EFFECTS OF THIACLOPRID IN COMBINATION WITH … · 2013). In addition, plant-pollinator interactions...

EFFECTS OF THIACLOPRID IN

COMBINATION WITH TEBUCONAZOLE

ON DIFFERENT DEVELOPMENTAL

STAGES OF OSMIA CORNUTA Word count: 12175

Ruben Vanderhaegen Student number: 01101110

Promotor: Prof. dr. ir. Guy Smagghe Tutor: ir. Maxime Eeraerts Master’s Dissertation submitted to Ghent University in partial fulfilment of the requirements for the degree

of Master of Science in Environmental Sanitation and Management.

Academic year: 2016 - 2017

EFFECTEN VAN THIACLOPRID IN

COMBINATIE MET TEBUCONAZOOL

OP VERSCHILLENDE

ONTWIKKELINGSSTADIA VAN OSMIA

CORNUTA Aantal woorden: 12175

Ruben Vanderhaegen Stamnummer: 01101110

Promotor: Prof. dr. ir. Guy Smagghe Tutor: ir. Maxime Eeraerts Masterproef voorgelegd voor het behalen van de graad Master of Science in de milieusanering en het

milieubeheer

Academiejaar: 2016 - 2017

i

Preface

“People worry about rhinos and polar bears, but they rarely are too bothered by the fact that we are losing some of our smaller wildlife quite rapidly, and actually they should be really

worried about that”

Dave Goulson

Bees are our main providers of fruits and vegetables. They are the source of the picnics’ divine characteristics: strawberries and apple juice. Their extraordinary plant pollinating capacities reach worldwide. Their buzz can give some a creep, others a feeling of sensation. They are friendly, not resembling that one imaginary black and yellow like creature with a stinger. They are part of the invisible, but crucial for our own well-being. Don’t expel bees, but discover and cherish them.

Sadly, bees face great difficulties worldwide. A lack of suitable flowers leave them defenseless against death and reproductive failure, and the remaining flowers in their small habitat patches are likely to be contaminated with all kinds of chemicals. It is not even certain that indigenous plants you buy in your own gardener center, specifically meant for bees, are free of pesticides. It has been proven that a multitude of pesticides are deadly for bees. Adult bees consume the nectar and pollen of flowers which are treated with pesticides, and feed these nectar and pollen to their progeny. Imagine you are a larvae of a solitary bee. You are in a completely dark cell. Your only option to survive is to consume the pollen your mother bee prepared for you. Spin a safe cocoon and wait. Several months. Develop. Let the magic proceed. Give the pesticides a chance to attack. You might emerge. You might find flowers and mate. You might find a nest to secure your progeny. Hopefully you are not defenseless.

Ironically, in orchards, bees are necessary to pollinate the fruit trees, but at the same time pesticides are sprayed which possibly kill these bees. Therefore, it is crucial to test the effects of those pesticides on bees. This is done in toxicity testing. This toxicity testing has already led to several laws and regulations which limit the use of harmful pesticides. This is not only good news for the bees’ health, but also for all other animals’, including ourselves’ health. However, we can do better. At this moment, new testing protocols and schemes are being developed to improve toxicity testing. This includes testing new pesticides and mixtures of pesticides on different bee species. It is an attempt to reveal the invisible. I want to learn more about the invisible, and that is how this study, which is a contribution to the growing field of toxicity testing, came into being.

This study would not have been possible without the help and provisions of the Laboratory of Agrozoology (Crop protection) of the Faculty of Bioscience Engineering at Ghent University. While professor Guy Smagghe supervised the study, Maxime Eeraerts guided me through the toxicity tests, assisted in analyzing the data and provided general feedback. Thank you both, I am very grateful. I also thank Sander Ostyn, my colleague during a part of this study, for assisting in setting up the adult oral toxicity test. In addition, I would like to thank my Snoebie and father for proofreading my manuscript.

iii

Table of contents

Preface ......................................................................................................................................... i

List of abbreviations ................................................................................................................... v

Abstract .....................................................................................................................................vii

Samenvatting ............................................................................................................................ viii

1. Introduction ........................................................................................................................ 1

2. Material and methods ........................................................................................................ 4

2.1. Study species ............................................................................................................... 4

2.2. Pesticides ..................................................................................................................... 6

2.3. Available protocols for adult and larval oral toxicity testing on solitary bees ............ 8

2.3.1. Available protocols for adult oral toxicity testing ................................................ 8

2.3.2. Available protocols for larval oral toxicity testing ............................................... 9

2.4. Experimental procedure ............................................................................................ 10

2.4.1. Protocol for adult oral toxicity testing ............................................................... 10

2.4.2. Protocol for larval oral toxicity testing ............................................................... 12

2.5. Statistical analyses ..................................................................................................... 16

2.5.1. Mortality ............................................................................................................. 16

2.5.2. Normalized food consumption........................................................................... 17

3. Results .............................................................................................................................. 18

3.1. Adult oral toxicity test ............................................................................................... 18

3.1.1. Mortality ............................................................................................................. 18

3.1.2. Normalized food consumption........................................................................... 18

3.2. Larval oral toxicity test .............................................................................................. 18

4. Discussion ......................................................................................................................... 22

4.1. Results and protocol of adult oral toxicity test ......................................................... 22

4.2. Protocol for larval toxicity testing ............................................................................. 23

5. Conclusion and future research ....................................................................................... 27

6. Bibliography ...................................................................................................................... 29

Appendix A ............................................................................................................................... 34

Appendix B ............................................................................................................................... 40

v

List of abbreviations

EFSA European Food Safety Authority

ERA Ecological Risk Assessment

EU European Union

ICPPR International Commission for Plant Pollinator Relationships

PPP Plant Protection Products

vii

Abstract

Global crop production is for 35% dependent on pollinators. A prominent group of these

pollinators are solitary bees. Unfortunately, over the last decades solitary bee populations

have declined, partly due to the use of pesticides. Pesticides are commonly applied worldwide

to reduce crop pests. Subsequently, nectar and pollen of crops are major potential sources of

pesticide exposure to different developmental stages of bees. Effects of pesticide

combinations especially seem to cause detrimental effects on bees and their larvae. In the

ecological risk assessment framework concerning plant protection products in the EU

(Regulation (EC) 1107/2009), no official test protocols are available for solitary bees. Hence, a

working group of the International Commission for Plant Pollinator Relationships – the “non-

Apis” working group – was established in 2013, with their main goal set to develop

standardized testing methods for non-Apis bees. To date, protocols of toxicity tests on solitary

bees are already described, but further ring-testing and validation has to be done.

