Research Article Received: 9 January 2009
Revised: 23 March 2009
Accepted: 18 May 2009
Published online in Wiley Interscience: 26 June 2009
(www.interscience.wiley.com) DOI 10.1002/ps.1809
Biological control of Lycoriella ingenua (Diptera: Sciaridae) in commercial mushroom (Agaricus bisporus) cultivation: a comparison between Hypoaspis miles and Steinernema feltiae Stephen Jessa∗ and Heinrich Schweizerb Abstract BACKGROUND: Mushroom cultivation may be adversely affected by insect pests, including sciarids (Lycoriella spp.), which were previously controlled by application of chemical pesticides. However, owing to food safety and environmental concerns, availability of pesticides for use during mushroom cultivation has diminished. Consequently, it is imperative to investigate alternative control strategies, not reliant on chemical pesticides, which may be used in an integrated pest management system. RESULTS: Application of the predatory mite Hypoaspis miles Berlese to commercial mushroom-growing beds at the beginning of spawn run or just prior to casing (830 mites m−2 ) significantly reduced immature sciarids, Lycoriella ingenua (Dufour), in the growing substrate and also adult activity towards the conclusion of cropping. A trend towards lower sciarid emergence from substrates and reduced adult sciarid activity was observed following the application of Steinernema feltiae (Filipjev) (1.5 × 106 nematodes m−2 ) at casing. No significant treatment effects on mushroom yield were observed. However, contamination of the mushroom crop by adult sciarids increased in untreated controls. Application of H. miles required a 12-fold increase in labour when compared with application of S. feltiae. CONCLUSION: Contingent upon the development of an effective application system, H. miles has potential for the biological control of sciarids in commercial mushroom production. c 2009 Society of Chemical Industry Keywords: Agaricus bisporus; mushroom culture; Lycoriella spp.; Sciaridae; Hypoaspis miles; biological control
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INTRODUCTION
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Previously, insect pest control within mushroom cultivation was relatively uncomplicated, involving the application of chemical pesticides to the compost or casing substrates.13 However, there are a number of problems associated with chemical control strategies, including insect pest resistance and environmental and food safety issues. This accentuates the need for novel, sustainable approaches to insect pest control during mushroom cultivation that integrate biological, cultural and chemical control strategies.14 – 16 Commercial use of the entomopathogenic nematode Steinernema feltiae (Filipjev) for biological control of sciarids during mushroom cultivation began in the mid-1990s and is now widely
∗
Correspondence to: Stephen Jess, Applied Plant Science, Agri-Food and Biosciences Institute, Newforge Lane, Belfast BT9 5PX, UK. E-mail:
[email protected]
a Applied Plant Science, Agri-Food and Biosciences Institute, Newforge Lane, Belfast BT9 5PX, UK b Department of Applied Plant Science, Queen’s University of Belfast, Agriculture and Food Science Centre, Newforge Lane, Belfast BT9 5PX, UK
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Cultivation of the common mushroom Agaricus bisporus (Lange) Imbach is commonly affected by two major classes of insect pest, sciarids and phorids.1 Sciarids Lycoriella spp. (Diptera: Sciaridae) are a frequent problem, particularly in production systems in which compost is transferred from composting facilities into mushroomgrowing houses before it is colonised by the mycelium (spawn running occurs in the mushroom-growing house).2,3 Subsequent to taxonomic reclassification, two species, Lycoriella ingenua (Dufour) (= L. mali = L. solani) and L. castanescens (Lengersdorf) (= L. auripila) are considered mushroom pests.4 – 6 Phorids Megaselia spp. (Diptera: Phoridae) commonly invade compost that is already colonised by the mycelium.7,8 Phorids are considered to be a seasonal, sporadic pest, with increased importance in production systems where the mushroom mycelia develop in the compost (spawn running occurs at compost manufacture) before it is transferred into the growing houses.9 The major economic impact of Megasalia halterata has been attributed to adults vectoring fungal pathogens, especially Verticillium spp.10 – 12
