Evidence Project Final Report - Defra,...

EVID4 Evidence Project Final Report (Rev. 10/14) of 1

General Enquiries on the form should be made to:

Defra, Strategic Evidence and Analysis E-mail: [email protected]

Evidence Project Final Report

Note

In line with the Freedom of Information Act 2000, Defra aims to place the results of its completed research projects in the public domain wherever possible. The Evidence Project Final Report is designed to capture the information on the results and outputs of Defra-funded research in a format that is easily publishable through the Defra website An Evidence Project Final Report must be completed for all projects.

This form is in Word format and the boxes may be expanded, as appropriate.

ACCESS TO INFORMATION

The information collected on this form will be stored electronically and may be sent to any part of Defra, or to individual researchers or organisations outside Defra for the purposes of reviewing the project. Defra may also disclose the information to any outside organisation acting as an agent authorised by Defra to process final research reports on its behalf. Defra intends to publish this form on its website, unless there are strong reasons not to, which fully comply with exemptions under the Environmental Information Regulations or the Freedom of Information Act 2000.

Defra may be required to release information, including personal data and commercial information, on request under the Environmental Information Regulations or the Freedom of Information Act 2000. However, Defra will not permit any unwarranted breach of confidentiality or act in contravention of its obligations under the Data Protection Act 1998. Defra or its appointed agents may use the name, address or other details on your form to contact you in connection with occasional customer research aimed at improving the processes through which Defra works with its contractors.

Project identification

1. Defra Project code PS2154

2. Project title

Alternative plant protection methods: Combined applications of microbial bio-insecticides and chemical pesticides for Integrated Pest Management of key insect pest species

3. Contractor organisation(s)

The University of Warwick

Coventry CV4 7AL

ADAS UK Ltd, Pendeford House, Pendeford Business Park, Wobaston Road, Wolverhampton, WV9 5AP

54. Total Defra project costs £103,601

(agreed fixed price)

5. Project: start date ................ 01/12/2013

end date ................. 28/02/2015

6. It is Defra’s intention to publish this form.

Please confirm your agreement to do so. ................................................................................... YES NO

(a) When preparing Evidence Project Final Reports contractors should bear in mind that Defra intends that they be made public. They should be written in a clear and concise manner and represent a full account of the research project which someone not closely associated with the project can follow.

Defra recognises that in a small minority of cases there may be information, such as intellectual property


or commercially confidential data, used in or generated by the research project, which should not be disclosed. In these cases, such information should be detailed in a separate annex (not to be published) so that the Evidence Project Final Report can be placed in the public domain. Where it is impossible to complete the Final Report without including references to any sensitive or confidential data, the information should be included and section (b) completed. NB: only in exceptional circumstances will Defra expect contractors to give a "No" answer.

In all cases, reasons for withholding information must be fully in line with exemptions under the Environmental Information Regulations or the Freedom of Information Act 2000.

(b) If you have answered NO, please explain why the Final report should not be released into public domain

Executive Summary

7. The executive summary must not exceed 2 sides in total of A4 and should be understandable to the intelligent non-scientist. It should cover the main objectives, methods and findings of the research, together with any other significant events and options for new work.

Insect pests significantly impair the production and quality of a wide range of food crops. At the moment,

pest control in most field-grown crops is based around applications of synthetic chemical insecticides.

These chemicals are very important for crop protection, but their excessive use can result in a number of

problems including damage to the environment, harm to non-target insects and the evolution of heritable

resistance in target pest populations. At the same time, the availability of conventional chemical pesticides

has declined markedly in the last ten years as many products have been taken off the market as a result

of increasingly strict regulations governing their authorization and use. It is generally accepted that the

way forward is to use Integrated Pest Management (IPM), a systems approach in which different crop

protection tools are used together with chemical pesticides in complementary ways. This should reduce

the pressure on pesticide use and prevent damage to the environment. IPM tools to be used in addition to

chemical pesticides include biological, cultural and physical controls, natural products, and breeding crops

for pest resistance. IPM forms a key part of new EU legislation on the Sustainable Use of Pesticides,

Directive 2009/128/EC. Under the Directive, there is now a legal obligation on farmers and growers to use

IPM where effective and practical. The Directive requires that biological controls and other non-chemical

methods should be preferred over conventional pesticides if they give satisfactory levels of control.

This project focused on the use of entomopathogenic fungi (“entomopathogenic” = “insect pathogenic”,

EPF) and entomopathogenic nematodes (EPNs), which are natural enemies of insects and can be used

as “biopesticides” of insect pests. The number of these products on the market is increasing. They have a

number of potential benefits including operator and bystander safety and lack of toxic residue. However

they are usually less efficacious than fully effective conventional insecticides and they can be adversely

affected by environmental conditions. The challenge is to make best use of the attractive properties of

entomopathogens while minimising their downsides. One way to do this could be to combine different

agents together in an IPM programme, for example by using two different entomopathogens together or by

combining an entomopathogen with a partially effective chemical insecticide, e.g. one that some insects in

a population may be resistant to, or one used at a reduced dose rate Some growers are already starting to

do this on some protected (glasshouse) crops. However, there is a severe lack of scientific information

available about how these different agents interact and how to use them in complementary ways,

particularly for pests that affect crops grown in the UK.

The purpose of this project was to investigate how combination treatments of biopesticides (EPF and

EPNs) and pesticides affect UK insect pests. We focused on the cabbage root fly (CRF), which is a major

pest of horticultural brassicas. The larvae of this pest feed on the roots of brassica plants. It is very

damaging and can be difficult to control even with conventional chemical insecticides. The current

pesticide used to control it, chlorpyrifos, is being phased out on safety grounds and is being replaced with

spinosad. However relying on one active ingredient for control of CRF increases the risk of the

development of pesticide resistance. Two biopesticides (the EPF Metarhizium brunneum, and the EPN

Steinernema feltiae) are available on the UK market and are being used as control agents of some

horticultural pests, mainly on protected crops but there is also interest in using them against CRF. These

biopesticides would be applied to compost modules used to raise brassica plants in the propagation

glasshouse before being transplanted to the field


The first part of the project consisted of a series of tests to investigate the effects of insecticides on the

survival and development EPF and EPNs, and also to study the compatibility of EPNs and EPF with each

other. For the experiments with insecticides, we started off by exposing EPF and EPNs to insecticides in

“in vitro” laboratory tests, and then investigated the effect of insecticides in compost and soil (which would

be more representative of insecticide exposure under propagation and field conditions). We used three

insecticides: chlorpyrifos, spinosad, and a coded product that was recently identified as having potential

for CRF control in a screening programme (‘SCEPTRE’) funded by the Agriculture and Horticulture

Development Board. There were some effects of insecticide treatment on M. brunneum in experiments

done in vitro, but the effect depended on the type of insecticide, its concentration, and the method by

which the fungus was exposed to it. When the germination of M. brunneum spores was quantified on an

agar medium supplemented with insecticides, spinosad caused a reduction in germination of up to a third

at 100% to 200% of the manufacturer’s recommended concentration, but it was not affected by

chlorpyrifos or the coded insecticide. In an experiment designed as a laboratory simulation of “tank

mixing” M. brunneum with an insecticide, the coded insecticide reduced germination and colony formation

in a concentration-dependent manner, but there was no effect of chlorpyrifos or spinosad. There was no

evidence of an adverse effect of any of the insecticides on M. brunneum in compost. A laboratory based

simulation of “tank mixing” S. feltiae with insecticides showed that spinosad and the coded insecticide had

no effect of S. feltiae viability at the manufacturer’s recommended concentration. However chlorpyrifos at

100, 50 and 25% of the manufacturer’s recommended concentration significantly reduced the number of

live nematodes, and therefore we would conclude that this insecticide is unlikely to be compatible with S.

feltiae in a tank mix, although more testing would be required to provide definitive proof. Adverse effects of

insecticides were also observed against S. feltiae when they were held together with the EPN in compost,

depending on the type of insecticide and its concentration. There was a significant negative effect of

chlorpyrifos on S. feltiae at 100% and 50% of the manufacturer’s recommended concentration, but not at

5% of the recommended concentration. There were also significant reductions in nematode viability when

S. feltiae was kept in compost with spinosad and the coded insecticide at 100% of the manufacturer’s

recommended concentration, but not at 50% or 5% of the recommended concentration. The reduction in

S. feltiae viability observed with spinosad in our experiment was 35% at 100% of the manufacturer’s

recommended concentration. We would have reservations about using S. feltiae with these insecticides

without further, more detailed testing. Finally, we found that M. brunneum and S. feltiae did not have any

adverse effects on each other when incubated together.

The second part of the project consisted of a series of glasshouse experiments to investigate the effects

on CRF of (i) combining M. brunneum and S. feltiae together; and (ii) combining each of the biopesticides

with a low dose application of the chemical insecticide spinosad. The work started with a pilot experiment.

This was followed by research to investigate the dose response of M. brunneum, S. feltiae and spinosad

against CRF larvae using conventional and organic brassica seed. There were then two relatively large

experiments to investigate the effect of combined applications of M. brunneum, S. feltiae and spinosad

against CRF. In the first of these, the control agents were applied to plants that had been sown directly

into pots: M. brunneum was incorporated directly into compost at the time of planting, while S. feltiae and

spinosad were applied as drenches immediately before the application of CRF eggs to the plants. In the

second experiment, the control agents were applied to brassica plants grown in small modules before

transplanting them to 1 litre pots (to simulate field conditions) after which CRF eggs were applied. The

effects of the treatments were evaluated by quantifying any reductions in numbers of CRF larvae and

pupae compared to a control, and in some experiments we also assessed plant vigour, weight, height and

root damage by CRF. We did not find evidence that combining M. brunneum or S. feltiae with a low dose

application of spinosad affected the level of CRF control compared to spinosad on its own. The spinosad

treatments generally gave good control on their own, although the amount of control varied from

experiment to experiment. We found that combining the biopesticides M. brunneum and S. feltiae together

could give significant control of CRF, whereas these agents did not control CRF when used on their own.

However this effect was not consistent, as it was only observed in two out of three experiments. It is likely

that the outcome of the interaction between M. brunneum and S. feltiae was affected by external factors

that differed between experiments, such as compost type (we suspect that different composts may affect

the retention of biopesticides in the root zone). This illustrates a key challenge in conducting experiments

on biologically-based IPM, in that we are dealing with a more complex system than that using conventional

chemical pesticides, involving multiple variables The combination of M. brunneum and S. feltiae was less

effective than using the chemical insecticide spinosad. However, our findings do provide proof of concept

that biopesticide combination treatments can give improved pest control of CRF.