In this study, oral toxicity tests on adult females and larvae of the solitary bee Osmia cornuta

were conducted, using the neonicotinoid thiacloprid and the fungicide tebuconazole as testing

compounds. The aims were to assess effects of exposure to thiacloprid, tebuconazole and to

their combination on both adult females and larvae of Osmia cornuta. Thereby also

contributing to the development of a standardized protocol for both adult and larval oral

toxicity testing on solitary bees.

This study found evidence of sublethal effects caused by exposure to the combination of

thiacloprid and tebuconazole in the adult oral toxicity test. Exposure to only thiacloprid and

tebuconazole showed no lethal nor sublethal effects in the adult oral toxicity test. This finding

pressurizes the regulatory risk assessment framework to also include toxicity tests on mixtures

of pesticides. No results of the larval oral toxicity test were obtained as this test was still

running at the end of this study.

This study also indicated that the protocol used for the adult oral toxicity test resulted in high

O. cornuta mortality during the first three days of exposure in all treatments, including all

control groups. This high mortality was probably caused by the non-feeding behavior of bees,

resulting in starvation. The protocol used in this study should therefore be revised to

overcome this non-feeding problem.

Furthermore, the protocol for larval oral toxicity testing in this study proved to be successful.

However, it also indicated that the major problem in designing a standardized protocol for

larval oral toxicity testing includes administration of equal concentrations of the test solutions

to each larva. Therefore, efforts should be made to further optimize homogenous

administering of the test solutions to each larva.

In conclusion: optimization of the protocols on toxicity testing is required to ensure the

protection of our pollinators, and ourselves.

Keywords: Osmia cornuta, insecticide, fungicide, oral toxicity test, larvae, laboratory rearing

viii

Samenvatting

Bestuivers zijn verantwoordelijk voor 35% van de wereldwijde productie van gewassen. Van

deze bestuivers zijn solitaire bijen een belangrijke groep. Helaas zijn solitaire bijen populaties

de laatste decennia sterk in aantal afgenomen, mede veroorzaakt door het gebruik van

pesticiden. Pesticiden worden wereldwijd toegepast ter bescherming van de gewassen.

Hierdoor zijn nectar en pollen van gewassen grote potentiële bronnen van blootstelling aan

pesticiden voor de verschillende ontwikkelingsstadia van bijen. Het lijken vooral effecten van

combinaties van pesticiden te zijn die schadelijk zijn voor bijen en hun larven. In het kader van

de ecologische risicobeoordeling van gewasbeschermingsmiddelen in de Europese Unie

(Regulation (EC) 1107/2009) zijn er nog geen officiële protocollen voor toxiciteitstesten op

solitaire bijen beschikbaar. Daarom is er een werkgroep van de ‘International Commission for

Plant Pollinator Relationships’, de “non-Apis” werkgroep, opgericht in 2013. Deze groep heeft

als hoofddoel het ontwikkelen van een gestandaardiseerde toxiciteitstest voor niet-Apis bijen.

Er zijn al protocollen van toxiciteitstesten op solitaire bijen beschreven, maar meer ringtesten

zijn nodig.

In deze studie werden orale toxiciteitstesten op volwassen vrouwtjes en larven van de solitaire

bij Osmia cornuta (gehoornde metselbij) uitgevoerd met het neonicotinoïde ‘thiacloprid’ en

het fungicide ‘tebuconazool’. Het doel was om effecten van blootstelling van thiacloprid,

tebuconazool en hun combinatie op volwassen vrouwtjes en larven van Osmia cornuta vast te

stellen. Hierdoor werd ook bijdragenen aan de ontwikkeling van een gestandaardiseerd

protocol voor toxiciteitstesten op zowel het volwassen als larvaal stadium van solitaire bijen.

In deze studie werd er een sublethaal effect aangetoond op volwassen bijen dat veroorzaakt

werd door blootstelling aan de combinatie van thiacloprid en tebuconazool in de volwassen

orale toxiciteitstest. Noch lethale, noch sublethale effecten op volwassen bijen werden

gevonden na blootstelling aan enkel thiacloprid en tebuconazool. Dit resultaat benadrukt dat

toxiciteitstesten op combinaties van pesticiden opgenomen zouden moeten worden in het

ecologische risicobeoordelingskader.

Helaas konden geen resultaten van de orale toxiciteitstest op het larvale stadium bekomen

worden aangezien deze test nog lopende was aan het einde van deze studie.

Het protocol dat in deze studie gebruikt werd voor de orale toxiciteitstest op volwassen bijen

veroorzaakte een hoge mortaliteit tijdens de eerste drie dagen van blootstelling in alle

behandelingen, inclusief de controles. Deze hoge mortaliteit werd waarschijnlijk veroorzaakt

doordat bijen zich niet leken te voeden, waardoor ze verhongerden. Daarom zou dit protocol

zo aangepast moeten worden dat bijen zich wel gaan voeden.

Het gebruikte protocol voor de orale toxiciteitstest op het larvale stadium in deze studie

toonde aan succesvol te zijn. Echter, het gaf ook weer dat het grootste probleem in het

standaardiseren van het protocol omtrent larvale toxiciteitstesten het toedienen van gelijke

concentraties van de testoplossing aan elke larve is. Daarom moeten er extra inspanningen

ix

geleverd worden om een protocol te ontwikkelen dat een toediening van gelijke concentraties

aan elke larve omvat.

In conclusie: optimalisatie van de protocollen in toxiciteitstesten is noodzakelijk om de

bescherming van onze bestuivers, en onszelf, te garanderen.

Trefwoorden: Osmia cornuta, insecticide, fungicide, orale toxiciteitstest, larven, opkweken

solitaire bijen

1

1. Introduction

Global crop production is for 35% dependent on pollinators (Klein et al., 2007), which provides

a total economic value of € 153 billion, representing 9.5% of the value of the global agricultural

production for human consumption in 2005 (Gallai et al., 2009). This global crop production is

furthermore enhanced by high pollinator diversity (Garibaldi et al., 2011; Garibaldi et al.,

2013). In addition, plant-pollinator interactions are of vital importance for the preservation of

the functional integrity of terrestrial ecosystems (Kearns et al., 1998; Ollerton et al., 2011), as

the reduction in pollinators may lead to reductions in pollinated plant species (Biesmeijer et

al., 2006).

