The Small Hive Beetle Trap

External attractant trap for small hive beetle

by D M Leemon, R A Hayes, B A Amos, S J Rice, D K Baker, K McGlashan February 2018

External attractant trap for small hive beetle

D M Leemon, R A Hayes, B A Amos, S J Rice, D K Baker, K McGlashan

February 2018

AgriFutures Australia Publication No 18/062 AgriFutures Australia Project No PRJ-009334

ISBN 978-1-76053-018-1 ISSN 1440-6845

External attractant trap for small hive beetle Publication No. 18/062 Project No. PRJ-009334

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Researcher Contact Details

Name: Dr Diana Leemon Address: Agri-Science Qld, Level 2AW, Ecosciences Precinct. GPO Box 267 Brisbane . 4001

Phone: +61 (7) 3708 8366 Email: [email protected]

In submitting this report, the researcher has agreed to AgriFutures Australia publishing this material in its edited form.

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Foreword

This research has developed a simple method for trapping small hive beetles before they enter hives and delivered the most comprehensive seasonal data on small hive beetle movement in Australia to date. Small hive beetle is now a serious pervasive pest of beekeeping in Australia in areas with mild winters and humid wet summers. Although a number of internal trapping systems are available, it is desirable to intercept small hive beetles before they enter a hive.

This research will benefit the Australian honey bee industry by providing a trapping system for small hive beetle including a simple trap and knowledge of when best to deploy traps and where best to locate the trap in respect of hives.

The key findings are that natural fermentation attractants while useful are subject to variation; thus development of a synthetic lure blended from the key attractant components in fermentation volatiles should be the ultimate goal. Results from investigations into the placement of traps and movement of small hive beetles throughout the year in Australia have provided invaluable ecological data to inform external trapping and monitoring strategies.

This study emphasises the importance of beekeepers managing the increase in numbers of small hive beetles potentially entering their hives during spring and after wet periods during late summer and early autumn.

This report is an addition to AgriFutures Australia’s diverse range of over 2000 research publications and it forms part of our Honey Bee and Pollination Program. The R&D program aims to support research, development and extension that will secure a productive, sustainable and more profitable Australian beekeeping industry and secure the pollination of Australia’s horticultural and agricultural crops into the future on a sustainable and profitable basis.

Most of AgriFutures Australia’s publications are available for viewing, free downloading or purchasing online at www.agrifutures.com.au.

John Harvey Managing Director AgriFutures Australia

Acknowledgments

We are very grateful for assistance and support from the Stradbroke Organic Beekeepers, Phill and Theresa Bowman, who allowed us to use their Gumdale apiary in our field studies. Phill often helped with the fieldwork and kindly starred in our educational video. We also greatly appreciate support from the Wheen Bee Foundation who provided financial and in kind support, especially allowing us to use the Wheen Bee Foundation research apiary in Richmond after it was launched early in 2017. The Wheen Bee Foundation CEO Fiona Chambers moved heaven and earth to get the research apiary ready for us to conduct our 2017 research trials. We thank Bruce White and the Hawkesbury Amateur Beekeepers Inc. who in a voluntary capacity carried out the 2017 fieldwork for us at the research apiary. We also thank Laura Rittenhouse and Frank for so enthusiastically helping us with earlier fieldwork from 2016 as well as hosting our visits to the Wheen Bee Foundation site. Queensland Beekeepers’ Association is acknowledged for its financial support provided for this project.

We would also like to thank other various Beekeepers who provided the brood and honeycomb samples used for the study of the geographical variation of hive products (Michael Duncan, Wayne Fuller, James Kershaw, Peter McMahon, Rob Stephens, John Zitgerman). We are also appreciative to a number of people who provided sites for testing traps (Chris Butcher of Corinda State High, Stephen Firth, Manon Griffiths, Lloyd Hancock, Wayne Jorgensen, Charles and Mary-Ann Millar, Helen Nahrung).

Dr David Mayer of the Queensland Department of Agriculture and Fisheries provided support for statistical analyses.

The late Dr Peter Teal and Dr Charlie Stuhl (USDA) for sharing their knowledge and experience with SHB chemical ecology.

All photos in this report were taken by the research team, most being taken by Diana Leemon or Andrew Hayes.

Abbreviations

DAF – Department of Agriculture and Fisheries (Queensland)

GC – Gas chromatography

MS - Mass spectrometry

NSW – New South Wales

QLD - Queensland

RH – Relative humidity

SHB – Small hive beetle

USA – United States of America

Contents

Foreword ...... 3 Acknowledgments ...... 4 Abbreviations ...... 5 Executive Summary ...... 13 Introduction ...... 17 Objectives ...... 20 Methodology...... 21 2.1 Colony ...... 21 2.2 Location of field studies ...... 21 2.3 Fermentation of hive products ...... 21 2.3.1 Geographical variation in source of hive materials ...... 22 2.3.2. Behavioural bioassays ...... 22 2.3.3 Chemical analysis ...... 23 2.4 Individual compounds attractive to SHB ...... 25 2.5 Synthetic lure development ...... 27 2.5.1. Carpophilus system evaluation ...... 27 2.5.2 Blend development and testing ...... 27 2.6 Aggregation pheromone ...... 30 2.6.1 Arena aggregation trials ...... 30 2.6.2 Chemistry of aggregation ...... 32 2.7 Volatile profiles of hives ...... 34 2.8 Trap design ...... 34 2.8.1 Trap evaluations ...... 35 2.9 Evaluation of efficacy of synthetic lure and trap ...... 37 2.9.1 Natural fermentate attractant investigations ...... 37 2.9.2 Ecological studies of SHB ...... 38 2.9.3 Synthetic lure testing ...... 43 2.10 Production of an educational video ...... 44 2.11 Statistical analyses ...... 45 Results ...... 46 3.1 Fermentation of hive products ...... 46 3.1.1 Behavioural bioassays ...... 46 3.1.2 Chemical analysis ...... 50 3.2 Individual compounds attractive to SHB ...... 65 3.3 Synthetic lure development ...... 66 3.3.1 Carpophilus system evaluation ...... 66 3.3.2. Blend testing ...... 66 3.4 Aggregation pheromone ...... 70 3.5 Volatile profile of hives ...... 77 3.6 Trap design ...... 83

3.7 Evaluation of efficacy of synthetic lures and traps ...... 85 3.7.1 Natural fermentate attractant investigations ...... 85 3.7.2 Ecological studies of SHB ...... 89 3.8 Educational video ...... 102 Implications ...... 103 Recommendations ...... 106 Glossary ...... 107 References ...... 109

Figures

Figure 1:SHB larvae converting honeycomb and brood (A) to slime (B)...... 22 Figure 2: Map of hive material sources and trial locations...... 23 Figure 3: Behavioural choice assay system in BugDorm tents. (A) Beetle trap (B) cage arena system (C) replicated cage arenas in bioassay...... 24 Figure 4: Volatile samples being collected for analysis from the air above fermenting slime in an Erlenmeyer flask...... 24 Figure 5: Gas chromatograph and mass spectrometer for analysis of volatile compounds in air samples...... 25 Figure 6: Single choice assay system for SHB volatile compound attractiveness testing. (A) individual arena (B) group of replicated arenas used in a bioassay...... 27 Figure 7: Aeration of (clustered) groups of SHB with saturated sucrose solution on crumpled paper provided for energy...... 33 Figure 8: Hive aeration sampling, at the Richmond (A) and Bellbowrie (B) field sites. Note the Tenax sample tube attached by plastic tubing to an air pump is inserted into the hive entrance...... 34 Figure 9: (A) Unitrap™ baited with slime, (B) Dowd trap baited with slime and (C) lantern trap with honey fermentate...... 35 Figure 10: Fibrous mat folded and placed on top of frames under the hive lid, with the fibrous side to the inside and vinyl to the outside. (A) Showing the positioning of the mat in the hive. (B) Mat removed from hive showing SHB trapped in the fibrous side. (C) Close up view showing how SHB become trapped in the mat when their legs get entangled in the fibres. . 39 Figure 11: (A) Trench traps being removed from a hive as part of the fortnightly hive check assessments of SHB burden, (B) SHB caught in the diatomaceous earth (DE) used inside a trench trap removed from a hive...... 39 Figure 12: Aspiration of SHB from frame during a hive validation to assess total number of SHB in a hive...... 41 Figure 13: Lantern trap modified for synthetic lure testing with a different odour release mechanisms. (A) Blend in falcon tube suspended over DE, (B) blend in Falcon tube suspended over vegetable oil and (C) blend in baggie suspended over DE...... 44 Figure 14: Sample radial plots (below) showing mean (+ SEM) proportion beetles attracted to each treatment for the same samples. Filled blue triangle showns mean proportion attracted, empty blue triangle is mean + SEM. Red triangle shows random attractiveness . 48 Figure 15: : Sample pie charts (above) showing mean proportion of small hive beetles attracted to treatments on each day from Bellbowrie, Maleny and Grafton. ( control brood honey slime not trapped)...... 49 Figure 16: Non-metric multidimensional scaling ordination showing clear differences between attractive and non-attractive samples. Each point in the ordination represents a single sample...... 50 Figure 17: Percentage of beetles attracted to fermenting hive products (slime) on days 18, 35 and 49 after inoculation with beetles across eight apiary locations. Columns headed with different letters are significantly different...... 51 Figure 18: Total ion chromatograms of an attractive (Canberra Day 18, top) and non- attractive (Canberra Day 35, bottom) sample showing differences in compounds detected. Key compounds are identified by number...... 64 Figure 19: Male clustering in aggregation arena with logarithmic trend line showing the increasing size of clusters with time...... 72 Figure 20: SHB showing clustering behaviour recorded over 15 min in an arena (glass Petri dish)...... 72 Figure 21: Total ion chromatogram showing differences in aerations of: pollen dough alone, SHB females before and after feeding on pollen dough. Compounds are identified below: . 76

Figure 22 Total ion chromatogram of an aeration of a hive from the Bellbowrie apiary (sampled 8 April 2016), air was sampled for one hour at approximately 500 ml/min onto a thermal desorption tube preloaded with Tenax TA (35/60 mesh) (Markes International Ltd.)...... 78 Figure 23: Non-metric multidimensional scaling ordination showing clear differences between samples from the two apiary sites. Each point in the ordination represents a single hive sample at a single sampling time. Ellipses added for ease of interpretation only...... 79 Figure 24: Non-metric multidimensional scaling ordination showing hive aeration samples collected across the sampling periods. There is clear clustering of samples collected from each sample period, with samples from early April significantly different from the other sampling intervals. Each point in the ordination represents a single hive sample at a single sampling time. Ellipses added for ease of interpretation only...... 79 Figure 25: Total ion chromatograms comparing aerations of a hive from the Bellbowrie apiary (lower) (sampled 24 May 2016), and from the Wheen Bee Foundation Research apiary (upper) (sampled 18 May 2016). Air was sampled for one hour at approximately 500 ml/min onto a thermal desorption tube preloaded with Tenax TA (35/60 mesh) (Markes International Ltd.). Key compounds important in distinguishing between the apiaries are identified by number, as in Figure 22 and their percentage contribution to the dissimilarity is shown below: ...... 80 Figure 26: Total ion chromatograms comparing aerations from different sampling times of a hive from the Bellbowrie apiary (sampled from bottom to top: 8 April, 27 April, 10 May and 24 May 2016). Air was sampled for one hour at approximately 500 ml/min onto a thermal desorption tube preloaded with Tenax TA (35/60 mesh) (Markes International Ltd.). Key compounds important in distinguishing between the apiaries are identified by number, as in Figure 22 and their percentage contribution to the dissimilarity is shown below...... 81 Figure 27: Non-metric multidimensional scaling ordination showing hive aeration samples collected across the sampling periods, showing only samples from the Bellbowrie apiary. There is, again, clear clustering of samples collected from each sample period, with samples from early April significantly different from the other sampling intervals. Each point in the ordination represents a single hive sample at a single sampling time. Ellipses added for ease of interpretation only...... 82 Figure 28: Non-metric multidimensional scaling ordination showing samples with respect to hive, only samples from the Bellbowrie apiary site are shown. There is no clustering of samples from within a hive, instead samples cluster with respect to sampling interval (see Figure 27). Each point in the ordination represents a single hive sample at a single sampling time...... 82 Figure 29: Comparison of attractiveness of honey and sucrose fermentate to SHB when used in lantern traps in the field...... 87 Figure 30: Representative chromatograms of the honey blend (A: 24 hrs; B: 144 hrs) and the sugar blend (C: 24 hrs; D: 120 hrs), with the peaks of the common components numbered. Ethanol (1), ethyl acetate (2), isobutanol (3), isopentanol (4), and 2-methyl butanol (5). Note that the scale of abundance is around 9x greater for graphs A and B than in C and D...... 88 Figure 31: Correlation between the visual + mat counts of SHB and fortnightly hive check counts at Bellbowrie...... 91 Figure 32: Correlation between the visual and fibrous mat counts of SHB and fortnightly hive check counts at Gumdale...... 91 Figure 33: Mean number of SHB (+SE) caught in lantern traps with a natural fermentation attractant at different distances from the Richmond apiary...... 92 Figure 34 : Mean number of SHB (+SE) caught in lantern traps with a natural fermentation attractant at different distances from the Gumdale apiary...... 93 Figure 35: Mean number of SHB (±SE) trapped per trap per day in attractant yeast traps at different distances from the Bellbowrie apiary over different seasons. (A) Later summer to early autumn (23/12/2016-31/3/2017). (B) Autumn through winter to early spring (1/4/2017 – 26/9/2107). (C) Spring (2/9/2017 – 22/11/2017)...... 95

Figure 36: SHB trapped in external attractant traps (lantern traps) and in the hives (hive check) in the (A) Bellbowrie and (B) Gumdale apiaries over 15 months...... 97 Figure 37 : Relationship between total numbers of SHB trapped inside hives (through hive check) and in external attractant traps the (A) Bellbowrie and (B) Gumdale apiaries and minimum daily temperature...... 98 Figure 38: Relationship between total numbers of SHB trapped inside hives (through hive check) and in external attractant traps at the (A) Bellbowrie and (B) Gumdale apiaries and daily rainfall...... 99 Figure 39: Total ion chromatograms of headspace of blend #4 in agar (1.5% in H2O) at time zero (above) and after 48 hours (below) showing differences in compounds detected. Key compounds are identified by number...... 100

Tables

Table 1: The ten most important compounds in distinguishing between attractive and non- attractive slime samples, including their known role in insect chemical ecology ...... 26 Table 2: Components of blends tested in laboratory assays and in field trial ...... 29 Table 3: Groups of SHB investigated for clustering indicative of aggregative behaviour in arena assays, including number and type of SHB used and number of replicates for each investigation...... 31 Table 4: Aerations of groups of SHB of different sex and mating status with 20 SHB per group, the number of replications of each assay is also shown...... 32 Table 5: Aerations of groups of “young” SHB of different sex, mating status and feeding status with 20 SHB per group, the number of replications of each assay is also shown...... 33 Table 6: Different trap designs and attractants tested for SHB trapping near apiary and in non-apiary sites between December 2014 and December 2016...... 36 Table 7: Dates when fortnightly assessments of SHB numbers in hives (relative measure) and total hive counts of SHB (validation) conducted ...... 40 Table 8: Details of traps deployed for collecting field data on SHB movement around three different apiaries (Bellbowrie, Gumdale and Richmond) ...... 42 Table 9: Percentage (± SEM) of small hive beetles attracted to slime. In virtually all samples, there was a significant effect of treatment on beetle choice, but only those samples where the slime was significantly more attractive than brood comb are shaded...... 47 Table 10: The most important volatile compounds to distinguish between attractive and non- attractive samples, including the percentage contribution they make to distinguish the groups. Several of the compounds detected have been shown to be detected by the antennae ...... 65 Table 11: Attractiveness of Catcha® system components to the small hive beetle, individually and in combination...... 66 Table 12: Attractiveness of synthetic blends to the small hive beetle in the individual arena bioassay...... 67 Table 13: Attractiveness of synthetic blends against a blank control to the small hive beetle in the BugDorm bioassay...... 68 Table 14: Comparison between attractiveness to the small hive beetle of blank control and two concentrations of synthetic blends (100% and 1%) in the BugDorm bioassay...... 69 Table 15: Comparison between attractiveness to the small hive beetle of synthetic blends compared to a blank control in the BugDorm bioassay...... 70 Table 16: Aggregative behaviour of SHB as measured in arena assays by percentage of time spent in a cluster (≥ 5 beetles)...... 73 Table 17: Commonly occurring compounds (mean % abundance ± SEM) in aerations of SHB by treatment (only replicate aerations wherein compounds were present included). ... 74 Table 18: Commonly occurring compounds (mean % abundance ± SEM) in aerations of SHB by feeding status treatment (only replicate aerations wherein compounds were present included)...... 75 Table 19: Results of trials to evaluate different trap designs between December 2014 and November 2016...... 84 Table 20: Mean percent SHB trapped per choice in four BugDorm arena assays. Mean percent un-trapped beetles is the summed percentage of all choices subtracted from 100. Means with the same superscript within rows were not significantly different...... 85 Table 21: Comparing fermentates with different ratios of water to honey for attractiveness to SHB in two-choice individual arena assays. The split column shows relative numbers of beetles per choice...... 85 Table 22: Comparison of the attractiveness to SHB of fermentate (75:25 water/honey with yeast) at different ages (0 – 7 days old) in two-choice individual arena assays. The split column shows relative numbers of beetles per choice...... 86

Table 23: Relative abundance of the five common components seen in the representative chromatograms (Figure A) of the honey and sucrose fermentates after 24 hrs and 120/144 hrs of fermentation...... 88 Table 24: Percentage of SHB in a hive accounted for through the fortnightly relative measures (visual, mat or trench traps with DE) of SHB. Hive check = all three relative measures added together...... 90 Table 25: Mean number of SHB trapped per trap per trapping interval at different distances from hives. Letters in brackets after the mean denote significance...... 94 Table 26: Weekly trapping rates of SHB (SHB/trap/week) trapped in external attractant traps at three locations from late 2016 until November 2017...... 96 Table 27: Results of large tent two choice-assays (vs. control) with adult SHB responding to baggie mechanism of blend (#10) release comparative to positive control (yeast in lantern trap)...... 101 Table 28: The percentage composition of each compound in blend #10, and levels detected after five days in traps with glycerine ...... 101

Executive Summary

What is the report about?

This report describes the findings from a three-year investigation into the development of an external attractant trap for the small hive beetle (Aethina tumida). It describes investigations into the development of a synthetic attractant odour blend for use in the external trap. The research provides strategies to monitor movement of the beetles both in the field and in hives, and has the potential to assist in small hive beetle control in an apiary.

Who is the report targeted at?

This report is targeted at the Australian beekeeping industry for the better management of small hive beetle.

Where are the relevant industries located in Australia?

Key regions affected by small hive beetle include mostly coastal areas ranging from far north Queensland down to central New South Wales. Beekeepers operating in regions with mild winters and warm, moist summers are most at risk from this pest which, under ideal conditions, can destroy a hive in less than a fortnight.

Background

The Australian honey bee industry contributes at least $101 million annually to the Australian economy through honey products and an estimated $1.7 billion annually through pollination services to crops and horticulture. The small hive beetle (SHB) Aethina tumida Murray is a serious pest of European honey bees outside of its native range in sub-Saharan Africa. Small hive beetles were first reported in Australia in 2002 from Richmond, New South Wales. Within ten years SHB had spread along the east coast of Australia, to Mareeba in the north and the Melbourne CBD in the south. Adult SHB have been reported to travel kilometres to locate a honey bee hive, through olfactory cues. They are opportunistic scavengers taking advantage of situations that suit their reproductive strategy. The larval stage of SHB is the destructive stage causing damage to hives when they feed on bee brood and pollen stores and ferment honeycomb. Small hive beetles carry a yeast which is primarily responsible for the fermentation of hive products associated with larval development. The resulting fermented honey is rejected by honey bees and cannot be marketed by the beekeeper. Heavy infestations can result in total hive collapse when large numbers of SHB larvae feed voraciously on the hive protein stores of pollen and brood, and cause the fermentation of the honeycomb into what is colloquially and expressively termed “slime”.

Small hive beetle is now the predominant apiary pest in the warm, moist regions of eastern Australia. Warm temperatures and wet summers provide ideal conditions for populations of this beetle to build with several generations possible between spring and autumn. A conservative estimate of hive losses in Queensland alone ranged from a peak of $11 million in one wet humid season to just under $2 million in a season preceded by a very dry spring. SHB also appear to be able to breed in and destroy the hives of native stingless bees. Anecdotal evidence suggests that the unmanaged bees responsible for much of the “free” pollination services are disappearing in areas of heavy beetle infestation.