www.soci.org used among commercial mushroom growers in Europe.17 Currently, it is the only available biological control agent, and thus the standard for biological pest control in this crop in Europe. Although S. feltiae is effective against sciarids, no commercially relevant experiment has demonstrated any convincing controlling effects on phorid populations.17 Bacillus thuringiensis (Berliner) var. israeliensis, which is successfully used against sciarids in northern America, is not approved for use on mushroom crops in the major mushroom production countries in Europe. Consequently, there is a requirement to investigate alternative biological agents that may provide effective control of sciarids and phorids during mushroom cultivation. Predatory mites from the genus Hypoaspis (Acari: Hypoaspidae) have demonstrated some potential for control of sciarids and phorids in small-scale laboratory trials, and some semi-field trials have provided encouraging but inconclusive results.13,15,18,19 It has been demonstrated that the level of control achieved with H. miles Berlese may be comparable with that of S. feltiae.13 Hypoaspis miles is commercially available as a biological agent to control sciarids (Bradysia spp.) in glasshouse crops (Biological Crop Protection Ltd, Ashford, Kent, UK). Therefore, considering the commercial availability of H. miles, its efficacy in sciarid control and, furthermore, its potential to control phorids, there is merit in further evaluation of this predatory mite as a biological control agent of the principal insect pests on a commercial scale of mushroom cultivation. During previous semi-commercial studies involving the use of Hypoaspis spp. mites during mushroom cultivation, weed moulds (Mucor spp.) and concomitant yield reductions associated with mite applications were observed.19 During these studies, and in common with recommended applications in glasshouses, mites were applied by distribution of the carrier material (vermiculite–peat mixture) across the mushroom-growing substrate.19 Subsequent tests indicated that the mould inoculum most probably originated from direct contact between the carrier material and the mushroom-growing substrate. Considering that mites will disperse some distance over the growing substrates, it was considered that point releases from containers (e.g. vials, sachets) may minimise the contact between mite carrier and mushroomgrowing substrate and thus reduce the weed mould and associated problems.15,20,21 The strategy proposed specifically for this experiment involved placing waxed paper sachets containing the peat–vermiculite carrier and the mites onto the substrate at a density of one sachet per block of compost (40 × 60 cm). The sachets were designed to facilitate the escape and dispersal of mites throughout the mushroom-growing substrate. This study compares the performance of single applications of H. miles (830 mites m−2 ) and S. feltiae (1.5 ×106 m−2 ) in the biological control of sciarids during commercial mushroom cultivation.
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MATERIALS AND METHODS
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2.1 Mushroom cultivation system The experiment was conducted at a commercial mushroom production farm in County Tyrone, Northern Ireland (Monaghan Mushrooms Ltd, Monaghan, Ireland). Cultural procedures followed an industry standard schedule with Phase II compost (Monaghan Mushroom Compost Ltd, Ireland) prepared from wheat straw and chicken litter providing the growing medium. Mushrooms were grown inside insulated polythene tunnels with 9 × 34 m ground floor dimension. Each tunnel contained three stacks of three
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shelves (each shelf 30 m long by 1.2 m wide) providing a total growing surface area of 324 m2 tunnel−1 . At the beginning of the growing cycle, shelves were filled with blocks (22 kg; 60×40×18 cm) of Phase II compost inoculated with A. bisporus spawn (Sylvan L81 ; Sylvan Spawn Ltd, UK) at a rate of 0.5% (w/w). Immediately after filling the shelves with the blocks, the top cover of the polythene block-wrap was removed and the compost was mechanically ruffled so that the remaining plastic wrap on the sides of the blocks was shredded and the surface over the entire shelf became uniformly flat. The surface of the compost was covered with moistened tissue paper to prevent desiccation. After a spawn-running period of 14 days with a mean compost temperature of 25 ◦ C, the paper was removed and the compost was again mechanically ruffled. The following day, a 5 cm layer of casing (a limestone and sphagnum peat mixture, 15 kg limestone 300 L peat−1 ; McDon Peat Ltd, Ireland) was applied to the surface of the compost. The casing layer was mechanically ruffled, and a case-run period with 22–23 ◦ C air temperature was initiated. Following 7 days of case running, the air temperature was reduced to 17 ◦ C and mushrooms were harvested in accordance with standard harvesting practices. The culture cycle, from the beginning of spawn running, including case running, crop initiation and harvesting of three flushes, was