A prominent group of these pollinators are bees, consisting of (managed) honey bees (Apis

spec.), bumblebees and solitary bees. Over the last decades bee populations have declined –

sometimes severely –, hereby posing economic and ecological implications. Managed honey

bee colonies have experienced substantial losses in North-America (Lee et al., 2015; Seitz et

al., 2016; Vanengelsdorp et al., 2008), South-Africa (Pirk et al., 2014) and Europe (Potts et al.,

2010a). Wild pollinators such as bumblebees have undergone substantial range contractions,

population declines and (localized) extinctions in Europe (Goulson et al., 2008; Kosior et al.,

2007), North-America (Goulson et al., 2008; Williams and Osborne, 2009), South-America

(Schmid-Hempel et al., 2014) and Asia (Inoue et al., 2008; Xie et al., 2008). Furthermore,

declines in solitary bee species in Europe have also been reported (Biesmeijer et al., 2006;

Carvalheiro et al., 2013).

Multiple drivers of bee declines have been put forward. The most important ones include

habitat loss and fragmentation (with consecutive limitation in floral resources and nesting

sites), parasites and pathogens, climate change, introduction of invasive alien species and the

use of pesticides (Goulson et al., 2015; Potts et al., 2010b and references therein). It should

not surprise that several of these drivers can act in combination.

Pesticides, including insecticides, herbicides and fungicides, are commonly applied worldwide

to reduce crop pests. Crop pests cause a reduction in potential crop yield of on average 35%

worldwide (Oerke, 2006). It can be stated that the correct use of pesticides leads to beneficial

socio-economic and environmental effects, but incorrect use may cause increased pest

resistance, adverse human health effects and loss of pollinators such as bees (Popp et al.,

2013). Furthermore, it is known that bees are exposed to cocktails of pesticides during their

different developmental stages (Mullin et al., 2010). Hence, the challenge remains to

understand and control the impact of pesticides on non-target species such as bees.

Neonicotinoids, a major class of insecticides, are used extensively to control crop pests by

spraying, as seed coatings and as soil additives (Blacquiere et al., 2012; Jeschke et al., 2011).

Their systemic nature results in translocation of the neonicotinoid to all plant tissues.

Consequently, in combination with their high toxicity to insects, they protect all parts of the

plant for a long period of time (van der Sluijs et al., 2013). Furthermore, neonicotinoids are

highly toxic to insects because they are neurotoxins that act as agonists of the insect nicotinic

2

acetylcholine receptors (nAChR), hereby causing overstimulation, paralysis and death

(Goulson, 2013). Next to the systemic nature of neonicotinoids, they are also very persistent

in the environment. Therefore, they are found in nectar and pollen of treated crops

(Blacquiere et al., 2012; David et al., 2016). Subsequently, nectar and pollen are major

potential sources of neonicotinoid exposure to different life stages of bees. Indeed, all bees

primarily feed on nectar and pollen and provide, in varying amounts, their offspring with it

(Peeters et al., 2012a). For example, solitary bee larvae feed almost exclusively on pollen,

which might result in very high exposure to neonicotinoids.

Several studies have reported on the acute and chronic toxicity of neonicotinoids (Blacquiere

et al., 2012 and references therein), as well as on their potential to cause sublethal effects on

bee species. For example, neonicotinoid exposure reduced wild bee density (Henry et al.,

2012; Rundlof et al., 2015), solitary bee reproductive success (Rundlof et al., 2015; Sandrock

et al., 2014), bumblebee (Bombus terrestris) colony growth, reproduction (Rundlof et al.,

2015; Whitehorn et al., 2012) and foraging performance (Gill et al., 2012) and honey bee (Apis

mellifera) foraging success (Henry et al., 2012). It was this kind of research that highlighted

the detrimental effects of neonicotinoids on wild bees and led to the banning of three

neonicotinoids in the EU from 1 December 2013 onwards (EU, 2013).

Besides insecticides such as neonicotinoids, fungicides are also widely applied to crops, with

the potential of exposure to bees. Especially synergistic effects of neonicotinoids and

fungicides seem to cause detrimental effects on honey bees and their larvae (Iwasa et al.,

2004; Schmuck et al., 2003), rather than the sole effect of fungicides. However, there is a lack

of research on these synergistic effects, especially on non-Apis bees. Therefore, understanding

the effects of simultaneous exposure to neonicotinoids and fungicides remains a major

challenge.

Honey bees are used in the ecological risk assessment (ERA) framework concerning the placing

of plant protection products (PPPs) on the market in the EU (Regulation (EC) 1107/2009). In

this regulatory framework, they also serve as surrogates for non-Apis pollinator bees, although

recently this has been greatly challenged (Arena and Sgolastra, 2014; Uhl et al., 2016). In the

recent guidance document of the European Food Safety Authority (EFSA), risk assessment of

PPPs on honey bees, bumblebees and solitary bees was included (EFSA, 2013). However, no

official test protocols are available for non-Apis bees. Hence, a working group of the

International Commission for Plant Pollinator Relationships (ICPPR) – the “non-Apis” working

group – was established in 2013. The main goal of the working group is to develop

standardized testing methods for bumblebees and solitary bees and thereby fill the gap in the

current tier testing systems. These standardized testing methods include both methods for

toxicity testing on adult bees as on larvae. The Laboratory of Agrozoology of the Faculty of

Bioscience Engineering at Ghent University is an active member of this ICPPR working group.

In addition, the regulatory risk assessment framework only focuses on single pesticides,

whereas effects of mixtures of pesticides are not included. This despite the evidence of bees

experiencing synergistic effects caused by simultaneous exposure to multiple pesticides (Gill

3

et al., 2012; Iwasa et al., 2004; Schmuck et al., 2003). Furthermore, toxicity testing, both in

laboratory and semi-field settings, should aim for exposing bees to field realistic

concentrations. Otherwise, debates could arise on the validity and relevance of the toxicity

test (David et al., 2016).