Project Report to Defra

8. As a guide this report should be no longer than 20 sides of A4. This report is to provide Defra with details of the outputs of the research project for internal purposes; to meet the terms of the contract; and to allow Defra to publish details of the outputs to meet Environmental Information Regulation or Freedom of Information obligations. This short report to Defra does not preclude contractors from also seeking to publish a full, formal scientific report/paper in an appropriate scientific or other journal/publication. Indeed, Defra actively encourages such publications as part of the contract terms. The report to Defra should include:

the objectives as set out in the contract;

the extent to which the objectives set out in the contract have been met;

details of methods used and the results obtained, including statistical analysis (if appropriate);

a discussion of the results and their reliability;

the main implications of the findings;

possible future work; and

any action resulting from the research (e.g. IP, Knowledge Exchange).

Background

Insect pests significantly reduce the yield and quality of a wide range of crops. At the moment, pest control

in most field-grown crops is based around applications of synthetic chemical insecticides. These chemicals

are an important tool for crop protection, but excessive use of insecticides can result in a number of

problems including damage to the environment, harm to non-target insects and the evolution of heritable

resistance in target pest populations. Within the European Union, concerns about the safety of some

chemical pesticides has led to the introduction of more stringent safety criteria for pesticide authorization,

and this has resulted in a significant number of products being withdrawn from the market (Chandler et al.,

2011a). At the same time, the rate of discovery of new chemical actives has declined markedly. This

means that farmers and growers are relying on an increasingly narrow range of insecticides.

A more sustainable approach to managing insect pests is needed. Most experts agree that the way to

achieve this is through Integrated Pest Management (IPM). This is a systems approach in which different

pest management tools are combined with careful monitoring of pests and their natural enemies. The aim is

to use conventional insecticides only when necessary, and to integrate them with other, complementary

methods in order to: (i) reduce the chances of resistance evolving; (ii) prevent harm to the environment; and

(iii) keep pests below the economic injury level in a cost effective way. IPM tools to be used in addition to

conventional chemical pesticides include biological, cultural and physical controls, natural products, and

breeding crops for pest resistance. IPM forms a key part of new EU legislation on the Sustainable Use of

Pesticides, Directive 2009/128/EC. Under the Directive, there is now a legal obligation on farmers and

growers to use IPM if effective and practical The Directive requires that biological controls and other non-

chemical methods should be preferred over conventional pesticides if they give satisfactory levels of

control.

For this project, we were concerned with investigating the use of biological controls based on

entomopathogenic fungi (EPF) and nematodes (EPNs) in IPM. An increasing number of EPF and EPN

products (commonly referred to as “biopesticides”) are available within the EU, mainly for use on

horticultural crops (Chandler et al., 2011a). The attractions of using entomopathogens include operator and

bystander safety, lack of toxic residue, narrow pest activity spectrum, a short interval between application

and harvest, and potential for some self-sustaining control (Bailey et al., 2010). Given these characteristics,

entomopathogens should fit in well with the aims of the Sustainable Use Directive. However,

entomopathogens are usually less efficacious than fully effective conventional insecticides, they are slower

acting, they tend to be more expensive to purchase and they can be adversely affected by environmental

conditions such as low temperatures (for a review see Chandler et al., 2011a). The challenge is to make

best use of the attractive properties of entomopathogens while minimising their downsides. One way to do

this could be to combine different agents together in an IPM programme, for example by using two different

entomopathogens together or by combining an entomopathogen with a partially effective chemical

insecticide, e.g. one that some insects in a population may be resistant to, or one used at a reduced dose In

previous work, we reviewed the scientific literature published on using entomopathogens in this way

(Chandler et al., 2011b). Biopesticide agents are normally developed by companies as if they were stand-


alone treatments, rather than considering them from the outset as IPM tools. In this respect, the

“developmental model” used for biopesticides mirrors that for conventional chemical pesticides. Despite

this, a number of studies have been published showing that combinations of entomopathogens, or

entomopathogens and insecticides, can have synergistic or potentiating effects (Quintela & McCoy, 1997;

Roditakis et al., 2000; Furlong & Groden, 2001; Wraight and Ramos, 2005; Ye et al., 2005; Santos et al.,

2007; Ansari et al., 2008; Shah et al., 2008; Chandler et al., 20011b). Use of entomopathogens can also

slow the evolution of pesticide resistance and reduce the expression of resistance after it has evolved

(Raymond et al., 2006; 2007; Gassmann et al., 2006; 2008; Farenhorst et al., 2009, 2010). However - with

the exception of research done at the University of Wales Swansea on combined applications of EPNs and

EPF for the biocontrol of vine weevil (Ansari et al., 2008) - most of this research has been done using insect

pests that do not occur in the UK. Moreover, much of the published work has been done at the laboratory

scale, with less research at the glasshouse or field scale. However, when we interviewed UK IPM

consultants, we found that combinations of biopesticides, or biopesticides and insecticides, are already

being using on some commercial protected crops. This is being done mainly on the basis of anecdotal

evidence of improved control. There are significant knowledge gaps about using biopesticides in

combinations for IPM for UK pests, therefore, and we believe that this is holding back the development of

IPM for UK farmers and growers.

The aim of this project was to help to develop a new understanding of how combination treatments affect

UK insect pests. We focused on the cabbage root fly, Delia radicum (CRF). This insect is the major pest of

horticultural brassicas in northern temperate regions. It is very damaging and can be difficult to control even

with conventional chemical insecticides (Finch 1989). Eggs are laid on or just below the soil surface close to

the stem of a host plant and the larvae feed on root tissue including the tap root. Attacked plants may be

stunted, leaves turn bluish, often with reddish leaf margins and if damage is severe plants may wilt and die.

In Britain there are two and in some areas three generations (late April to May, late June to July and August

to early September) of the fly each year. In a recent review (Defra Project IF01100) it was estimated that

CRF causes losses of around £15 million/year. It is estimated that without the use of insecticides and

relying on currently available non-pesticide control measures there would be a 62% reduction in the

production of horticultural brassicas. We chose CRF as a target pest because it is economically important,

is difficult to control, there are currently limited options for using conventional chemical pesticides and

hence there is a need to investigate new IPM approaches for it. As such, CRF illustrates many of the

problems facing the development of sustainable pest management. Control of CRF on conventionally-

grown leafy and flowerhead brassicas is currently based on drenching modules with the organophosphate

insecticide chlorpyrifos (e.g. Dursban WG) before planting, with over 90% UK plants treated. However,

there is an operator exposure risk associated with handling modules drenched with chlorpyrifos. As a

result, the approval for this use of chlorpyrifos is being withdrawn, and spinosad (Tracer) now has approval

as an alternative module drench. Relying on one active ingredient for control of CRF increases the risk of

the development of pesticide resistance. There are particular concerns about spinosad, given its importance

in other IPM systems but with resistance problems already in serious pests such as WFT. Several studies

have investigated the potential of using non-chemical controls including the use of EPF such as

Metarhizium brunneum (available commercially as the product Met52) and the EPN Steinernema feltiae.

Research at Warwick showed that larvae of CRF and the closely related onion fly are susceptible to a range

of EPF including Met52 in laboratory and glasshouse experiments (Davidson & Chandler, 2005; Chandler &

Davidson, 2005). However a high concentration of fungal spores was needed for a commercially acceptable

level of control. Met52 now has an EAMU for use in module-raised brassicas for control of CRF. EPNs are

also being sold for CRF control, currently for the home garden market, and we know that there is interest

from the manufacturers in using EPNs for CRF control in professional brassica production.

The first stage of the current project was based on investigating the compatibility of EPNs, EPF and

insecticides. Chemical insecticides can be harmful to entomopathogens and hence it is important to

determine their effects for IPM (Jaronski, 2010). There is some evidence of mutual inhibition of M.

brunneum with two species of symbiotic bacteria associated with entomopathogenic nematodes,

Photorhabdus luminescens and Xenorhabdus poinarii (Ansari et al., 2005). However at present there is

limited information in the public domain on the direct effects of chemical insecticides on M. brunneum and

entomopathogenic nematodes. The manufacturers / suppliers of EPNs and EPF provide technical

information sheets to growers stating whether the entomopathogens are compatible with chemical

insecticides, but they generally do not provide details of the methods used to test compatibility. Moreover,

the categorization of “compatible” and “incompatible” used by the biopesticide companies tends to follow

the IOBC scheme where an insecticide is defined as “harmless” towards a biological control agent if it


causes less than 25% reduction in control capacity, which does not provide the level of accuracy required

for our purposes. The second stage of the project quantified the effects of M. brunneum, S. feltiae and

spinosad when used in pairs against CRF. To our knowledge, combination treatments involving

biopesticides and chemical insecticides have not been investigated before against CRF. We reasoned that

they warranted attention on the basis that once chlorpyrifos has been withdrawn for use, control of CRF will

be reliant on spinosad (with a risk of resistance developing) or biopesticides, and that only partial control is

likely with individual biopesticides.

Aims & Objectives

The aim of this project was to provide a new understanding of the effects on insect pests of combining

entomopathogens and chemical insecticides as an Integrated Pest Management approach. The long term

goal of this area of work is to be able to provide more effective and sustainable crop protection for growers.

The project had two objectives:

Objective 1: Quantify the in vitro effects of selected insecticides on entomopathogenic fungi and

nematodes.

Objective 2. Analyse the effect of combination treatments of M. brunneum, S. feltiae and spinosad

against cabbage root fly

The research was done with the EPF Metarhizium brunneum, which is used in the commercial biopesticide

product Met52 (Novozymes UK Ltd.), and the EPN Steinernema feltiae, which is used as the product

Nemasys (BASF UK Ltd.). The first part of the research was to investigate the compatibility of EPF and

EPNs with insecticides used for CRF control. Clearly, if an insecticide was harmful to an EPN or EPF then it

could not be used as a combination treatment in IPM. We also studied the compatibilty of M. brunneum and

S. feltiae together. In the second part of the project, we investigated the effects of combined applications of

(M. brunneum + S. feltiae), (M. brunneum + spinosad), and (S. feltiae + spinosad) on populations of

cabbage root fly larvae feeding on brassica plants in a series of glasshouse experiments.

Objective 1. Quantify the in vitro effects of selected insecticides on entomopathogenic

fungi and nematodes.

The aim of this Objective was to evaluate: (i) the compatibility of different chemical insecticides with M.

brunneum and S. feltiae, and (ii) the compatibility of M. brunneum and S. feltiae with each other. This was

done using a series of experiments to quantify entomopathogen germination, growth or survival following

treatment. The research started with s set of ‘in vitro’ experiments in which M. brunneum and S. feltiae were

treated with insecticides in water or a laboratory growth medium, before moving on to measuring the effect

of insecticides on M. brunneum and S. feltiae in compost.

