). Once the infective structure has firmly contacted the nematode, all of these fungi directly penetrate the nematode cuticle and then grow through and consume the nematode body. c M. glutinosporum produces zoospores that usually encyst near the parasitized nematode. Each encysted spore forms a bud which adheres to the surface of passing nematodes (Barron, 1976). b
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Fig. 2. Detection frequency of nematophagous fungi in arenas of experiments 1 and 2. Each arena contained nonheat-treated soil and no wax moth larva or a wax moth larva parasitized with Heterorhabditis marelatus isolate k (experiment 1) or Steinernema glaseri (experiment 2). Wax moth larvae were exposed to entomopathogenic nematodes on day 0 and placed in arenas on day 4; experiments 1 and 2 were ended on day 35. Detection frequency indicates the proportion of 10 replicates (trials 1 and 2 combined) with the indicated fungus. Open bars and striped bars indicate data from arenas without and with nematodeparasitized larvae, respectively.
density was 207G39 propagules/g soil (meanG1 standard error) or 17G2 propagules/g soil in arenas with or without H. marelatus-parasitized wax moth larvae. In contrast to A. oligospora, M. glutinosporum was not detected unless H. marelatus-parasitized wax moth larvae were present (Fig. 2); and even in the presence of H. marelatus-parasitized wax moth larvae, M. glutinosporum was detected in only 13 of 48 total arenas and never when soil water was at the driest level tested (Fig. 3B). When detected, M. glutinosporum was abundant: averaged across trials and soil water contents, and including arenas both with and without M. glutinosporum (nZ48), M. glutinosporum population density was 582G245 or 0G0 propagules/g soil in arenas with or without
H. marelatus-parasitized wax moth larvae. Including only those arenas in which M. glutinosporum was detected (nZ13), the mean population density of M. glutinosporum was 2148G768 propagules/g soil. Detection of fungi that form adhesive branches (Gamsylella gephyropaga) or constricting rings (Drechslerella brochopaga) increased in the presence of H. marelatusparasitized moth larvae (Fig. 2), but population densities (averaged over all water levels) remained less than 20 propagules/g soil (data not shown). Detection of fungi other than A. oligospora that form adhesive networks (A. eudermata, A. musiformis, and A. paucispora) was either unaffected or possibly suppressed by wax moth larvae (Fig. 2), and population densities of these fungi (averaged
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Fig. 3. Population densities of Arthrobotrys oligospora and Myzocytium glutinosporum in nonheat-treated soil as affected by the presence or absence of nematode-parasitized insects and soil water content. Experiment 1 (A and B) was done with Heterhorhabditis marelatus isolate k, and experiment 2 (C and D) was done with Steinernema glaseri. Solid bars and open bars indicate data from arenas without and with nematode-parasitized larvae, respectively. Values are the meansC1 standard error of 10 replicates (trials 1 and 2 combined).
over all water levels) was less than 2 propagules/g soil (data not shown). The total population density of nematode trapping fungi was 812 or 23 propagules/g soil in arenas with or without H. marelatus-parasitized larvae. In the former case, M. glutinosporum made up 72% and A. oligospora made up 25% of the total. We frequently observed A. oligospora conidiophores and conidia, which are distinctive (Cooke and Godfrey, 1964), on the soil surface and the insect cadaver. To quantify this sporulation, the soil surface and cadaver were examined with reflected light and a dissecting microscope (35–70!magnification), and sporulation was rated on a scale from 0 to 3 (0, no A. oligospora conidiophores; 1, fewer than 10 A. oligospora conidiophores per arena; 2, 10–75 A. oligospora conidiophores per arena; 3, O75 A. oligospora conidiophores per arena). In the presence of H. marelatus-parasitized wax moth larvae, the percentage of arenas rated 0, 1, 2, or 3 was 54, 35, 10, and 0%, respectively. Conidiophores like those produced by A. eudermata, G. gephyropaga, or D. brochopaga, were infrequently observed. None of these structures was observed in arenas without nematodeparasitized insects. Large numbers of H. marelatus (143,000G7000) dispersed through soil and into the moats of arenas receiving H. marelatus-parasitized wax moths, and this number was