The inclusion of solitary bees in this regulatory framework concerning the placement of PPPs

is of major importance. This is because effects of pesticides on solitary bees might be more

pronounced than on honey bees (Rundlof et al., 2015), and solitary bees are more important

pollinators than honey bees (Breeze et al., 2011; Klein et al., 2007). The solitary bees of the

mason bee genus (Osmia spec.) are identified as being suitable testing species in the

regulatory risk assessment of PPPs (EFSA, 2013; Sgolastra et al., 2015). Both contact and oral

toxicity tests on Osmia spec. are described, but further ring-testing and validation has to be

done (EFSA, 2013), as data of toxicity tests on solitary bees is scarce. This is especially the case

for toxicity tests on the larval stadia, but see for example (Abbott et al., 2008; Becker and

Keller, 2016; Konrad et al., 2008; Sgolastra et al., 2015). Toxicity tests on solitary bee larvae

require further improvement, as it may be that larvae, compared to adult bees, are more

susceptible to adverse effects of pesticides, ultimately leading to bee declines.

When it comes to systemic pesticides, such as neonicotinoids, the oral route of exposure,

compared to (in)direct contact, is the most likely. In addition, recent studies indicate that oral

exposure is often the most relevant for bees (EFSA, 2012).

In this study, oral toxicity tests on adult females and larvae of the solitary bee European

orchard bee (Osmia cornuta) were conducted, using the neonicotinoid thiacloprid and the

fungicide tebuconazole as testing compounds. The aims were

(1) to test the proposed protocol of the “non-Apis” working group for acute and

chronic oral toxicity testing for adult solitary bees, and contribute to the

development of standardized toxicity tests

(2) to contribute to the development of a standardized protocol for larval toxicity

testing on solitary bees

(3) to assess effects of exposure to thiacloprid, tebuconazole and to their

combination on both adult females and larvae of Osmia cornuta

4

2. Material and methods

2.1. Study species

This study was performed on the solitary bee species European orchard bee (Osmia cornuta;

Hymenoptera: Megachilidae). It mostly occurs in the southern parts of Europe but also in

North-Africa and the Middle-East (Peters, 1977). The species nests in pre-established cavities,

is univoltine (one generation per year) and gregarious, meaning that individuals live a solitary

life but can nest next to each other, hereby forming a population even though there is no

mutual cooperation or antagonism between individuals (Krunić and Stanisavljević, 2006).

Furthermore, O. cornuta is polylectic. They consume nectar and collect pollen from orchard

trees (e.g. apple, pear, etc), strawberry, blackberry, raspberry, several meadow plants (e.g.

Muscari botryoides, Taraxacum officinalis) but also trees such as oak (Krunić and

Stanisavljević, 2006 and personal observations).

Morphologically, O. cornuta (Figure 2B) is characterized by sexual dimorphism. Females are

larger than males, have shorter antennae, two horns and dark hairs on their face and dense

orange hairs on their abdomen. Males have white hairs on their face and clypeus and long

antennae. Female weight ranges from 0.06 g to 0.2 g (N = 342; mean weight = 0.13 g).

Individuals of O. cornuta emerge from their cocoons with the onset of warmer weather and

blooming of plants, mostly in early spring (Krunić and Stanisavljević, 2006). Males emerge

several days earlier than females and, attracted by their pheromones, they await the females’

emergence at the entrance of the nest (Krunić and Stanisavljević, 2006). Upon emergence, the

females mate immediately with several males (Krunić et al., 1995). After mating, females start

searching for suitable nesting places, which can take up to two days, meanwhile visiting

flowers for their own consumption (Krunić and Stanisavljević, 2006). Personal observations

suggested that mated females prefer south orientated nesting cavities exposed to direct

sunlight, although they might also prefer nesting cavities not exposed to direct sunlight (Krunić

and Stanisavljević, 2006 and Eeraerts M., pers. communication).

In the nesting cavities, females construct a series of cells separated with mud partitions (Bosch

et al., 2008), with the final plug, which conceals the cavity, being significantly thicker than the

other partitions (Stanisavljević, 2000). Once the first partition is built, the female transports

pollen and nectar to the nest and deposits it into the cell (Figure 2C). Personal observations

showed that two females sometimes entered the same nesting cavity, which resulted in a

short but intense fight, after which one took off. It was unclear whether or not she returned

to the same nesting cavity later on. On top of the pollen and nectar mass, hereafter referred

to as ‘provision’, the female lays one egg, as can be seen on Figure 2D. Larger provisions are

supplied to cells that are closer to the base of the nesting cavity, compared to cells closer to

the exit (Bosch et al., 2008). This is because females lay fertilized eggs, from which females

will develop, closer to the base of the nesting cavity, whereas unfertilized eggs, from which

males will develop, are laid closer to the exit (Krunić and Stanisavljević, 2006). Average fresh

weight of female and male provisions in O. cornuta are 0.5 g and 0.25 g, respectively (Bosch

5

and Vicens, 2002). Cocoons vary in size and shape, and O. cornuta cocoons from which females

will develop, are longer and heavier compared to cocoons from which males will develop

(Stanisavljević, 2000 and also see Box 1). After construction of the nests, adult females die and

leave their progeny to develop alone. The total period of activity for adult females of O.

cornuta is three to four weeks (Krunić and Stanisavljević, 2006; Sgolastra et al., 2015).

Eggs hatching to larvae takes only several days (Stanisavljević, 2000). O. cornuta has five instar

larvae. The first larval instar emerges from the egg membrane but does not feed on the

provision. Consumption of the provision and defecation starts only in the second and fifth

larval instar, respectively (Figure 2E) (Krunić and Stanisavljević, 2006). Temperature and other

factors directly determine the duration of the larval stage (Stanisavljević, Lj. Z., personal

communication). In natural conditions, larval consumption takes up to 60 days, after which

the fifth instar transforms into the prepupa (Figure 2F), which spins a cocoon in four to six

days (Figure 2G) (Sgolastra et al., 2015; Stanisavljević, 2000). After up to 50 days, the prepupa

metamorphoses into a pupa which develops by late summer into an adult. This adult

overwinters in the cocoon as a dormant stage (diapause) and emerges again in early spring

(Bosch et al., 2008; Krunić and Stanisavljević, 2006).

Box 1

In this study, an additional small scale experiment was conducted to investigate potential

differences in weight and length of female and male cocoons of O. cornuta. For this,

cocoons were weighted (± 0.001 g) and measured along the longest side of the cocoon (±

0.01 cm), and individually incubated in petri dishes. After emergence, the sex of the adult

bee was linked to the weight and length of its cocoon. Analyses showed that cocoons with

females were significantly longer (Kruskal-Wallis; χ² = 37.48, P < 0.0001, ♀: mean = 1.45 cm,

range = 1.25 – 1.65 cm; ♂: mean = 1.3 cm, range = 1.15 - 1.5 cm) and heavier (Kruskal-Wallis;

χ² = 33.069, P < 0.0001; ♀: mean = 0.2011 g, range = 0.1412 – 0.2674 g; ♂: mean = 0.1481

g, range = 0.0936 – 0.2387 g) compared to cocoons with males (Figure 1). Cocoon weight

was also strongly correlated with cocoon length (N = 87; Pearson: rho = 0.84, P < 0.0001).