Materials & Methods

Task 1.1. EPF, EPNs and insecticides used in the study

Metarhizium brunneum is used in the EPF product Met52 (Novozymes UK Ltd. Original insect host = Cydia

pomonella (Lepidoptera : Tortricidae). Country of origin = Germany). The experiments done in Objective 1

used an isolate of M. brunneum from the Warwick Crop Centre collection of entomopathogenic fungal

cultures that is also used as the active ingredient in Met52. The isolate is maintained in cryopreservation

(Chandler, 1994). The Warwick Crop Centre code for this fungal isolate is 275.86. A laboratory stock

culture of M. brunneum 275.86 was grown on Sabouraud Dextrose agar (SDA) from cryopreserved conidia

and then stored at 4°C. The stock culture was then used to grow individual “secondary” cultures for use in

experiments. These were grown on SDA at 23°C, in the dark, for 10 days before use. Conidia were

harvested in 10ml of sterile 0.05% Triton X-100 by agitating gently with a sterile spreader. The conidia were

enumerated using an Improved Neubauer haemocytometer and conidia suspensions were then adjusted to

the concentration required for the experiment.

For Objective 1, Steinernema feltiae was obtained from BASF UK as the product Nemasys (note that, for

the pilot glasshouse study done in Objective 2.2, we used S. feltiae as the product Exhibitline sf supplied by

Syngenta Bioline, but for subsequent experiments we used Nemasys). The product was stored at 4°C prior

to use. Nematode suspensions were made up on the day of the experiment according to the

manufacturer’s instructions. Aliquots were diluted as required in tap water and nematode concentration and

viability was assessed by counting numbers of motile and non-motile nematodes in a Doncaster dish or a

Hauxley counting chamber.


Three chemical insecticides were used, all supplied by Dow Agrosciences: chlorpyrifos (used as the

product Dursban), spinosad (used as the product Tracer), and HDC coded product 198 (supplied as a

formulated test product; this product is not yet available commercially and hence we are not allowed to

reveal its identity). Chlorpyrifos and spinosad are currently available for CRF control by growers. HDC198

was identified in the Sceptre LINK project (http:.scpetre.hdc.org.uk/) as a potentially valuable new

insecticide with activity against CRF larvae and hence it was decided to include it in this part of the project.

Task 1.2. Quantifying the effect of insecticides on EPF: Direct exposure of M. brunneum conidia on agar

medium supplemented with insecticides

For this experiment, each insecticide was incorporated into SDA at seven different concentrations

(including an untreated control) from 0 – 200% of the manufacturers’ recommended concentration (as

calculated from recommended field rates). The insecticides were prepared in sterile distilled water and then

added to SDA that had been autoclaved and cooled to 45°C. This was then poured into Petri plates (90mm

diameter, 45 ml SDA per plate). Controls consisted of SDA without insecticides. Aliquots of a conidia

suspension (20l; 1 x 107 conidia ml

-1) of M. brunneum 275.86 were pipetted onto each of three areas

(circles approximately 1cm diameter) on each plate. Plates were sealed with Nescofilm and incubated in

the dark at 23 ± 1°C. Sampling was carried out after 24 hours by pipetting a drop of lactophenol methylene

blue inside each circle to stop conidia germination. Plates were sealed and stored at 4°C before

examination under the light microscope (magnification x200). The numbers of germinated and

ungerminated conidia were recorded for a sample of approximately 100 conidia per circle. Germination

was defined as the point when the length of an emerging germ tube was equal to, or longer than, the length

of the conidium. The experiment used three pseudo-replicate plates per treatment and was repeated three

times. The percentage germination was compared to the controls using an ANOVA (Genstat, 2000).

Task 1.3. Quantifying the effect of insecticides on EPF: Measurement of M. brunneum colony formation on

agar medium supplemented with insecticides

Aliquots of 100µl of a conidia suspension (1 x 104 conidia ml

-1) of M. brunneum 275.86 were spread evenly

over the surface of SDA plates in which insecticides had been incorporated at six different concentrations

(see above). Plates were incubated at 23 ± 1°C in the dark. After 6-7 days, the number of colonies per

plate was counted. The experiment was repeated three times and the average number of colonies per

plate was compared to the control using an ANOVA (Genstat, 2000).

Task 1.4. Quantifying the effect of insecticides on EPF: Germination and colony formation of M. brunneum

following immersion of conidia in insecticide solution

A suspension of conidia of M. brunneum 275.86 (10mls, 1 x 107 conidia ml

-1) was combined with a solution

of each insecticide at seven different concentrations (including an untreated control) from 0 – 100% of the

manufacturers’ recommended concentration. Suspensions were incubated for 24 hours at 23 ± 1°C.

Aliquots (20l) of the conidia / insecticide suspension were then pipetted onto three previously marked

circles (approximately 1cm diameter) onto two SDA plates per treatment. The plates were sealed and

incubated in the dark at 23 ± 1°C for 24 hours, after which germination was terminated by pipetting a drop

of lactophenol methylene blue inside each circle. Plates were sealed and stored at 4°C before examination

under the light microscope and germination quantified as described previously. In addition, 100l aliquots

of the conidia / insecticide suspension were diluted in 9.9ml SDW (a 1/100 dilution) and 100l of the

resulting suspension spread evenly over the surface of three SDA plates. These plates were incubated at

23 ± 1°C, in the dark, and after 6 - 7 days the number of colonies grown on each plate was counted. This

data was used to estimate the concentration of viable conidia in the suspension plated onto the SDA. The

experiment was repeated three times and the average concentration of viable conidia per treatment was

compared to the control using an ANOVA (Genstat, 2000).

Task 1.5. Quantifying the effect of insecticides on EPNs: measuring the viability of S. feltiae following

immersion in insecticide solution

A suspension of Steinernema feltiae infective juveniles (IJs) was prepared from the EPN biopesticide

Nemasys by dispersing the nematodes in tap water according to the manufacturer’s instructions. The

suspension was counted and adjusted to a concentration of 1.2 x 104 infective juveniles (IJ) ml

-1 and then

10 ml aliquots were combined with a 10ml solution of each insecticide at seven different concentrations

(including an untreated water control) from 0 – 100% of the manufacturers’ recommended concentration.

There were three replicates per treatment. The suspensions were incubated for 16 hours at 22 ± 1°C in the

dark. Nematodes were allowed to settle out during incubation and excess water was removed from each

suspension by pipette. The nematodes were then washed twice in 30ml tap water and observed under a


binocular microscope. The numbers of viable (= movement in response to mechanical stimulation) and

non-viable (= no movement in response to mechanical stimulation) nematodes were counted in 1ml sub-

samples at an expected concentration of 120 IJ ml-1

. Effect of treatments on the proportion of viable

nematodes in test populations was compared to the control using an ANOVA.

Task 1.6. Quantifying the effect of insecticides on EPNs and EPF: measuring the viability of nematodes

and fungi following exposure to insecticides in soil.

Peat-based compost, used by commercial propagators for brassica plant raising, was treated with either

Met52 or Nemasys. Met52 was applied as commercially formulated granules (supplied by Fargro Ltd) that

were mixed by hand into compost at the manufacturer’s recommended rate (0.5kg m-3

). Aliquots (20ml) of

the treated compost were then added to individual Universal tubes (20ml is equivalent to the amount of

compost contained in a commercial brassica plug in a “216” module tray used for plant raising). Solutions

of each insecticide were prepared at four different concentrations (including an untreated water control) and

1ml aliquots pipetted onto the compost surface of each Universal tube followed by addition of 1 ml tap

water to give final concentrations from 0 – 100% of the manufacturers’ recommended rate (2ml tap water

was added to the controls). There were three replicates per treatment. Nemasys was prepared as a tap

water suspension at a concentration of 3.5 x 104 IJ ml-1 and 1ml aliquots were added to individual

Universal tubes, each containing 20ml brassica compost. This was followed immediately by application of

1ml aliquots of insecticide solution to each Universal tube at four different final concentrations from 0 –

100% of the manufacturers’ recommended rate (1ml of Nemasys plus 1ml of water was added to the

controls). There were three replicates per treatment. Samples of compost were taken from controls at time

zero, while samples from all treatments were collected on day five. Samples from Met52 treatments

consisted of 1g compost taken from the top of each Universal tube. These were suspended in 0.05% triton

X-100 wetter solution, vortex mixed, serially diluted, and then aliquots (0.2ml) plated onto a agar medium

that is selective for M. brunneum (Potato dextrose - yeast extract (PDAY) agar supplemented with

chloramphenicol (0.5g l-1

), cyclohexamide (0.25 g l-1

) thiabendazole (0.004g l-1

) , rose Bengal (0.01g l-1

)).

Plates were incubated at 23 ± 1°C, in the dark, and after 6 - 7 days the number of colonies grown on each

plate was counted. Samples of S. feltiae were collected by adding the compost from each Universal tube to

a modified Baermann funnel lined with a milk filter, applying 500 ml water per funnel until water covered

the compost and incubating overnight. Nematode IJs that had migrated through the filter were then

collected from the funnel reservoir. Collected IJs were observed under a binocular microscope and the

number of IJs collected was counted as described previously. Three sub-sample counts were made per

replicate.

Task 1.7. Quantifying compatibility of EPNs and EPF.

Suspensions of conidia of M. brunneum 275.86 were prepared at five concentrations (10ml volume, 0.05%

Triton X-100 wetting agent) from 1 x 104 to 1 x 10

8 conidia ml

-1 and then added to an equal volume of a

suspension of S. feltiae lJs (1.2 x 104 IJs ml

-1) and incubated in the dark at 23 ± 1°C for 24 hours. Each

suspension was then shaken by hand and 10 ml pipetted into each of two separate Universal tubes. For

one of each pair of tubes, the S. feltiae IJs were allowed to settle out, then removed by pipetting, washed in

milli-Q water and incubated at 23 ± 1°C for a further 24 hours. Aliquots of 5ml IJs were then transferred to a

Doncaster dish and examined under a binocular microscope: the numbers of viable (= movement in

response to mechanical stimulation) and non-viable (= no movement in response to mechanical

stimulation) nematodes were counted. An aliquot (20l) was removed from each of the remaining tubes

and pipetted onto three previously marked circles (approximately 1cm diameter) on SDA plates. The plates

were sealed with nescofilm and incubated in the dark at 23 ± 1°C, after which germination was terminated

by pipetting a drop of lactophenol methylene blue inside each circle. Plates were sealed and stored at 4°C

before examination under the light microscope and germination quantified as described previously.

Results

Task 1.2. Quantifying the effect of insecticides on EPF: Direct exposure of M. brunneum conidia on agar

medium supplemented with insecticides

Germination of M. brunneum 275.86 conidia in the control plates ranged from 71.2% to 93.6% (mean ± SE = 87.6% ± 1.891). Neither chlorpyrifos nor HDC198 had any significant effect (P> 0.05) on conidia germination. Spinosad significantly (P< 0.05) reduced the number of conidia that germinated when tested at the manufacturer’s recommended concentration (28% reduction) and double the recommended concentration (20% reduction) (Table 1).