Adult SHB have been found to be attracted to a range of hive odours including the odours of adult worker bees, fermenting hive products, and a honey bee alarm pheromone. The odours associated with fermenting hive products or “slime” have been found to be especially

attractive to adult SHB. A range of in-hive control options are now being deployed by many beekeepers, most of which rely on the bees chasing SHB into a harbourage trap. These vary greatly in trap design and the method of lure presentation. However, there is a need for additional out of hive control measures such as an external attractant trap that can stop SHB entering hives. An external attractant trap would also be useful for biosecurity as a surveillance tool for monitoring SHB in areas of uncertain infestation or to demonstrate area freedom in areas where SHB are not yet known to occur.

Aims/objectives

The project aimed to develop an external trap for the small hive beetle, based on the attractive odours associated with fermenting hive products to assist commercial and hobbyist beekeepers in the monitoring, management and control of this apiary pest.

Methods used

Laboratory studies were used to investigate the attractiveness of a range of fermenting hive products to SHB. Hive products were provided by beekeepers in Queensland and New South Wales. The chemical composition of the fermented hive products was elucidated through the use of gas chromatography- mass spectrometry (GC-MS). Behavioural bioassays with SHB were used to confirm the attractiveness of individual compounds identified as attractive to SHB as well as blends of the individual compounds. Further behavioural bioassays were undertaken to assess aggregating behaviour of SHB. Chemical analysis of air samples from aggregating SHB using GC-MS was conducted to search for aggregation pheromones. Triggers to adult SHB emergence from soil were also investigated.

Field research was carried out in three apiaries at three different locations and involved external trapping and in-hive monitoring of beetle populations. Different trap designs and trap placement were tested across the apiary sites and at non-apiary sites. The seasonal movement and in-hive population changes of SHB were monitored over 15 months. The volatile profiles of hives in two different apiaries were investigated for possible correlation with SHB burden.

Results/key findings

The chemical composition and hence attractiveness of fermenting hive products to SHB was found to vary with source of hive products. The variation was used to identify ten compounds which were key to this attraction. These compounds have been tested in the laboratory in behavioural bioassays individually at a variety of concentrations, several of which were determined to be attractive. Synthetic blends were created with these compounds and again tested for attraction. Laboratory tests identified the best four synthetic blends for field testing.

Field tests at three apiary sites in Queensland and New South Wales examined the attractiveness of these blends, and in addition tested a variety of trap designs, odour release mechanisms and killing agents. One trap, readily available from hardware stores, was the most promising of the trap designs and was used at three apiary sites to gather seasonal data on SHB movement. However, a trap design customised for trapping and killing SHB is required. Trap placement studies using the simple traps with a natural fermentation attractant showed large numbers of SHB could be trapped close to hives (within 6 m and up to 185 m away). In two apiaries a set of traps 100 m from hives caught more SHB than a set placed 20 m from hives, although the difference was not statistically significant (p ≤0.05). The trapping efficacy varied at different distances and it is highly likely that wind direction is a key factor that needs further investigation. It is likely that the mechanism of the attractant used helps the traps intercept SHB flying towards an apiary.

Behavioural studies demonstrated that small hive beetles aggregate, and that this behaviour is mediated by volatile chemical signals. To date, we have been unable to detect the chemical signal of a putative aggregation pheromone, but this warrants further research.

Hive aerations sampled the volatile profile of hives in two different apiaries, however there was not enough difference in the SHB populations to establish a relationship between hive odour profile and SHB load. These investigations did generate background data on the volatile profile of healthy hives. This information provides a starting point for future odour studies investigating the volatile profile of hives carrying disease or even undergoing changes in relation to queen pheromone.

Ecological studies into the seasonal changes in SHB movement in the field and SHB populations in hives and moving demonstrated that temperature is a key factor. There was little movement through winter when temperatures were low. Small hive beetles began to move during spring after daily minimum temperatures increased above 12°C. The movement in autumn appeared to decrease once average daily temperatures started to drop below 20°C. During summer and early autumn, rainfall appeared to have a key role in initiating SHB movement. Beetles ceased to move during prolonged dry spells, but large numbers were trapped in the days following a significant rainfall event. These results provide the most comprehensive seasonal data on SHB movement in Australia to date.

Implications for relevant stakeholders

Much of the previous work conducted in field surveys of small hive beetle has used a variant of fermenting hive products, whether it be the use of inoculated pollen dough or a mixture of honey and brood comb itself. Our research demonstrates that not all fermenting hive products are the same, and that odours produced by these materials vary with the starting products. Effective monitoring and control of the beetle pest require a standardised lure to enable accurate comparison between capture rate across time and location. This research has progressed the development of such a lure, determining many of the components that such a blend will contain. An optimised synthetic blend of attractive chemicals will provide such a lure, and such an optimisation will be a vital part of future research.

Laboratory studies alone are insufficient to validate the true effectiveness of such an attractant. Vagaries of weather (temperature, moisture, wind), floral availability and the specific layout of an apiary will all impact on the efficacy of a trap. Trap design, odour release mechanism and killing agent are all areas that would benefit from further research. The entire trapping system must be considered when determining timing and placement of deployed traps.

The current study has demonstrated aggregative behaviour, but has not yet determined the nature of an aggregation pheromone. An assessment as to whether further investigations into an aggregation pheromone are viable will depend upon a cost: benefit analysis as to whether or not this level of investment is appropriate in further SHB research. This study has provided invaluable recommendations as to the timing of SHB trapping with a natural fermentation based attractant solution. We have provided suggestions of when it is appropriate to deploy traps, and when trap use will be ineffective. Use of these traps for approximately 12 months resulted in the interception and trapping of nearly 4,000 beetles. The educational video prepared as part of this project will allow beekeepers to prepare their own SHB traps to manage this pest: [https://www.youtube.com/watch?v=YHUmK5SlzXU&t]

Recommendations

The following recommendations are targeted at beekeepers, decision-makers and researchers:

 This project demonstrated that commercially available lantern traps with a simple yeast based attractant can be deployed strategically from spring to autumn to intercept and trap SHB flying towards an apiary. An educational video on how to prepare and deploy this trap has been produced to disseminate the information to beekeepers: [https://www.youtube.com/watch?v=YHUmK5SlzXU&t]

 Seasonal data on the weekly and fortnightly changes in numbers of SHB trapped in the field and in hives suggests these changes are primarily influenced by temperature with rainfall as an important influence through the hot seasons. This information is the most comprehensive Australian ecological data on SHB generated yet and can inform strategic trapping programs using the simple yeast traps currently available or a customised SHB trap with synthetic attractant lure once developed. Furthermore, it has been demonstrated that traps placed anywhere up to 200 m from hives can be effective at intercepting SHB flying towards an apiary. Further research is needed to investigate distances greater than 200 m from hives and gather more data on the effect of direction and the effect of prevailing winds on trap efficacy.

 Further funding is required to continue research to refine and test the blend of compounds attractive to SHB to produce a synthetic lure. Research also needs to be undertaken to produce a release mechanism to optimise the release rate of the components of the synthetic lure with additional research warranted to customise a trap that incorporates a killing system for SHB and the synthetic lure.

Introduction

The Australian honey bee industry contributes at least $101 million annually to the Australian economy through honey products and an estimated $1.7 billion annually through pollination services to crops and horticulture (ABARES, 2014-2015 data). The small hive beetle (SHB) Aethina tumida Murray (Coleoptera: Nitidulidae) is a pest of European honey bees, Apis mellifera L. (Hymenoptera: Apidae) (Murray, 1867). The beetle is native to sub-Saharan Africa where it is a minor pest of little economic importance restricted to infesting weak, stressed, or diseased African honey bee colonies (Lundie, 1940, Neumann and Elzen, 2004a, Ellis and Hepburn, 2006). Outside of its native Africa, SHB has proven to be far more destructive. SHB was officially recorded in Florida, USA in 1998 (Elzen et al., 1999). By 2004, SHB had spread from the south east of the USA to 30 states and was estimated to have caused US $3 million in losses to the beekeeping industry (Hood, 2004).

The first report of SHB in Australia was from an apiary in Richmond, New South Wales (Somerville, 2003). By 2011, SHB had spread along the east coast of Australia, to Mareeba in the north and the Melbourne CBD in the south (Lamb and Leemon, 2011). Adult SHB have been reported to travel kilometres to locate a honey bee hive, by using olfactory cues (Somerville, 2003, Hood, 2004). They are opportunistic scavengers that take advantage of situations that suit their reproductive strategy (Somerville, 2003). The SHB larval stage can cause extensive damage to hives and stored comb as they feed on bee brood and pollen stores and leave behind waste. Fermentation of honey in the comb has been associated with the presence of large numbers of SHB larvae and a yeast (Kodamaea ohmeri), which has been isolated from the fermenting combs and, in the laboratory, from all of the small hive beetle’s life stages (Torto et al., 2007c, Benda et al., 2008, Leemon, 2012). The resulting fermented honey is rejected by honey bees and cannot be marketed by the beekeeper. Heavy larval infestations may also result in total hive collapse after the queen ceases to lay eggs and the bees abscond from their hive (Hepburn and Radloff, 1998, Hood, 2004).

Small hive beetle is now the predominant apiary pest in warm, moist regions of eastern Australia. Warm temperatures and wet summers provide ideal conditions for populations of this beetle to increase, with several generations possible between spring and autumn. Under the right environmental conditions during hot and humid weather, large numbers of adult beetles can initiate a mass oviposition in a hive. Although bees can remove small numbers of SHB larvae from a hive they are unable to manage the resulting large number of emerging larvae. Total hive collapse occurs after these larvae feed voraciously on the hive protein stores of pollen and brood, and cause the fermentation of the honeycomb into what is colloquially and expressively termed “slime”. Usually all that is left of a once flourishing hive is a number of dead bees surrounded by a mass of gorging SHB larvae in an odorous fermented slime (Leemon, pers. obs., 2007).

The only recent data available on the extent of damage caused by SHB was collected through voluntary surveys of registered Queensland beekeepers between 2008 and 2016 (Leemon et al. unpublished, 2017). Estimates of damage were calculated only on the losses reported by the survey respondents, with an average response rate to the surveys of 44%. A conservative estimate of annual hive losses during this period (not including nucleus hive losses) peaked at $5.5 million during the 2010-2011 summer. Although the losses decreased in the following years, the losses for 2015-2016 were still estimated at just under $1 million. Moreover, in the last five years, there has been a steady uptake of various methods of SHB control, and the spring leading into the 2015-2016 summer was unusually dry and thus likely to slow SHB population increases.

Small hive beetle also appear to be able to breed in and destroy the hives of native stingless bees (Heard, 2016). Anecdotal evidence suggests that the unmanaged/feral bees

responsible for much of the “free” pollination services are disappearing in areas of heavy beetle infestation.

A range of in-hive control options are now being deployed by many beekeepers, most of which rely on the bees chasing SHB into a harbourage trap. These vary greatly in trap design and the method of lure presentation. Trapping systems currently in use for adult SHB are all in-hive and include Apithor™ (Levot and Somerville, 2012), modified bottom boards and trench traps among others based on beetle behaviour inside a honey bee colony (Elzen et al., 2002). However, there is a need for additional out-of-hive control measures such as an external attractant trap that can stop SHB entering hives. An external attractant trap would also be useful for biosecurity as a surveillance tool for monitoring SHB in areas of uncertain infestation or to demonstrate area freedom in areas where SHB are not yet known to occur.

Olfactometer and wind tunnel investigations have shown that adult beetles are attracted to a range of hive odours, including the odours of adult worker bees, fermenting hive products, and a honey bee alarm pheromone (Suazo et al., 2003, Torto et al., 2005, Nolan and Hood, 2008). The fermentation of honey has been noted to occur in association with the development of larval SHB and the fermentation odours have been found to be especially attractive to adult SHB (Suazo et al., 2003, Torto et al., 2005, Nolan and Hood, 2008, Hayes et al., 2015). Torto et al. (2007c) and Arbogast et al. (2009) reported the presence of the yeast Kodamaea ohmeri in fermented hive products. K. ohmeri has been isolated from all SHB life stages and it dominates in the slime associated with SHB (Benda et al., 2008, Leemon, 2012). A lure using a strain of this yeast and pollen dough has been patented as an attractant for use in an in-hive trap and investigated for out-of-hive traps (Torto et al., 2007b, Arbogast et al., 2009, Teal et al., 2011). Hayes et al. (2015) demonstrated how these fermentation odours became more complex and attractive to adult SHB over time. This research suggested that some of the component odours of the fermenting hive products would be an ideal place to search for specific compounds highly attractive to adult SHB which can be blended to produce a SHB lure for use in an external trapping system.

An external trap must rely on SHB flying behaviour and have a lure strong enough to at least intercept the SHB emerging from pupation in nearby soil or entering the apiary, as SHB are highly attracted to odours emitted from honey bee hives (Arbogast et al., 2007, Teal et al., 2007). An attractive lure for the orchard pest beetles, Carpophilus spp. (which belong to the same Nitidulid family of insects as the SHB) is based on a synthetic aggregation pheromone working synergistically with a food co-attractant (Bartelt et al., 1992, Hossain et al., 2006, Bartelt and Hossain, 2010). This system is commercially available as the Catcha® Trapping System (Bugs for Bugs©). During its development beginning in the 1990s, a number of factors affected the performance of the trap in the field such as trap type and trap placement, including height above the ground and lure formulation (Bartelt and Hossain, 2010). The system uses Magnet™ funnel traps containing the pheromone impregnated septum and food blend to lure the Carpophilus beetles and an insecticide (Dichlorvos) to kill the insects once inside the trap (Hossain et al., 2007).

Hayes et al. (2015) demonstrated that a number of mixed-sex adult SHB were attracted to traps baited with the slime associated with SHB larval growth and that the attractiveness increased through time as the slime aged and fermentation developed. This study provided strong evidence that SHB find the volatiles in the slime highly attractive and that the volatile profile becomes more attractive with time. In the study, there were no visual or tactile cues to distinguish between traps, and very few beetles made a random choice to enter a trap without odour attractant (7 out of 520 = 1.35% across all reps in all trials), suggesting that the choice to enter a trap was directed by odour. A key driver in the change of attractiveness through time was likely to have been ethanol, a well-known attractant compound for a variety of insects from many different orders (Montgomery, 1983, Byers, 1992) including other members of the Nitidulidae family (Bartelt and Hossain, 2006). The levels of ethanol

increased significantly through time, and may well explain at least the early increases in attraction. As the ethanol levels reached a plateau around day 12 of larval development, further increases in attractiveness were likely to have been as a result of the increasing production of other fermentation products. Most of the compounds detected were typical compounds produced by yeast fermentation including a variety of alcohols, aldehydes and acetate esters. These types of fermentation products have been found to be highly attractive to Carpophilus davidsoni (Bartelt and Hossain, 2006). Many of these compounds (e.g. ethanol, isobutanol, isopentanol, isopentyl acetate) have previously been reported arising from either honey bees (Torto et al., 2005) or an artificial substance known as pollen dough, inoculated with the yeast K. ohmeri (Torto et al., 2007c). In both cases these compounds were shown to be attractive to SHB. Interestingly, one of the detected compounds was isopentyl acetate, a compound known since the 1960s to be a honey bee alarm pheromone (Boch et al., 1962). This substance has previously been reported to elicit an electrophysiological response from SHB (Torto et al., 2007c), and it could be a key cue for beetles searching for a stressed hive.

Fermenting hive products (slime) are clearly more attractive to SHB than the unfermented hive products. Honeycomb has previously been shown to be highly attractive to SHB (Suazo et al., 2003), therefore it may well be that in the field these products will also be more attractive to incoming adult SHB than the hive. However Hayes et al. (2015) demonstrated the inherent variation in the rate of production of these volatile mixes. They suggest that the volume and rate of emission of fermentation volatiles are affected by the level of protein and carbohydrate available in the hive for the initial SHB larval development; which in turn influences the development of the yeast driving the fermentation. These levels will vary throughout the year, and are likely influenced by the local flowering vegetation. In addition, the number of SHB larvae present as well as abiotic conditions such as temperature and humidity will also affect the development of the attractive volatiles. The enormous potential for variation in natural products will limit their use in attractant traps for SHB. However, suitable blends of synthetic compounds, based on selected fermentation volatiles show potential for a lure with minimal variation, suitable for deployment in an effective out-of-hive trap for this pest.

As demonstrated with the successful attract and kill Catcha® trapping system, aggregation pheromones combined with blends of key fermentation compounds can be synergistic and essential for maximising trap efficacy (Bartelt and James, 1994, Bartelt and Hossain, 2006, Hossain et al., 2008, Bartelt and Hossain, 2010). It is possible that SHB may produce an aggregation pheromone, thus further work in this area is warranted (Teal, pers. comm., 2014). Torto et al. (2007c) reported that they were not able to demonstrate whether SHB produce either sex or aggregation pheromones, however, they did not give any details of how they investigated this. Stuhl and Teal (pers. comm., 2014) described how SHB come together to form clusters and thought it highly likely that SHB use an aggregation pheromone. An investigation into the presence of an aggregation pheromone in small hive beetles is warranted because of the enormous potential for improvement of an attractant lure if SHB produce such a pheromone. Anecdotally, many apiarists have noted a huge variation in the numbers of beetles in hives within the one apiary and some have proposed a variation in the volatiles emitted by the hives, such as the bee alarm pheromone, as a reason for this. However, it may be possible that the large numbers of beetles in certain hives are due to beetles in those hives feeding on a preferred food substrate (e.g. pollen) giving off an aggregation pheromone. Thus the attractiveness of hives to SHB could be due to both the status of the available food in the hive as well as a pheromone signal to other SHB.

The aim of this project was to develop an external attractant trap for SHB based on the attractive odours associated with fermenting hive products and search for an aggregation pheromone that could be combined with the fermentation odours to enhance trap efficacy.

Objectives

The overall aim of the project is the development of an external trap for the small hive beetle, to assist commercial and hobbyist beekeepers in the monitoring, management and control of this apiary pest. The specific objectives are:

1. To determine which individual component compounds from fermenting hive products are attractive to small hive beetle (laboratory)

2. To determine the optimum blend of the above compounds, in terms of attractiveness and longevity (laboratory)

3. To develop a suitable attractant lure (using the synthetic blend identified above) for use in an external trap for the small hive beetle (laboratory)

4. To investigate the occurrence of an aggregation pheromone in the small hive beetle, the addition of which would enhance trap success (laboratory / field)

5. To examine differences in hive volatile profiles between hives carrying very high and low numbers of SHB (field/laboratory)

6. Preliminary investigation to find suitable trap design for field evaluation of the lure (laboratory/field)

7. To evaluate the efficacy of the external SHB trap in apiaries located in a variety of locations in NSW and Queensland, over two summer seasons (field).

8. To produce an educational video suitable for explaining the functioning of the lure and traps to the commercial beekeeping industry and hobbyist beekeepers.

Methodology

2.1 Colony

Small hive beetles used in assays were cultured in an insectary at the Ecosciences Precinct, Dutton Park, Queensland, using the method of Cribb et al. (2013). Larvae were reared on a diet of honeycomb and brood comb in plastic bags and pupated in a bed of moist sand in complete darkness. Adults were kept in ventilated containers with sugar and moist paper toweling under a light regime of 12:12 L/D. The insectary was maintained at 27°C and 65% relative humidity. Field-collected adults were regularly added to the colony to preserve genetic diversity. Rearing of individual, unmated beetles was based on Neumann et al. (2013). Final instar larvae were retrieved and placed in vials (5 ml) filled with sandy soil (7 g) (moistened to 5% w/w) and incubated in darkness until an adult was visible above the soil whereupon they were sexed. Beetles were sexed by gently squeezing the abdomen to cause the eversion of the ovipositor of females and protrude the tergite in males. 2.2 Location of field studies

Studies were carried out in three apiaries, two in Queensland at Bellbowrie and Gumdale and one in New South Wales at Richmond. The Bellbowrie apiary (27.55°S, 152.89°E) had six hives and was the primary apiary used by the project to supply hive materials for the SHB colony and production of slime for assays and trials unless stated otherwise. The Gumdale apiary (27.49°S, 153.16°E) owned by commercial beekeepers had 25 hives. The Richmond apiary (33.60°S, 150.75°E) was the Wheen Bee Foundation research apiary with 20 hives. Other sites were used as non-apiary sites paired with apiary sites for some of the trapping trials, these included Chapel Hill (27.52°S, 152.96°E) paired with Bellbowrie and Carina Heights (27.51°S, 153.09°E) paired with Gumdale. 2.3 Fermentation of hive products

This study investigated the odours emanating from fermenting hive products (slime) over time. Production of slime was standardised in the laboratory for use in assays. Pieces of honeycomb with a combined weight of 300 g and pieces of brood comb with a combined weight of 150 g were placed in an opaque plastic bag (190 × 190 mm) with a 100 mm flap (Figure 1). Approximately 3 ml of tap water was sprayed into the bag using an atomizer to promote slime development. The bag was placed in a transparent plastic container (190 × 190 mm), and 40 unsexed, sexually mature SHB adults were added. The containers were sealed with a perforated plastic lid and incubated at 27 °C and 65% relative humidity in darkness. The bag contents were sprayed with water three times per week.