Figure 1: Boxplots showing that O. cornuta cocoons with females are significantly longer and heavier

compared to cocoons with males. Boxplots show the median (thick black line) and the range (whiskers or dots)

of cocoon weight (left) and length (right) for females and males.

6

Figure 2: O. cornuta lifecycle at the Laboratory of Agrozoology, Faculty of Bioscience Engineering at Ghent

University. A: nest box with layers of wooden blocks consisting of holes at the joining surfaces. B: male and female

O. cornuta. C: detail of the nest box with unfinished and one finished (final plug of mud) nesting cavities. Two O.

cornuta females are still in the act of constructing nests. Yellow pollen on the female is clearly visible. D: O.

cornuta eggs on top of provision in a nesting cavity. E: provision consuming larva. F: cocoon spinning prepupa.

G: completed cocoon.

2.2. Pesticides

In this study, adult and larval O. cornuta bees were exposed to two different systemic

pesticides. These included thiacloprid, a neonicotinoid insecticide, and tebuconazole, a

triazole fungicide. Both pesticides have previously been detected together in pollen collected

from bees and plants, as well as on individual bees (David et al., 2015; David et al., 2016; Mullin

et al., 2010; Roszko et al., 2016). Characteristics of both pesticides are given in Table 1.

7

Table 1: Characteristics of the insecticide thiacloprid and the fungicide tebuconazole.

Product

name

Active

component

Concentration active

component

Class Current usage

Calypso Thiacloprid 480 g/L Neonicotinoid Spray

insecticide

Tebusip Tebuconazole 250 g/L Demethylation

inhibitors

Spray fungicide

Adult and larval oral toxicity tests on O. cornuta were conducted with field realistic exposure

concentrations of thiacloprid and tebuconazole. Therefore, a literature search was performed

to check for residue concentrations on plants, bee colonies, pollen and nectar in the field

(Table 2).

Table 2: Literature overview of residue concentrations of thiacloprid and tebuconazole on plants, bee colonies,

pollen and nectar in the field (OSR = oil seed rape).

Pesticide Species Part of plant Concentration

range (ppb)

Authors

Thiacloprid OSR Nectar 6.5 - 500 Pohorecka et al., 2012; EFSA, 2013

OSR Pollen 2.1 - 900

Botias et al., 2015; David et al., 2016;

EFSA, 2013; Pohorecka et al., 2012

Wildflowers Pollen 0.3 - 0.6 David et al., 2016

Honey bees Pollen loads 30 Skerl et al., 2009

Honey bees Pollen (hive) 49.1 - 326 Rennich et al., 2012

Honey bees Pollen 1.7 - 136 Mullin et al., 2010; Roszko et al., 2016

Tebuconazole OSR Pollen 5.2 David et al., 2016

Wildflowers Pollen 3.3 – 7 David et al., 2016

Honey bees Pollen loads 6.5 (mean) Chauzat et al., 2009

Honey bees Pollen 0.9-64.6 Roszko et al., 2016

Based on these residue concentrations in the field, a range of five concentrations for both

thiacloprid, tebuconazole and a mixture of thiacloprid and tebuconazole were selected for the

adult oral toxicity test on O. cornuta (Table 3). For the larval oral toxicity test, only the highest

concentrations of thiacloprid, tebuconazole and their combination were used (Table 3).

It has to be noted that the concentration range for tebuconazole was chosen to be significantly

higher than the field realistic concentrations. This is because fungicides such as tebuconazole

are sprayed during full bloom of bee attractive crops, when O. cornuta bees are active.

Consequently, there is a realistic probability of exposure to high concentrations of

tebuconazole.

8

2.3. Available protocols for adult and larval oral toxicity testing on solitary

bees

As mentioned before, Osmia species are suitable testing species in the regulatory risk

assessment of PPPs (EFSA, 2013; Sgolastra et al., 2015). Oral toxicity tests on both adult and

larval Osmia species are already described and conducted, but further ring-testing and

validation has to be performed (EFSA, 2013). To test for potential effects of pesticides on

solitary bees such as Osmia species, individual feeding (i.e. exposure) is required.

2.3.1. Available protocols for adult oral toxicity testing

Ladurner et al. (2003) were the first researchers who developed an individual feeding method

where adult bees can be exposed to known amounts of pesticides with a very high rate of

success. This method was called the ‘Flower method’, and was proposed to serve as a

standardized oral acute toxicity test for a multitude of bee species (Ladurner et al., 2003), from

which an LD50 – the dose (μg of active substance per bee) inducing 50% mortality of the tested

bees after a specified test duration – can be calculated.

In this method, a plastic ampoule (inner diameter = 2 mm, height = 5 mm) is inserted into the

calyx of a flower (e.g. Prunus avium L. or Convolvulus arvensis L.) (Figure 3). This plastic

ampoule is switched for the reproductive column of the flower, which is removed with a pair

of forceps. Into this ampoule, the test solution is provided (10 μL) from which bees can

consume. Flowers and bees are housed individually in ice cream cups made of waxed

cardboard (diameter = 8 cm, height = 5 cm) covered with a petri dish lid to which a wired mesh

is attached to provide sufficient aeration. Flowers are positioned on an inverted glass vial

stopper (Figure 3). Crucial is that flowers used, are not treated with chemicals. Disadvantages

of this method are its labor intensiveness and the fact that it is designed to administer only a

single dose of the test substance to an individual bee.

Subsequent studies used this ‘Flower method’, but adjusted several aspects to contribute in

standardizing this method. Ladurner et al. (2005) used artificial flowers instead of natural

flowers, where they used color, odor and shape cues (of the natural flowers visited by their

study species Osmia lignaria, Apis mellifera and Megachile rotundata). They also tested if

training the bees improved feeding success, as bees in nature memorize flower cues (color,

odor and shape) after being associated with a food reward. Despite their efforts, they

concluded that the ‘Flower method’ with natural flowers as described by Ladurner et al. (2003)

is anyways more effective. This result was confirmed by Hinarejos et al. (2015), who compared

the performance of artificial flowers against the ‘Petal method’, a simplification of the ‘Flower

method’ that only uses one petal of the flower instead of the whole calyx of the flower as in

the ‘Flower method’. The ‘Petal method’ also resulted in high feeding success, is less labor

intensive than the ‘Flower method’ and therefore a good candidate as a standardized method

for oral toxicity testing. A study that build upon the ‘Petal method’ was conducted by Heard

et al. (2017). These authors performed oral toxicity tests on O. bicornis where they held

individual bees in 500 ml plastic cages with a ventilated lid. They fed bees with a 5 ml Leur

9

centric syringe with the tip cut off. This tip was UV colored cued and carried an artificial yellow

silk petal. A major advantage of this method is that it allows for continuous feeding and

exposure (e.g. 10 days in Heard et al. (2017)).