Table 1: Mean (back-transformed) % germination of M. brunneum 275.86 on SDA supplemented with insecticides (transformed data in parenthesis).

Insecticide application rate (% of manufacturer’s recommended concentration)

Insecticide 0% 5% 10% 25% 50% 100% 200%

chlorpyrifos 87.6 (68.26)

94.3 (76.32)

94.2 (76.11)

96.9 (80.57)

94.7 (76.78)

95.2 (77.39)

95.8 (78.28)

HDC198 87.6

(68.26) 85.9

(69.38) 86.9

(69.31) 91.2

(73.69) 90.0

(73.03) 90.8

(73.69) 86.1

(68.76) Spinosad 87.6

(68.26) 81.3

(65.18) 86.0

(68.48) 84.5

(66.89) 79.7

(63.46) 62.9*

(52.54) 70.1*

(56.93) LSD (p < 0.05; df= 40) = 8.115

* significantly different from the control

Task 1.3. Quantifying the effect of insecticides on EPF: Measurement of M. brunneum colony formation on

agar medium supplemented with insecticides

None of the insecticides examined had any significant effect (P > 0.05) on the number of colonies formed on SDA (Table 2). Table 2: Mean (back-transformed) colony forming units of M. brunneum 275.86 produced after conidia were plated onto SDA supplemented with insecticides


Insecticide 0% 5% 10% 25% 50% 100% 200%


337.3 (2.55)

347.3 (2.53)

371.1 (2.56)

421.1 (2.61)

374.3 (2.56)

433.6 (2.63)

HDC198 424.4

(2.62) 358.7 (2.54)

428.4 2.62)

350.0 (2.53)

363.6 (2.55)

395.8 (2.58)

386.2 (2.57)

Spinosad 424.4

(2.62) 412.2 (2.61)

388.9 (2.59)

409.8 (2.60)

432.4 (2.62)

452.2 (2.65)

384.2 (2.57)

LSD (p < 0.05; df= 39) = 0.110

Task 1.4. Quantifying the effect of insecticides on EPF: Germination and colony formation of M. brunneum

following immersion of conidia in insecticide solution

Germination of conidia on the control plates ranged from 87.6% to 96.8% (mean ± SE = 93.6% ± 0.948).

Neither chlorpyrifos or spinosad had a significant (P> 0.05) effect on conidia germination, but HDC198

significantly (P < 0.05) reduced the number of conidia that germinated when tested at 50% of the

manufacturer’s recommended concentration (12% reduction) and 100% of the recommended concentration

(15% reduction) (Table 3). HDC198 also caused a significant (P < 0.05) reduction in the number of colony

units formed per plate at 12.5% and 25% of the manufacturer’s recommended concentration (Table 4).

Table 3: Mean (back-transformed) % germination of M. brunneum 275.86 following exposure to insecticides. Fungal conidia were immersed for 24 h in insecticide solution, then 20μl aliquots plated onto SDA (transformed data in parenthesis).


Insecticide 0% 2.5% 5% 12.5% 25% 50% 100%


95.2 (77.80)

94.8 (77.10)

94.2 (76.12)

95.0 (77.39)

89.2 (70.99)

94.1 (75.95)

HDC198 94.7

(76.95) 92.7

(74.41) 93.7

(75.65) 91.9

(73.75) 90.7

(72.69) 89.7*

(71.54) 89.3*

(70.89) Spinosad 92.4

(75.75) 94.2

(76.11) 94.7

(76.75) 93.7

(75.64) 93.5

(75.36) 92.4

(74.21) 91.3

(72.92) LSD (p < 0.05; df= 41) = 5.124

* significantly different from the control


Table 4: Mean (back-transformed) colony forming units per plate of M. brunneum 275.86 following exposure to insecticides. Fungal conidia were immersed for one hour in insecticide solution, then diluted and plated onto SDA (transformed data in parenthesis).


Insecticide

0% 2.5% 5% 12.5% 25% 50% 100%


213.3 (2.32)

216.6 (2.33)

222.4 (2.34)

253.9 (2.37)

213.5 (2.30)

243.4 (2.36)

HDC198 318.9

(2.41) 257.4 (2.40)

220.2 (2.34)

224.1* (2.32)

199.0* (2.27)

249.1 (2.37)

253.4 (2.38)

Spinosad 270.3

(2.37) 249.1 (2.39)

264.0 (2.42)

276.7 (2.41)

247.9 (2.37)

275.9 (2.41)

265.7 (2.39)

LSD (p < 0.05; df= 29) = 0.077

Task 1.5. Quantifying effect of insecticides on EPNs: measuring the viability of S. feltiae following

immersion in insecticide solution

In the water control, a mean of 85.3 live nematodes ml-1

was recorded. A significant (P<0.001) reduction in nematode viability was observed with chlorpyrifos at 100, 50 and 25% of the manufacturer’s recommended concentration (MRC), with a clear dose response (Fig. 1): there were reductions of 79% (for 100% MRC), 39% (for 50% of MRC) and 15% (for 25% of MRC). Spinosad (Tracer) and the coded insecticide HDC198 had no significant effect on nematode viability (Fig. 1).

Figure 1: Mean number of live nematodes per 1ml sub-sample following direct exposure to chlorpyrifos (Dursban WG), spinosad (Tracer) and a coded insecticide (HDC198) at different rates compared to the water control (ANOVA: F (8,16) =38.12, P<0.001)

Task 1.6. Quantifying effect of insecticides on EPNs and EPF: measuring the viability of nematodes and

fungi following exposure to insecticides in soil.

There was no significant effect (P> 0.05) of any insecticide on the number of colony forming units per plate

of M. brunneum recovered from insecticide-treated compost (Table 6).

For S. feltiae, a mean of 105.4 live nematodes ml-1

was recorded in the water control. A significant (P<0.01) reduction in nematode viability was observed with chlorpyrifos at 100% and 50% of the manufacturer’s recommended concentration (MRC) (Fig. 2): there were reductions of 54% (for 100% MRC) and 37% (for 50% of MRC). However there was no reduction at 5% of MRC for chlorpyrifos. Significant (P< 0.05) reductions were also observed for spinosad (Tracer) at 100% MRC (a reduction of 35%) and the coded insecticide HDC198 at 100% MRC (a reduction of 27%) but not at 50% MRC or 5% MRC (Fig. 2).


Table 6: Mean (back-transformed) colony forming units per plate of M. brunneum 275.86 isolated from compost supplemented with insecticides (transformed data in parenthesis).


Insecticide

0% 5% 50% 100%

chlorpyrifos 4 (0.60) 4.17 (0.59) 2.83 (0.37) 2.17 (0.32)

HDC198 4 (0.60) 2.2 (0.28) 3.5 (0.49) 6.75 (0.81)

Spinosad 4 (0.60) 4.5 (0.64) 3.17 (0.48) 4.83 (0.61)

LSD (p < 0.05; df=21) =0.3641

Figure 2: Mean number of S. feltiae per ml following exposure in compost to chlorpyrifos (Dursban WG), spinosad (Tracer) and a coded insecticide(HDC198) at different rates compared with the water control. *LSD 5% (26.404) for comparison of treatments means with the control. Control confidence limit: mean

±15.24, treatment confidence limit: mean ±21.56.

Task 1.7. Quantifying compatibility of EPNs and EPF

Nematode survival ranged from 44% to 75.6% in the untreated samples (mean = 62.4% ±4.06). There was no significant difference in nematode survival observed when the nematodes were submerged in any of the rates of M. brunneum 275.86 tested. Fungal germination ranged from 93 to 96% (mean = 94.9% ±0.50). There was no significant difference in fungal germination when spores were submerged in any of the rates of nematodes tested (Table 5).

Table 7: Mean (back-transformed) % S. feltiae survival and % germination of M. brunneum 275.86 after the two biopesticides were kept together in a mixture. Suspensions of S. feltiae and M. brunneum were mixed together and incubated for 24 h, after which S. feltiae survival and M. brunneum germination were measured (transformed data in parenthesis).

Concentration of S. feltiae and M. brunneum (% of manufacturer’s recommended concentration)

% nematode survival % fungal germination

0% 62.4 (51.50) 94.8 (76.95) 2.5% 63.9 (51.93) 94.9 (77.13) 5% 66.1 (53.19) 94.1 (76.21) 12.5% 65.0 (53.72) 96.6 (79.73) 25% 63.9 (52.75) 94.8 (77.54) 50% 65.2 (53.63) 97.2 (80.25) 100% 65.0 (53.86) 94.6 (76.61) df

42

12

LSD (p < 0.05) (3.445) (4.666)


Conclusions and discussion

This part of the project consisted of a series of tests to investigate the effects of insecticides on EPF

and EPNs, and also to study the compatibility of EPNs and EPF with each other. For the experiments

with insecticides, the intention was to start off by exposing EPF and EPNs to insecticides in “in vitro”

laboratory tests, and then investigate the effect of insecticides in compost (which would be more

representative of insecticide exposure under field conditions). Some insecticide compatibility tests done

in vitro have been known to give false-positive results in which inhibition observed under laboratory

condition does not translate to field scale effects. This can occur as a result of compartmentalization of

the chemical pesticide within soil or plant tissue, lower pesticide concentrations encountered under field

conditions, or – in the case of above-ground environments - drying of pesticide residues on foliar

surfaces (Jaronski, 2010; Inglis et al., 2001; Cuthbertson et al., 2005). Therefore the results of these

insecticide compatibility experiments need to be interpreted carefully.

For experiments with M. brunneum:

There were some effects of insecticide treatment on M. brunneum 275.86 in experiments done in vitro,

but the effect depended on the type of insecticide, its concentration, and the method by which the

fungus was exposed to it. There was no evidence of an adverse effect of insecticides on M. brunneum

in compost, in an experiment designed to mimic field conditions.

In our first experiments (1.2, 1.3) we exposed M. brunneum to insecticides incorporated into SDA

medium used to culture the fungus. We considered this to be the most extreme form of exposure of the

fungus to insecticide in our series of experiments. Germination of M. brunneum conidia on SDA

supplemented with chlorpyrifos or HDC198 was not affected at any of the insecticide concentrations

tested (1.2). Spinosad caused a reduction in germination of up to 28% at 100% to 200% of the

manufacturer’s recommended concentration. The formation of M. brunneum colonies on SDA

supplemented with insecticides was not affected by any of the insecticides tested (1.3). This might

indicate that fungal growth is less sensitive to inhibition by spinosad than fungal germination.