Figure 1:SHB larvae converting honeycomb and brood (A) to slime (B). 2.3.1 Geographical variation in source of hive materials

The chemical composition of hive products (honey, pollen and wax) is assumed to vary with floral source throughout Australia. To investigate whether the differences in the starting products influence the attractiveness and volatile compounds produced during slime production, we obtained hive products from across the geographical range of the beetle. Two frames each of honeycomb and brood comb were sourced from each of five apiaries in Queensland and three apiaries in New South Wales (Figure 2). The frames of honeycomb and brood comb were transported overnight to the Ecosciences Precinct and stored at approximately 5°C. Hive products were allowed to reach room temperature before use in assays. The slime (fermenting hive products) from each of the eight different locations was sampled on days 18, 35 and 49 after introduction of the beetles for chemical analysis and use in behavioural assays. It was noted when SHB found the slime samples highly attractive and when SHB did not find the slime samples attractive. The chemical profiles of the attractive and non-attractive samples were analysed and compared using multivariate statistical techniques so that individual compounds present when the slime is attractive to SHB could be identified.

2.3.2. Behavioural bioassays

The comparative attractiveness of the slime samples to adult SHB was assayed. In each assay, four beetle traps containing either honeycomb, brood comb, slime, or nothing (control) were placed in each of six replicate cage arenas (600 × 600 × 600 mm) made of white mesh and plastic (BugDorm insect tent 2120, Australian Entomological Supplies Pty. Ltd., Coorabell, NSW) (Figure 3). Beetle traps consisted of cylindrical plastic vials (108 × 44 mm) with 50 mm funnels made of fibre-glass insect-screen (1 × 0.5 mm pore size) inserted into their openings, allowing beetles entry but not exit easily. At the back of each trap was placed 5 ml of a test material, followed by a 30 mm2 piece of cotton wool, which was roughly mixed with the test material, and then a 150 mm2 piece of crumpled paper towel. Control traps contained cotton wool and paper towel only. Aluminium foil was wrapped around the bases and sides of the traps, to darken their interiors thus increasing their attractiveness to beetles. The traps were secured to the floor in the corners of the cages using Blu-Tack™ (Bostick Australia; Thomastown, Victoria) and were orientated diagonally with their openings facing towards the centre of the cage floor. The cages were placed in a controlled environment room (27°C, 65% RH, L/D 13:11). Forty mixed-sex adult A. tumida held temporarily in a 75 ml plastic vial were released into the centre of the cage floor at the start of the assay. After 19 hrs, traps were removed and the beetles in each were counted. Any beetles remaining outside the traps were recovered and their numbers recorded.

Figure 2: Map of hive material sources and trial locations.

2.3.3 Chemical analysis

Volatile compounds emanating into the air above samples (headspace) of the slime from each replicate apiary were analysed to examine changes in these volatiles as the slime aged. Laboratory air was pulled through a charcoal trap then over the sample of slime, honeycomb and brood comb (approximately 5 ml) in a glass Erlenmeyer flask (250 ml) at a flow rate of 250 ml/min for 24 hrs (Figure 4). After passing over the sample, the air passed through a thermal desorption tube preloaded with Tenax TA (35/60 mesh) (Markes International Ltd.).

Air samples were released from the tubes by means of thermal desorption using a TD-100 thermal desorption unit (Markes International Ltd.) and introduced into a gas chromatograph (GC) (Agilent 6890 Series) coupled to a mass spectrometer (MS) (Agilent 5975) and fitted with a silica capillary column (Agilent, model HP5-MS, 30 m × 250 µm internal diameter × 0.25 μm film thickness) (Figure 5). Data were acquired under the following GC conditions: carrier gas He at 51 cm/s, split ratio 13:1, transfer-line temperature 280°C, initial temperature 40°C, initial time 2 min; rate 10°C/min, final temperature 260°C, final time 6 min. The mass spectrometer was held at 230°C in the ion source with a scan rate of 3.89 scans/s. Peaks that were present in blank (control) samples were discarded from analysis in test samples. Tentative identities were assigned to peaks with respect to the National Institute of Standards and Technology (NIST) mass spectral library. Mass spectra of peaks from different samples with the same retention time were compared to ensure that the compounds were the same.

Figure 3: Behavioural choice assay system in BugDorm tents. (A) Beetle trap (B) cage arena system (C) replicated cage arenas in bioassay.

Figure 4: Volatile samples being collected for analysis from the air above fermenting slime in an Erlenmeyer flask.

Figure 5: Gas chromatograph and mass spectrometer for analysis of volatile compounds in air samples.

2.4 Individual compounds attractive to SHB

The ten compounds identified in the above investigations (Table 1) as most important in distinguishing between attractive and non-attractive slime were tested in single SHB choice assays under controlled laboratory conditions.

A suitable solvent was identified as 100% ethanol, given that all compounds are soluble in it and it is itself on the list of the attractive compounds. Following this, all compounds were tested in 2-choice behavioural bioassays against both a blank and control and ethanol. Previously used BugDorm bioassays (see section 2.3.2) were complemented by individual arena bioassays in order to discern and mitigate a potential aggregation factor in results.

Plastic containers (170 × 115 × 70 mm) were used as arenas for individual adult A. tumida. These had six holes punched in the lid (< 1 mm in diameter) and one hole (28 mm diameter) in each end to fit 20 ml (54 × 26 mm) plastic vials. These vials were fitted with 50 mm funnels made of fibre-glass insect-screen (1 × 0.5 mm pore size) inserted into their openings, allowing beetles entry but not exit. Aluminium foil was wrapped around the bases and sides of these traps, to darken their interiors thus increasing their attractiveness to beetles (Figure 6). All traps contained approx. 30 mm2 piece of crumpled paper towel. Treatment vials contained one of three concentrations of the compound (10%, 2%, 1%) in ethanol (500 µL) and control traps contained an empty plastic zip lock bag (40 × 50 mm) or ethanol (500 µL) in a zip lock bag which was then heat sealed. One adult SHB was placed in the middle of the arena, the lid replaced and incubated at 27°C, 70% RH for 18 hrs. All SHB used were previously unused in any bioassays and were ≤ 21 days since emergence from the soil. Each compound was tested in two choice assays against both a blank control and an ethanol control (30 replicates per concentration) (Figure 6).

Table 1: The ten most important compounds in distinguishing between attractive and non-attractive slime samples, including their known role in insect chemical ecology

Compound Class Compound Comment Alcohol 2,3-butanediol Known attractant for other beetles1,2 ethanol Common insect attractant, including other Nitidulidae3 Found in Catcha® system isobutanol Known attractant for other Nitidulidae4 Found in Catcha® system isopentanol Known attractant for A. tumida5 and other Nitidulidae4 Found in Catcha® system phenylethyl alcohol Known attractant for A. tumida5 and other Nitidulidae6 Ester ethyl acetate Known attractant for other Nitidulidae4 Found in Catcha® system isopentyl acetate Honey bee alarm pheromone7 Known attractant for A. tumida5 phenylethyl acetate Known attractant for other beetles8,9 Ketone acetoin Known attractant for other beetles1,2 acetone Known attractant for other beetles3 Carboxylic acid acetic acid Known attractant for other beetles3

1Rochat et al. (2002), 2Rochat et al. (2000), 3El-Sayed (2018), 4Bartelt and Hossain (2006), 5Torto et al. (2007a), 6Zilkowski et al. (1999), 7Boch et al. (1962), 8Yang et al. (2017), 9Shepherd and Sullivan (2013).

Figure 6: Single choice assay system for SHB volatile compound attractiveness testing. (A) individual arena (B) group of replicated arenas used in a bioassay.

A positive control was conducted for the individual arena bioassay design using day 13 slime roughly mixed with approximately 10 mm3 cotton wool. This was conducted in three temperature and humidity controlled locations which were henceforth used as concentration variables. The optimum treatment volume (compound in solvent) was determined by testing three volumes of ethanol (10 replicates of each volume).

A further positive control was conducted to determine the effect of an inducement to remain in the trap after a choice was made. Inaccessible slime, to provide a positive control attractive odour but not a potential food source, was tested against a control and accessible slime. Slime was rendered inaccessible by separating the cotton wool from the crumpled paper and entry tunnel by a piece of gauze. Following this, each compound and Blend #1 (Section 2.2.3.2) were tested again against an ethanol control with a food reward, this being the crumpled paper towel coated in a saturated sugar solution. 2.5 Synthetic lure development

It was noted that some of the individual compounds identified as attractive to SHB are also part of the Carpophilus Catcha® Trapping System (Bugs for Bugs©). Moreover, both SHB (Aethina tumida) and Carpophilus beetles are in the family Nitidulidae, therefore the Catcha® system was tested first for attractiveness to adult SHB. Later, ten different blends of the individual compounds identified above were developed and tested first in individual arenas then in BugDorm assays to select the best/optimal blend for testing in the field.

2.5.1. Carpophilus system evaluation

Both the Catcha® liquid component (parts A + B) (500 μL; proportions as per manufacturers’ instructions) and pheromone component (part C) (~ 0.015 g piece of pheromone- impregnated rubber septum - approximately 1/64 of whole septum) of the system were tested in individual arenas (50 replicates) and BugDorm assays (6 replicates). One component of the liquid (acetaldehyde) (part B) was used in subsequent blends, given assay results.

2.5.2 Blend development and testing

The compounds used in the ten blends are summarised in Table 2, and details of the proportions of compounds and how they were tested are outlined below.

Blend #1 was made up of all ten compounds in the proportion in which they appeared in attractive Day 18 slime, averaged across all sites: phenylethyl alcohol (28.01%), isopentanol (14.71%), phenylethyl acetate (10.98%), ethyl acetate (10.13%), acetoin (9.89%), 2,3- butanediol (5.60%), ethyl isobutyrate (7.40%), isobutanol (4.71%) and isopentyl acetate (3.92%) in ethanol solvent. This blend was tested at three different concentrations (100%, 1% and 0.1%) in the individual arena bioassay described above (50 replicates each concentration) and at two concentrations (100% and 0.1%) in the BugDorm bioassay (6 replicates) against a blank control.

Blend #2 was made up of the compounds which had statistically significant positive responses in the individual compound testing bioassays, and in the concentration at which this occurred excluding phenylethyl acetate: isobutanol (10%), ethyl acetate (2%), 2,3- butandiol (1%) and acetoin (1%) in ethanol solvent. This blend was tested in both the individual arena bioassay described above (84 replicates) and in the BugDorm bioassay (6 replicates) against a blank control. Testing of Blend #2 was repeated using cotton wicks (500 µL on 10 mm-long wick in individual arenas) (1 ml on 20 mm-long wick in bugdorm) instead of plastic zip lock bags (50 and 6 replicates respectively).

Blend #3 was made up of statistically significant positive responses in the individual compound testing bioassays, and in the concentration at which this occurred: isobutanol (10%), phenylethyl acetate (10%), ethyl acetate (2%), 2,3-butandiol (1%) and acetoin (1%) in ethanol solvent. This blend was tested in both the individual arena bioassay described above (50 replicates) and in the BugDorm bioassay (6 replicates) against a blank control using the zip-lock bag methodology outlined above.

Blend #4 was made up of statistically significant positive responses in individual compound testing bioassays with a food reward as well as important compounds identified by colleagues at the United States Department of Agriculture (USDA) (Teal, pers. comm., 2014). It contained ethyl acetate (45%), ethanol (20.6%), isopentyl acetate (17.4%), ethyl caproate (10.7%), nonanal (4.8%), decanal (1.4%) and acetaldehyde (0.1%). This blend was tested at four different concentrations (100%, 1%, 0.1%, 0.05%) in the individual arena bioassay described above (50 replicates each concentration) and at three concentrations (100%, 1%, 0.1%) in the BugDorm bioassay (6 replicates each concentration) against a blank control using the zip-lock bag methodology outlined above. A three-choice test in the BugDorm bioassay compared blank against two concentrations (100% and 1%) of the blend (6 replicates). A modified version of this blend (blend #4a) was developed with a higher concentration of acetaldehyde (1%) and a compensatory reduction in the concentration of ethanol. A three-choice test in the BugDorm bioassay compared blank against two concentrations (100% and 1%) of the blend (6 replicates).

Blend #5 was based on volatile proportions detected in a fermentate generated from yeast and honey fermentation (see 2.9.1). It contained ethanol (55.7%), isopentanol (14.5%), isobutanol (13.3%), 2-methylbutanol (12%), ethyl acetate (2%), propanol (1.5%) and acetic acid (1%). A three-choice test in the BugDorm bioassay compared blank against two concentrations (100% and 1%) of the blend (6 replicates).

Blend #6 was a simplified version of blend #5, focusing only on the major components. It contained ethanol (55%), isopentanol (15%), isobutanol (15%) and 2-methylbutanol (15%). This blend was tested at two different concentrations (100%, 1%) in the individual arena bioassay described above (50 replicates each concentration). A three-choice test in the BugDorm bioassay compared blank against two concentrations (100% and 1%) of the blend (6 replicates).

Blend #7 was based on blend #5 with the addition of acetaldehyde, which appears to be an important component of the commercially available Carpophilus system (see 2.5.1). It

contained ethanol (55%), isopentanol (14.5%), isobutanol (13%), 2-methylbutanol (12%), ethyl acetate (2%), propanol (1.5%), acetic acid (1%) and acetaldehyde (1%). A three-choice test in the BugDorm bioassay compared blank against two concentrations (100% and 1%) of the blend (6 replicates).

Blend #8 was based on blend #6 with the addition of acetaldehyde, which appears to be an important component of the commercially available Carpophilus system (see 2.5.1). It contained ethanol (55%), isopentanol (15%), isobutanol (15%), 2-methylbutanol (14%) and acetaldehyde (1%). A three-choice test in the BugDorm bioassay compared blank against two concentrations (100% and 1%) of the blend (6 replicates).

Blend #9 was based on volatile proportions detected in a fermentate generated from yeast and sucrose fermentation (see 2.9.1). It contained ethanol (91%), isobutanol (4%), isopentanol (3%) and ethyl acetate (2%). A three-choice test in the BugDorm bioassay compared blank against two concentrations (100% and 1%) of the blend (6 replicates).

Table 2: Components of blends tested in laboratory assays and in field trial

Blend # 1 2 3 4 5 6 7 8 9 10 Comp’d ethanol Y Y Y Y Y Y Y Y Y Y ethyl acetate Y Y Y Y Y Y Y Y isopentanol Y Y Y Y Y Y Y isobutanol Y Y Y Y Y Y Y Y Y isopentyl acetate Y Y phenylethyl alcohol Y phenylethyl Y Y acetate acetoin Y Y Y 2,3-butanediol Y Y Y ethyl isobutyrate Y acetaldehyde Y Y Y Y ethyl caproate Y nonanal Y decanal Y propanol Y Y acetic acid Y Y 2-methylbutanol Y Y Y Y

Blend #10 was based on blend #9 with the addition of acetaldehyde, which appears to be an important component of the commercially available Carpophilus system. It contained ethanol (90%), isobutanol (4%), isopentanol (3%), ethyl acetate (2%) and acetaldehyde (1%). A three-choice test in the BugDorm bioassay compared blank ethanol and 100% blend #10 (6 replicates). A three-choice test in the BugDorm bioassay compared blank against two concentrations (100% and 1%) of the blend (6 replicates).

A matrix of two-and three-choice bioassay tests in the BugDorm compared the attractiveness of all possible combinations of blends #4, #6, #8 and #10 (6 replicates of each combination). 2.6 Aggregation pheromone

Behavioural assays were used to first assess whether SHB would aggregate and the conditions under which they aggregated. Later air samples were collected above different groups of SHB and analysed with GC-MS to search for unique compounds that may indicate an aggregation pheromone.

2.6.1 Arena aggregation trials

A series of bioassays were conducted to investigate and quantify aggregative behaviour in SHB (Table 3). In each bioassay, 16 SHB (single or mixed sex) were sedated with CO2 and placed in a 15 cm diameter Petri dish (the arena) divided into eight sectors via a marked template underneath. Two SHB were placed in each sector, but were then free to move around the arena, during which time they were under observation to assess aggregative behaviour. The arena was lit by two overhead lamps to eliminate shadows and filmed, with the distribution of SHB recorded every 30 seconds for 15 min. Aggregative groups were defined as beetles being within one beetle length (approx. 6 mm) of each other and a cluster was defined as a group of at least five beetles.

Five different conditions for clustering that might indicate aggregative behaviour were investigated with mixed sex and single sex SHB (Table 3). For all single sex investigations males and females were separated from the opposite sex within two days of emergence from the soil and kept isolated until needed for a bioassay.

The initial investigation looked for clustering behaviour in either a single sex group of males, a single sex group of females or a mixed sex group with both males and females.

The second investigation examined whether males or females instigated the “clustering” behaviour. Groups of eight adult male SHB were shaken briefly in a small amount of powdered iridescent dye, enough to colour their carapace but not to impede movement, and placed in an arena with eight unmarked females. One marked male and one unmarked female were placed in each sector. This bioassay was then repeated with marked females and unmarked males.

Table 3: Groups of SHB investigated for clustering indicative of aggregative behaviour in arena assays, including number and type of SHB used and number of replicates for each investigation.

Treatment Number and type of beetle per Replicates (n) group of each assay

16 male SHB 3 Untreated (separated from 16 female SHB 3 opposite sex within 2 days of emergence from soil) 16 mixed sex SHB (males and 3 females together)

8 marked male SHB + 8 2 unmarked female SHB Marked using iridescent dye 8 marked female SHB + 8 2 unmarked male SHB

8 male SHB + 8 A. diaperinus 3 With darkling beetles (Alphitobius diaperinus) 8 female SHB + 8 A. diaperinus 3

8 live unmated female SHB + 8 10 dead colony* mixed sex SHB With dead SHB 8 live colony* female SHB + 8 10 dead colony* mixed sex SHB 16 male colony* SHB with 9 antennae removed After removal of antennae 16 female colony* SHB with 9 antennae removed

(* colony beetles consisted of male and female SHB maintained together, their “mated” status was unknown)

The third investigation explored whether clustering behaviour was guided by visual clues alone. Single sex groups of eight males or eight female SHB were placed in the arenas with eight mixed sex darkling beetles (Alphitobius diaperinus), with one beetle of each species per sector. A. diaperinus were chosen because they are of a similar size and colouring to SHB.

The fourth investigation studied whether SHB would still include other dead SHB (conspecifics) in any clusters. A group of mixed sex SHB were first euthanised by exposure to -20 °C, then stored at this temperature until use. One dead SHB plus either one unmated female or one female from a mixed colony were placed in each sector.

The fifth investigation tested whether clustering would still occur in SHB with impaired sensory detection. A group of males and a group of females had their antennae removed (antennectomised) and were kept at 27 °C and 65% RH for 3hrs prior to use in bioassays. Then two SHB were placed in each sector for each group.

The clustering investigations are summarised in the Table 3 along with the number of SHB used and the number of times each investigation was replicated.

2.6.2 Chemistry of aggregation

To investigate whether unique volatiles are produced in association with observed aggregative behaviour, air samples were collected from groups of SHB (20 per group) and analysed (Tables 4, 5). SHB were placed in an Erlenmeyer flask (250 ml) and aerated at a flow rate of 250 ml/min for 24 hrs. After passing over the insects, the air passed through a thermal desorption tube preloaded with Tenax TA (35/60 mesh) (Markes International Ltd.) (Figure 7) for analysis using GCMS (conditions as per Section 2.3.3). Various treatments of SHB were tested based on conditions previously noted as required for pheromone production and emission in other insects such as Carpophilus spp. (Bartelt and Hossain, 2010). Beetles ≤ 4 weeks since emergence from soil are referred to as young; unmated refers to individually raised beetles that have had no contact with SHB of the opposite sex; sucrose refers to saturated sucrose solution on paper towel (≈ 30 mm2); protein feed refers to Bee Build Protein Sausage™ (produced by L. & P. Dewar, Kalbar QLD); starved refers to 24 hrs without access to food. Control aerations were also collected for each treatment. SHB were allowed to feed for 24 hrs before aerations.

Table 4: Aerations of groups of SHB of different sex and mating status with 20 SHB per group, the number of replications of each assay is also shown.