unaffected by soil water content or trial; no H. marelatus were found in moats from arenas lacking wax moth larvae. Soil water content was less at the end of the experiment than at the start. Averaged over both trials, soil water content decreased from 14.0 to 11.2%, 20.0 to 16.2%, 26.0 to 22.6%, 32.0 to 27.6%, and 38.0 to 33.7%. One arena in trial 2 was discarded because the soil had become saturated; apparently water from the moat splashed into the soil when the arena was moved. 3.2. Experiment 2 (with S. glaseri) Data for experiment 2, which used S. glaseri- rather than H. marelatus-parasitized wax moth larvae, were similar to those of experiment 1 (Figs. 2 and 3), except that addition of S. glaseri-parasitized wax moth larvae in experiment 2 caused a much greater increase in the population density of A. oligospora than did addition of H. marelatus-parasitized larvae in experiment 1 (Fig. 3A vs C); the increase in A. oligospora was unaffected by soil water content or trial. Averaged across trials and soil water contents, A. oligospora population density was 5095G556 or 23G3 propagules/g soil in arenas with or without S. glaseri-parasitized wax moth larvae. Note that some estimates of A. oligospora population density and variance in experiment 2 may be low because some arenas with S. glaseri-parasitized wax moth larvae generated dilution plates that were all positive for
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A. oligospora. To use the most probable number program for these data, we assumed that an additional dilution would have generated all negative plates. Of the 10 arenas for each water level (five for each of two trials), the number requiring this assumption were zero at 14% water, one at 20% water, two at 26% water, three at 32% water, and two at 38% water. Arenas without S. glaseri-parasitized wax moth larvae did not require this assumption. As with H. marelatus-parasitized larvae in experiment 1, M. glutinosporum increased from nondetectable numbers to large numbers in the presence of S. glaseri-parasitized larvae (Figs. 2 and 3D). Averaged across trials and soil water content, and including arenas both with and without M. glutinosporum (nZ49), M. glutinosporum population density was 0G0 propagules/g soil in the absence of wax moth larvae and 187G101 propagules/g soil in the presence of S. glaseri-parasitized wax moth larvae. Including only those arenas in which M. glutinosporum was detected (nZ14), the mean population density of M. glutinosporum was 655G329 propagules/g soil. The total population density of nematode trapping fungi was 5370 or 29 propagules/g soil in arenas with or without S. glaseri-parasitized larvae. In the latter case and in contrast to experiment 1, M. glutinosporum made up only 3% and A. oligospora made up 94% of the total. In the absence of S. glaseri-parasitized wax moth larvae, no A. oligospora sporulation was observed on the surface of the soil. In the presence of S. glaseri-parasitized wax moth larvae, the percentage of arenas rated 0, 1, 2, or 3 for A. oligospora sporulation on the soil surface was 14, 19, 12, and 4%, respectively. When sporulation was heavy, A. oligospora conidiophores occurred on the cadaver and covered the surface of soil, even 5 cm from the cadaver. Large numbers of S. glaseri (24,980G1537) dispersed through soil and into the moat of arenas receiving S. glaseriparasitized wax moths, and this number was unaffected by soil water content or trial; no S. glaseri were found in moats from arenas lacking wax moth larvae. Soil water content was less at the end of the experiment than at the start, but the difference was much less that in experiment 1. Averaged over both trials, soil water content decreased from 14.0 to 13.7%, 20.0 to 19.5%, 26.0 to 24.9%, 32.0 to 31.3%, and 38.0 to 36.7%. The incubator contained more arenas during experiment 1 than experiment 2, and this may explain the greater water loss in experiment 1. One arena in trial 2 of experiment 2 was discarded because the soil had become saturated. 3.3. Experiment 3 In contrast to experiments 1 and 2, all arenas in experiment 3 received nematode-parasitized moth larvae. In nonheat-treated soil, A. oligospora was detected in most replicates but other fungi were not (Fig. 4). Detection seemed unaffected by nematode species or nematode isolate, except that D. brochopaga may have been detected
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Fig. 4. Detection frequency of nematophagous fungi in nonheat-treated soil as affected by entomopathogenic nematode in experiment 3. Arenas received wax moth larvae parasitized by Heterhorhabditis marelatus isolate b, Heterhorhabditis marelatus isolate k, or Steinernema glaseri. Detection frequency indicates the proportion of replicates of 18 total replicates (trials 1–3 combined) with the indicated fungus.