Experiment 1.4 was designed as laboratory simulation of “tank mixing” M. brunneum with an insecticide.

In this experiment, conidia of M. brunneum 275.86 were suspended in a solution of insecticide for 24h

before 20μl aliquots were plated onto SDA (any insecticide in the 20μl aliquot would have diffused into

the SDA, with a dilution factor of approximately 1 / 2000 v/v, meaning that any residual effect of

insecticide on fungal spores would be unlikely). Conidia germination was measured after 24 hours

incubation on the SDA. The number of colony forming units produced on the SDA was also counted.

There was no effect of chlorpyrifos or spinosad on M. brunneum, but coded insecticide HDC198

reduced germination and colony formation in a concentration-dependent manner. Before doing the

experiment, we hypothesised that this would produce less exposure to the insecticide than in 1.2 and

1.3, so the observation of a reduction in germination and colony formation with HDC198 in 1.4, but not

in 1.2 and 1.3, was unexpected. The reason for the result is not known. One possible explanation is that

susceptibility to insecticide could depend on the level of metabolic activity of M. brunneum conidia.

Conidia of M. brunneum do not germinate if held just in water, as they require an external source of

carbohydrate as a signal for cell activation and enzymatic degradation of the conidia wall, resulting in

germination. In experiment 1.4, M. brunneum conidia were held for 24 h in an insecticide solution made

up in water before being plated onto SDA. It is possible that the conidia were less metabolically active

when held in water than when on the SDA. This could affect their ability to detoxify an insecticide (for

example through the action of cytochrome P450s).

Experiment 1.6 investigated the effect of insecticides on M. brunneum 275.86 in compost. In this case,

exposing the fungus to insecticides in compost over five days caused no significant reduction in the

concentration of M. brunneum conidia in compost (as measured by counts of colony forming units). We

considered this experiment to be more representative of insecticide exposure under field conditions.

Experiment 1.7 investigated the compatibility of S. feltiae and M. brunneum with each other. There was

no indication that these two agents were harmful to each other.

For experiments with S. feltiae:

Experiment 1.5 was designed to give a laboratory based simulation of tank mixing S. feltiae with

insecticides. In this experiment, spinosad (Tracer) and the coded insecticide HDC198 had no effect of

S. feltiae viability at the manufacturer’s recommended concentration and thus we would conclude that

these insecticides products are probably safe to tank mix with S. feltiae. However, chlorpyrifos


(Dursban WG) significantly reduced the number of live nematodes at 100, 50 and 25% of the

manufacturer’s recommended concentration, and therefore we would conclude that this insecticide is

unlikely to be compatible with S. feltiae in a tank mix, although more testing would be required to

provide definitive proof (see above).

In experiment 1.6, EPNs were exposed to insecticides in compost over five days. There was a

significant negative effect of chlorpyrifos on S. feltiae at 100% and 50% of the manufacturer’s

recommended concentration, but not at 5% of the recommended concentration. There were also

significant reductions in nematode viability when S. feltiae was kept in compost with spinosad and the

coded insecticide HDC198 at 100% of the manufacturer’s recommended concentration, but not at 50%

or 5% of the recommended concentration. The reduction in S. feltiae viability observed with spinosad in

our experiment was 35% at 100% of the manufacturer’s recommended concentration. We would have

reservations about using S. feltiae with these insecticides without further, more detailed testing. Two of

the main manufacturers of EPN products say that spinosad is considered to be “harmless” to S. feltiae,

based on an IOBC scheme in which “harmless” is defined as causing less than 25% reduction in control

capacity (no information is available for the other two insecticides).

Objective 2. Analyse the effect of combination treatments of M. brunneum, S. feltiae and

spinosad against cabbage root fly

The aim of this objective was to investigate the effect of dual combinations of M. brunneum, S. feltiae and

spinosad against CRF under glasshouse conditions. The work started with a small-scale pilot experiment

(see 2.2 below). This was followed by research to investigate the dose response of M. brunneum, S. feltiae

and spinosad against CRF larvae (2.3) and a relatively large experiment to investigate the effect of

combined applications of M. brunneum, S. feltiae and spinosad against CRF (2.4). Finally, a large

glasshouse experiment was done to study the effects of combined applications of M. brunneum, S. feltiae

and spinosad against CRF using application methods likely to be done by commercial growers.

Materials & Methods

Task 2.1. Insect rearing

Cultures of the cabbage root fly, Delia radicum, were reared according to the method of Finch & Coaker

(1969). Adult flies were kept in mesh cages (35 x 35 x 35 cm, ca. 300 flies per cage) within a controlled

temperature room (18 1°C, 16:8 L:D). Each cage contained a drinking station (a filter paper wick

protruding from a 100 ml jar filled with water), plus a supply of carbohydrate (10% sucrose solution soaked

onto absorbent cotton wool in a Petri dish lid (90 mm diameter)), and amino acids (yeast extract sandwich

spread smeared onto a Petri dish lid and sprinkled with brewer’s yeast powder (30% brewer’s yeast in soya

flour)). Eggs were collected 12 days post emergence using an oviposition site (a cube of swede (ca. 2cm

diameter) on damp sand in a Petri dish lid) placed within the cage for 24 h. Eggs were washed from the

sand, floated on water, and collected by sieving through a cotton sheet. Larvae were reared on whole

swede kept in damp sand within pots (150 mm diameter) covered by muslin. Ca. 300 eggs were placed at

the base of each swede, then maintained within a controlled temperature room (18 1°C, 16:8 L:D) for 30

d. Pupae were washed from the sand, collected by floating on water, and then transferred in batches of ca.

1000 to dampened vermiculite in glass jars (300 ml). Pupae were stored for up to one month at 4°C before

transfer to adult fly cages (see above), one jar of pupae per cage. Adult flies began to emerge 2 – 3 d after

pupae were placed in cages.

Task 2.2. Pilot glasshouse experiment: effect of combining M. brunneum + spinosad, and M. brunneum +

S. feltiae, on populations of cabbage root fly larvae feeding on brassica plants

This pilot glasshouse experiment investigated the effect of combined applications of different control agents

on populations of CRF larvae feeding on potted cauliflower plants. Two different combinations were

investigated: (i) M. brunneum with spinosad; and (ii) M. brunneum with S. feltiae. The treatments used in

the experiment are given in Table 8. The S. feltiae treatment was applied as the product Exhibitline sf

supplied by Syngenta Bioline. Spinosad was applied as Tracer supplied by Dow Agrosciences. The M.

brunneum treatment was applied as Met52 granules supplied by Novozymes UK Ltd. Met52 was mixed into

Levington M2 compost (68% water content) at the manufacturer’s recommended rate (0.5kg m-3

) using a St

Moritz paddle compost mixer with a 300 L capacity. The treated compost was then added to 1 litre plant

pots. Samples (1g) were taken from each pot in order to determine the concentration of M. brunneum

conidia (determined as number of colony forming units (cfu) g-1

compost) as described in (1.6). Untreated

compost was also added to 1 litre plant pots for treatments that did not use M. brunneum. Cauliflower


seeds, Brassica oleracea, cv Skywalker, were sown in wetted vermiculite in a controlled environment room

(18 1°C, 16:8 L:D) and then newly germinated seedlings transplanted into the M. brunneum treated

compost / untreated compost pots, one seedling per pot. The seed was supplied as a standard pellet form

with a fungicide coating (thiram (as ProSeed), iprodione (as Rovral Aquaflo) and metalaxyl-M (as Apron

XL350). The plants were grown for four weeks in a glasshouse maintained at a minimum temperature of

18°C and vented at 23°C. At this point, compost samples were taken from each pot to determine the

concentration of M. brunneum conidia as described previously. The spinosad and S. feltiae treatments were

then applied as 5 ml drenches pipetted onto the compost surface to the whole of the base of the stem of the

plant in each pot (for combination treatments, S. feltiae drenches were applied before spinosad application).

The S. feltiae treatment was applied at a rate of 3.5 x 104 IJ nematodes per plant. This was followed

immediately by or before the addition of cabbage root fly eggs adjacent to stems, 1cm below the soil

surface, 15 eggs per plant. There were nine replicate pots per treatment and the pots were arranged in a

randomised complete block design with a three pot blocking structure. The plants were maintained in the

glasshouse for 28 days (4 weeks), after which CRF larvae and pupae were washed from each pot and

counted.

Table 8: Treatments used in pilot glasshouse experiment (2.2) to investigate effect of combining M.

brunneum + spinosad, and M. brunneum + S. feltiae, on populations of cabbage root fly larvae

Treatment No. Description

1 Untreated (water) control

2 M. brunneum applied at manufacturer’s recommended rate

3 S. feltiae at recommended rate

4 Spinosad applied at 5% of manufacturer’s recommended rate

5 Spinosad at 10% of recommended rate

6 M. brunneum at recommended rate + S. feltiae at recommended rate

7 M. brunneum Met52 at recommended rate + spinosad 5% of recommended rate

8 M. brunneum at recommended rate + spinosad at 10% of recommended rate

Task 2.3. Investigation of dose response of M. brunneum, S. feltiae and spinosad applied against cabbage

root fly larvae feeding on brassica plants in a glasshouse experiment

This experiment investigated the effect of different concentrations of M. brunneum, S. feltiae and spinosad

applied against CRF fly larvae feeding on potted brassica plants in a glasshouse. Cauliflower seeds,

Brassica oleracea, cv Skywalker (supplied as pelleted seed coated with a fungicide, see 2.2), were sown

directly into 0.7 l pots containing either untreated or Met 52 incorporated Levington F2 compost at five

concentrations (25% - 200% of the recommended rate for field use) and grown for four to six weeks in the

glasshouse. Plants were then drenched with 5ml of Spinosad +/- S. feltiae (supplied as Nemasys from

BASF UK Ltd) at five test concentrations (1% -200% of the recommended rate for field use). Control plant

received only water treatment. Application of the treatments was followed immediately by the addition of

CRF eggs adjacent to stems, 5mm below the soil surface, 20 eggs per plant. The treatments were

arranged in a randomised block design, with ten blocks. Each block consisted of all treatments with one

plant per treatment. Four weeks after the application of eggs, the plants were harvested and weighed and

larvae and pupae were washed from the pots. This experiment was then repeated for Met52 incorporated

compost only, tested at five different concentrations from 25% to 200% of the manufacturer’s recommended

concentration, but this time using cauliflower seed (Brassica oleracea, cv Skywalker) supplied as organic

seed that had not been coated with a fungicide

Task 2.4. Effect of combining M. brunneum + spinosad, and M. brunneum + S. feltiae, on populations of

cabbage root fly larvae feeding on brassica plants, quantified in a glasshouse experiment

This experiment was done to evaluate the effect of combined applications of M. brunneum + spinosad, and

M. brunneum + S. feltiae, applied against CRF fly larvae feeding on potted brassica plants in a glasshouse,

using concentrations of each control agent that were chosen on the basis of the previous experiment.