completed in 52 days. 2.2 Experimental design Four treatments were compared: (1) untreated control; (2) S. feltiae immediately after casing (standard biological control) 1.5 × 106 m−2 (SCIA-RID ; Koppert, Wadhurst, East Sussex, UK) applied in 1 L water m−2 ; (3) H. miles at the beginning of spawn running, applied 1 day after filling the shelves with compost, 830 mites m−2 ; (4) H. miles applied 1 day before casing, 830 mites m−2 . The mites were applied by placing onto the surface of each block of compost (60 × 40 cm) one sachet (6 cm ×11 cm) which contained approximately 200 mites in a peat and vermiculite carrier material (Hyposure(M) ; Biological Crop Protection Ltd, Ashford, Kent, UK). To allow the mites to escape, each sachet was torn open across the entire length of one of the 6 cm sides at the time of application. Each experimental unit comprised an entire growing house with nine shelves (1.2 × 30 m) and a 324 m2 growing area containing approximately 25 t of compost. On each of 4 days in 1 week, one house was filled with compost. Each of these houses was allocated one of the four treatments described, and the four houses were considered as one replicate. The experiment comprised four replicates (4 weeks ×4 houses = 16 houses). Each treatment was allocated once to the first, once to the second, once to the third and once to the fourth house filled in a particular week, resulting in a balanced design conforming to a Latin square. 2.3 Data collection Adult sciarid populations were monitored with four light traps in each house. Each trap consisted of an 11 W fluorescent lowenergy bulb (Sylvania Mini-Lynx, colour 827/600 lumens) in a light baton fitted into a neutral grey-coloured plastic plant pot (18.0 cm; Richard Sankey & Son Ltd, Nottingham, UK). A doublesided yellow sticky trap (10 × 21 cm area; Agralan Ltd, Swindon, Wiltshire, UK) suspended below the pot by means of wire hooks trapped flying insects. Monitoring commenced 6–9 days after the beginning of spawn run and continued for 6 weeks until the end of cropping. After an initial trapping period of 11 days, sticky traps were changed weekly. Sticky traps were examined under a stereo
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Biological control of sciarids in commercial mushroom culture microscope, and, owing to the high numbers of insects per sticky card, subsamples of four strips (two per side) of 2 cm width and 10 cm length were examined and the numbers of adult sciarids were recorded. Populations of immature sciarids in the compost or casing were assessed by collecting substrate samples immediately before casing (1–4 days pre-casing), during the early part of the harvest (10–13 days post-casing) and late during the harvest (24–27 days post-casing). The initial samples were taken in advance of firstgeneration adult emergence and considered to reflect the level of ingressing populations. Samples were collected from the centre of the growing tunnels (at least 10 m from the front or rear end of the tunnel). At the two side stacks, one sample was taken from each shelf; at the middle stack, two samples were taken from each shelf. Therefore, on each occasion, 12 samples were collected from each house. Each sample consisted of approximately 200 mL of substrate material excavated from the top 5 cm of the mushroomgrowing bed. In the laboratory, the samples were incubated at 24 ◦ C for 21 days in disposable plastic food containers (400 cm3 ) with discs of yellow sticky traps (5 × 5 cm) attached to the inside of the lid in order to trap and record emerging adult flies. After the incubation (fly emergence) period, the substrate from the samples was dried at 75 ◦ C to determine the dry weight of material collected. For logistical reasons, the change of the light-trap sticky cards and collection of the substrate samples occurred on the same day for all four houses that were filled during a particular week. As the four houses were filled on four consecutive days, trapping periods and sampling days in relation to the cropping cycle differed accordingly for the four houses. Exact days relative to the casing day are displayed in Fig. 1. In addition to monitoring sciarid populations, total mushroom yields from each house were obtained from the grower.
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Table 1. Effect of different biological control treatments on mushroom yield Mean yield (95% confidence limits) (kg m−2 )
Treatment Untreated control S. feltiae 1 day after casing H. miles 1 day after house filling H. miles 1 day before casing
16.77 (15.37–18.30) 16.39 (15.02–17.88) 16.61 (15.22–18.13) 17.41 (15.95–19.00)
appropriate analysis of variance (PROC GLM; SAS, 1999) with the week of filling the house, the day of filling the house and the treatment as effects.