more frequently with H. marelatus-b than with H. marelatus-k or S. glaseri. Heat-treatment of soil eliminated nematophagous fungi from all arenas of trials 2 and 3. In trial 1, two of 18 arenas with heat-treated soil and H. marelatus-b parasitized wax moth larvae contained A. oligospora (10 propagules/g soil) and one arena with heat-treated soil and S. glaseriparasitized wax moth larvae contained A. oligospora (1199 propagules/g soil) (Fig. 5A). In an analysis of A. oligospora population density that only included data from nonheat-treated soil, the effects of trial and nematode isolate were significant. But because the interaction of trial and isolate was not significant, the data from the three trials were combined to simplify the presentation. All three nematode isolates supported large A. oligospora population densities, but A. oligospora population density was 11–17 times greater with S. glaseriparasitized wax moth larvae than with H. marelatusparasitized wax moth larvae (Fig. 5A). As in experiment 1, M. glutinosporum was the most common fungus after A. oligospora in experiment 3. It was not detected in heat-treated soil, but its overall population density in nonheat-treated soil was 185 propagules/g of soil if all replications were included and O500 propagules/g soil if arenas without M. glutinosporum were excluded (Fig. 5B). In nonheat-treated soil, the total population density of nematode-trapping fungi was 500, 712, or 4811 propagules/g soil when moth larvae were parasitized with H. marelatus-b, H. marelatus-k, or S. glaseri, respectively. The percentages of the total made up of M. glutinosporum or A. oligospora were 13 and 83% with H. marelatus-b, 60 and 38% with H. marelatus-k, and 1 and 98% with S. glaseri.
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Fig. 5. Population densities of Arthrobotrys oligospora and Myzocytium glutinosporum in heat-treated soil (solid bars) and nonheat-treated soil (open bars) as affected by nematode-parasitized insects (experiment 3). Values are the meansC1 standard error of 18 replicates (trials 1–3 combined).
In arenas containing nonheat-treated soil, A. oligospora sporulated on the soil surface in 28, 22, and 28% of the arenas containing H. marelatus-b, H. marelatus-k, and S. glaseri-parasitized wax moth larvae, respectively. But sporulation on the soil surface was seldom heavy and was not a good indicator of population density in soil. For example, 13 of 18 arenas with S. glaseriparasitized larvae exhibited no A. oligospora sporulation but contained 3168G747 A. oligospora propagules/g of soil. The strength of top-down effects of trapping fungi and other natural enemies on nematodes was measured as the fraction of dispersing nematodes that reached the moat in arenas with heat-treated vs nonheat treated soil. We excluded the three arenas of heat-treated soil that contained A. oligospora. H. marelatus-k dispersal was 22% less (P!0.05) and S. glaseri dispersal was 29% less (PZ0.08) in nonheat-treated soil than in heat-treated soil (Fig. 6A and C). In contrast, H. marelatus-b dispersal was greater in nonheat-treated than in heat-treated soil (Fig. 6B). Many more H. marelatus than S. glaseri were recovered from moats (Fig. 6A and B vs C). Most S. glaseri recovered from moats had dispersed there by day 18, but most H. marelatus did not disperse to moats until day 25 or 32.
Fig. 6. Cumulative numbers of entomopathogenic nematodes recovered from arena moats (experiment 3) as affected by heat treatment of soil and species of nematode used to infect the insects. Wax moth larvae were exposed to entomopathogenic nematodes on day 0 and placed in arenas on day 4; experiment 3 was ended on day 45. A (Heterhorhabditis marelatus isolate k), B (H. marelatus isolate b), and C (Steinernema glaseri). Values are the meansG1 standard error of 18 replicates (trials 1–3 combined).
After they emerged from the insect cadaver, S. glaseri but not H. marelatus infective juveniles aggregated and remained on the surface of the cadaver for several days. Steinernema glaseri-parasitized cadavers were more degraded than H. marelatus-parasitized cadavers at the end of experiment 3; the percentage degradation was 64G2% for H. marelatus-parasitized cadavers, regardless of isolate or soil treatment, whereas the percentage degradation for S. glaseri-parasitized cadavers was 78G4% in heat-treated soil and 92G1% in nonheat-treated soil. In nonheat-treated soil, more fungal hyphae (unidentified) surrounded S. glaseriparasitized cadavers than H. marelatus-parasitized cadavers.
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The biomasses of H. marelatus and S. glaseri infective juveniles were calculated using a published formula (Andra´ssy, 1956) and published morphometric data (Liu and Berry, 1996; Poinar, 1978; Stock, 1997). The calculated biomass of a H. marelatus or S. glaseri infective juvenile was 0.32 or 1.47 mg, respectively. If the mean numbers of nematodes in the moats (about 175,000 for H. marelatus and 25,000 for S. glaseri) were used to estimate the number of nematodes produced per wax moth larva, then each larva produced about 56 mg of H. marelatus or about 37 mg of S. glaseri. Nematode and fungus numbers were uncorrelated when the analyses included data from both heat-treated and nonheat-treated soil. When the analyses included data only from nonheat-treated soil, numbers of H. marelatus-k in the moat were positively correlated with the population density of A. oligospora (PZ0.04); other correlations were not significant. Soil water at the end of experiment 3 was unaffected by nematode isolate or the interaction of trial and soil treatment, but was significantly affected by trial and soil treatment. Averaged over soil treatments, soil water, initially at 32.0%, was 31.2, 28.0, and 25.1% at the end of trial 1–3, respectively. Averaged over the three trials, soil water content was 28.6% in nonheat-treated soil and 27.6% in heat-treated soil. One arena in trial 3 of experiment 3 was discarded because the soil had become saturated.