Assay Treatment Sex of beetles Number of Replicates

A Male 2 “Young” colony SHB B Female 4 C Male 4 “Old” colony SHB D Female 6 E Male 9 “Young” unmated SHB F Female 9 “Young” unmated SHB with Male 5 G sucrose

Table 5: Aerations of groups of “young” SHB of different sex, mating status and feeding status with 20 SHB per group, the number of replications of each assay is also shown.

Assay Sex of Treatment Feeding status n beetles H before protein feed Male 5 Unmated I after protein feed Male 5

J before protein feed Female 9 Unmated K after protein feed Female 9

L before protein feed Male & 1 Unmated M after protein feed Female N before protein feed Male 5 Unmated after protein feed 5 O Male and water P before protein feed Female 5 Unmated after protein feed 5 Q and water Female

after protein feed 2 R Male and water Unmated, starved after protein feed 5 S and water Female

Figure 7: Aeration of (clustered) groups of SHB with saturated sucrose solution on crumpled paper provided for energy.

2.7 Volatile profiles of hives

To investigate whether the volatile profile of a hive correlated with SHB burden, aerations were collected from European honey bee hives. Samples of volatiles from four hives in the Bellbowrie apiary were collected onto thermal desorption tubes preloaded with Tenax TA (35/60 mesh) (Markes International Ltd.) using a vacuum pump to suck the air from inside the entrance to the brood box (sampling time = one hour, flow rate approx. 500 ml/min) (Figure 8). Samples were collected from each hive on four separate occasions (8th April 2016, 24th April 2016, 10th May 2016 and 24th May 2016). Aerations were also conducted on four hives from the Wheen Bee Foundation Research Apiary (on 17th May 2016).

Samples were thermally desorbed from the tubes as described in section 2.3.3. Differences between samples were analysed using multivariate statistical analyses.

Figure 8: Hive aeration sampling, at the Richmond (A) and Bellbowrie (B) field sites. Note the Tenax sample tube attached by plastic tubing to an air pump is inserted into the hive entrance.

2.8 Trap design

A range of trap designs was evaluated in the laboratory and in the field for SHB trapping. In lieu of a synthetic lure that was still in development, two different “natural” attractants were deployed in the traps. Slime was used as the attractant in most of the initial investigations, except for one trial using a synthetic blend (blend #2). Later a simple fermentate produced by the fermentation of honey and sugar with baker’s yeast was adopted to provide better consistency than the slime (section 2.9.1). The Unitrap™ (Figure 9A) was first evaluated because of its functional similarity to the Magnet™ funnel trap successfully used in the Catcha® system. After some initial, though limited, success field trapping SHB with Unitraps™, the research moved to using the Dowd traps successfully used for trapping Carpophilus in the US (Bartelt and Hossain, 2010). This particular design allowed the trap to orient with wind direction to optimise dispersal of the attractant (Figure 9B). Dowd traps were built to the specifications outlined in Dowd et al. (1992). Later, simple lantern traps (Figure 9C), readily available through many retail outlets as flytraps, baited with fermentate were trialled. Other in-house trap designs were considered, although preliminary investigations showed these to be impractical.

Figure 9: (A) Unitrap™ baited with slime, (B) Dowd trap baited with slime and (C) lantern trap with honey fermentate.

2.8.1 Trap evaluations

Ten trials to evaluate different trap designs were conducted from December 2014 to November 2016 before the lantern trap was successfully adopted to provide data on field movement of SHB (Table 6). Traps were set out in a range of locations to include sites near apiaries and sites away from any known apiaries. The first trial also included three sites near native stingless bee hives (Tetragonula carbonaria).

Table 6: Different trap designs and attractants tested for SHB trapping near apiary and in non-apiary sites between December 2014 and December 2016.

Trial Time Trap tested and Locations attractant

1 19/12/14 - Unitrap™ Apiary (2 sites), native stingless 6/1/15 hives (3 sites) and non-apiary (3 Slime & control sites) around Brisbane 2 10/2/15 – Unitrap™ Apiary (3 sites) and non-apiary(3 26/2/15 sites) around Brisbane Slime & control 3 13/3/15 – Unitrap™ Apiary(3 sites) and non-apiary (5 30/3/15 sites) around Brisbane Slime & control 4 7/10/15 – Unitrap™ Bellbowrie apiary 3/11/15 Slime & control 4 distances from hives 5 13/11/15 – Unitrap™ Bellbowrie apiary 27/11/15 Blend #2 & control 6 19/12/15 – Unitrap™ Apiary (1 site) and non-apiary (2 1/1/16 sites) around Brisbane Slime, Catcha® attractant & control 7 Feb 2016 Unitrap™, Dowd traps, Apiary (2 sites) and non-apiary (2 2 “in house” design sites) around Brisbane traps Slime , yeasts, honeycomb & control 8 30/8//16 – Dowd traps Gumdale apiary 20/12/16 Slime replaced every fortnight & control 9 31/8/2016 – Dowd traps Bellbowrie apiary 22/11/16 Slime replaced every fortnight & control 10 5/9/16 – Dowd traps Richmond apiary 29/11/16 Slime replaced every fortnight & control 11 10/11/16 Lantern traps Bellbowrie and Gumdale apiaries onwards (see Table 8)

The Unitraps™ and Dowd traps contained fermented hive products as the attractant (18 days old at deployment), that was not replaced for the duration of trapping, unless otherwise noted. Unitraps™ also used Fipronil-treated paper as the killing agent (SureFire Vista™ 200SC, 1.5 ml/L). The design of the Dowd trap obviated the need for a killing agent. The lantern traps contained fermentate (fermenting honey water and yeast, see section 2.9.1) which served as both the attractant and killing agent as most SHB drowned in this liquid.

The slime baits were prepared from fermented hive products of approximately 18 days old when the slime is known to be highly attractive. In each trial the slime material was placed in vials which were sealed until deployment, at which time gauze lids replaced the sealed lids to allow volatiles to escape and contain any SHB larvae in the slime. It was noted that having active larvae in the slime helped the slime attractant last longer. For the Unitraps™ approximately 60 cm lengths of paper towel were wetted with the killing agent. Once dry the paper was scrunched and placed around the outer wall of the trap in a circular fashion providing a harbourage for incoming insects. The lures were placed in the middle of the treated paper and the trap lids were secured. Control Unitraps™ contained only insecticide- treated paper. All traps were labelled and placed at their various locations in the shade (where possible) and approximately 1-1.5 m above the ground or at a similar height to the hive entrance. Slime baits were wetted with water at least once a week. Traps were deployed for approximately 14 days and upon retrieval the contents were counted and recorded. 2.9 Evaluation of efficacy of synthetic lure and trap

Successfully testing a lure and trap in the field requires not only a suitable trap design and odour release mechanism but knowledge of when and where to deploy the traps and a method to assess the efficacy of the traps. Although there has been information published on the biology of SHB (Annand, 2011, Neumann et al., 2013) there is very little information on the ecology of this pest; mostly the effect of temperature on different SHB stages and effect of soil moisture on SHB pupation and emergence (de Guzman and Frake, 2007, Meikle and Patt, 2011, Neumann et al., 2013). Annand (2011) presented some limited ecological data on seasonal movement of SHB. This study developed a simple reliable trapping system using a natural fermentation based attractant with a commercially available trap to record SHB movement over a year in three different apiary sites. Traps were also set out to provide data on the effect of distance from hives on trapping efficacy. In addition, a simple relative measure of SHB load in hives was investigated so that population changes in hives could be estimated.

2.9.1 Natural fermentate attractant investigations

While a synthetic lure was being developed and tested in the laboratory a more consistent fermentation odour source to slime was also developed for use in trap testing and gathering field trapping data. The alternative odour source was based on a natural fermentate of commercially available, dehydrated bakers’ yeast (Saccharomyces cerevisiae) (Lowan® Whole Foods Instant Dried Yeast), honey and water and/or sucrose. Various mixes were tested in a series of laboratory behavioural bioassays and chemically analysed to choose the best fermentate for use in the field.

2.9.1.1 Behavioural bioassays

The attractiveness of honey-based fermentates produced from different ratios of honey to water was compared in behavioural choice bioassays with adult SHB. Later the different fermentates were compared to slime in the larger Bug Dorm arenas. A constant amount of yeast (1.5% of final blend by mass) was added to each of three ratios of water to honey by mass (75:25, 60:40, 50:50). The mixtures were allowed to ferment at 27 °C before use.

Individual arena assays were conducted as per Section 2.2.3.1, except 1 ml of the fermentate was applied to cotton wool per vial and 50 replicates were used; Bugdorm arena assays were conducted as per section 2.3.2, except 2 ml of fermentate was applied to cotton wool per vial.

The relative attractiveness of fermentates from a 75:25 honey water mix at different ages (0, 1, 2, 3, 4, 5, 6 and 7 days old) was tested in behavioural choice assays in individual arenas (as above) with adult SHB.

The attractiveness of five-day-old fermentates from either a 75:25 ratio of water and honey or water and sucrose was assessed in individual arenas (as above) and large Bug Dorm arenas (as above).

The comparative attractiveness of the honey and sugar fermentates to SHB in the field was assessed in the Bellbowrie apiary. Pairs of traps with the two fermentates were hung in the shade approximately one metre apart and approximately 1.5 m from the ground at four locations (10, 40, 100 or 200 m from the hives). Traps were deployed for six weeks and collected every one to three days to remove and count SHB. The fermentates were replenished every three days.

A fermentate based on a combination of sucrose and honey was assessed using lantern traps (Envirosafe fly trapTM) in the large Bug Dorm arenas (section 2.3.2) and in larger tent arena assays. In large tent assays, two lantern traps containing either 290 ml of fresh blend (250 ml tap water / 20 g honey / 40 g granulated sucrose / 5 g yeast) or nothing (control) were placed in diagonally opposite corners in each of four replicate cage arenas (2050 × 1500 × 1030 mm) made of white mesh and plastic (Speed Mosquito Net, Equip Health Systems Pty LTD, O’Conner, Western Australia). Aluminium foil was wrapped around the bases and sides of the traps, to darken their interiors thus increasing their attractiveness to beetles. The cages were placed in a controlled environment room (27 °C, 65% RH, L/D 13:11). Fifty mixed sex adult A. tumida, held temporarily in a 75 ml plastic vial were actively released into the centre of the cage floor at the start of the assay. After 19 h, traps were removed and the beetles in each were counted. Any beetles remaining outside the traps were recovered and their numbers recorded. A subsequent assay was conducted as above, but where the traps were suspended on wire hooks approximately 800 mm from the floor of the cage.

2.9.1.2 Chemical analysis

Volatile components in the headspace above both a honey based fermentate and sugar fermentate were analysed as the fermentates aged. Up to four replicates of each type were incubated at 27°C and 65% RH and sampled by extracting air (500 ml) from above the blend onto a thermal desorption tube preloaded with Tenax TA (35/60 mesh) (Markes International Ltd.). Samples were taken at 0, 6, 24, 44, 74, 95, 144, and 172 h for the honey fermentate and at 0, 6, 24, and 120 h for the sugar fermentate. Analysis was conducted as per section 2.3.3.

2.9.2 Ecological studies of SHB

Studies were designed to gather ecological data on SHB population changes throughout the year in response to temperature and rainfall. In addition, triggers in soil moisture changes were investigated. A simple method of estimating SHB numbers in a hive was designed and tested.

2.9.2.1 Relative measure of SHB burden in hives

A simple relative measure of the number of SHB in hives was made using both trapping and visual assessments. Two simple traps were used. A vinyl mat with fibrous backing folded with the fibrous material to the interior was placed on top of the frames of the uppermost honey super, under the hive lid (referred to as “fibrous mat”) (Figure 10). Trench traps (AJs beetle eater™) filled with diatomaceous earth (DE) (Mt Sylvia absorbacide) were placed between the frames in each honey super (referred to as “DE traps”) (Figure 11). Each fortnight the number of SHB seen as the fibrous mat was peeled back was recorded along with the number of SHB trapped in the fibrous mat and the DE traps. Both the fibrous mat and DE traps were replaced each fortnight. The fibrous mats were placed in the freezer to kill any live SHB or bees trapped in the fibres before a SHB count was made. Likewise, the DE traps were placed in the freezer to kill any live SHB, after which the DE was sieved to recover and count the SHB.

Figure 10: Fibrous mat folded and placed on top of frames under the hive lid, with the fibrous side to the inside and vinyl to the outside. (A) Showing the positioning of the mat in the hive. (B) Mat removed from hive showing SHB trapped in the fibrous side. (C) Close up view showing how SHB become trapped in the mat when their legs get entangled in the fibres.

Figure 11: (A) Trench traps being removed from a hive as part of the fortnightly hive check assessments of SHB burden, (B) SHB caught in the diatomaceous earth (DE) used inside a trench trap removed from a hive.

Initially a folded disposable cleaning cloth (Chux™) was placed under the queen excluder on top of the brood box to trap SHB. This method was being promoted as a simple but effective method of trapping SHB by some beekeepers. The principle being that the bees attacked the cloth and “chewed” it into a fibrous tangle that trapped SHB trying to move through it. However, after four lots of hive checking (8 weeks) this method was discontinued as it

proved to be far too variable. While this method did trap large numbers of SHB where the cloth was well chewed into a fluffy and fibrous mass, not all hives reacted the same way. In some hives the bees thoroughly chewed up the cloth and removed it from the hive within a fortnight and other hives did not chew the cloth but coated the cloth with bands of wax and propolis that were ineffective for trapping SHB.

The relative measures were conducted in each of four hives in the apiaries in Bellbowrie, Gumdale and Richmond for varying periods between September 2016 and November 2017 (Table 7).

Table 7: Dates when fortnightly assessments of SHB numbers in hives (relative measure) and total hive counts of SHB (validation) conducted

Apiary Site Trapping dates Number of Weeks Validation fortnightly dates checks

17/08/16 - 08/06/17 22 44 17/08/16

18/01/17 Bellbowrie 11/05/17

18/07/17 - 22/11/17 10 22 11/10/17

30/08/16 - 09/05/17 19 38 11/04/17 Gumdale 30/08/17 - 14/11/17 6 12 14/11/17

05/09/16 - 29/11/016 7 14 -

Richmond 25/03/17 - 05/05/17 4 8 05/05/17

05/10/17 - 20/11/17 4.5 9 20/11/17

A validation of the relative measure of SHB in hives was carried out by counting all of the SHB in each of the trial hives, using a method modified from Annand (2011). In total eight validations were carried out; four in the Bellbowrie apiary; and two each in the Gumdale and Richmond apiaries. Validation was achieved by dismantling a hive, one box at a time and frame by frame and using a small vacuum pump to aspirate every visible SHB into a holding container (Figure 12). Once each frame was thoroughly inspected for SHB it was placed in a spare box placed on a lid to prevent escape of any undetected SHB. Once a box was emptied after inspection it was placed on top of the box now containing inspected frames and the process repeated for the next box until the brood box was completed. Particular care was taken inspecting the brood box to not dislocate the queen. Once the brood box was completed each frame was again inspected for SHB as it was returned to the hive. A different holding container was used for each hive. The holding containers were placed into a freezer to kill the SHB, so that they could be counted.

Figure 12: Aspiration of SHB from frame during a hive validation to assess total number of SHB in a hive.

2.9.2.2 Field movement of SHB

Paired control and treatment lantern traps were hung in the shade (where possible) at three different apiary sites (Table 8). Treatment traps contained an attractant mixture of honey (60 ml/3 tablespoons), water (250 ml/1 cup) and bakers’ yeast (2.83 g/ 1 teaspoon). The yeast was suspended in the water then the honey was mixed in until it dissolved. Control traps contained water (290 ml). The pairs were placed ideally 1-2 metres apart at different positions around the apiaries (Table 8). At Gumdale and Richmond traps were positioned at four different ordinal/directional points around each apiary at two different distances. It was not possible to achieve this configuration of traps at Bellbowrie, instead traps were hung at a greater variety of distances and at greater distances from the apiary than at Gumdale and Richmond (Table 8). In addition, two traps (attractant & control) were deployed at non- apiary sites paired with Bellbowrie (Chapel Hill) and Gumdale (Carina Heights) to assess the capacity of the traps to monitor SHB away from hives. The attractant mixture was regularly checked for SHB and replaced with a fresh mixture, every one to three days at Bellbowrie, twice weekly at Richmond and weekly at Gumdale.

Table 8: Details of traps deployed for collecting field data on SHB movement around three different apiaries (Bellbowrie, Gumdale and Richmond)

Location Trapping Dates Trapping Details 4 attractant traps + controls 11/11/16 – 22/12/16 Placed 6 – 90 m from hives, various directions 7 attractant traps + controls 23/12/16 – 26/03/17 Placed 6 – 200 m from hives, various directions Bellbowrie 8 attractant traps + controls 27/3/17 – 08/06/17 Placed 6 – 200 m from hives, various directions 8 attractant traps 08/07/17 – 22/11/17 Placed 6 – 200 m from hives, various directions 1 attractant trap + 1 control Chapel Hill 07/12/16 – 11/05/17 Non-apiary site paired with Bellbowrie 8 attractant traps + controls 29/11/16 – 09/05/17 4 traps next to hives, 4 traps 20 m from hives directions NW, NE, SE, SW from hives Gumdale 8 attractant traps 4 traps 20 m & 4 traps 100 m 30/08/17 – 14/11/17 from hives directions: NW, NE, SE, SW from hives 1 attractant trap + 1 control Carina Heights 07/12/16 – 02/05/17 Non-apiary site paired with Bellbowrie 8 attractant traps + controls 16/03/17 – 05/05/17 4 traps 20 m & 4 traps 100 m from hives directions: N, S, E, W from hives Richmond 8 attractant traps + controls 30/09/17 – 18/11/17 4 traps 20 m & 4 traps 100 m from hives directions: N, S, E, W from hives

2.9.2.3 Emergence trigger investigations

When trapping SHB in the field it was noted that there was often a surge in the number of SHB trapped after rain, particularly if the rain occurred after a spell of hot, dry weather. This suggests that rain may be a trigger for SHB to emerge from the soil and seek out hives. Therefore, the importance of rain as an emergence trigger for A. tumida was investigated in a series of laboratory assays. One hundred final instar larvae were rinsed in water and allowed to walk over damp paper towel to remove material from the cuticle. Individual larvae were placed in a plastic vial (15 × 97mm.) containing 13 g of soil that had been wetted to 5% moisture. The larvae were placed on top of the soil and allowed to burrow down to pupate. The vials were covered with fine gauze and lid with a single perforation (10mm dia.). Vials were then randomly allocated to four treatment groups (25 per group). Group A soil moisture was maintained at 5% for the duration of the assay. Group B soil moisture was maintained at 5% until day 18, when it was brought to 15% to simulate a rain event, and then unaltered for the remainder of the assay. Group C soil moisture was maintained at 5% until day 11, when it was allowed to dry before being brought to 15% at day 18 and then unaltered for the

remainder of the assay. Group D soil moisture was maintained at 5% until day 11, when it was allowed to dry before being brought to 15% at day 25 and then unaltered for the remainder of the assay. All vials were incubated at 27°C and 65% RH in continuous darkness. After 48 hrs, the pupation depth of all visible pupae was recorded. Every 72 hrs, vials were weighed and soil moisture was adjusted as per the treatment regime under red light. All adult beetles were weighed and sexed upon emergence. The assay continued until all adult beetles had emerged. The entire assay was conducted twice.

2.9.3 Synthetic lure testing

To match the release rate of the synthetic blends to that of the natural fermentate attractant, different release mechanisms for the blends were investigated in preparation for field trials.

To use in the field, the lantern traps were modified to accommodate different odour release mechanisms and killing agents inside the traps (Figure 13).

2.9.3.1 Laboratory testing

Because of the success of the lantern trap with honey/sugar fermentate attractant, the trap was modified with a wire frame to hold Falcon tubes for containing test blend (Figure 13A, B). The blend being tested (30 ml either concentrated or diluted in H2O to 50%) was placed in the Falcon tube (50 ml) and a cotton wick (120 cm) was submerged in the fluid reaching above the top of the tube (1 cm) and secured via a hole (≈1 cm diameter) in the lid. This system was tested for attractiveness to adult SHB and blend release rate in laboratory trials. Other materials were also tested this way as medium for blend release: water absorbing crystals (Eden® Water Storage Crystals), agar (1.5% in H2O) and glycerine. In order to mitigate the high volatility of the some of the components of the blend (See Section 2.2.3.2), plastic zip lock bags used in compound testing (Section 2.2.3.1) were also tested. Plastic bags have been used in several insect lure-trapping systems, including for Cerambycid species (Mitchell et al., 2015, Collignon et al., 2016).