The protocol that was used for adult oral toxicity testing in this study was based on the

abovementioned studies as well as on proposed protocols for acute (Appendix A, only digitally

available) and chronic oral toxicity testing on solitary bees as discussed in the “non-Apis”

working group.

Figure 3: Feeding unit for the flower method. Cross-section drawing and photograph of alfalfa leafcutter bee

(Megachile rotundata) female on test flower. The plastic ampoule containing the test solution can be seen at the

center of the corolla. (am: ampoule, ca: flower calyx, co: flower corolla, st: stopper (flower holder), ts: test

solution) (Ladurner et al., 2003).

2.3.2. Available protocols for larval oral toxicity testing

In order to conduct toxicity tests on larvae of Osmia species, eggs and larvae must be collected

from artificial nests in the field. These artificial nests can be provided to existing Osmia

populations or positioned in areas with sufficient and appropriate blooming plant species

where adults can be released as loose cocoons in, for example, a cardboard box. Artificial nests

can consist of reed canes, cardboard tubes, wooden blocks with paper straws positioned in

drilled holes or layers of wooden blocks with holes at the joining surfaces (Figure 2A) (Abbott

et al., 2008; Becker and Keller, 2016; Konrad et al., 2008; Sgolastra et al., 2015). Individual

canes, straws or tubes can be collected upon nest completion (sealed with a final plug) and

brought to the laboratory where they can be carefully dissected with a sharp razor (Becker

and Keller, 2016). A nest box consisting of layers of wooden blocks with holes at the joining

surfaces must be transferred and dissected as a whole. This is disadvantageous because one

cannot select for larvae of specific ages. After dissection, the provisions with eggs or larvae

can be individually weighted and transferred with a forceps to a 24- or 48-well plate (Becker

10

and Keller, 2016; Konrad et al., 2008; Sgolastra et al., 2015). Well plates should be covered

with a paper towel to allow air flux (Becker and Keller, 2016). Eggs and larvae can be sexed

based on their provision mass and position in the nest, but it is recommended that their sex is

confirmed after emergence of its cocoon in the next year (EFSA, 2013; Sgolastra et al., 2015).

Abbott et al. (2008) introduced two methods for application of the test solution. In their first

method (‘own pollen’), they injected 10 µl of the test solution in the provision, whereas in

their second method (‘new pollen’), they removed the eggs or larvae from the provisions and

replaced the provisions with a pre-blended pollen mixture of the appropriate amount of the

test solution. Even though their second method allows for an equally concentrated exposure

of the test solution within the provision, this method might not be preferred as this

manipulation could be destructive for the eggs and larvae (Sgolastra et al., 2015). Therefore,

Abbott et al.’s (2008) first method more widely served as a basis for further improvements on

toxicity testing on Osmia larvae. Subsequently, testing solution is most often delivered into a

longitudinal fissure or hole in the provision (EFSA, 2013; Konrad et al., 2008; Sgolastra et al.,

2015). The amount of the desired concentration of the test solution that should be added to

the provision equals around 50 µL per gram of provision (EFSA, 2013; Konrad et al., 2008;

Sgolastra et al., 2015). Well plates are suggested to be stored in darkness at 22 ± 2°C and 55-

65% RH for good development (Becker and Keller, 2016; Konrad et al., 2008; Sgolastra et al.,

2015).

Once the larval toxicity test is set up, larval development and mortality are observed daily until

cocoon spinning (EFSA, 2013; Konrad et al., 2008; Sgolastra et al., 2015). When all bees

reached the adult stage in the cocoon, O. cornuta bees should be transferred to 12-14°C for

7-15 days followed by a wintering period of 150-180 days at 2-4°C (EFSA, 2013; Sgolastra et

al., 2015). After the wintering period, cocoons should be incubated at 25 ± 5°C in individual

cages and checked daily for emergence and mortality (Abbott et al., 2008; EFSA, 2013; Konrad

et al., 2008; Sgolastra et al., 2015). Based on these larval toxicity testing methods, the

following endpoints can, for example, be recorded: pre-emergence mortality (total number of

dead bees before emergence), larval period (total number of days from egg/larva until cocoon

spinning), emergence time (total number of days required to emerge) and post-emergence

longevity (total number of days the bee lived after emergence) (Sgolastra et al., 2015).

2.4. Experimental procedure

2.4.1. Protocol for adult oral toxicity testing

Cocoons were obtained in January 2017 from Inbuzz in the Netherlands

(http://www.inbuzzextra.nl/) as well as from an artificial nest box with cupboard tubes from

a private garden in Londerzeel, Belgium. All cocoons were stored at 1.5°C to secure completion

of the diapause. After completion of their diapause, cocoons were incubated at 22°C and 50-

60% RH and checked daily. Emerged females were transferred and stored at 3°C (no longer

than 10 days) until enough females were obtained for the experiment. Prior to the start of the

experiment, individual bees were weighted (± 0.001 g) and transferred at random to an

http://www.inbuzzextra.nl/

11

artificial ventilated box that served as testing cage (Figure 4). This test cage contained a small

hole at the side through which a 1 mL Leur centric syringe was placed. The tip of this syringe

was cut off to provide a ± 4 mm drinking hole. Efforts were made to slope the syringe in such

a way that the tip touched the bottom of the artificial box (Figure 4). This syringe contained

the test solution and could easily be replaced with a new syringe if required. The syringe did

not contain any flower cue (i.e. odor, color or shape cue) as was described in the ‘Flower-‘ and

‘Petal method’ (Hinarejos et al., 2015; Ladurner et al., 2003). This is because a previous oral

toxicity tests with O. bicornis indicated that adult females consumed the test solution

according to the method described above (Eeraerts M., unpublished data from 2015 and

2016). Prior to the real experiment, bees were habituated to the test cage for a two-day period

where they could feed ad libitum on a 50% sugar water solution. Bees were incubated at 22°C

and 55% RH under an artificial light (16h) – dark (8h) rhythm.