4. Discussion As predicted, the population density of several nematode-trapping fungi increased substantially when nematodeparasitized insects were added to BMR soil. The strongest numerical response was by A. oligospora; in the presence of parasitized insects, this fungus frequently ‘bloomed’ in that its population density increased 10–100 times and sometimes exceeded 10,000 propagules/g of soil. In some arenas the bloom was literal: A. oligospora conidiophores and conidia covered the soil surface. Three other fungi (M. glutinosporum, D. brochopaga, and G. gephyropaga) were seldom detected unless nematode-parasitized insects were present. Numbers of D. brochopaga and G. gephyropaga remained relatively small, but M. glutinosporum often increased to numbers as large as those of A. oligospora. The resources underlying this strong numerical response are unclear because the nematode-parasitized insect comprises several potential resources and because A. oligospora in particular is thought to be a ‘facultative’ trapper of nematodes, i.e. one that uses nematodes as a nitrogen source but decomposing organic matter as a carbon and energy source (Cooke, 1963; Barron, 1992). Additional research is needed to determine whether A. oligospora and the other fungi responded to the entomopathogenic nematodes, the dead insect, bacterivorous nematodes stimulated by the dead
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insect, or to some combination of these. We expect that other substrates at BMR (e.g. decomposing lupine leaves, mammal feces, isopod feces, dead arthropods not killed by nematodes, etc.) will also elicit numerical responses, but we doubt that these responses will be as strong as those documented in this study. Some trapping fungi were not enhanced by nematodeparasitized insects, and these were A. eudermata, A. musiformis, and A. paucospora. All three are similar to A. oligospora in that they use adhesive nets to capture nematodes. One possible explanation for this result is that the three species could not increase and were even suppressed because their niches overlap with that of A. oligospora. Koppenho¨fer et al. (1997) provided some evidence that A. oligospora competes with and suppresses A. eudermata and A. paucospora. In contrast to A. oligospora, M. glutinosporum has been infrequently studied. Its morphology and how it infects nematodes have been described (Barron, 1976), but its population biology in soil has not. The current study suggests that even though its zoospores swim poorly and encyst rapidly to form adhesive structures (Barron, 1976), M. glutinosporum prefers high soil water potentials. The dominant fungus in this study, A. oligospora, responded to both species of entomopathogenic nematode but much more to S. glaseri than to H. marelatus. Gross differences in nematode susceptibility cannot explain this result, because both nematodes are highly susceptible to the fungus on agar (Koppenho¨fer et al., 1996). But differences in susceptibility too small to detect on agar might be accentuated in soil. In particular, Heterorhabditis spp. retain their second-stage cuticle longer than Steinernema spp., and the extra cuticle can increase the chances of escaping a fungal trap (Timper and Kaya, 1989; Timper et al., 1991; Timper and Brodie, 1995). Other possible differences underlying the stronger response to S. glaseri than to H. marelatus include nematode biomass produced per insect, the timing of nematode emergence, the behavior of nematodes after emergence, the effect of nematode infection on the insect cadaver, and the effect of the symbiotic bacteria. Differences in nematode biomass, however, cannot explain the stronger effect of S. glaseri because S. glaseri produced less rather than more nematode biomass per insect. S. glaseri did emerge from the cadaver sooner than did H. marelatus, but how this might result in stronger enhancement of the fungi is unclear. Behavior after emergence seems a better explanation for the greater response of fungi to S. glaseri: unlike H. marelatus, S. glaseri aggregated on the cadaver for several days after emerging, and this may have increased exposure to adjacent fungi. Such behavior has been described for S. glaseri (Lewis and Gaugler, 1994) but not for H. marelatus. The effect of the nematode on the cadaver may also be important, because S. glaseri-parasitized cadavers degraded much faster and seemed leakier than H. marelatus-parasitized cadavers. Perhaps materials