A number of materials were tested for efficacy as killing agents in the lantern traps: propylene glycol, diatomaceous earth (DE) and canola oil. The agent needs to be effective in killing and/or drowning SHB which are well-adapted to walking on sticky and waxy surfaces which occur in the hive. The agent also needs to either be effectively odourless or possess odour not repellent to SHB as it may interfere with the attractant lure.

Figure 13: Lantern trap modified for synthetic lure testing with a different odour release mechanisms. (A) Blend in falcon tube suspended over DE, (B) blend in Falcon tube suspended over vegetable oil and (C) blend in baggie suspended over DE.

2.9.3.2 Field Testing

Blends #6 and #8 were chosen for field testing at both the Gumdale and Bellbowrie apiary sites. Blend #6 (30 ml 100% or 50% in H2O) was released via a cotton wick (see above) with DE (20 mm depth) in the bottom of the trap. Traps were placed in the same positions as paired honey/yeast and control lantern traps at Gumdale; 2 × 100% (NW and SW), 2 × 50% (NE and SE) in March/April 2017, replaced every week for four weeks. This was repeated with Blend #8 during April/May 2017.

Blend #10 was tested at three sites (one apiary: Bellbowrie; two non-apiary: Chapel Hill 27.52°S, 152.96°E and Camira 27.64°S, 152.92°E) using glycerine as a release mechanism during October 2017. Four treatment traps (100% blend, 1:1 blend: glycerine, 1:2 blend: glycerine and 100% glycerine) were deployed for five days and monitored for loss in mass.

Blends #4 and #10 were tested at Gumdale and Bellbowrie apiary sites using plastic bag methodology developed in section 2.4.1. At Gumdale traps (× 3 including water control) were placed at each of 8 locations: north, east, south, west both inner and outer. Traps had DE (15 g) in the bottom as the killing agent; traps were replaced once a week for two weeks during November 2017. At Bellbowrie traps (× 3 including water control) were placed at 4 locations (as per Section 2.3) and were replaced once a week for two weeks during November and December 2017. 2.10 Production of an educational video

An educational video detailing the method for making and deploying the yeast-based external attractant trap was developed for the beekeeping community. The video features a commercial beekeeper (Phillip Bowman, Stradbroke Island Organic Honey) and is suitable for uploading to YouTube. The video was shot at the Gumdale apiary site with a Canon EOS 700D and post edited in VEGAS Movie Studio 14 Platinum (Magix Software GmbH).

2.11 Statistical analyses

Behavioural choice assays in individual arenas were analysed using the Chi-squared method. All other behavioural choice assays with two groups were analysed using the Mann- Whitney U test. Behavioural choice assays with a minimum of three groups were analysed using either a Kruskall-Wallis one-way, non-parametric Analysis of Variance or a generalised linear model (GLM) for a binomial distribution with the logit-link function. Probabilities from the GLM were estimated using deviance ratios. Means were estimated via back- transformation of the link function and the differences between means were compared using Fisher’s Protected LSD test. In aggregation choice assays, Mann-Whitney U analysis was used to test sex differences in the amount of time beetles spent in clusters. A non-linear regression was used to model the amount of time beetles took to form clusters of five or more.

An analysis of covariance (ANCOVA) was used to examine time to emergence between soil moisture and simulated rainfall treatments with the covariate of larval mass, as this was significantly different between groups. A Pearson’s rank correlation was used to examine the relationship between pupation depth and time to emergence. A one-way Analysis of Variance (ANOVA) was used to compare between mean number of SHB trapped at different distances from hives within the Bellbowrie apiary with the differences between means compared using Fisher’s Protected LSD test.

To determine whether chemistry of fermenting hive products were significantly different from each other, an analysis of similarity (ANOSIM) was used. The ANOSIM tests are a range of Mantel-type permutations of randomisation procedures, which make no distributional assumptions. These tests depend only on rank similarities, and thus are appropriate for these types of data. We used a ‘similarity percentages’ (SIMPER) analysis to ascertain the relative contribution of each of the components to assign the leaves to a priori determined groups, to determine differences among groups and to assess similarity among individuals within each group. The software used for the multivariate analysis was Primer 7 for Windows (V 7.0.13, PRIMER-e, Clarke and Gorley (2001)). All other analyses were performed in Genstat (V 16.1.0.10916, Payne et al. (2015)).

Results

3.1 Fermentation of hive products

The fermented hive products (slime) were found to vary in attractiveness to SHB with the source location of the hive products and through time as the fermentation developed. This variation was used to look for attractive compounds suitable for use in a synthetic lure for SHB. Ten putative compounds for consideration in a lure were identified through a subtractive analysis of the compounds always present in the slime when it was attractive to SHB and those that were not present when the slime was not attractive to SHB.

3.1.1 Behavioural bioassays

There were differences in the attractiveness of the different fermentation products for small hive beetles (Table 9, Figures 14, 15). On Day 18, for the majority of sampling sites, slime was the most attractive treatment, although there was some variability between sites with respect to this attractiveness.

Significant differences between attraction to the various treatments within a site by beetles on Day 18, 35 and 49 after inoculation with beetles was examined using a generalised linear model (GLM). Differences between sites within a day were also examined. On Day 18, the percentage of beetles attracted to the fermented hive products from each site was significantly higher than that attracted to brood comb for seven of the eight sites (Table 9, Figures 14, 15).

After that, the pattern became more variable, with attractiveness having completely dropped off at some sites, while others were still attractive after seven weeks. Overall there are three general patterns observed (Figure 14), those where all samples are attractive, regardless of the time since inoculation with beetles (e.g. Cairns, Townsville, Bellbowrie, Richmond), those that were initially attractive on Day 18, but this attractiveness decreased and on Days 35 and 49 it was not significantly different to random (e.g. Bundaberg, Maleny, Goulburn) and the Grafton sample where fermenting hive products were never significantly attractive.

Table 9: Percentage (± SEM) of small hive beetles attracted to slime. In virtually all samples, there was a significant effect of treatment on beetle choice, but only those samples where the slime was significantly more attractive than brood comb are shaded.

SHB Attraction to slime Location Day 18 Day 35 Day 49

66.1 ± 3.1% 63.1 ± 3.1% 71.9 ± 2.9% Cairns (χ2 = 118.36, P < 0.001) (χ2 = 101.86, P < 0.001) (χ2 = 134.76, P < 0.001)

74.7 ± 2.8% 46.9 ± 3.2% 64.5 ± 3.1% Townsville (χ2 = 156.91, P < 0.001) (χ2 = 68.63, P < 0.001) (χ2 = 119.55, P < 0.001)

56.4 ± 3.2% 32.0 ± 3.1% 26.5 ± 2.9% Bundaberg (χ2 = 112.49, P < 0.001) (χ2 = 103.20, P < 0.001) (χ2 = 118.0, P < 0.001)

68.9 ± 3.0% 33.3 ± 3.1% 24.9 ± 2.9% Maleny (χ2 = 127.19, P < 0.001) (χ2 = 65.02, P < 0.001) (χ2 = 33.14, P < 0.001)

84.8 ± 2.3% 70.9 ± 3.0 % 55.8 ± 3.3% Bellbowrie (χ2 = 212.61, P < 0.001) (χ2 = 164.07, P < 0.001) (χ2 = 86.39, P < 0.001)

31.0 ± 3.0% 11.4 ± 2.1% 4.3 ± 1.3% Grafton (χ2 = 59.14, P = 0.007) (χ2 = 84.24, P < 0.001) (χ2 = 117.81, P < 0.001)

78.8 ± 2.6% 53.2 ± 3.3% 63.8 ± 3.1% Richmond (χ2 = 172.14, P < 0.001) (χ2 = 95.54, P < 0.001) (χ2 = 118.35, P < 0.001)

83.3 ± 2.4% 14.0 ± 2.3% 14.1 ± 2.3% Goulburn (χ2 = 196.61, P < 0.001) (χ2 = 122.47, P < 0.001) (χ2 = 93.87, P < 0.001)

Slime significantly 7 / 8 3 / 8 4 / 8 attractive

Bellbowrie Day 18 Day 49

Maleny Day 18 Day 49

Grafton Day 18 Day 49

Figure 14: Sample radial plots (below) showing mean (+ SEM) proportion beetles attracted to each treatment for the same samples. Filled blue triangle showns mean proportion attracted, empty blue triangle is mean + SEM. Red triangle shows random attractiveness

Bellbowrie Day 18 Day 35 Day 49

Maleny Day 18 Day 35 Day 49

Grafton Day 18 Day 35 Day 49

Figure 15: : Sample pie charts (above) showing mean proportion of small hive beetles attracted to treatments on each day from Bellbowrie, Maleny and Grafton. ( control brood honey slime not trapped).

3.1.2 Chemical analysis

Gas chromatography-mass spectrometry (GC-MS) allowed us to compare between the volatiles above samples from each of the sample sites, and also to see how this chemical composition changed as the slime aged. We detected 246 different peaks across all samples, but many of these are likely not important in beetle attractiveness. We detected large differences in the suite and concentration of compounds detected in samples shown from the bioassay to be attractive compared to those that were not significantly attractive (Figure 16).

A Bray-Curtis similarity matrix was prepared on fourth-root transformed data to compare between samples. This matrix takes account not only of presence/absence of the compounds detected in the aeration above the slime, but also some measure of their relative abundance. Using multidimensional scaling ordination (MDS) and Analysis of Similarity (ANOSIM), we compared between attractive and non-attractive samples, and found a significant difference between them (ANOSIM: Global R = 0.429, P = 0.001) (Figure 16).

Stress: 0.1 Attractive

Non-attractive

Figure 16: Non-metric multidimensional scaling ordination showing clear differences between attractive and non-attractive samples. Each point in the ordination represents a single sample.

Cairns Townsville Bundaberg Maleny Bellbowrie Grafton Richmond Goulburn

100

90 a a ab 80 ab x ab A ab 70 AB x x bc x 60 AB ABC 50

40 BCD c BCD y 30 y

Percentage ofbeetles attracted slime to 20 CD yz D 10 z

0 Day 18 Day 35 Day 49 Days after inoculation with beetles

Figure 17: Percentage of beetles attracted to fermenting hive products (slime) on days 18, 35 and 49 after inoculation with beetles across eight apiary locations. Columns headed with different letters are significantly different.

Abundance

6 TIC: 011114_02.D\data.ms

4e+07 6e+07 TIC: 141014_02.D\data.ms (*)

3.5e+07 5.5e+07 2 8 4 3e+07 5e+07

2.5e+07 4.5e+07 5 3 9 2e+07 4e+07

1.5e+07 3.5e+07

1e+07 3e+07 7 1 0

abundance 500000 2.5e+070 11 1 2e+070

1.5e+07 1 0 1e+07

5000000 9

3 6 7 8 0 2.00 4.00 6.00 8.00 10.00 12.00 14.00 16.00 18.00

Time--> retention time (min)

Figure 18: Total ion chromatograms of an attractive (Canberra Day 18, top) and non-attractive (Canberra Day 35, bottom) sample showing differences in compounds detected. Key compounds are identified by number.

1 ethanol 7 2,3-butanediol 2 ethyl acetate 8 isopentyl acetate 3 isobutanol 9 phenylethyl alcohol 4 acetoin 10 dihydrocarvone 5 isopentanol 11 phenylethyl acetate 6 ethyl isobutyrate

A Similarity Percentage (SIMPER) analysis was conducted to ascertain which compounds were most important in distinguishing between attractive and non-attractive samples. We detected an 81.9% dissimilarity between the groups, and determined those fourteen compounds which contributed most to this dissimilarity. Several of these components are part of the commercial Catcha® lure for Carpophilus beetles, and some components have also been reported to be able to be detected electrophysiologically by the small hive beetle through EAD (Table 10).

Table 10: The most important volatile compounds to distinguish between attractive and non-attractive samples, including the percentage contribution they make to distinguish the groups. Several of the compounds detected have been shown to be detected by the antennae

Mean (± SEM) Mean (± SEM) % abundance % abundance % Compound per gram for per gram for EAD active? †, ‡ Contribution attractive non-attractive samples samples

phenylethyl alcohol 13.64 ± 2.46 4.78 ± 1.99 2.24 Y

isopentanol 8.18 ± 1.92 4.66 ± 1.39 1.73 Y

acetoin 4.82 ± 0.87 1.83 ± 0.73 1.49

phenylethyl acetate 5.02 ± 1.46 3.12 ± 1.34 1.42

2,3-butanediol 4.28 ± 1.62 1.12 ± 0.45 1.39

ethyl acetate 4.32 ± 0.93 3.13 ± 1.14 1.37 Y

isobutanol 3.17 ± 1.28 0.67 ± 0.31 1.31 Y

ethanol 2.75 ± 0.73 1.62 ± 0.62 1.22 Y

acetic acid 2.39 ± 0.57 0.74 ± 0.33 1.09

acetone 1.58 ± 0.67 1.40 ± 0.60 1.07

isopentyl acetate 2.12 ± 0.55 1.07 ± 0.72 1.06 Y

† Torto et al. (2005), ‡ Torto et al. (2007a)

3.2 Individual compounds attractive to SHB

Initial assays to optimise test conditions showed that harbourage paper soaked in a saturated sucrose solution was significantly more attractive than paper soaked in water alone (χ2 = 4.9, P = 0.002), and no combination of dry sugar or dry paper was attractive (χ2 = 0.167, P = 0.563; χ2 = 0.048, P = 0.758 respectively). Sugar soaked paper with accessible fermenting hive products was preferred to sugar soaked paper alone (χ2 = 3.38, P = 0.009), but if the slime was inaccessible, then the food reward of sugar-soaked paper was preferred (χ2 = 5.04, P = 0.002). These results informed subsequent assays, which always contained the sugar-soaked paper in both treatment and control, to ensure that once a choice had been made for a trap, beetles would remain within the paper.

When compounds were tested individually, there was no significant preference for treatment over blank (empty bag) for four of the compounds (isobutanol, isopentanol, isopentyl acetate and phenylethyl alcohol) at any tested concentration. Five of the other compounds (2,3- butanediol, acetic acid, acetoin, acetone and ethyl acetate) were significantly attractive when compared to blank at one concentration, but not at the others; and one compound (phenylethyl acetate) was significantly avoided at a concentration of 10%. Ethanol was significantly preferred to a blank vial (χ2 = 3.38, P = 0.009).

3.3 Synthetic lure development

3.3.1 Carpophilus system evaluation

The percentage of beetles attracted to each component of the commercially available Catcha® system, individually and in combination, are shown (Table 11). The only treatments that attracted significant numbers of beetles were those containing part B (acetaldehyde). These results are important, if the Catcha® attractant developed for the related Nitidulid Carpophilus beetles also worked for SHB then this research could have moved straight to field testing once a suitable trap was evaluated. However, this result showed that further work on a SHB specific attractant lure was warranted.

Table 11: Attractiveness of Catcha® system components to the small hive beetle, individually and in combination.

Test Statistic (χ2) Components vs. % beetles (Significance) control responding Catcha® part A 61.4% 1.14 (0.132)

Catcha® part B 54.5% 0.182 (0.546)

Catcha® part C 52.4% 0.048 (0.758)

Catcha® parts A + B + C 64.6% 2.04 (0.043)

Catcha® parts A + B 66.7% 2.50 (0.025)

Catcha® parts A + C 33.3% 2.33 (0.031)*

Catcha® parts B + C 76.9% 5.65 (0.003)

* Combination is repellent.

3.3.2. Blend testing

Response of beetles to blends at a variety of combinations in the individual arenas showed that blend #1 was attractive at any concentration below 100%, at none of the tested combinations were either blend #4 nor blend #6 attractive, and blends #1 and #4 at 100% were significantly avoided (Table 12).

Table 12: Attractiveness of synthetic blends to the small hive beetle in the individual arena bioassay.

Blend # vs. control % beetles Test statistic responding (χ2) (significance)

Blend #1 (100%) 10.9% 14.09 (1.11 × 10-7) *

Blend #1 (1%) 66.7% 4.67 (0.031)

Blend #1 (0.1%) 67.6% 8.80 (0.003)

Blend #4 (100%) 25.0% 7 (0.008) *

Blend #4 (1%) 50.0% 0 (1)

Blend #4 (0.1%) 61.8% 1.88 (0.17)

Blend #4 (0.05%) 39.0% 1.98 (0.16)

Blend #6 (100%) 43.6% 0.641 (0.423)

Blend #6 (1%) 42.9% 0.857 (0.354)

* Blend is repellent at this concentration.

Initial tests in the BugDorm bioassay tested several of the blends against a blank control, and found that blend #1 was not attractive at any concentration tested, while blend #4 was significantly attractive, but only at a concentration of 100% (Table 13).

Table 13: Attractiveness of synthetic blends against a blank control to the small hive beetle in the BugDorm bioassay.

Blend # vs. Test statistic control (significance)

Blend #1 (100%) U = 7.5 Control a (P = 0.106) Blend #1 100% a

Blend #1 (0.1%) U = 9.0 Control a (P = 0.165) Blend #1 0.1% a

Blend #4 (100%) U = 2.0 Control a (P = 0.009) Blend #4 100% b

Blend #4 (1%) U = 10.5 Control a (P = 0.264) Blend #4 1% a

Blend #4 (0.1%) U = 8.0 Control a (P = 0.123) Blend #4 0.1% a

The blends were then tested in the BugDorm assay in a three-choice test, between a blank control and two concentrations of the blend (100% and 1%). Blend #6, #8 and #10 were significantly attractive at 100% concentration (Table 14).

Table 14: Comparison between attractiveness to the small hive beetle of blank control and two concentrations of synthetic blends (100% and 1%) in the BugDorm bioassay.

Blend # Test statistic (significance)†

Blend #4 χ2 = 3.30 Control a (P = 0.037) Blend #4 1% a Blend #4 100% a Blend #4a χ2 = 8.39 Control a (P < 0.001) Blend #4a 1% a Blend #4a 100% a Blend #5 χ2 = 2.28 Control a (P = 0.102) Blend #5 1% a Blend #5 100% a Blend #6 χ2 = 51.73 Control a (P < 0.001) Blend #6 1% a Blend #6 100% b Blend #7 χ2 = 3.60 Control a (P = 0.037) Blend #7 1% a Blend #7 100% a Blend #8 χ2 = 118.76 Control a (P < 0.001) Blend #8 1% a Blend #8 100% b Blend #9 χ2 = 3.01 Control a (P = 0.049) Blend #9 1% a Blend #9 100% a Blend #10 χ2 = 105.82 Control a (P < 0.001) Blend #10 1% a Blend #10 100% b † Significance below 0.05 indicates that the proportion of beetles trapped overall was significant, but pairwise comparisons indicate if there was a significant difference between treatments.

Comparing between blends in all possible combinations of two-and three blend choices suggested that although the order of attractiveness was complicated, blend #4 and blend #10 warranted further testing (Table 15).

Table 15: Comparison between attractiveness to the small hive beetle of synthetic blends compared to a blank control in the BugDorm bioassay.

Blend # Test statistic (significance)† Blend #4 vs #6 χ2 = 0.86 Control a (P = 0.421) Blend #4 a Blend #6 a Blend #4 vs #8 χ2 = 4.92 Control a (P = 0.007) Blend #4 a Blend #8 a Blend #4 vs #10 χ2 = 14.34 Control a (P < 0.001) Blend #4 b Blend #10 ab Blend #6 vs #8 χ2 = 15.89 Control a (P = 0.002) Blend #6 b Blend #8 ab Blend #6 vs #10 χ2 = 53.54 Control a (P < 0.001) Blend #6 a Blend #10 b Blend #8 vs #10 χ2 = 29.79 Control a (P < 0.001) Blend #8 b Blend #10 b Blend #4 vs #6 vs #8 χ2 = 0.92 Control a (P = 0.429) Blend #4 a Blend #6 a Blend #8 a

Blend #4 vs #6 vs χ2 = 5.14 Control a #10 (P = 0.001) Blend #4 a Blend #6 a Blend #10 a Blend #6 vs #8 vs χ2 = 1.9 Control a #10 (P = 0.126) Blend #6 a Blend #8 a Blend #10 a

† Significance below 0.05 indicates that the proportion of beetles trapped overall was significant, but pairwise comparisons indicate if there was a significant difference between treatments. 3.4 Aggregation pheromone

Behavioural studies showed that SHB have a strong tendency to aggregate under the conditions tested and there was evidence to suggest that a volatile cue could be involved. However further studies are needed to find putative aggregation pheromones.