Figure 4: Artificial ventilated test cage with a 1 mL Leur centric syringe touching the bottom of the box with its

tip. Each cage housed one adult female. Dimensions of the cage are indicated on the picture.

After the two-day habituation period in the testing cage, adult females were assigned to a

treatment (Table 3). These treatments included the range of five concentrations of thiacloprid,

tebuconazole and the mixture of thiacloprid and tebuconazole, as well as three control groups

(50% sugar water solution, 50% sugar water + 2.5% acetone solution and 50% sugar water +

5% acetone solution). The ‘50% sugar water solution’ control group was used to control for

potential effects of acetone, whereas the ‘50% sugar water + 5% acetone solution’ control

group tested for the innocence of acetone. Sample size for each treatment can be found in

Table 3.

Thiacloprid and tebuconazole test solutions were prepared by dissolving Calypso (480 g/L) and

Tebusip (250 g/L) in a 50% sugar water + 2.5 % acetone solution. Subsequent dilution series in

a 50% sugar water + 2.5 % acetone solution were performed to reach the desired

concentrations. All dilutions were stored in volumetric flasks at 3°C throughout the whole

experiment.

12

The duration of the experiment was 24 days. During the first 12 days of the experiment, O.

cornuta bees were continuously orally exposed to the test solutions (± 1 mL). For the last 12

days, all bees were continuously orally exposed to untreated 50% sugar water solution. The

experiments’ timespan represents ± 3/4th of an individual bees’ lifespan as their total period

of activity is three to four weeks. This test can therefore be considered as a chronic oral toxicity

test. Endpoints used in the toxicity test were mortality – which was assessed daily (24h ±3h) –

and food consumption (sublethal effect) – which was assessed every three days.

Throughout the whole experiment (24 days), syringes were replaced every three days (72 h ±

3 h). This was done to counteract potential degradation of the pesticides (first 12 days), to

minimize absorption of the pesticides in the syringes (first 12 days) and to provide fresh

untreated 50% sugar water (last 12 days). Upon replacement, old and newly filled syringes (±

1 mL) were weighted, providing data on ‘total three-day consumption’ in grams (g) per bee

for each treatment. This ‘total three-day consumption’ measure was corrected for

evaporation of the test solution over the course of those three days. Therefore, additional

syringes (N = 10), which were solely filled with a 50% sugar water solution, were incubated at

22°C and 55% RH to calculate the average three-day evaporation of the test solution from the

syringes. This average three-day evaporation was then subtracted from the ‘total three-day

consumption’ to achieve the ‘real three-day consumption’ per bee for each treatment.

Furthermore, it can be expected that differences in body weight attribute to variability in food

consumption. For example, heavier bees might eat more compared to thinner bees.

Therefore, ‘real three-day consumption’ was normalized to body weight by dividing each bee’s

food consumption (g) by its own body weight (g) according to the following formula: food consumed (g)

body weight (g). A value of ‘one’ means that the bee consumed its own body weight in three

days. This new metric, hereafter referred to as ‘normalized food consumption’ was used for

all analyses.

2.4.2. Protocol for larval oral toxicity testing

O. cornuta eggs and larvae were collected from two types of nest boxes. First, a wooden nest

box consisting of layers with holes at the joining surfaces (Figure 2A). This wooden nest box

was positioned at Campus Coupure, Ghent University, in a flower and blossom rich

environment, where upon blooming, cocoons were released in a cupboard box on top of the

nest box. Second, a triangular nest box consisting of individual cupboard tubes (Figure 5). This

nest box was positioned in a private garden in Londerzeel, Belgium, rich in flowers and

blossoms where wild O. cornuta bees nested.

The wooden nest box and eleven individual cardboard tubes of the triangular nest box were

transferred to the lab. The wooden nest box was dissected layer by layer whereas the

individual cardboard tubes were carefully opened with a sharp razor. Eggs and larvae attached

to their own provision were transferred with a forceps to an individual well (diameter 15 mm)

in a 24-well plate with flat bottom. Prior to this transfer, a ± 1 mm layer of grinded Biobest

honeybee pollen was placed on the bottom of each well of this 24-well plate (Figure 6A). This

13

was done to stabilize the provision in the well – with the aim to prevent eggs and larvae to

detach from their provision – as well as to provide larvae that detached from their provision

with extra pollen – to increase their chances of successful survival. Indeed, a small pilot study

indicated that larvae were prone to detach from their own provision in wells (diameter 15

mm) with flat bottom, even after successful transfer to the well, which resulted in mortality

(Figure 6B). This detachment was probably caused by our handling or breaking of the

provision. 24-well plates were bought sterile and opened individually before transfer of the

eggs and larvae. Upon transfer, eggs and larvae were sexed – based on their position in the

nesting cavity and their provision mass – and aged (See box 2). Larvae older than second instar

(N = 58) were discarded from the toxicity test experiment, but were anyways reared to use

them in describing the protocol of the oral larval toxicity test. Finally, a small longitudinal hole

was made in each provision in which the test solution was injected (Figure 6C). In between the

lid and the bottom of the well plate, a paper towel was placed to allow for some airflow. Eggs

and larvae were incubated in darkness at 24°C and 60% RH and checked every two days to

determine detachment from the provision, larval mortality and time to cocoon spinning.

Figure 5: Triangular nest box filled with individual cardboard tubes.

Figure 6: A: O. cornuta egg attached to its provision on top of a ± 1 mm layer of grinded Biobest pollen on the

bottom of a well of a 24-well plate. B: a detached O. cornuta larva two days after successful transfer of the larva

with its provision from the nest to a well during a small pilot study. C: a small longitudinal hole in the provision

of an O. cornuta larvae in which the test solution was injected.

14

Box 2: Aging of larvae of O. cornuta for larval oral toxicity test

Aging was based on descriptions and depictions of O. cornuta and O. bicornis (Figure 7)

(Peeters et al., 2012b; Rust et al., 1989 and Bosch, J. and Stanisavljević, Lj. Z. personal

communication)

Figure 7: A: O. cornuta egg at the start of development. B: developed O. cornuta egg with visible segments. C:

first instar larva emerged from the egg membrane, not feeding on the provision. D: second instar larva feeding

on the provision. The yellowish provision is clearly visible in its digestive tract. E: fifth instar larva with clearly

visible faeces. F: prepupa right before spinning its cocoon.