3
RESULTS
Adult emergence from substrate samples taken before casing was similar across all treatments (Fig. 1A). Hypoaspis miles applied immediately before casing significantly reduced immature sciarids in the substrate during the early part of the mushroom harvest (P < 0.1) (Fig. 1A). Hypoaspis miles applied at the beginning of spawn running also decreased adult sciarid emergence (P < 0.05), and with S. feltiae as casing drench, a trend towards reduced emergence of adult flies was observed on this sampling occasion (Fig. 1A). Adult emergence from the final substrate samples increased considerably compared with earlier sampling occasions with no differences observed between treatments. Adult sciarid activity towards the end of the cropping cycle exhibited a similar pattern, with significant controlling effects of both H. miles treatments and a trend towards reduced activity associated with the application of S. feltiae (Fig. 1B). No treatment effects on mushroom yield were observed (Table 1). However, anecdotal information from the grower indicated that adult sciarids in the untreated control were contaminating the crop to such a degree as to result in downgrading of some mushrooms, with associated monetary losses. In one growing house, where mites were applied immediately before casing, patches of a weed mould (Mucor spp.) were observed on the casing. The location of the patches tended to coincide with the location of mite sachets. In this house, the vigour of mycelial growth during spawn was substandard, and therefore the casing was ruffled deeper than usual in order to mix some of the already spawn-run compost into the casing. No other problems associated with mite applications, such as contamination of mushrooms or nuisances to workers, were reported. However, the labour required to apply the mite sachets to the compost surface was substantially higher (approximately 3 man-hours house−1 ) than for applying the nematodes (approximately 0.25 man-hours house−1 ).
4
DISCUSSION
In this experiment, which was conducted in commercial A. bisporus cultures, the performance of the predatory mite H. miles as a biological control agent of L. ingenua was equal or superior to that of the nematode S. feltiae at the specified application rates. Adult emergence from early substrate samples, taken before casing, provides some measure of ingressing sciarid population and was similar across all treatments (Fig. 1A). Therefore, comparing the
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2.4 Quantitative data analysis The numbers of flies from the four sticky card subsamples of each light trap were adjusted to obtain the numbers of flies per trap during the trapping period in each house. The numbers of emerged sciarids from each of the 12 substrate samples were adjusted according to the dry weight of the samples and combined from each sampling occasion and growing house to obtain the number of flies emerged kg−1 dry substrate. The insect counts conformed to repeated-measures designs (six sampling periods of light traps per house, three sampling occasions of substrate samples per house) superimposed to the Latin square design of allocating the growing houses to the treatments. Therefore, following square root transformation (y = x 1/2 ) to obtain independence of variance from the means, the count data were analysed using mixed linear models (PROC MIXED; SAS, 1999).22 The treatment, sampling time (trapping period for light traps, sampling occasion for substrate) and their interaction were included as fixed effects. The week of filling the house, the day of filling the house (Latin square design) and, owing to repeated measures, the house were specified as random effects. Treatment effects were examined by comparing the least-square means within each sampling time. To test whether any of the three treatment methods reduced the numbers of sciarids, each one was compared with the untreated control. Consequently, for each sampling time, three planned one-tailed comparisons were made, with the significance levels adjusted according to Sidak.23 The yield data conformed to a simple Latin square design and, after logarithmic transformation, were analysed by means of an
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Emergence of adult L. ingenua from substrate samples (number of adults per kg dry weight)
A
S Jess, H Schweizer
1400 1200 1000
Untreated control S. feltiae one day after casing H. miles at begin spawn running H. miles one day before casing
800 600 400
(*) *
200 0 1 - 4 days before casing
10 - 13 days after casing
24 - 27 days after casing
Time of substrate collection
Adult L. ingenua per four light traps
B
25000
20000
15000 **
10000
5000
0 (-8, -5) to (3, 6)
(3, 6) to (10, 13)
(10, 13) to (17, 20)
(17, 20) to (24, 31)
(24, 27) to (31, 34)
(31, 34) to (38, 41)
Trapping period [days relative to casing (precasing times are negative)] Figure 1. Effect of different biological control treatments on the sciarid population in commercial mushroom-growing houses: (A) immature sciarids in compost and casing; (B) adult sciarid activity. Average values with 95% confidence limits are plotted. The sampling times and beginning and end of trapping periods relative to casing time vary by 4 days. Thus, in B, (−8, −5 to 3, 6) indicates that in one house of each replicate the period lasted from day 8 before casing until day 3 after casing, in another house it lasted from day 7 before casing until day 4 after casing, etc. Asterisks indicate the statistical significance of three planned one-tailed comparisons. The tested hypothesis was as follows: is the fly level reduced in the biological control treatment compared with the untreated control? ∗ P < 0.05, (∗ ) P < 0.01, levels adjusted according to Sid´ak23 for the three comparisons per sampling time or period.