3.4.1 Aggregative behaviour investigations: arena trials

Small hive beetles consistently demonstrated aggregative behaviour under the given treatments. Specifically, SHB clustering behaviour does not appear to be sex specific as both sexes were observed in clusters and no apparent cluster initiation was observed for either sex. Small hive beetle clustering behaviour appears to be species specific given that

clustering SHB largely ignored a similarly sized black beetle (A. diaperinus). SHB did not cluster with dead conspecifics. Although the time at least one beetle spent in contact with a dead beetle was relatively high, no cluster was observed to include a dead conspecific. SHB still cluster after antennae have been removed indicating that the behaviour may be mediated by both volatile and non-volatile chemical communication.

SHB consistently formed clusters (group of 5 or more beetles) within one to two minutes. Female SHB formed clusters slightly faster than male SHB, the mean time (± SE) for a cluster of female SHB to form was 1:00 min. (± 0:17) and 1:30 min. (± 0:35) for a male SHB cluster to form. A cluster of mixed sex SHB took slightly longer to form than single sex clusters at 1:50 min (± 0:20), although the difference between these three groups was not significant (Kruskall-Wallis: H = 2.247, P > 0.100). In general cluster size increased throughout the recorded period (Fig 19); however, the clusters of beetles were in a constant state of flux, with single beetles leaving and re-entering the cluster throughout the recorded period even if the net cluster size did not change. Once an initial cluster formed SHB would often move in and out of more than one cluster, at least one cluster remained for between 84 – 87% of the recorded time for single and mixed sex trials (Table 16). There was no significant difference in the percentage of the total time males, females and mixed sex SHB spent in clusters (Kruskall-Wallis: H = 0.3041, P > 0.100). Over time SHB clusters tended to grow so that all SHB were in one large cluster (Figure 20).

6 Cluster size Cluster

4 y = 4.2684ln(x) - 0.1073 R² = 0.6943 2

Time (min)

Figure 19: Male clustering in aggregation arena with logarithmic trend line showing the increasing size of clusters with time.

0:00 15:00

Figure 20: SHB showing clustering behaviour recorded over 15 min in an arena (glass Petri dish).

Table 16: Aggregative behaviour of SHB as measured in arena assays by percentage of time spent in a cluster (≥ 5 beetles).

Treatment Sex SHB used Replicates Time to Clustering initiate behaviour cluster (min) (mean % of time in (mean ± cluster ± SEM) SEM) Untreated male 3 1:30 ± 0:35 84.95 ± 4.69 (separated from opposite sex within female 3 1:00 ± 0:17 87.10 ± 4.93 2 days of emergence from male + female 3 1:50 ± 0:20 84.95 ± 3.88 soil)

marked male + No evidence of sex- 2 - specific clustering Marked using female iridescent dye marked female No evidence of sex- 2 - + male specific clustering No cluster included With darkling male 3 - A. diaperinus beetles (Alphitobius No cluster included diaperinus) female 3 - A. diaperinus No cluster included unmated female 10 - dead SHB With dead SHB No cluster included colony* female 10 - dead SHB After antennae male colony* 9 4:07 ± 1:04 61.29 ± 7.99 removed female colony* 9 4:57 ± 0:57 54.84 ± 8.88 (* colony beetles consisted of male and female SHB maintained together, their “mated” status was unknown)

Studies with iridescent dye showed that the formation of clusters appeared not to have any sex-specific nature; males clustered with other males, females clustered with other females, and they both clustered with each other. The cue for clustering appeared to be more than visual as no SHB cluster included any of the similarly sized and coloured darkling beetles (A. diaperinus). Neither did the SHB form clusters that included any dead SHB. However both the unmated and mixed mating status females did “investigate” the dead SHB, with both groups spending the same amount of time (68 - 72%) in contact with at least one dead beetle.

There was a difference in the clustering behaviour of SHB with and without antennae. Male and female antennectomised SHB took the same amount of time to form a cluster as each other (Mann-Whitney: U = 33.5, P = 0.556), however it took significantly longer for beetles without antennae to form a group of at least five beetles than it did for intact beetles (Mann- Whitney: U = 13.0, P < 0.001). There was no significant difference in overall percentage of time spent in clusters for antennectomised male and female adults (Mann-Whitney: U= 35.5, P = 0.0.682). Male SHB without antennae spent 61.29 ± 7.99% of their time in clusters and the antennectomised female SHB spent 54.84 ± 8.88% or their in time in clusters. Although there was a significant difference in the percentage of total time in a cluster between intact and antennectomised beetles (Mann-Whitney: U = 19.0, P < 0.001), this is due to latency to form a cluster in the first place for beetles without antennae. Once a cluster was formed in any group of beetles, the cluster persisted for the majority of the remainder of the observation

period (84.28 ± 3.62%), and there was no significant difference between intact and antennectomised beetles (Mann-Whitney: U = 51.5, P = 0.126).

These investigations demonstrate that SHB exhibit a strong aggregative behaviour which is most likely initiated and driven by sensory cues other than or in addition to visual cues. The increased latency to form a cluster exhibited by SHB with antennae removed suggests that the signal to aggregate could be mediated by a volatile agent that is detected by both sensory hairs typically covering the body of insects as well as the antennae. Once aggregated, other sensory modalities (e.g. thigmotaxis) mean that cluster size is maintained, but a volatile chemical signal appears to be vital for rapid aggregation.

3.2.4.2 Chemistry of aggregation

Ten compounds which commonly occurred across treatments were identified and their percentage abundance was determined: isopentanal, acetoin, benzaldehyde, acetophenone, 2-ethylhexanol, octanal, nonanal, decanal, carophyllene and (Z)-9-tricosene. The treatments which were deemed to be the most biologically relevant in terms of beetle behaviour and biology are included below (Tables 17 and 18).

The compounds identified from these aerations included many typical fermentation products, such as acetoin, and a variety of aldehydes including benzaldehyde, isopentanal, octanal, nonanal and decanal. Aldehydes are common attractant compounds to a wide variety of insect groups (El-Sayed, 2018). Three of these aldehydes (octanal, nonanal and decanal) have previously been determined to be attractive to SHB in dual choice wind tunnel bioassays (Torto et al., 2005). Although there is no indication that these compounds are produced by the beetles themselves.

Table 17: Commonly occurring compounds (mean % abundance ± SEM) in aerations of SHB by treatment (only replicate aerations wherein compounds were present included).

Commonly occurring compounds (mean % abundance ± SE) Assay

treatment

tricosene

ethylhexanol

benzaldehyde 2 octanal nonanal decanal (Z) A Young, 2.47 ± 0.28 5.76 ± 3.49 ± 25.13 ± 16.68 ± 46.47 ± colony males 2.8 0.21 6.75 2.35 6.72 B Young, 3.56 ± 0.41 8.5 ± 1.8 3.85 ± 50.61 ± 19.64 ± 27.67 ± colony 1.94 6.18 3.04 18.72 females G Young, 4.25 3.45 ± 3.58 ± 21.67± 9.42 ± unmated 0.26 1.07 2.59 1.42 males + sucrose Control: 3.11 4.73 1.98 sucrose (n = 1)

Table 18: Commonly occurring compounds (mean % abundance ± SEM) in aerations of SHB by feeding status treatment (only replicate aerations wherein compounds were present included).

Commonly occurring compounds (mean % abundance ± SE)

Assay

Treatment

isopentanal acetoin benzaldehyde acetophenone 2 ethylhexanol octanal nonanal decanal caryophyllene N Males 7.44 5.66 10.69 before ± ± ± 2.06 feeding 2.25 2.85 O Males 2.01 4.99 5.13 0.67 1.32 0.57 4.39 ± 2.98 0.73 after feeding ± ± ± ± ± 0.92 ± 1.3 ± 0.84 1.11 4.34 0.03 0.21 0.36 P Females 0.75 1.25 2.6 5.75 17.56 5.66 before ± ± ± ± ± feeding 0.18 0.49 0.08 2.32 0.68 Q Females 1.36 8.19 0.85 0.69 1.46 4.98 1.08 0.85 after feeding ± ± ± ± ± ± ± 1.34 0.19 0.12 0.69 0.81 0.14 0.1 pollen dough 0.3 0.74 0.76 1.08 0.6 5.68 1.6 0.26 (n = 1)

Figure 21: Total ion chromatogram showing differences in aerations of: pollen dough alone, SHB females before and after feeding on pollen dough. Compounds are identified below:

1. isopentanal 4. ethyl hexanal 7. decanal 2. acetoin 5. acetophenone 3. benzaldehyde 6. nonanal

Small hive beetle distribution among honey bee colonies has been reported to be non- random with colony phenotype and size not influencing infestation levels (Neumann and Elzen, 2004b, Spiewok et al., 2007). Stuhl & Teal, (pers. comm.) thought it highly likely that SHB use an aggregation pheromone to come together to form the clusters which they had observed. The presence of such pheromones have been reported in other Nitidulid beetles, including many species in the genus Carpophilus (Bartelt and Hossain, 2010). These aggregation pheromones are now known to be mostly produced by males and only in the presence of a food source (Cossé and Bartelt, 2000, Bartelt and Hossain, 2010). Pheromone emission from Carpophilus males was found to begin one to four days after placement of adult males on food (Bartelt and James, 1994). Mating can also influence pheromone production with emission from one Carpophilus species decreasing by up to 90% after mating (Nardi et al., 1996).Twenty-two male specific tetraene and triene hydrocarbons were identified from ten Carpophilus species through research into the chemical ecology of Carpophilus spp. over five decades. Nine of these putative pheromones were found to be relatively abundant so were then isolated and synthesised for application in Carpophilus beetle control programs with co-attractant food odours (Bartelt and Hossain, 2010).

Aerations of various treatments of groups of SHB yielded complex volatile profiles. Although several compounds were found to be common between treatments, none were in the tetraene or triene class of hydrocarbons. It was also deemed that the compounds found in this study were not unique enough to constitute a conspecific targeted pheromone (El-Sayed, 2018). Access to a protein-based food source did not appear to initiate the production and release of a volatile pheromone for SHB under the conditions tested. It is possible that further feeding scenarios need to be explored, such as allowing SHB males to be starved for longer, or feed for longer, or use hive products rather than the manufactured protein source, or feed alone rather than in groups. Hence more work is required to determine the exact conditions

under which SHB could release a volatile aggregation pheromone. Although no putative, SHB aggregation compounds have been found, the results of this study do contribute to the broader knowledge of SHB aggregative behaviour and provide a foundation for further investigations. 3.5 Volatile profile of hives

Air samples from the hives exhibited a variety of chemicals in the hive odour, including many (such as the long-chain waxes tricosane, pentacosane and heptacosane) that would be expected from a hive containing wax. Also detected were plant and floral volatiles (such as α- pinene, β-pinene, acetophenone, methyl benzoate, α-copaene and α-gurjunene) (El-Sayed AM 2018) which are again not unexpected in the hive of an insect that collects food resources from flowers (Figure 22).

Interestingly, hive aerations also detected odours linked to fermentation, such as ethanol, 2,3-butanedione, 1,2-butanediol and acetoin. These compounds could be due to hive product fermentation induced by Kodamaea ohmeri (all hives examined had some level of small hive beetle infestation), but they may also be due to other fermentation processes occurring as a normal process within the hive.

There was a statistically significant difference between samples taken from the two apiary sites examined (R = 0.458, P = 0.007), regardless of the time of year when the samples were collected (Figure 23). The compounds important in distinguishing between the apiaries were determined by a SIMPER analysis, and are shown below (Figure 25). The sampling time did, however, have a significant effect upon the volatile composition of the hive, (Global R = 0.392, P = 0.005) (Figure 24), with samples from early April being significantly different to those from late April or May. There were no differences between samples for any of the other sampling periods (Figure 24). The most important compounds (and the numbers used to highlight them in Figure 26) in distinguishing early April from the other sampling periods are: acetoin (3), acetyl valeryl (6), diisobutyl carbinol (8) and pentadecane (21) (SIMPER).

Considering only the samples from Bellbowrie, the pattern for differentiation of the sampling time held (R = 0.406, P = 0.002) (Figure 26, 27) but there was no pattern of individual hives (R = 0.035, P = 0.325), with sampling period being more important than hive in determining the volatile profile of a hive (Figure 28).

1. ethanol 18. -copaene 2. 2,3-butanedione 19. tetradecane 3. acetoin 20. -gurjunene 4. 1,2-butanediol 21. pentadecane 5. 2,3-butanediol 22. L-calamenene 6. acetyl valeryl 23. hexadecane 7. -pinene 24. heptadecane 8. diisobutyl carbinol 25. nonadecane 9. benzaldehyde 26. eicosane 10. -pinene 27. heneicosane 11. cis--ocimene 28. heneicosane 12. acetophenone 29. tricosene 13. methyl benzoate 30. tricosane 14. nonanal 31. pentacosane 15. dodecane 32. pentacosane 16. decanal 33. heptacosene 17. tridecane 34. heptacosane

Non-metric MDS Transform: Square root Resemblance: S17 Bray-Curtis similarity 2D Stress: 0.1 Apiary Bellbowrie Richmond

Figure 23: Non-metric multidimensional scaling ordination showing clear differences between samples from the two apiary sites. Each point in the ordinationNon-metric represents MDS a single hive sample at a single sampling time. Ellipses added for ease of interpretationTransform: only. Square root Resemblance: S17 Bray-Curtis similarity 2D Stress: 0.1 Sample time early April late April early May late May

Figure 24: Non-metric multidimensional scaling ordination showing hive aeration samples collected across the sampling periods. There is clear clustering of samples collected from each sample period, with samples from early April significantly different from the other sampling intervals. Each point in the ordination represents a single hive sample at a single sampling time. Ellipses added for ease of interpretation only.

Figure 25: Total ion chromatograms comparing aerations of a hive from the Bellbowrie apiary (lower) (sampled 24 May 2016), and from the Wheen Bee Foundation Research apiary (upper) (sampled 18 May 2016). Air was sampled for one hour at approximately 500 ml/min onto a thermal desorption tube preloaded with Tenax TA (35/60 mesh) (Markes International Ltd.). Key compounds important in distinguishing between the apiaries are identified by number, as in Figure 22 and their percentage contribution to the dissimilarity is shown below:

2. 2,3-butanedione (1.73%) 20. -gurjunene (1.86%) 3. acetoin (5.73%) 22. L-calamenene (1.74%) 4. 1,2-butanediol (1.62%) 30. tricosane (2.59%) 5. 2,3-butanediol (2.33%) 31. pentacosene (1.96%) 6. acetyl valeryl (4.43%) 32. pentacosane (2.90%) 7. -pinene (4.87%) 34. heptacosane (3.28%) 8. diisobutyl carbinol (2.08%) 35. unknown 1 (2.64%) 14. nonanal (2.25%) 36. unknown 2 (5.00%)

Figure 26: Total ion chromatograms comparing aerations from different sampling times of a hive from the Bellbowrie apiary (sampled from bottom to top: 8 April, 27 April, 10 May and 24 May 2016). Air was sampled for one hour at approximately 500 ml/min onto a thermal desorption tube preloaded with Tenax TA (35/60 mesh) (Markes International Ltd.). Key compounds important in distinguishing between the apiaries are identified by number, as in Figure 22 and their percentage contribution to the dissimilarity is shown below.

3. acetoin (3.79%) 6. acetyl valeryl (3.69%) 8. diisobutyl carbinol (4.92%) 21. pentadecane (2.94%)

Non-metric MDS Transform: Square root Resemblance: S17 Bray-Curtis similarity 2D Stress: 0.1 Sample time early April late April early May late May

Figure 27: Non-metric multidimensional scaling ordination showing hive aeration samples collected across the sampling periods, showing only samples from the Bellbowrie apiary. There is, again, clear clustering of samples collected from each sample period, with samples from early April significantly different from the other sampling intervals. Each point in the ordination represents a single hive sample at a single sampling time. Ellipses added for ease of interpretation only.

Transform: Square root Resemblance: S17 Bray-Curtis similarity 2D Stress: 0.1 Hive B1 B2 B3 B4

Figure 28: Non-metric multidimensional scaling ordination showing samples with respect to hive, only samples from the Bellbowrie apiary site are shown. There is no clustering of samples from within a hive, instead samples cluster with respect to sampling interval (see Figure 27). Each point in the ordination represents a single hive sample at a single sampling time.

The initial impetus for this hive odour research was anecdotal observations by beekeepers that the wide variation in SHB load across the hives in an apiary may be related to a difference in the odour profile of the different hives. In this study, there was little variation in SHB load within the apiaries where the volatiles were collected, thus we were unable to clearly address any relationship between SHB load and hive volatile profile. There is likely a wide range of factors influencing the volatile profile of any given hive including floral source, current bee population and makeup (i.e. number of drones and number and age of workers), disease status and queen status. In the Bellbowrie apiary the SHB burden varied through time, but at any one time the burden was quite consistent across the apiary. It was hard to judge if changes in the volatile profiles of the hives changed through time due to climatic change with the seasons, or SHB load.

This work is the first report of an investigation into the volatiles generated from a hive and has delivered some very complex data from which any clear trends and conclusions may be difficult to make. However, the positive outcomes include the development of a methodology for sampling volatiles in a hive. The data generated provides background information on the volatile profile for healthy hives that could be used in the future for comparison with hives with a particular pest or disease load. There may even be potential for comparisons with the volatiles coming from a hive producing queen pheromone or one that has gone queen-less. 3.6 Trap design

The first trap designs evaluated (Unitraps™, Dowd traps), though able to trap some SHB when baited with slime, proved unsuitable because of low trapping efficacy. Lantern traps which are economical and readily available from hardware stores proved to be suitable for trapping SHB when baited with a simple honey- yeast fermentate. Unitraps™ baited with slime were able to trap SHB with SHB trapped both near honey bee hives and native bee hives and in locations away from any known hives, though in very low numbers (Table 19). More SHB (26) were trapped during late December 2014, at least a fortnight after the summer rains commenced, than during February 2015 (7) in a period of hot dry weather. No small hive beetles were trapped in the third trial in March (Table 19). All traps baited with slime trapped a range of other insect species including ants, flies, moths, cockroaches and other beetles. A smaller range of insects including ants, flies and moths were caught in some of the unbaited control traps. One issue with the Unitraps™ could have been the dark green colour. de Guzman et al. (2011) showed a significant higher number of SHB trapped in white traps compared to dark traps. They reasoned that higher reflectance might be important for insects such as SHB that fly both before and after dusk.

Despite the great success using the Dowd traps to trap Carpophilus beetles (Bartelt and Hossain, 2010), when they were baited with slime they were ineffective at catching SHB (Table 19). The Dowd traps were white so colour could not have been a negative issue. This design may not suit the flight dynamics of SHB, and the release of the attractant may also have been impaired compared to that in the Unitraps™ and later, the lantern traps. There was a stark contrast in SHB trapping success once the lantern traps with the honey-yeast fermentate were deployed. Although this system in itself can still be optimised it was the breakthrough that was needed to be able to begin designing trapping trials to gather data on the movement of SHB and effect of things such as distance of trap from apiary. Experience with the lantern traps suggests that the design needs to be customised for SHB by increasing the capacity for volatiles to exit the trap and prevent any SHB exiting the trap once they have entered. Although many SHB appeared to have drowned in the liquid in the traps, once the traps were opened a number of live SHB hiding in the dark coloured inner contours of the lid could be seen scurrying off to hide. Occasionally a SHB would escape from a trap when the trap was being collected. A design modification to either kill all SHB entering the trap or at least prevent the exit of any SHB once they have entered is desirable.

Table 19: Results of trials to evaluate different trap designs between December 2014 and November 2016.

Trial # Trap tested and date Comments

26 SHB caught at 8 locations. 19 SHB were caught near Unitrap™ hives at Bellbowrie (18 near Apis mellifera hives, 1 near 1 19/12/14 - 6/1/15 Tetragonula hive); 7 SHB were caught at 2 non-apiary sites near forest (6 + 1). No SHB caught at 4 sites. 7 SHB caught at 4 locations. 3 SHB caught at 2 apiary Unitrap™ 2 sites (2 + 1) and 4 SHB caught at 2 non-apiary sites. No 10/2/15 – 26/2/15 SHB caught at 1 apiary site and 1 non-apiary site. Unitrap™ No SHB caught 3 13/3/15 – 30/3/15 Unitrap™ No SHB caught 4 7/10/15 – 3/11/15 No SHB caught with Blend #2 or slime, but did catch lots Unitrap™ 5 of other insects, typically flies and other beetles. The blend 13/11/15 – 27/11/15 caught more beetles than the slime. 7 SHB caught in slime (5+1+1), no SHB caught with Unitrap™ 6 Catcha® lure. Lots of Carpophilus beetles caught at each 19/12/15 – 1/1/16 location with the Catcha® lures. Unitrap™, Dowd, 2 “in No SHB caught in any traps 7 house” design traps Feb 2016 Dowd Only 2 SHB caught in this period at Gumdale 8 30/8//16 – 20/12/16 Only 2 SHB caught 22/11/16 – by which time lantern traps Dowd 9 also deployed and caught 8 SHB on same date at 31/8/2016 – 22/11/16 Bellbowrie. Dowd Only 1 SHB caught during this period at Richmond 10 5/9/16 – 29/11/16 Lantern traps Large numbers of SHB, see Table 26 11 10/11/16 onwards

3.7 Evaluation of efficacy of synthetic lures and traps

3.7.1 Natural fermentate attractant investigations

3.7.1.1 Behavioural bioassays

All three ratios of the water to honey fermentates were at least as attractive to SHB as slime in the four-choice BugDorm assays (Table 20). The fermentate from the 75:25 ratio of water to honey was the most attractive when compared against the other ratios in both the BugDorm (Table 20) and two- choice individual arena assays (Table 21). Consequently, all further investigations were based on fermentates from a 75:25 ratio of water to honey (and/or sucrose).