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leaking from the cadaver supported saprophytic growth of nematode-trapping fungi. Differences in cadaver degradation could result from how the nematodes emerged or from differences in the species of symbiotic bacteria or their antibiotics (Zhou et al., 2002). To quantify nematode mortality caused by trapping fungi and other natural enemies, we compared numbers of nematodes that emerged and moved from parasitized insects through at least 5 cm of heat-treated (natural enemies removed) or nonheat-treated soil (natural enemies present) and into a moat. Mortality was 22% with H. marelatus-k and 29% with S. glaseri. In the case of H. marelatus-b, dispersal was greater in nonheat-treated than in heat-treated soil, suggesting an artifact or inadequate replication relative to the variance; a previous study reported a large variance in the number of H. marelatus produced per ghost moth or wax moth (Strong et al., 1996). Regardless, the data failed to support the hypothesis that trapping fungi kill substantial proportions of nematodes (O50%) emerging from parasitized insects (Strong, 1999). Given the very strong bottom-up effects of nematodeparasitized insects on trapping fungi, the relatively weak top-down effects described in the previous paragraph are difficult to explain. Perhaps the fungi responded too slowly, i.e. perhaps the numerical response occurred only after many nematodes had already dispersed. Perhaps the rapid emergence and dispersal of many nematodes simply overwhelmed the traps. Perhaps A. oligospora and M. glutinosporum are inefficient at trapping [for more data on the failure of A. oligospora to trap many nematodes in soil, see Jaffee (2002, 2003, 2004) but very efficient at converting captured nematodes into fungal propagules. Finally, these fungi may have used substrates other than or in addition to nematodes. The use of wax moth rather than ghost moth larvae requires discussion. As noted previously, we used wax moths in place of the natural ghost moths because ghost moth larvae are often unavailable. While sharing many similarities, wax moths and ghost moths also differ. For example, wax moths are more susceptible to H. marelatus infection (Strong et al., 1996); this difference, however, is probably unimportant because H. marelatus-parasitized ghost moths are abundant at BMR and because they produce at least as many nematodes as do parasitized wax moths of similar size (Strong et al., 1996). In contrast, moth larva size is probably important. Ghost moth larvae that hatch from eggs and move through soil to roots are smaller than last instar wax moth larvae and don’t become large until after they penetrate roots. It is reasonable to expect that small, nematode-parasitized ghost moth larvae in the soil will produce fewer nematodes and elicit a smaller fungal response than did last instar wax moths. But ghost moth larvae in roots, which can be as large or larger than the wax moths used in this study, are also parasitized by nematodes and are exposed to trapping fungi. This happens because roots penetrated by the larvae often
crack (unpublished observations, D.R. Strong). When the cracking is extensive, the large, nematode-parasitized ghost moth larvae are fully exposed to soil and to nematodetrapping fungi. It follows that data generated with nematode-parasitized wax moths may be similar to data generated from large, nematode-parasitized ghost moths. But because ghost moth larvae in the field range from small to large and because ghost moths may differ from wax moth in other important ways, future research should compare fungal responses to wax moths and ghost moths and should determine the significance of moth size. Future research should also explore the presence and activity of trapping fungi in the penetrated lupine roots for two reasons. First, as insects move through soil, they can superficially acquire the conidia of trapping fungi (Fowler and Garcia, 1989), and we hypothesize that ghost moths in lupine root galleries are frequently contaminated with A. oligosopora. Second, A. oligospora has cellulolytic activity (Barron, 1992; Jaffee, 2004), and the combination of dead lupine wood and nematode-parasitized insects might induce both strong numerical responses and abundant trapping of nematodes. In addition to being relevant to the community ecology of BMR, the current data are also relevant to the general ecology of nematode-trapping fungi and to biological control of plant-parasitic nematodes. From the perspective of fungal ecology, the very large numerical response to nematode-parasitized wax moths is exciting and worth understanding. That trapping fungi can be enhanced by organic amendments to soil is well documented (Stirling, 1991), but to the best of our knowledge no amendment has elicited so large a response as that reported here by nematode-parasitized insects. And from the perspective of biological control, at least some researchers who study natural enemies of plant-parasitic nematodes will want to understand this numerical response, how it relates to nematode mortality, and how it might be used to manage pest nematodes. Two important questions are: ‘Which resources are the fungi using?’ and ‘Will other soils exhibit both strong bottom-up effects on trapping fungi and strong top-down effects on nematodes?’ Acknowledgements We thank Harry Kaya and Francine Farrell for comments on an earlier version of this paper. References Andra´ssy, I., 1956. The determination of volume and weight of nematodes. Acta zoologica (Hungarian academy of science) 2, 1–15, in: Zuckerman, B.B., Brzeski, M.W., Deubert, K.H. (Eds.), English Translation of Selected East European papers in Nematology, 1967. University of Massachusetts, Cranberry Experiment Station, East Wareham, Massachusetts, pp. 73–84. Barron, G.L., 1976. Nematophagous fungi: three new species of Myzocytium. Canadian Journal of Botany 22, 752–762.
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