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treatments within each of the later sampling times is considered to reflect the efficacy of the different control measures. The potential of H. miles, especially when applied immediately before casing, is evident from the reduction in immature sciarids emerging from the substrate collected during the early part of the mushroom harvest (Fig. 1A) and the lower adult sciarid activity towards the end of the crop (Fig. 1B). Late mite applications, immediately before casing, tended to perform better than early applications, at the beginning of spawn running, which contrasts with findings from small-scale laboratory experiments.15,19 However, in these laboratory experiments, early mite applications were applied to the casing surface, and consequently the movement of the predatory mite towards immature sciarids in the compost may have been somewhat impeded by the casing layer.24 In the present study,
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mites were released onto the compost shortly before application of the casing layer, which may have afforded the mites greater access to immature sciarids in the compost. Additionally, predatory mites applied directly onto the compost would have greater access to the first generation of sciarids than the nematodes, which may have contributed to the improved efficacy compared with the nematodes. The increased adult sciarid emergence from final substrate samples and the absence of treatment effects suggest a lack of persistence across all treatments. While later-occurring sciarid infestations are less important in this three-harvest production system, there may be implications for potential infestation of subsequent crops. This lack of persistence of both nematodes and mites has been observed previously.13,25
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Biological control of sciarids in commercial mushroom culture
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The predatory mite H. miles applied immediately before casing performed remarkably well in controlling sciarid pests of mushroom cultures. Neither adverse effects on quantitative or qualitative mushroom yield nor any nuisance to workers were observed. Contingent upon the development of a cost-effective application method, H. miles may hold true potential as a biological control agent against sciarids in mushroom cultures.
ACKNOWLEDGEMENTS The authors are grateful to Monaghan Mushroom Ltd, and in particular Mr Conor McGarvey, for provision of facilities and management of the crop during the experiment. Biological Crop Protection Ltd supplied the predatory mites and requisite packaging. Ms Heather Knowles and Mr John Anderson provided essential technical assistance during this study. Discussions with John Dale (Biological Crop Protection Ltd) and Bernard Desrumeaux (Provincial Centrum voor Land- en Tuinbouw Proefcentrum voor de Champignonteelt vzw, Roeselare, Belgium) provided valuable input for the design of this study. Funding for the project was provided by a European Union CRAFT award QLK5-CT-2002-70693.