Table 20: Mean percent SHB trapped per choice in four BugDorm arena assays. Mean percent un-trapped beetles is the summed percentage of all choices subtracted from 100. Means with the same superscript within rows were not significantly different.

Control Honey Slime 50:50 60:40 75:25 GLM

a ab c bc 2 0 7.8 55.8 35.5 - - χ = 96.91(3,20) (SE±1.61) (SE±6.26) (SE±5.12) P < 0.001

a a b c 2 0 1.7 32.1 - 64.95 - χ = 136.29(3,20) (SE±0.33) (SE±2.32) (SE±2.35) P < 0.001

a a b b 2 0.4 2.1 57.02 - - 40.4 χ = 119.21(3,20) (SE±0.17) (SE±0.48) (SE±5.21) (SE±5.70) P < 0.001

a b b c 2 0 - - 20.5 22.2 52.1 χ = 72.05(3,20) (SE±3.43) (SE±3.27) (SE±3.21) P < 0.001

Table 21: Comparing fermentates with different ratios of water to honey for attractiveness to SHB in two-choice individual arena assays. The split column shows relative numbers of beetles per choice.

Ratios split Preference χ2 P 60:40 vs 75:25 17 vs 31 75:25 4.08 0.043 50:50 vs 75:25 8 vs 39 75:25 20.4 6.13 x 10-6 50:50 vs 60:40 6 vs 39 60:40 24.2 8.68 x 10-7

When the attractiveness of different ages of fermentate based on a 75:25 ratio of water to honey was compared, all ages of the blend were attractive to beetles although the attractiveness varied with time. The freshly prepared fermentate was least attractive, attractiveness increased after day one to a peak around day five then decreased to a level similar to the fresh fermentate by day seven (Table 22).

Table 22: Comparison of the attractiveness to SHB of fermentate (75:25 water/honey with yeast) at different ages (0 – 7 days old) in two-choice individual arena assays. The split column shows relative numbers of beetles per choice.

Blend ages Split Preference χ2 P day 0 vs blank* 90 vs 3 day 0 81.4 1.86 x 10-19 day 1 vs blank 49 vs 0 day 1 49 2.56 x 10-12 day 2 vs blank 48 vs 2 day 2 42.32 7.75 x 10-11 day 3 vs blank 46 vs 2 day 3 40.33 2.14 x 10-10 day 4 vs blank 45 vs 3 day 4 36.75 1.34 x 10-9 day 5 vs Blank 49 vs 1 day 5 46.08 1.14 x 10-11 day 6 vs Blank 45 vs 2 day 6 39.34 3.56 x 10-10 day 7 vs blank 42 vs 4 day 7 31.39 2.11 x 10-8

day 0 vs day 1 15 vs 32 day 1 6.15 0.013 day 0 vs day 2 10 vs 35 day 2 13.89 0.002 day 0 vs day 3 26 vs 26 neither 0.532 0.466 day 0 vs day 4 5 vs 39 day 4 26.27 2.96 x 10-7 day 0 vs day 5 6 vs 39 day 5 24.2 8.68 x 10-7 day 0 vs day 6 13 vs 31 day 6 7.36 0.007 day 0 vs day 7 24 vs 23 neither 0.021 0.884

day 1 vs day 3 24 vs 21 neither 0.2 0.655 day 1 vs day 4 22 vs 21 neither 0.023 0.879

day 7 vs day 4 22 vs 27 neither 0.51 0.475 day 7 vs day 5 15 v 33 day 5 6.75 0.009 * Pooled data from two repeat assays.

When sucrose was used in the fermentate instead of honey there was no difference in the attractiveness to SHB in laboratory assays. Five day old fermentates based on either a 75:25 ratio of water to honey or water to sugar were both attractive to SHB when tested in both individual arena (χ2 = 0.083, P = 0.773) and Bugdorm (χ2 =87.01, P < 0.001) assays.

However, in the field significantly more SHB were trapped in lantern traps with the honey fermentate than in paired lantern traps with the sucrose fermentate (χ2 = 37.13, P = 1.1 × 10-09). Furthermore, for every pair of traps at different distances from the hives, there was a significant difference between the number of SHB trapped in the honey fermentate compared to the sucrose fermentate (10 m: χ2 = 14.53, P = 0.0001, 40 m: χ2 = 9, P = 0.0027; 100 m: χ2 = 7.71, P = 0.005; 200 m: χ 2= 7.74, P = 0.005) (Figure 29).

Honey 2.5 Sucrose

1.5

0.5 Mean SHB caught per trap trap per caught SHB Mean

0 10 40 100 200 Trap distance from apiary (m)

Figure 29: Comparison of attractiveness of honey and sucrose fermentate to SHB when used in lantern traps in the field.

A fermentate based on a mixture of honey and sucrose was found to be significantly attractive to SHB when tested in lantern traps in the laboratory. This mixed fermentate was tested in two-choice large tent arena assays with the lantern traps both on the ground (U:1, P < 0.001) and suspended approximately 80cm from the ground (U:0, P < 0.001).

3.7.1.2 Chemical analysis

Analysis of the volatile components of the honey and sucrose fermentates showed that although their odour profiles had some common components, the honey based fermentate was more complex with a greater abundance of components (Figure 30, Table 23). The common components were ethyl acetate, isobutanol, isopentanol, 2-methyl butanol, and ethanol, with the latter being the most abundant in both of the fermentates. Propanol and acetic acid were also present in the honey fermentate, albeit at very low levels. Although the common components appeared to diminish faster in the honey fermentate there was still a greater abundance of most components after 144 hrs than in the sucrose fermentate at 120 hrs (Table 23).

Figure 30: Representative chromatograms of the honey blend (A: 24 hrs; B: 144 hrs) and the sugar blend (C: 24 hrs; D: 120 hrs), with the peaks of the common components numbered. Ethanol (1), ethyl acetate (2), isobutanol (3), isopentanol (4), and 2-methyl butanol (5). Note that the scale of abundance is around 9x greater for graphs A and B than in C and D.

Table 23: Relative abundance of the five common components seen in the representative chromatograms (Figure A) of the honey and sucrose fermentates after 24 hrs and 120/144 hrs of fermentation.

24hrs 120hrs 144hrs compound sugar honey sugar honey ethanol (1) 48,012,758 175,445,193 48,148,995 38,044,948 ethyl acetate (2) 1,093,814 30,881,662 526,004 1,053,843 isobutanol (3) 3,166,527 42,184,104 536,672 8,195,844 isopentanol (4) 503,659 42,400,320 293,002 9,095,369 2-methyl butanol (5) 200,921 36,052,499 233,431 11,341,773

Based on the results above it was decided to use a combination of both honey and sucrose to produce a natural fermentation attractant for use in lantern traps in the field. The mix of honey and sucrose provided an economical mix for use in the field studies to gather data on trap placement and monitoring the movement of SHB.

3.7.2 Ecological studies of SHB

Before the efficacy of a synthetic lure and trap can be evaluated, knowledge of when best to deploy traps and where best to site traps is crucial, as well as a way of measuring the change in SHB numbers in a hive. If there are no SHB flying around then even the most attractive SHB lure possible will fail to trap any SHB. To date there is little data documenting the seasonal and intra-seasonal field movement of SHB in Australia. Annand (2011) recorded SHB movement into and out of four hives in Richmond, NSW, over 24 hrs periods once a month, for 14 months. This study showed that SHB mostly fly around dusk and peak movements occurred in October and April at this location. The data was only collected for 24 hrs once a month so did not show what SHB movement was occurring for the rest of each month. Ecological studies have been conducted on the SHB range in the southeast of the USA (Arbogast et al., 2007, Arbogast et al., 2009, de Guzman et al., 2010, de Guzman et al., 2011). Arbogast et al. (2007) investigated the effect of shade and distance from hives on trapping efficiency. de Guzman et al. (2010) explored seasonal dynamics of SHB in two different apiary sites, while de Guzman et al. (2011) studied the effect of trap colour and height in two different apiaries throughout 2009.

There is no Australian data showing the movement of SHB into hives on a monthly basis at different locations. There are many anecdotal observations on SHB movement (pers. comm. with different beekeepers) but these have not been systematically documented. Hence, this study needed to find a basic trapping system capable of regular SHB trapping to document ecological data such as the movement patterns of SHB throughout the year, triggers for SHB movement at the beginning of spring and during the warmer months to inform when to deploy traps.

Knowing where to set traps in relation to an apiary is also important. Although Arbogast et al. (2009) conducted studies in this area, their conclusions are confusing in light of the results presented. Finding and counting all SHB in a hive is challenging as they can move very quickly once a hive is opened and are known to hide under clusters of bees, in empty cells and in crevices (Leemon, unpubl., 2009). The current method of assessing the total number of SHB in a hive as outlined in Annand (2011) and de Guzman et al. (2006) is both time consuming and extremely disruptive to a hive. A less invasive method that can be easily and regularly carried out is needed to track changes in the population of SHB in a hive. This is essential if the efficacy of a lure and trap is to be gauged against the number of SHB that manage to enter a hive.

This study developed an easy method for estimating the number of SHB in a hive and thus any changes in SHB numbers in a hive that might be influenced by either a large influx of SHB or successful interception of SHB with an external trap. Trapping data (in hive and external) showed the seasonal changes in SHB numbers at three locations in Australia in response to temperature, and suggest that intra-seasonal changes in numbers are influenced by rainfall. The results of this study suggest that an effective trapping strategy would involve putting out attractant traps for SHB during September when the minimum temperatures begin to increase above 12 °C. This appears to also coincide with a max daily temperature around 27 °C. This agrees with de Guzman et al. (2010) who found an increase in SHB populations in hives when day temperatures were ≥ 27 °C. Their suggestion that increased SHB populations will result from the accelerated developmental cycles at higher temperatures is supported by several laboratory investigations (Schmolke, 1974, Neumann et al., 2001, de Guzman and Frake, 2007, Meikle and Patt, 2011).

Trapping during extended hot dry periods is not supported by this study, as it appears that SHB do not move about under these conditions during summer. However, as soon as it rains after a period of hot dry weather traps should be deployed. This study also showed no significant difference between

trap catch in traps placed up to 185 m from hives compared to traps placed within 20 m of hives in spring and 5 m in summer. Trap placement more than 185m from hives was not assessed.

3.7.2.1 Relative measures in hives

The relative measures of SHB (visual counts, SHB trapped in a mat, SHB trapped in trench traps) were carried out fortnightly in three different locations for 15 months at Bellbowrie and Gumdale with a break during winter. At Richmond the relative measures were conducted in three periods of 2.5, 1.5 and 1.5 months. The relative measures were validated by pulling a hive apart and counting all SHB a total of eight times across the three locations. All three relative measures added together constitutes a hive check value. The percentage of SHB in a hive was estimated by dividing a relative measure (e.g. mat, trench trap, visual or hive check) by the total number of SHB in a hive (hive validation + hive check) (Table 24). Using this method, the hive check value represents 38.9% (± 0.4) of the SHB in the hive (Table 24).

Table 24: Percentage of SHB in a hive accounted for through the fortnightly relative measures (visual, mat or trench traps with DE) of SHB. Hive check = all three relative measures added together.

Comparison Average

Hive Check/Total 38.9 ± 3.8%

Visual/Total 11.42 ± 1.7%

Trench Trap/Total 11.61± 2.6%

Mat /Total 15.88± 3.3%

Mat + Visual/Total 27.3± 4.0%

The visual measure (numbers of SHB seen under mats as they are lifted) showed the least variation. When the visual score is added to the mat score, the trend follows that of the hive check even more closely. This suggests that these two scores combined are the least variable of the three scores over time (Figures 31 & 32). With a fibrous mat placed on top of the frames in the top super it is no extra work to take both a visual count and count the number of SHB trapped in the mat. This simple relative measure can be carried out quickly and with little disruption to the hive; it only involves opening a hive, counting the SHB seen as the mat is lifted, replacing the mat, and counting the SHB trapped in the removed mat. Although the number counted will only give an estimate of the number of SHB in the hive at 27.3± 4.0% (Table 24), this method allows the changes in SHB numbers in a hive to be tracked while an external trapping system is being evaluated.

500 Total Mat + Visual 450 400 350 300 250 200 150

100 Number SHB counted SHB Number 50 0

Date Figure 31: Correlation between the visual + mat counts of SHB and fortnightly hive check counts at Bellbowrie.

180 Total Mat + Visual 160 140 120 100 80 60 40

Number SHB counted SHB Number 20 0

Date

Figure 32: Correlation between the visual and fibrous mat counts of SHB and fortnightly hive check counts at Gumdale.

3.7.2.2 Field movement of SHB

3.7.2.2.1 Emergence Triggers

The soil moisture and simulated rainfall regimes significantly affected the time to emergence of SHB adults (F3, 154 = 19.56, P < 0.01). The mean days to emergence were 17.78, 19.05, 20.30 and 21.25 for groups A-D, respectively, and all treatment groups were significantly different from each other. In treatments A and B the pupation soil was kept at a constant 5% moisture until day 18, when the moisture level was increased to 15% for treatment B and kept at 5% for treatment A until all SHB had emerged. As the average time for emergence in treatment A was less than 18 days, it might be interpreted that a sudden increase in moisture (equivalent to a rain downpour in the field) might delay emergence of any SHB that are still in the soil. For soils dried after day 11 (treatments C & D) the data suggests that a sudden increase in moisture, akin to rainfall in the field, may stimulate the emergence. Further research is needed to provide data to support field observations in this area.

Time to emergence did not differ between sexes (F1, 149 = 1.41, P = 0.24), nor did pupation depth (F84, 64 = 1.19, P = 0.308). There was no correlation between pupation depth and time to emergence (r = 0.015, n = 84, P = 0.893).

Bernier et al (2014) found that SHB sex ratio was influenced by soil water content, with one female to every male in dry to intermediate soils, but three females to one male in wet soils. However further investigation is required to fully understand what happens when soils dry out at different stages of pupation and whether some SHB pupae can undergo eclosion to adults but will wait in dry soil until a rain event occurs before emergence. de Guzman and Frake (2007), when looking at the effect of temperature on development, observed that some teneral SHB adults spent days in their pupation cells and only moved up through the soil gradually. They also observed that some newly emerged adults went back underneath the soil. This behaviour was observed incidentally in our laboratory assays and SHB colony.

3.7.2.2.2 Trap placement

Distance from hive did not appear to have a consistent influence on trapping efficacy with large numbers of SHB trapped at various distances up to 185 m from hives. However different patterns were seen depending on how the trapping was carried out (i.e. only 2 distances compared or a range of distances compared) and when the trapping occurred (i.e. different seasons). When traps were placed at either 20m or 100 m from the hives more SHB were trapped further from the hives. However, there was no significant difference between the number caught in the lantern traps with a natural fermentation attractant at either Richmond (P = 0.306) (Figure 33) or Gumdale (P = 0.182), (Figure 34).

4.5 4 3.5 3 2.5 2 1.5 1

Mean SHB caught per trap per caught SHB Mean 0.5 0 20 100 Trap distance from apiary (m)

Figure 33: Mean number of SHB (+SE) caught in lantern traps with a natural fermentation attractant at different distances from the Richmond apiary.

4.5 4 3.5 3 2.5 2 1.5 1

Mean SHB caught per trap per caught SHB Mean 0.5 0 20 100 Trap distance from apiary (m)

Figure 34 : Mean number of SHB (+SE) caught in lantern traps with a natural fermentation attractant at different distances from the Gumdale apiary.

When attractant traps were deployed over a range of distances (5 m – 185 m from Bellbowrie hives) there was an overall significant difference in the number of SHB trapped at different distances depending on the time of year (Table 25, Figure 35). There was no significant difference in the numbers of SHB trapped either closest to the hives (5 m) or furthest from the hives (185 m) regardless of season. During spring, most SHB were trapped at these two distances. However, in late summer the highest numbers were trapped at two intermediate distances (50 m and 125 m), though not at 75 m. In winter, there was no significant difference in the numbers of SHB caught over all distances from 5 m to 185 m from the hives (Table 25, Figure 35). Trapping at all three sites showed that SHB can be easily trapped either close to hives or further away. Arbogast et al. (2009) concluded that trapping frequency decreased with distance from hives, however they did not comment on their trapping efficacy with yeast inoculated pollen dough. Although the results presented suggest that they trapped more SHB more than 75 m from the hives than at less than 50 m from the hives. They did note that traps in the shade trapped far more SHB than traps in partial shade. The traps at all sites were hung in trees to capture the shade, where possible. Trap efficacy is most likely affected by wind direction. Traps downwind of hives may have intercepted and trapped more SHB following a hive odour trail. This is one aspect that would benefit from more data. In this study, SHB were trapped further from hives than in the Arbogast et al. (2009) study (185 m compared to 160 m). Another difference between the studies was the number of SHB trapped over a similar time period (61 weeks); Arbogast et al. (2009) reported trapping 508 SHB across 12 traps, in this study 2989 SHB were trapped in up to 8 traps in the Bellbowrie apiary.

Lantern traps with the honey fermentate attractant located at Chapel Hill caught 19 SHB over 18 weeks, while those at Carina Heights trapped three SHB over 17 weeks. In the same time none of the lantern traps without the attractant at the same location caught any SHB. This demonstrates that SHB can be trapped at distances much greater than 185 m from managed hives, albeit at very low trapping efficiency and frequency. This also suggests that while some SHB can be attracted to the traps with the honey fermentate a long way from hives, the presence of hives is likely to increase the movement of SHB into a general area. It can be speculated that the fermentate trapping system, and perhaps even a synthetic lure works best when positioned to intercept SHB attracted to an apiary site. Further research in this area is warranted.

Table 25: Mean number of SHB trapped per trap per trapping interval at different distances from hives. Letters in brackets after the mean denote significance.

23/12/16-31/3/17 1/4/17-26/9/17 27/9/17-22/11/17 Distance from Summer to Early Autumn – Early Spring (56 days) hives(m) Autumn (98 days) Spring (178 days)

5 2.15 (ab) 1.47 5.19 (b) 10 1.43 (a) 1.04 3.76 (a) 20 2.87 (bc) 2.07 2.05 (a) 50 4.33 (d) 1.20 2.38 (a) 75 2.72 (bc) 1.09 2.24 (a) 125 3.65 (cd) 0.96 3.76 (ab) 185 2.85 (bc) 1.60 5.91 (b) F 4.55 1.02 2.52 P P < 0.001 P = 0.414 P = 0.017

Figure 35: Mean number of SHB (±SE) trapped per trap per day in attractant yeast traps at different distances from the Bellbowrie apiary over different seasons. (A) Later summer to early autumn (23/12/2016-31/3/2017). (B) Autumn through winter to early spring (1/4/2017 – 26/9/2107). (C) Spring (2/9/2017 – 22/11/2017).

3.7.2.2.3 Seasonal trapping of SHB

The SHB trapped at the different sites varied with time of year and with location (Table 26). There was a hierarchy in the number of SHB trapped at the three locations with the highest numbers trapped at Bellbowrie and lowest at Richmond. The Lantern traps deployed near the Bellbowrie apiary trapped 2,989 SHB over 46 weeks and those at the Gumdale apiary trapped 533 over 33 weeks (Table 26). Lantern traps deployed at Richmond trapped 118 SHB during two seven-week periods in early autumn and spring of 2017 (Table 26). Combining the in hive check counts (Figure 32) with the lantern trap counts, the total number of SHB trapped in and around the Bellbowrie apiary was 5148 over a 15-month period, while 1327 were trapped in the Gumdale apiary over the same period. The overall number trapped at Richmond (255) was much lower, and the trapping rate was also much lower. These data suggest that SHB populations around each of the three apiaries were very different. Despite this difference the overall seasonal patterns of SHB trapping were similar.