There were seven treatments in total (Table 9). Cauliflower seedlings, Brassica oleracea, cv Skywalker,

were sown directly into 0.7 litre pots containing either untreated or Met 52 incorporated Levington F2

compost (at 100% of manufacturer’s recommended concentration) and grown for four to six weeks in the

glasshouse (minimum temperature 18°C, venting set at 23°C). Plants were drenched with 5ml of spinosad

(supplied as Tracer from Dow Agrosciences) +/- S. feltiae (supplied as Nemasys from BASF UK Ltd) at a

concentration of 1% (Spinosad) and 100% (Nemasys) of the recommended field rate, which was

determined from the results of (2.3). This was followed immediately by the addition of CRF eggs adjacent to

stems, 5mm below the soil surface, 15 eggs per plant. Controls were untreated plants. The treatments

were assessed in a randomised block design, with 40 blocks. Each block consisted of all treatments with


one plant per treatment. Two weeks (14 days) after the application of the eggs, the plants were assessed

for plant vitality. Each plant was given a score where 0= healthy and 3 = dead. Four weeks (28 days) after

the application of eggs, the plants were harvested and weighed and larvae and pupae were washed from

the pots. This experiment was then repeated, but this time it was done using cauliflower seed (Brassica

oleracea, cv Skywalker) supplied as organic seed that had not been coated with a fungicide.

Table 9: Treatments used in glasshouse experiment (2.4) to investigate effect of combining M. brunneum +

spinosad, and M. brunneum + S. feltiae, on populations of cabbage root fly larvae

Treatment No.

Description


2 M. brunneum

3 S. feltiae

4 Spinosad

5 M. brunneum plus S. feltiae

6 M. brunneum plus Spinosad

7 S. feltiae plus Spinosad

Task 2.5. Effect of a combination of entomopathogens and Spinosad against CRF larvae (field simulation)

This experiment evaluated the effect of combined applications of (i) M. brunneum + spinosad, (ii) S. feltiae +

spinosad and (iii) M. brunneum + S. feltiae, applied against CRF fly larvae feeding on potted cauliflower

plants in a glasshouse on a commercial brassica propagation nursery (average air temperature set at

20°C). The experiment followed a similar strategy to that of (2.4) but it was modified to reflect the methods

and conditions by which brassica plants are raised by commercial propagators. There were seven

treatments in total (Table 9). Metarhizium brunneum was applied to compost in the plugs before sowing as

Met52 granules (supplied by Fargro UK Ltd) at the manufacturer’s recommended rate (0.5kg m-3

, EAMU

1568/2011). It was mixed into brassica growing compost using the grower’s compost mixer. The S. feltiae

treatment was applied to compost in the plugs before transplanting as a drench treatment of Nemasys

(supplied by BASF UK) at the manufacturer’s recommended rate (3.5 x 104 IJs per plant in 1 litre water per

1000 plants). Spinosad was applied to compost in the plugs before transplanting as a drench treatment of

Tracer (supplied by Dow Agrosciences) at 1% of the manufacturer’s recommended rate (0.12 ml per 1l

water per 1000 plants). The experiment was done as follows: Immediately before sowing, three modular

trays (216 modules per tray), used by professional propagators to raise plug plants, were filled with Met52

treated compost (treatments 2, 5 & 6) (Table 3). The other trays (treatments 1, 3, 4 & 7) were filled with

untreated compost. Ten samples of compost (1g each) were taken from the Met52 treated compost in order

to determine the concentration of M. brunneum conidia (determined as number of colony forming units (cfu)

g-1

compost) as described in (1.6). Cauliflower seed, Brassica oleracea cv. Freedom, was sown into the

modular trays by the commercial brassica propagator. The plants were raised up to the transplanting stage

by the propagator under standard organic commercial conditions. No fungicides or insecticides were

applied to the plants during this time, although the grower did release a mix of six parasitoid species for

biological control of aphids, according to his normal commercial practice. When the plants had reached the

transplanting stage, the S. feltiae and spinosad treatments were applied as drench treatments as set out in

Table 9. The drenches were applied using a syringe to the compost around the whole of the base of the

stem of the plant in each module. For treatments 3 and 4, after drenching with 1ml of treatment per plant,

water was added at a further 1 ml per plant to be consistent with supplier’s recommendations. For

treatment 7 (S. feltiae plus spinosad) the S. feltiae was drenched first followed by the spinosad. For the

water controls and treatment 2 (Met52 as individual treatment), 2 ml water was drenched to the base of

each plant. One day after the application of the drench treatments, the plug plants were transplanted from

the modular trays into 1-litre pots using John Innes no. 2 compost to represent field soil. Each plant module

was placed in the pot of compost so that the top of the compost in the module was 3 cm below the level of

the compost in the pot. This is consistent with the depth modules are planted using a commercial planting

machine in the field. This was followed immediately by the addition of CRF eggs to the compost adjacent to

stems, 15 eggs per plant. Four weeks (26 days) after the application of eggs, the plant height and weight

was assessed, larvae and pupae were washed from the pots and counted and Root Damage Index (RDI)

was assessed. For this experiment, there were 40 replicate pots per treatment done according to a

randomised block design (40 blocks, each block being one plot of seven pots, each plot having one pot per

treatment). However, only 15 replicate pots per treatment were assessed, after statistical analysis

(ANOVA) showed significant effects of treatments without proceeding to assess the full 40 replicates.


Table 9: Treatments used in field experiment (2.5) to investigate effect of combining M. brunneum +

spinosad, and M. brunneum + S. feltiae, on populations of cabbage root fly larvae

Treatment No.

Description


2 M. brunneum applied at manufacturer’s recommended rate

3 S. feltiae applied at manufacturer’s recommended rate

4 Spinosad applied at 1% of recommended rate

5 M. brunneum (at recommended rate) plus S. feltiae (at recommended rate)

6 M. brunneum (at recommended rate) plus spinosad (at 1% of recommended rate).

7 S. feltiae (at recommended rate) plus spinosad (at 1% of recommended rate).

Results

Task 2.2. Pilot glasshouse experiment: effect of combining M. brunneum + Spinosad, and M. brunneum +

S. feltiae, on populations of cabbage root fly larvae feeding on brassica plants

Mean numbers of CRF (larvae plus pupae) per plant were not significantly reduced (P > 0.05) by the

application of M. brunneum or S. feltiae on their own compared to the control (Figure 3). Spinosad caused a

significant (P < 0.05) reduction in the mean number of CRF per plant at 10% of the manufacturer’s

recommended concentration (MRC) but not at 5% MRC. A combination of spinosad at 10% MRC plus M.

brunneum also significantly (P< 0.05) reduced the mean number of CRF per plant, but was not significantly

different to that of spinosad at 10% MRC on its own. Mean numbers of CRF per plant were significantly (P <

0.05) reduced by the combination of M. brunneum and S. feltiae (3.78 per plant). However this combination

treatment was not as effective as spinosad at 10% MRC (0.78 per plant) or spinosad at 10% MRC plus M.

brunneum (0.78 per plant).

Figure 3: Mean number of CRF larvae and pupae recovered per plant in Experiment 2.2. Treatment

number corresponds to that described in Table 8. Error bars represent the standard error of the mean

Task 2.3. Investigation of dose response of M. brunneum, S. feltiae and spinosad applied against cabbage

root fly larvae feeding on brassica plants in a glasshouse experiment

Two experiments were done in task 2.3. In the first experiment, the effect of using different concentrations

of spinosad, S. feltiae, and Met52 (applied both as a granular formulation and as a drench) against CRF

was investigated using conventional brassica seed that had been coated with a standard fungicide mix.

Mean numbers of CRF (larvae plus pupae) per plant were not significantly (P > 0.05) reduced by S. feltiae

or M. brunneum at any of the concentrations tested (Fig. 4). In contrast spinosad significantly reduced (P <

0.05) the number the number of CRF per plant at all concentrations tested (3.3 to 0 per plant) compared to

the water control (5.5 per plant) and there was a clear dose response (Fig. 4).


A B

C D

Figure 4: Mean number of CRF larvae and pupae recovered per plant from pots treated with (A) spinosad

(Tracer); (B) S. feltiae (Nemasys); (C) M. brunneum (Met52 incorporated compost); (D) M. brunneum

(Met52 drench). Applications were made at a range of concentrations from 1-200% of manufacturer’s

recommended concentration. The experiment was done using “conventional” brassica seed pelleted in a

fungicide-containing seed coat. Error bars represent the standard error of the mean.

In the second experiment, we measured the effect of different concentrations of M. brunneum, used as

Met52 compost incorporated granules, applied to plants grown from conventional seed (fungicide coated)

versus organic seed (no fungicide coating). The plants of both treatments did not grow enough root mass to

support the development of CRF larvae through to pupal stage. Therefore, the experiment was assessed

my measuring the effect of treatments on relative plant vitality score as an indication of CRF feeding

damage (Fig. 5). Plants were scored on a scale of 0 – 3, where 0 was classified as a healthy plant (no

wilting, fully expanded leaves, no discolouration) while 3 was classified as plant death. There was no

apparent pattern between plant vitality score and the concentration of Met52 applied to compost for the

conventional brassica seed. However there were indications of a dose response on the organic seed, with a

significant (P < 0.05) improvement in plant vitality score on organic seed with Met52 at 200% of the

manufacturer’s recommended concentration (Fig. 5).

A B

Figure 5: Plant vitality score after 2 weeks for (A) organic seed and (B) standard treated seed. Error bars

represent the standard error of the mean.


Task 2.4. Effect of combining M. brunneum + spinosad, and M. brunneum + S. feltiae, on populations of

cabbage root fly larvae feeding on brassica plants, quantified in a glasshouse experiment

This experiment investigated the effect of combined applications of M. brunneum (at 100% of

manufacturer’s recommended concentration, MRC) S. feltiae (100% MRC) and spinosad (used at 1% MRC)

against CRF. The experiment was done twice: firstly using conventional seed (to reflect standard grower

practice) and then using organic seed. In the first experiment, the plants did not grow enough root mass to

support the development of CRF larvae through to pupal stage. Therefore, the experiment was assessed

my measuring the effect of treatments on plant vitality score as an indication of CRF feeding damage (Fig.

6). There was no significant effect (P > 0.05) on plant vitality score with (i) M. brunneum alone (ii) S. feltiae

alone (iii) M. brunneum combined with S. feltiae (iv) spinosad in combination with S. feltiae. There was a

significant (P < 0.05) effect of spinosad alone, and spinosad in combination with M. brunneum, on plant

vitality score.