REFERENCES 1 Fletcher JT, White PF and Gaze RH, Mushrooms: Pest and Disease Control, 2nd edition. Intercept, Andover, Hants, UK (1989). 2 Hussey NW and Gurney B, Biology and control of the sciarid Lycoriella auripila (Winnertz) (Diptera: Lycoriidae) in mushroom culture. Ann Appl Biol 62:395–402 (1968). 3 Binns ES, Field and laboratory observations on the substrates of the mushroom fungus gnat Lycoriella auripila (Diptera: Sciaridae). Ann Appl Biol 96:143–152 (1980). 4 Menzel F, Sciaridae. Checklists of insects of the British Isles (new series). Part 1: Diptera. Handbk Ident Br Insects 12:20–24 (1998). 5 Menzel F and Mohrig W, Revision der palaarktischen Trauermucken (Diptera: Sciaridae) unter besonderer Beruckuchtigung der deutschen Fauna. Studia Dipterologica 3:663 pp. (1998). 6 Menzel F and Mohrig W, Revision der palaarktischen Trauermucken (Diptera: Sciaridae). Studia Dipterologica 6:1–720 (1999). 7 Hussey NW, Recent work on the control of the Worthing phorid. Mushroom Growers Assoc Bull 144:495–505 (1961). 8 Richardson PN and Hesling JJ, Laboratory rearing of the mushroom phorid Megaselia halterata (Diptera: Phoridae). Ann Appl Biol 88:211–217 (1978). 9 Scheepmaker JWA, Geels FP, van Griensven LGLD and Smits PH, Substrate dependent larval development and emergence of the mushroom pests Lycoriella auripila and Megaselia halterata. Entomol Exp Appl 79:329–334 (1996). 10 Gandy DG, A technique for screening bacteria causing brown blotch of cultivated mushrooms. 1967 Annual Report, Glasshouse Crops Research Institute, Littlehampton, UK, 1967, pp. 150–154 (1968). 11 Cross M and Jacobs L, Some observations on the biology of spores of Verticillium malthousei. Mushroom Sci 8:239–244 (1968). 12 White PF, Spread of the mushroom disease Verticillium fungicola by Megaselia halterata (Diptera: Phoridae). Prot Ecol 3:17–24 (1981). 13 Jess S and Kilpatrick M, An integrated approach to the control of Lycoriella solani (Diptera: Sciaridae) during production of the cultivated mushroom (Agaricus bisporus). Pest Manag Sci 56:477–485 (2000). 14 White PF, Biological control of mushroom pests: an evaluation. Proc Internat Cong on the Science and Cultivation of Edible Fungi. Vol. 2, Balkema, Rotterdam, The Netherlands, pp. 475–484 (1995). 15 Jess S and Bingham JFW, Biological control of sciarid and phorid pests of mushroom with predatory mites from the genus Hypoapsis (Acari: Hypoaspidae) and the entomopathogenic nematode Steinernema feltiae. Bull Entomol Res 94:159–167 (2004). 16 Smith JE, Challen MP, White PF, Edmondson RN and Chandler D, Differential effect of Agaricus host species on the population development of Megaselia halterata (Diptera: Phoridae). Bull Entomol Res 94:159–167 (2006).
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In relation to nematodes, limited production of subsequent generations of infective juveniles reduces recycling. This may be due to overinvasion of sciarid larvae by nematodes at high application rates.26 As a consequence, fewer nematodes reach adulthood, reducing subsequent generation of infective juveniles. Alternatively, the moisture content of the substrate may be limiting.27 Reduced moisture content of the substrate may also affect the activity of predatory mites. Further work is required to establish and resolve the issues relating to lack of persistence of these biological control agents. The observed reduction in sciarid populations attributed to the biological control agents was not associated with a corresponding increase in mushroom yield. Unfortunately, the farm management and the buyer of the crop considered that detailed economics of products were commercial-in-confidence. Therefore, no sound information was available to evaluate economics except in the broadest terms. Studies on L. castanescens Lengersdorf have demonstrated significant linear relationships between the mean number of sciarid larvae and the yield at all crop stages, and deduced an economic threshold of one larva 125 g−1 .28 However, it has been demonstrated that yield losses associated with L. ingenua infestation occurred only in late flushes, with no loss of total yield, and an injury threshold of 13 larvae 125 g−1 was determined.29 The levels of adult emergence recorded in the present study indicate that larval numbers considerably exceeded the injury threshold. In addition, reports from the grower indicate that the quality of mushrooms produced in the controls was adversely affected, resulting in monetary losses. Consequently, the reduced contamination of the crop with flies and the consequent increase in yield quality are sufficient motivation for growers to engage in sciarid control. The observed patches of weed mould (Mucor spp.) in one of the houses that received late mite application tended to coincide with the locations where the mite sachets were deposited. This suggested that the inoculum for the weed mould originated from the sachet. The deeper than normal ruffling, in order to incorporate some mushroom mycelium from the compost into the casing, is likely to have resulted in the shredding of the mite sachets, with concomitant spreading of the mite carrier material. However, in a subsequent attempt to reproduce the problem, by deep ruffling of the casing layer with mite sachets beneath, no evidence of weed moulds was observed. Possibly, in addition to shredded mite sachets due to deep ruffling, reduced vigour in A. bisporus mycelial growth may be required. Hence, under optimum growing conditions, weed mould may not be a significant problem. In addition, mushroom yields from the house were unaffected by the weed mould, which also indicates the relatively benign nature of the weed mould. The major disadvantage associated with the use of H. miles as a biocontrol method in this commercial-scale experiment was the relatively high labour requirement (3 h per application compared with 15 min for the application of nematodes). The distribution of the sachets was the main contributor to this high labour requirement. However, considering the dispersal potential of H. miles, it may be possible to reduce the density of mite release points considerably. In this study, one mite sachet was applied per 0.24 m2 surface area. By increasing the numbers of mites per sachet accordingly, the density of sachets might reasonably be reduced fivefold into the range of 1 sachet per 1.2 m2 surface area. This would not only reduce labour input, but, by decreasing the number of release stations, might also reduce the risk of weed mould contamination.