Table 26: Weekly trapping rates of SHB (SHB/trap/week) trapped in external attractant traps at three locations from late 2016 until November 2017.

Location Dates of trapping Time (weeks) # SHB trapped SHB/trap/week

11/11/16 - 22/12/16 6 356 14.83

23/12/16 - 26/03/17 11.5 1437 17.85 Bellbowrie 27/03/17 - 08/06/17 10.5 358 4.26

18/07/17 - 22/11/17 18 706 4.90

29/11/16 - 09/05/17 21 285 1.70 Gumdale 30/08/17 - 14/11/17 12 248 2.58

16/03/17 - 05/05/17 7 76 1.36 Richmond 30/09/17 - 18/11/17 7 42 1.29

The seasonal variation in numbers of SHB trapped externally with lantern traps at the three sites paralleled the variation in SHB numbers in the hives accounted for through hive checks in both the Bellbowrie and Gumdale apiaries (Figure 36). SHB populations built up during spring to a peak in late spring - early summer before declining then peaking again in late summer - early autumn at both the Bellbowrie and Gumdale sites (Figure 36). This is consistent with reports by de Guzman et al. (2011) who found that the numbers of SHB caught in their external traps built up to a peak in late spring, dipped in mid-summer, before peaking again in late summer, and then dropping off through autumn to a minimum through winter. A previous study by de Guzman et al. (2010) recording adult SHB in bee colonies found adult SHB most abundant in bee colonies in early autumn. Annand (2011) also found the numbers of SHB in hives built up at the end of summer to their highest peaks in autumn. These findings suggest that the optimal times for intensive trapping with an external trap are during early spring as new SHB move into hives and during late summer - early autumn as the SHB from the summer reproductive cycles begin emerging from the soil searching for bee colonies.

The relative proportion of SHB trapped externally to the number counted in hives (hive check) varied across the sites and time of year. In Gumdale, the proportion from 29/11/16 to 9/5/17 was 0.43, but from 30/8/17 to 14/11/17 improved to 0.88 (Figure 36). The increase in trap efficacy at Gumdale was most likely due to a change in trap placement. Two sets of traps were placed at four ordinal points

around the apiary but during the later period were moved to 20 m and 100 m from the apiary. The difference in Bellbowrie might reflect that trapping is more effective when the air temperatures are higher as this will help volatile release. At Bellbowrie from 22/11/16 to 8/6/17, the relative proportion of SHB trapped externally to in hives was 0.95, suggesting that the yeast fermentate traps were quite efficient (Figure 36). However, this proportion dropped from 18/7/17 to 22/11/17 to 0.51. The numbers trapped externally did not begin to increase again until late September early October when the daily temperatures started to increase. The dispersal rate of volatiles is affected by temperature. The Gumdale apiary is closer to the coast so experiences milder temperatures than Bellbowrie to the west of Brisbane.

500 Hive check Lantern traps 450 A 400 350 300 250 200 150

100 Number SHB counted SHB Number 50 0

Hive check Lantern traps 180 B 160 140 120 100 80 60

40 Number SHB counted SHB Number 20 0

Date

Figure 36: SHB trapped in external attractant traps (lantern traps) and in the hives (hive check) in the (A) Bellbowrie and (B) Gumdale apiaries over 15 months.

Temperature is a key abiotic factor influencing every stage of the SHB life cycle (Schmolke, 1974, Neumann et al., 2001, de Guzman and Frake, 2007, Meikle and Patt, 2011). Total numbers of SHB trapped in hives and externally over 12 months illustrated the relationship between temperature and seasonal pattern of SHB numbers (Figure 37). de Guzman et al. (2010) measured an increase in beetle population in hives during periods when ambient temperatures were ≥ 27°C. We compared maximum daily temperatures and minimum daily temperatures with the number of SHB trapped over 12 months (Figure 37) and found that minimum daily temperatures more closely matched the change in SHB numbers. SHB numbers appeared to begin to decline once the minimum daily temperature

starts to drop to consistently below 20°C, and then appeared to begin to increase when the minimum daily temperature started to climb above 12°C. We found this increase to coincide with daily maximum temperatures around 27°C corroborating the findings of de Guzman et al. (2010). In addition, low daily minimum temperatures will negatively affect the dispersal capacity of a volatile attractant lure, decreasing trap efficacy.

Total SHB Min temp 700 A 30

600 25

C) ° 500 20 400 15 300 10 200

5 ( temp min Daily Number SHB counted SHB Number 100

0 0

300 30 Total SHB Min temp

250 25

C) ° 200 20

150 15

100 10 Daily min temp ( temp min Daily

Number SHB counted SHB Number 50 5

0 0

Date Figure 37 : Relationship between total numbers of SHB trapped inside hives (through hive check) and in external attractant traps the (A) Bellbowrie and (B) Gumdale apiaries and minimum daily temperature.

B Laboratory emergence trigger investigations suggested that a rainfall event after dry periods will stimulate SHB to emerge from the soil. It was noted in the Bellbowrie apiary that few SHB were trapped once maximum daily temperatures climbed well above 30°C and the ground dried after a period without rain. However, once a downpour of rain occurred, a large number of SHB could be trapped in the following days. As is common for many insects, if it was raining in the afternoon into the night, no SHB appeared to be flying. The interplay between increase in SHB populations and rainfall events during summer into early autumn can be seen in data from both Bellbowrie and Gumdale (Figure 38).

Total SHB Rainfall 700 A 80 600 70 60 500 50 400 40 300 30 200

20 Daily rainfall (mm) rainfall Daily

Number SHB counted SHB Number 100 10

0 0

300 Total SHB Rainfall 80 70 250 B 60 200 50

150 40

30 100 20 (mm) Rainfall

Number SHB counted SHB Number 50 10

0 0

Date

Figure 38: Relationship between total numbers of SHB trapped inside hives (through hive check) and in external attractant traps at the (A) Bellbowrie and (B) Gumdale apiaries and daily rainfall.

The use of simple lantern traps with a natural fermentation attractant along with in-hive trapping have provided important seasonal data on the field movement of SHB along with changes in SHB populations in hives. This information is critical for developing future trapping strategies with an external trap and attractant customised for SHB. However more seasonal data would allow for an even better understanding of SHB ecology under Australian conditions.

3.7.3 Synthetic lure testing/evaluation

Based on the successful trapping of beetles in the laboratory with synthetic blends of pure compounds, a series of pilot studies of very limited extent were used to test their efficacy under field trapping conditions.

In four weeks each of trapping with blend #6 and #8, using wicks as the release mechanism, in lantern traps with diatomaceous earth as a killing agent, only one SHB was captured. This was in a

50% blend #6 at the Gumdale apiary. The efficacy of this trap/release rate combination was considered inadequate for further use.

These results suggested that further laboratory testing of the mechanism was required. Results of laboratory release rate studies of these blends using cotton wicks (blend #1, and #4), water absorbing crystals (Eden® Water Storage Crystals), agar (1.5% in H2O) (blend #4) and glycerine (blend #10) found that none of these methods supported an adequate release rate of the volatile components.

Over 76 hrs, release rate through the cotton wicks (blend #1 = 0.10 ± 0.02 g/h, blend #4 = 0.18 ± 0.03 g/h) was considered insufficient to produce enough of an odour source to be viable in a natural situation. Within twelve hours, all liquid had evaporated from the water-absorbing crystals.

After 48 hrs, while most of the original compounds of blend #4 were still detectable in the agar headspace, the concentration had dropped dramatically and relative proportion of the components had shifted (Figure 39). Using the BugDorm bioassay to assess the attractiveness of this mixture, the trapping rate was much lower than that seen in other assays (73.24 ± 2.83 % beetles untrapped). Of the trapped beetles, significantly more were trapped in control than treatment traps (Mann-Whitney: U = 1.5, P = 0.006).

Figure 39: Total ion chromatograms of headspace of blend #4 in agar (1.5% in H2O) at time zero (above) and after 48 hours (below) showing differences in compounds detected. Key compounds are identified by number.

1. ethanol 4. ethyl caproate 2. ethyl acetate 5. nonanal 3. isopentyl acetate 6. decanal

The positive control (yeast/honey mix) yielded a significantly stronger response in suspended lantern traps than Blend #10 in baggies (U = 11.0, P = 0.028) (Table 27).

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Table 27: Results of large tent two choice-assays (vs. control) with adult SHB responding to baggie mechanism of blend (#10) release comparative to positive control (yeast in lantern trap).

Release mechanism Mean % (± SEM) Test statistic Preference in treatment trap

baggies 45.81 (± 0.14) U = 1.0, P < 0.001 Blend #10

positive control 63.02 (± 4.2) U = 0.0, P < 0.001 Yeast mix (yeast/honey mix)

Aging of blend #10 in lantern traps for five days in the field completely changed the relative proportions of the various components (Table 28), with some of them now below the detection limit of the GC-MS. In addition, there were extra compounds detected in the headspace of these solutions that were not a part of the original blend. Without the original complexity of the blend, all of these release methods were also considered inadequate for extensive field testing.

Table 28: The percentage composition of each compound in blend #10, and levels detected after five days in traps with glycerine

Compound % in original blend % detected after 5 days in glycerine

ethanol 90.0 3.0

acetaldehyde 1.0 -

ethyl acetate 2.0 5.6

isobutanol 4.0 22.5

isopentanol 3.0 68.7

isopentyl acetate - 0.17

isopentyl isovalerate - 0.07

Although limited, the only synthetic blend that demonstrated promise was the release of blend #4 from zip lock baggies in lantern traps. Over two weeks of field testing in November 2017 at the Gumdale apiary, two small hive beetles were captured in traps with this combination. Although not very dramatic, it was only blend #4 lures that captured any beetles over this period, with no beetles trapped in either control traps or those containing blend #10. This limited success warrants further investigation and optimisation.

Even though a synthetic blend is shown to be attractive to SHB under laboratory conditions if the release mechanism does not allow for the synchronous release of the various components it will be

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ineffective in the field. Further development of a suitable release mechanism for the blends under investigation is critical for the success of an external attractant trap. 3.8 Educational video

An educational video was produced and has been uploaded for public viewing: https://www.youtube.com/watch?v=YHUmK5SlzXU&t=

The video describes how to make up and deploy a simple attractant trap to catch SHB, with Brisbane based beekeeper, Phil Bowman of Stradbroke Island honey describing and demonstrating the process. In the video Phil shows the trap, which is readily available from most hardware stores, and the ingredients, which are also readily available from grocery stores, then demonstrates how to mix the ingredients and add to the traps. How and where to hang traps in relation to an apiary is shown, then the collection of traps and removal of SHB from the traps is shown. In addition to Phil describing the recipe for the attractant, it is shown as an overlay on the video. This video should provide enough information for a keen beekeeper to be able to make up and deploy these simple traps to aid in the management of SHB.

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Implications

While eradication of SHB from Australia is now impossible it is essential that beekeepers are provided with the information and tools to maintain SHB populations in their hives at low levels. Hives carrying large numbers of SHB are at risk of a collapse when conditions are met for their mass reproduction. Subsequent infestation by thousands of destructive larvae with associated fermentation of honeycomb causes the entire hive to become slimed. Recent surveys conservatively estimated the annual damage from SHB to range from $2 million to $11 million in Queensland alone. The life cycle of SHB includes an external transition phase from fully developed larvae through pupae to adults. Small hive beetle numbers build up in hives through external recruitment. To maintain low numbers of SHB in hives it is imperative to stop as many as possible from entering hives.

As part of the development of an external trap information needed to be gathered about the seasonal movement of SHB including factors driving this movement. In addition, a suitable trap design and understanding of the optimal trap placement for SHB trapping was required.

Each of the eight objectives of this project has been met with some being exceeded. This project has developed an external trapping system that has laid the foundation for future research to refine this system into a more efficacious system for intercepting and trapping SHB flying into apiaries.

Fermentation of hive products sourced from a range of locations where SHB are known to cause damage showed that the starting materials affected the amount and type of volatiles produced. The amount and quality of protein and carbohydrate in the hive materials likely affect the fermentation rate, and thus the chemical composition of the resultant fermentation products including the volatiles. This variability will impact on SHB attractants using natural products such as fermented pollen dough or slime. Consistency in an attractant lure is important and will only be gained by using a blend of the pure compounds that are key to the attractiveness of the natural products to SHB.

Regardless of the source of the hive materials we found slime was only attractive when ten key compounds were present in specific proportions. Many of these compounds have previously been found to be attractive to insects, with some already used in attractive blends. Most, if not all, of these compounds are known to be products of yeast fermentation. However, maximal attractiveness to an individual insect species rests with the relative proportion of the compounds and the overall concentration of the blends.

Different insect species might find the same compounds (yeast derived) attractive but they will prefer the compounds in different proportions. In addition, some compounds can be attractive at one concentration but repulsive to the same insect if the concentration is increased. Finding the specific proportions of the key attractive compounds that SHB find most attractive could involve using a matrix. To examine all possible blends of the ten attractive compounds identified as well as ascertaining the concentration at which they are most attractive to SHB would involve an enormous number of possibilities, requiring years of work. Therefore, a targeted approach was taken to deliver results in a timely manner. A blend with all ten compounds will likely be far too expensive, therefore a lower number of compounds from the list of ten needed to be chosen without compromising attractiveness. A further consideration affecting the choice of compounds was the need to match compounds with compatible volatility. Chemical compounds vary widely in their volatility or the rate at which they will evaporate. The correct proportions of the selected compounds need to be maintained as they are emitted from the lure in the field. Combining compounds where one or two evaporate

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within days while others take weeks to evaporate will markedly change the nature of the odour signal received by the target insect. Working with a smaller number of compounds for a lure is not only more economical but makes it easier to match the vapour pressure of the compounds affecting the emission from the lure.

Informed choice was used to design the blends for testing, with iterative bioassays guiding the process. The choice of compound and proportion was determined by the proportions of compounds in highly attractive fermentation products, lures used commercially and research in the USA. A range of concentrations of each blend was then tested.

Considerable progress has been made towards finding an optimum blend of yeast compounds attractive to SHB and satisfying all the conditions required for a good synthetic lure. A variety of blends have been tested at different concentrations, and four blends attractive to SHB in the laboratory were chosen for field testing as lures. However, the jump from laboratory to field is difficult especially when laboratory testing is limited to distances less than four metres and field testing requires SHB attractive odours to be active over at least a kilometre. More work is needed to find a more attractive blend with greater field activity.

Both a suitable trap design and release mechanism for the attractant odours are critical for the testing of an attractant lure. The initial scope of this project did not cover developing a customised trap for SHB, therefore a number of readily available designs were investigated. The only design that showed promise was the lantern trap retailed for trapping house flies. However, experience with these traps suggested that the design needs to be customised for SHB by increasing the capacity for volatiles to exit the trap and preventing any SHB exiting the trap once they have entered. A design modification to either kill all SHB entering the trap or at least prevent the exit of any SHB once they have entered is particularly important for use of this trap style with a synthetic lure. In addition, the design needs to incorporate a suitable release mechanism to allow the synchronous release of the various components of the attractant lure. Six different release mechanisms were trialled but none of them appeared suitable for use with the trapping system tested.

Investigations into the occurrence of an aggregation pheromone produced by SHB clearly demonstrated that they consistently aggregate under a range of conditions. The research also supports volatile compounds being involved in the aggregation. While chemical analyses have not revealed any putative aggregation compounds more work is required in this area. The research to find the aggregation compounds used by Carpophilus beetles took many years and a lot of investment. This beetle has a very high economic impact across a number of high value orchard crops. Although there is potential to markedly increase the efficacy of an attractant trap with an aggregation pheromone, it is doubtful that the cost benefit for the small hive beetle will support years of research.

Through this project we developed a methodology to examine differences in hive volatile profiles to ascertain if there are differences which could be accounted for by very high numbers of SHB. We were unable to clearly identify any relationship between SHB load and hive volatiles because there was not enough variation in SHB load within the apiaries where the volatiles were collected. There is likely a wide range of factors influencing the volatile profile of any given hive including floral source, current bee population and makeup (i.e. number or drones and number and age of workers), disease status and queen status. The data generated in this project showed the complexity of hive volatiles and produced background information on the volatile profile for healthy hives that could be used in the future for comparison with hives with a particular pest or disease load. There may even be potential

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for comparisons with the volatiles coming from a hive producing queen pheromone or one that has gone queen-less.

Synthetic attractant lures for SHB were evaluated in two apiaries in Queensland but did not show the same attractiveness as they did in the laboratory. As noted above both the trap design and release mechanism could have impacted on this performance.

While making progress towards the synthetic attractant lure external attractant traps consisting of a natural fermentation attractant and simple lantern traps were used to gather important ecological data on SHB movement in three apiaries in Queensland and New South Wales. This study also developed an easy method for estimating the number of SHB in a hive and thus any changes in SHB numbers in a hive that might be influenced by either a large influx of SHB or successful interception of SHB with an external trap. Trapping data (in hive and external) showed that seasonal changes in SHB numbers at the three locations are primarily influenced by temperature with rainfall as an important influence through the hot seasons.

The results of this study suggest that an effective trapping strategy would involve deployment of external attractant traps for SHB from mid to late spring when maximum daily temperatures rise to 27°C and minimum daily temperatures rise above 12°C. The efficacy of attractant traps set out too early in spring will be compromised if temperatures are too low for volatiles to be released from the lure. Conversely, deployment of traps in late autumn will also be dependent on average daily temperatures. During summer, as the daily temperatures rise above 30°C rainfall has an impact on SHB movement via soil moisture levels. Small hive beetle movement decreases during periods of hot dry weather but immediately resumes in the days following a downpour of rain, when traps should again be deployed.

This research showed that traps can be deployed from close to hive or as far away as 200 m. It could be possible to trap even further from a hive in early spring but this was not tested in this project. Small hive beetles were trapped in areas away from any known hives although trapping efficacy was much greater within 200 m of hives suggesting that the traps act to intercept SHB heading to an apiary.

The trapping data from this project is essential for developing future trapping strategies with a synthetic attractant lure and customised external trap for SHB.

An educational video was produced to explain how to make up and use the lantern traps using a honey-yeast fermentate to trap SHB. This external attractant trap caught nearly 3,000 SHB over 12 months in one apiary during this project. While not the endpoint in the development of an external attractive trap using a synthetic lure for SHB, it has successfully intercepted beetles and prevented a substantial number entering hives.

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Recommendations

 Seasonal data on the weekly and fortnightly changes in numbers of SHB trapped in the field and in hives suggests these changes are primarily influenced by temperature with rainfall as an important influence through the hot seasons. This information is the most comprehensive Australian ecological data on SHB yet generated and can inform strategic trapping programs using the simple yeast traps currently available or a customised SHB trap with synthetic attractant lure once developed. Furthermore, it has been demonstrated that traps placed anywhere up to 185 m from hives can be effective at intercepting SHB flying towards an apiary. Further research is needed to investigate distances greater than 185 m from hives and gather more data on the effect of direction and the effect of prevailing winds on the trap efficacy.

 Further funding is required to continue research to refine and test the blend of compounds attractive to SHB to produce a synthetic lure. Research also needs to be undertaken to produce a release mechanism to optimise the release rate of the components of the synthetic lure with additional research warranted to customise a trap for small hive beetles that incorporates a killing system for SHB and the synthetic lure.

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Glossary

Abiotic Physical rather than biological, not derived from living organisms

Absorption The uptake of a substance by another

Adsorption The adhesion of a substance to a solid surface

Aeration The sampling of air for analysis

Aggregation Volatile compound interpreted by insects as a cue to form groups pheromone

Antennectomy The removal of antennae

Aspirate Drawing up an air sample via suction

Eclosion For an insect: the act of emerging from a pupal case or hatching from an egg

Emergence When insects come out of the ground as adults after their pupal stage

Eversion The process of turning inside-out

Fermentate A product made by fermentation

Headspace The air above samples

Instar An insect growth stage (between two successive moults) The number of instars in the development of insects varies with species.

Mass A device used to determine the masses of the compounds in a sample spectrometer

Olfactometer An instrument used to detect and measure odour dilution. In the application in these studies it is a device used to measure the reaction of insects to an odour source

Oviposition Egg-laying

Ovipositor A tubular organ through which a female insect deposits eggs

Phenotype The observable characteristics of an individual

Slime Fermenting hive-product, progressively altered by the action of small hive beetles and yeast over time

Teneral Adults freshly emerged from pupae, before their exoskeletons have hardened

Tergite A hardened plate usually on the back of an insect, especially of the abdominal section

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Thermal The process by which heat is used to remove compounds from a solid desorption substance

Volatile Compounds that can easily become vapours or gases compounds

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112 External attractant trap for small hive beetle by D M Leemon, R A Hayes, B A Amos, S J Rice, D K Baker, K McGlashan February 2018

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