Figure 6: Task 2.4. Plant vitality score after 2 weeks for plants treated with M. brunneum (Met52), S. feltiae

(Nemasys), spinosad (at 1% of manufacturer’s recommended concentration) and their combinations. Error

bars represent the standard error of the mean.

When the experiment was repeated with the organic seed, the plants had a better root system and were

able to support the cabbage root fly population throughout the experiment. Mean numbers of CRF larvae

plus pupae per plant were significantly reduced (P < 0.05) by spinosad alone (2.8 per plant) and in

combination with M. brunneum (1.9 per plant) or with S. feltiae (2.7 per plant) compared with those in the

water controls (9.1 per plant) (Fig. 7). Mean numbers of CRF larvae and pupae per plant were not reduced

by M. brunneum alone (10.2 per plant) or S. feltiae (9.7 per plant) alone or when the two agents were used

together (9.0 per plant) compared to the control.

The mean plant weight (g) was significantly (P< 0.05) increased by spinosad alone (53.78 g per plant ) and

in combination with M. brunneum (59.65) or with S. feltiae (55.45) compared to the control control (35.78g),

(Fig. 8). Mean plant weight (g) was not increased by M. brunneum (41.48g) or S. feltiae (35.39g) when used

alone. However, mean plant weight (g) was significantly increased by the combination of M. brunneum and

S. feltiae (44.47g), (P<0.05) although this treatment was not as effective as spinosad alone or the spinosad

combination treatments.


Figure 7: Task 2.4. Mean number of CRF larvae and pupae recovered per plant following treatment with M.

brunneum (Met52), S. feltiae (Nemasys), spinosad (at 1% MRC) and their combinations (organic brassica

seed). Error bars represent the standard error of the mean.

Figure 8: Task 2.4. Mean weight of brassica plants infested with CRF larvae and pupae and treated with M.

brunneum (Met52), S. feltiae (Nemasys), spinosad (at 1% MRC) and their combinations (organic brassica

seed). Error bars represent the standard error of the mean.

Task 2.5. Effect of a combination of entomopathogens and spinosad against CRF larvae (field simulation)

Numbers of CRF larvae and pupae per plant. Mean numbers of CRF larvae plus pupae per plant were

significantly reduced by spinosad alone (0.6 per plant) and in combination with M. brunneum (1.2 per plant)

or with S. feltiae (0.67 per plant) compared with those in the water controls (4.73 per plant), P<0.05, Figure

9. Spinosad alone or in combination with M. brunneum or S. feltiae were equally effective. Mean numbers

of CRF larvae and pupae per plant were not reduced by M. brunneum (5 per plant) or S. feltiae (4.27 per

plant) when used alone. However, mean numbers of CRF larvae and pupae were significantly reduced by

the combination of M. brunneum and S. feltiae (2.73 per plant), (P<0.05) although this treatment was not as

effective as spinosad or the spinosad combination treatments.


Figure 9. Task 2.5. Mean numbers of CRF larvae and pupae per plant following treatment with M.

brunneum (Met52), S. feltiae (Nemasys), spinosad (at 1% MRC) and their combinations. Bars with different

letters are significantly different (P<0.05).

Mean Root Damage Index (RDI). The Mean Root Damage Index measures the percentage of root damage

caused by larval feeding, e.g. 40% damage is given an RDI of 40 and this RDI value is considered as the

‘threshold’ above which CRF reduces cauliflower yield if other factors e.g. drought are not limiting. The

mean RDI was significantly reduced to below 40 by spinosad alone (27) and in combination with M.

brunneum (30.7) or with S. feltiae (17.0) compared with that in the water controls (96.3), P<0.05, Figure 2.

Spinosad alone or in combination with M. brunneum or S. feltiae were equally effective in reducing RDI. The

mean RDI was not reduced by M. brunneum (97.3) or S. feltiae (95.7) when used alone or in combination

(88.7).

Figure 11. Task 2.5. Mean RDI per plant following treatment with M. brunneum (Met52), S. feltiae

(Nemasys), spinosad (at 1% MRC) and their combinations. Bars with different letters are significantly

different (P<0.05).

Mean plant height. The mean plant height (cm) was significantly increased by spinosad alone (14.97) and in

combination with M. brunneum (14.87) or with S. feltiae (13.65) compared with that in the water controls

(5.84), P<0.05, Figure 3. Spinosad alone or in combination with M. brunneum or S. feltiae were equally

effective in increasing plant height. Mean plant height was not increased by M. brunneum (3.81) or S. feltiae

(5.22) when used alone or in combination (6.59).

Mean plant weight. The mean plant weight (g) was significantly increased by spinosad alone (20.57) and in

combination with M. brunneum (20.56) or with S. feltiae (19.04) compared with that in the water controls

(1.44), P<0.05, Figure 4. Spinosad alone or in combination with M. brunneum or S. feltiae were equally

effective in increasing plant weight. Mean plant height was not increased by M. brunneum (1.19) or S.

feltiae (1.86) when used alone or in combination (2.60).

Compost temperatures. The air temperature in the glasshouse was set at 20°C. Mean, maximum and

minimum compost temperatures recorded with dataloggers were within the activity range for S. feltiae (10-

30°C) throughout the experimental period (Figure 5). Minimum compost temperatures fell below 15°C

between 28 January and 12 February, which may have affected M. brunneum activity, but mean

temperatures were above 15°C throughout the experimental period. M. brunneum activity is fastest


between 20 and 30°C. Although maximum compost temperatures exceeded 20°C on many dates, mean

temperatures were 15-18°C which may have delayed infection during the 4-week experiment.

Figure 12. Task 2.5. Mean plant height (cm) following treatment with M. brunneum (Met52), S. feltiae

(Nemasys), spinosad (at 1% MRC) and their combinations. Bars with different letters are significantly

different (P<0.05).

Figure 13. Task 2.5. Mean plant weight (g) following treatment with M. brunneum (Met52), S. feltiae (Nemasys), spinosad (at 1% MRC) and their combinations. Bars with different letters are significantly different (P<0.05).

Figure 14. Daily mean, maximum and minimum compost temperatures during experiment 2.5. The black horizontal line indicates 15°C, which is considered to be the lower temperature for M. brunneum activity.


Conclusions and discussion

The purpose of Objective 2 was to investigate the effects on CRF of (i) combining two biopesticides

together (M. brunneum and S. feltiae) and (ii) combining each of the biopesticides with a low dose

application of the chemical insecticide spinosad.

We had established in Objective 1 that low dose applications of spinosad were unlikely to impair M.

brunneum or S. feltiae, and that M. brunneum and S. feltiae were also likely to be compatible when

used together. Objective 2 consisted of a series of glasshouse experiments against CRF larvae feeding

on potted brassica plants. The work started with a pilot experiment (Task 2.2). This was followed by

research to investigate the dose response of M. brunneum, S. feltiae and spinosad against CRF larvae

using conventional and organic brassica seed (Task 2.3). There were then two relatively large

experiments (Tasks 2.4 and 2.5) to investigate the effect of combined applications of M. brunneum, S.

feltiae and spinosad against CRF: (i) in task 2.4, the control agents were applied to plants that had

been sown directly into 0.7 litre pots (M. brunneum was incorporated directly into compost at the time of

planting, while S. feltiae and spinosad were applied as drenches immediately before the application of

CRF eggs to the plants); (ii) in Task 2.5, the control agents were applied to brassica plants grown in

small modules before transplanting them to 1 litre pots (to simulate field conditions).

We did not find evidence that combining M. brunneum or S. feltiae with a low dose application of

spinosad affected the level of CRF control compared to spinosad on its own. The spinosad treatments

generally gave good control on their own, although the amount of control varied from experiment to

experiment. For example, in Task 2.2, we observed no significant control with 5% spinosad, whereas

we did get significant control with 5% spinosad in Task 2.3. This variability may have been caused by

differences in application methods, plant vigour, susceptibility of the pest, or environmental conditions.

In Task 2.5, spinosad, used at 1% of the recommended rate, was effective in reducing numbers of CRF

larvae and pupae, root damage and in increasing plant size and weight.

We found that combining M. brunneum and S. feltiae together could give significant control of CRF,

whereas these agents did not control CRF when used on their own. However, the action of the

combined treatment was not consistent across all experiments: it was observed in Tasks 2.2 and 2.5

but not in task 2.4. The mechanism for the effect of the M. brunneum–S. feltiae combination in 2.2 and

2.5 is not known: given that neither M brunneum nor S. feltiae had an effect on CRF on their own, this

looks to be a case of mutual potentiation between the two agents in the combination. The reason for

the lack of control in the combination treatment in Task 2.4 is not known. We suspect that it may be

connected to the type of compost used in the experiments. In Tasks 2.2 and 2.5 the biopesticides were

applied to medium structure, peat-based composts (Levington M2 and brassica growing compost

respectively) while experiment 2.4 used fine structure compost (Levington F2). It is possible that the

medium grade composts were better able to retain M. brunneum conidia and S. feltiae IJs within the

root zone next to CRF larvae, whereas they could have been more susceptible to being washed from

the root zone by plant watering in the fine structure compost. Soil composition and structure are known

to influence the vertical movement of entomopathogenic fungi and nematodes, and this is an area that

may require investigation in future. This illustrates one of the key challenges of conducting experiments

on biologically-based IPM, in that we are dealing with a complex pest management system involving

multiple variables. Potential sources of variation in our study included: (i) biological factors (biopesticide

virulence, pest susceptibility to infection, plant vigour and resistance to CRF damage); (ii)

environmental factors (compost type, temperature, the presence of seed-coated fungicides); (iii)

experimental practice (e.g. the way in which the biopesticides were applied).

We found variation in plant vigour between experiments, which affected the ability of plants to support

CRF populations. In experiment no.1 done in Task 2.3, and experiment no. 1 in Task 2.4, the plants did

not produce sufficient root mass in the presence of CRF larvae to support the survival and development

of the larvae through to the pupal stage. As a result, we recovered low numbers of larvae and pupae

and were not able to analyse the data statistically. Instead, we recorded and analysed the effect of the

treatments using a plant vitality score. The reasons for variation in plant vigour between experiments is

not known, but it may have been affected by the time of year in which the experiments were done and

glasshouse conditions (e.g. temperature or supplementary lighting).