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www.soci.org 17 Jess S, Schweizer H and Kilpatrick M, Mushroom applications, in Nematodes as Biocontrol Agents, ed. by Grewal PS, Ehlers R-U and Shapiro DI. CAB International, Wallingford, Oxon, UK, pp. 191–213 (2005). 18 Lind R, Control of mushroom flies with the predatory mite, Hypoaspis miles. Contract Report Horticultural Development Council Project M9, Horticultural Development Council, West Malling, Kent, UK, 28 pp. (1993). 19 Jess S, Desrumaux B and Schweizer H, Predation of mushroom sciarids Lycoriella castanescens (Diptera: Sciaridae) and phorids Megaselia halterata (Diptera: Phoridae) by mites from the genus Hypoaspis (Acari: Hypoaspidae). Crop Prot (unpublished data). 20 Ydergaard S, Enkegaard A and Brdsgaard H, The soil-dwelling predatory mite Hypoaspis miles: temperature-dependent development, reproduction and longevity as well as the extent of migration to the above soil parts of plants. SP Rapport – Statens Planteavlsfors 4:305–316 (1996). 21 Workman PJ and Martin NA, Effect of pesticides on cymbidium orchid pollen-cap mite and its predator Hypoaspis sp. New Zealand Plant Prot Soc 55:380–384 (2002). 22 Sokal RR and Rohlf FJ Biometry: the Principles and Practice of Statistics in Biological Research, 3rd edition. WH Freeman and Company, New York, NY (1995).
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23 Sid´ak Z, Rectangular confidence regions for the means of multivariate normal distributions. J Am Statistical Assoc 62:626–633 (1967). 24 Cantelo WW, Movement of Lycoriella mali (Diptera: Sciaridae) through mushroom-growing medium. J Econ Entomol 81:195–200 (1988). 25 Scheepmaker JWA, Geels FP, Smits PH and van Griensven LGLD, Control of the mushroom pests Lycoriellaauripila (Diptera: Sciaridae) and Megaselia halterata ((Diptera: Phoridae) by Steinernema feltiae (Nematoda: Steinernematidae) in field experiments. Ann Appl Biol 131:359–368 (1997). 26 Grewal PS and Richardson PN, Effects of application rates Steinernema feltiae (Nematoda: Steinernematidae) on biological control of the mushroom fly Lycoriella auripila (Diptera: Sciaridae). Biocont Sci Technol 3:29–40 (1993). 27 Tomalak ML and Lippa JJ, Factors affecting entomophilic activity of Neoaplectana feltiae in mushroom compost. Entomol Exp Applic 59:105–110 (1991). 28 White PF, The effect of sciarid larvae (Lycoriella auripila) on cropping of the cultivated mushroom (Agaricus bisporus). Ann Appl Biol 109:11–17 (1986). 29 Kielbasa R and Snetsinger R, Life history of the sciarid fly L. mali and its injury threshold on the commercial mushroom. Bulletin 833, Pennsylvania State University, College of Agriculture, 11 pp. (1981).
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