Task 2.3 was intended to measure the effect of different doses of spinosad, M. brunneum and S. feltiae

on control of CRF. A clear dose response for all three agents would have enabled us to plot the effects

of the agents as isobolograms (a plot of the effect of one agent versus the effect of the other at different

doses). This allows the experimenter to determine whether the combination has a synergistic,


antagonistic or additive effect compared to the solo treatments. We obtained a dose response with

spinosad, but we did not obtain a dose response with M. brunneum or S. feltiae. The doses used for the

biopesticides were based around the manufacturers recommended concentration for M. brunneum and

S. feltiae against CRF. It is possible that the lack of dose response was connected with the type of

compost used in the experiment (Levington F1, see above). However we also found evidence of an

effect of the coating used on the pelletized seed. We repeated the multiple dose experiment for M.

brunneum using organic seed and measured the effect on plants using the plant vitality score method,

and here we did get a significant improvement in plant vitality using M brunneum at twice the

manufacturer’s recommended rate. This finding suggests that the efficacy of M. brunneum was

negatively affected by the fungicide coating used for the brassica seed in experiment No. 1 for Task

2.3. Despite this, we took the decision to use conventional seed in subsequent experiments (Tasks 2.4

and 2.5) since this reflects normal grower practice. However the effect of fungicide seed coating is an

issue that would be worth investigating in future. If there had been sufficient time, we would have

conducted more dose response studies to obtain a definitive dose response curve for M brunneum and

S. feltiae using conventional and organic seed. However, because we were on limited time, we took the

decision to investigate the effect of combination treatments in experiments 2.4 and 2.5 using the

manufacturer’s recommended concentrations for M. brunneum and S. feltiae. There is also an obvious

issue from this work that the manufacturers current recommended concentrations for M. brunneum and

S. feltiae may well not be high enough to give effective control of CRF, and this is also worth

investigating in future.

In Tasks 2.2 and 2.5, although the combined application of M. brunneum and S. feltiae caused a

significant reduction in CRF numbers, the effect was never as effective as using the insecticide

spinosad on its own. This is a useful observation, but it does not detract from the main objective of the

study, i.e. to obtain new information on the effects of combination treatments in IPM.

We also looked at some other indicators of treatment effects in the experiments. In experiment No. 2 in

task 2.4, we observed that mean plant weight was not increased by M. brunneum or S. feltiae when

used alone, but was significantly increased by the combination of M. brunneum and S. feltiae. In Task

2.4, we recorded root damage index, plant weight and height. There was no significant effect of M.

brunneum or S. feltiae alone or in combination on these indicators.

Overall conclusions and future work

We found that combining the biopesticides M. brunneum and S. feltiae together could give significant

control of CRF, whereas these agents did not control CRF when used on their own. However, the effect

was only observed in two out of three experiments, and it is likely that the outcome of the interaction

between M. brunneum and S. feltiae was affected by external factors such as compost type. The

combination of M. brunneum and S. feltiae was much less effective than using the chemical insecticide

spinosad. However, the research did provide proof of concept, i.e. that biopesticide combination treatments

can give improved pest control. As already mentioned, this approach to IPM is complex, and is difficult to

study experimentally, as it involves multiple variables. Nevertheless, it is an important area for study,

because the use of biopesticides is being promoted through the EC Sustainable Use Directive and –

because biopesticides are usually not as efficacious as fully effective chemical insecticides – it is likely that

different biopesticides will have to be used together or with other crop protection agents. We know from

previous work that this is already being done on some UK protected crops for other pest species, but at the

moment it is based on anecdotal evidence.

There are a number of areas that need to be investigated in future work. For the CRF pest system, there is

a requirement to do the following: (i) determine the dose response of M. brunneum and S. feltiae; (ii)

understand how compost type in brassica modules and use of fungicides affects biopesticide efficacy and

mediates the interaction of M. brunneum and S. feltiae; (iii) investigate whether using M. brunneum and S.

feltiae (or other biopesticides) can give economically beneficial levels of CRF control in commercial field

conditions (note that, at present, commercial propagators use peat-based compost for raising brassica

plants, but there is increasing pressure to use peat replacement in horticultural substrates). There is also a

requirement to investigate whether the combination treatments studied against CRF also have effects on

other pests. In particular, there is a need to investigate the effects of combination treatments on pests that

have evolved resistance to chemical insecticices, since there is evidence that biopesticides can slow the

evolution of pesticide resistance and reduce the expression of resistance after it has evolved. A highly

suitable pest for this would be western flower thrips, Frankliniella occidentalis, as it is currently economically

very damaging to a wide range of horticultural crops, shows widespread pesticide resistance, including to


spinosad, and is susceptible to both EPF and EPNs, both of which are being used by growers with varying

degrees of success. Note that we originally proposed including western flower thrips in the project, but this

was removed in order to reduce the cost and timeframe of the project, with a view to investigating the

approach with other pests later.

Acknowledgements We are grateful for advice received during the project from Novozymes UK Ltd., BASF UK, and Fargro Ltd.

References to published material

9. This section should be used to record links (hypertext links where possible) or references to other published material generated by, or relating to this project.

References

Ansari, M. A., Tirry, L. & Moens, M. (2005). Antagonism between entomopathogenic fungi and bacterial

symbionts of entomopathogenic nematodes. BioControl, 50, 465 – 475.

Ansari, M.A., Shah, F.A., & Butt, T.M. (2008). Combined use of entomopathogenic nematodes and

Metarhizium anisopliae as a new approach for black vine weevil, Otiorhynchus sulcatus, control.

Entomologia Experimentalis et Applicata 129, 340-347.

Bailey, A. S., Chandler, D., Grant, W. P., Greaves, J., Prince, G. & Tatchell, G. M. (2010). Biopesticides:

Pest management and regulation. CABI, Wallingford UK, 232 pp.

Chandler, D. & Davidson, G. (2005). Evaluation of the entomopathogenic fungus Metarhizium anisopliae

against soil dwelling stages of the cabbage maggot, Delia radicum (L.) (Diptera: Anthomyiidae), in

glasshouse and field experiments, and the effect of fungicides on fungal activity. Journal of

Economic Entomology, 98, 1856 – 1862.

Chandler, D, Bailey, A.S., Tatchell, G. M., Davidson, G., Greaves, J. & Grant, W. P. (2011a). The

development, regulation and use of biopesticides for Integrated Pest Management. Philosophical

Transactions of the Royal Society B 366, 1987 - 1998.

Chandler, D., Pope, T. & Bennison, J. (2011b) A desk study of current knowledge on the combined use of

microbial biopesticides and chemical pesticides in Integrated Pest Management. Defra final report

PS2135, 20 pp.

Davidson, G. & Chandler, D. (2005). Laboratory evaluation of entomopathogenic fungi against larvae and

adults of the onion maggot (Diptera: Anthomyiidae). Journal of Economic Entomology, 98, 1848-

1855.

Farenhorst, M., Mouatcho, J.C., Kikankie, C.K., Brooke, B.D., Hunt, R.H., Thomas, M.B., Koekemoer, L.L.,

Knols, B.G.J., and Coetzee, M. (2009). Fungal infection counters insecticide resistance in African

malaria mosquitoes. Proceedings of the National Academy of Sciences of the USA 106, 17443-

17447.

Farenhorst, M., Knols, B.G.J., Thomas, M.B., Howard, A.F.V., Takken, W., Rowland, M., and N'Guessan,

R. (2010). Synergy in efficacy of fungal entomopathogens and permethrin against West African

insecticide-resistant Anopheles gambiae mosquitoes. Plos One 2010;5:e12081. doi:

10.1371/journal.pone.0012081.

Finch, S. 1989. Ecological considerations in the management of Delia pest species in vegetable crops.

Annual Review of Entomology, 34, 117 –137.

Furlong, M.J., and Groden, E. (2001). Evaluation of synergistic interactions between the Colorado potato

beetle (Coleoptera : Chrysomelidae) pathogen Beauveria bassiana and the insecticides,

imidacloprid, and cyromazine. Journal of Economic Entomology 94, 344-356.

Gassmann, A.J., Stock, S.P., Carriere, Y., and Tabashnik, B.E. (2006). Effect of entomopathogenic

nematodes on the fitness cost of resistance to Bt toxin Cry1Ac in pink bollworm (Lepidoptera :

Gelechiidae). Journal of Economic Entomology 99, 920-926.

Gassmann, A.J., Stock, S.P., Sisterson, M.S., Carriere, Y., and Tabashnik, B.E. (2008). Synergism

between entomopathogenic nematodes and Bacillus thuringiensis crops: integrating biological

control and resistance management. Journal of Applied Ecology 45, 957-966.

Jaronski, S. T. (2010). Ecological factors in the inundative release of fungal entomopathogens. BioControl,

55, 159 – 185.

Quintela, E.D., and McCoy, C.W. (1997). Pathogenicity enhancement of Metarhizium anisopliae and

Beauveria bassiana to first instars of Diaprepes abbreviatus (Coleoptera: Curculionidae) with

sublethal doses of imidacloprid. Environmental Entomology 26, 1173-1182.


Raymond, B., Sayyed, A.H., and Wright, D.J. (2006). The compatibility of a nucleopolyhedrosis virus

control with resistance management for Bacillus thuringiensis: Co-infection and cross-resistance

studies with the diamondback moth, Plutella xylostella. Journal of Invertebrate Pathology 93, 114-

120.

Raymond, B., Sayyed, A.H., Hails, R.S., and Wright, D.J. (2007). Exploiting pathogens and their impact on

fitness costs to manage the evolution of resistance to Bacillus thuringiensis. Journal of Applied

Ecology 44, 768-780.

Roditakis, E., Couzin, I.D., Balrow, K., Franks, N.R., and Charnley, A.K. (2000). Improving secondary pick

up of insect fungal pathogen conidia by manipulating host behaviour. Annals of Applied Biology

137, 329-335.

Santos, A.V., de Oliveira, B.L., and Samuels, R.I. (2007). Selection of entomopathogenic fungi for use in

combination with sub-lethal doses of imidacloprid: perspectives for the control of the leaf-cutting

ant Atta sexdens rubropilosa Forel (Hymenoptera : Formicidae). Mycopathologia 163, 233-240.

Shah, F. A., Gaffney, M., Ansari, M. A., Prasad, M. & Butt, T. M. (2008) Neem seed cake enhances the

efficacy of the insect pathogenic fungus Metarhizium anisopliae for the control of black vine

weevil, Otiorhynchus sulcatus (Coleoptera: Curculionidae). Biological Control 44, 111-115.

Wraight, S.P., and Ramos, M.E. (2005). Synergistic interaction between Beauveria bassiana- and Bacillus

thuringiensis tenebrionis-based biopesticides applied against field populations of Colorado potato

beetle larvae. Journal of Invertebrate Pathology 90, 139-150.

Ye, S.D., Dun, Y.H., and Feng, M.G. (2005). Time and concentration dependent interactions of Beauveria

bassiana with sublethal rates of imidacloprid against the aphid pests Macrosiphoniella sanborni

and Myzus persicae. Annals of Applied Biology 146, 459-468.

Evidence Project Final Report - Defra,...

Documents

Transcript of Evidence Project Final Report - Defra,...