A B

C D

15

Box 2: continued.

Figure 7: Continued.

Eggs/larvae were randomly assigned to a different treatment. Treatments consisted of one

control (50% sugar water), one thiacloprid (2000 ppb), one tebuconazole (12500 ppb) and one

combined thiacloprid and tebuconazole (2000 + 12500 ppb) group. An overview of the

treatments and the respective sample sizes are found in Table 3.

To obtain the aimed concentrations, thiacloprid and tebuconazole test solutions were

prepared by dissolving 1 mL of Calypso (480 g/L) and Tebusip (250 g/L), respectively in 9 mL

of a 50% sugar water solution. Subsequently, dilution series with a maximum dilution factor

of ten were performed in 50% sugar water to reach the required concentrations. 10 µL of the

obtained test solutions was administered to the eggs/larvae by injection in the longitudinal

hole in their provision (Figure 6C). It was estimated that provisions weighted 0.10 g, as some

larvae were already in their second larval instar stage, thus consuming their provision.

F E

16

Table 3: Overview of the different treatments for the adult and larval oral toxicity tests. Concentrations of each treatment and sample sizes at the start of the adult and larval oral toxicity tests are given.

Treatment Concentration Sample size adults Sample size larvae

Control 1 50% sugar water 20 26

Control 2 50% sugar water + 2.5% acetone 20 /

Control 3 50% sugar water + 5% acetone 20 /

Thiacloprid 1 2000 ppb 16 18

Thiacloprid 2 800 ppb 18 /




Tebuconazole 1 12500 ppb 18 15

Tebuconazole 2 5000 ppb 19 /




Mixture 1 2000 + 12500 ppb 18 21

Mixture 2 800 + 5000 ppb 15 /

Mixture 3 320 + 2000 ppb 18 /

Mixture 4 128 + 8000 ppb 18 /

Mixture 5 50 + 320 ppb 18 /

2.5. Statistical analyses

The larval oral toxicity test was still running at the end of this study. With some exceptions (N

= 5), no larvae started spinning their cocoon, and only five larvae died during the experiment.

Therefore, no statistical analyses could be performed for the larval oral toxicity test and all

analyses described below are performed for the adult oral toxicity test. For this, all datasets

were graphically and statistically checked for normality (Shapiro – Wilk test), homoscedasticity

(Bartlett's test) and outliers. Non-parametric tests were used when required. Statistical

analyses were performed in R (R Development Core Team 2015).

2.5.1. Mortality

To assess potential effects of treatment on mortality for each day of exposure, generalized

linear models (GLM) with binomial distribution were used. Mortality (modeled as a binary

variable with 0 = dead and 1 = alive) and treatment were modeled as the dependent and

independent variable, respectively. For this test, only data about bees which consumed the

test solution were used. Therefore, (1) mortality data about deceased bees during the first

two days of exposure of which no consumption data was available and (2) mortality data about

bees with a normalized food consumption of less than 0.2 g/g between day zero and three

which died within five days were discarded from analyses (N = 90). The 0.2 g/g cut-off value

was chosen arbitrarily.

17

Due to the high mortality during the first three days of exposure in all treatments, it was

concluded that not exposure to the test solution but another factor was responsible for the

observed mortality. Therefore, a second test was conducted to investigate if normalized food

consumption affected mortality after three (Day 3) days. For this test, normalized food

consumption of bees that died and were still alive at Day 3 was compared. This was done by

conducting a Kruskal-Wallis test at day 3 with mortality and normalized food consumption

modeled as the independent and dependent variable, respectively.

2.5.2. Normalized food consumption

To assess potential effects of treatment on normalized food consumption, normalized food

consumption after three (Day 3), six (Day 6), twelve (Day 12) and twenty-four (Day 24) days

was compared between treatments. For this, normalized food consumption per bee was

summed to obtain normalized food consumption data per bee after six, twelve and twenty-

four days. Kruskal-Wallis and One-way ANOVA tests were conducted to test for significance in

normalized food consumption between treatments. Nemenyi’s post hoc (with dist="Tukey")

for pairwise multiple comparisons of the ranked data and Tukey’s post hoc tests were used

after Kruskal-Wallis and One-way ANOVA, respectively.

18

3. Results

3.1. Adult oral toxicity test

3.1.1. Mortality

Mortality data of O. cornuta bees during the adult oral toxicity test for each day are

represented in Appendix B, table B1 (only digitally available) and illustrated in Figure 8. Figure

8 only depicts the three control groups together with the highest and lowest concentrations

of each treatment group to simplify the figure, but all treatments showed the exact same

trend. This figure shows that mortality was very high (up to 60%) in all treatments, including

control groups, during the first three days of exposure, but remained relatively constant

throughout the rest of the experiment.

Treatment had no significant effect on mortality of O. cornuta bees at each day (GLM: all P >

0.3635). However, normalized food consumption was significantly lower for dead (N = 32)

compared to living (N = 207) bees at Day 3 (Kruskal-Wallis; χ² = 35.382, P < 0.0001; Figure 9).

3.1.2. Normalized food consumption

Treatment had a significant effect on normalized food consumption after Day 3 (N = 238;

Kruskal-Wallis; χ² = 31.353, P = 0.01809), Day 6 (N = 192; Kruskal-Wallis; χ² = 51.266, P <

0.0001), Day 12 (N = 183; ANOVA; F = 3.539, P < 0.0001) and Day 24 (N = 169; Kruskal-Wallis;

χ² = 45.426, P < 0.0001). Sample size per treatment ranged from five to 17.

Despite a global significance after Day 3, Nemenyi’s post hoc test could not identify which

treatments significantly differed from each other. The significant p-values (<0.05) of all post

hoc test for Day 6, Day 12 and Day 24 are given in table 4, and depicted in Figure 10.

3.2. Larval oral toxicity test

The larval oral toxicity test was followed up for circa three weeks but was still running at the

end of this study. The aim was to determine larval detachment from the provision, larval

mortality and time to cocoon spinning. During this experiment, no larvae detached from its

provision, overall mortality was very low (N = 5) and five larvae started spinning their cocoon.

Due to these low sample sizes, statistical tests could not be performed to relate mortality and

time to cocoon spinning to treatment. The protocol of the larval oral toxicity test is further

discussed in the discussion section ‘Protocol for larval toxicity testing’.

19

Figure 8: Mortality trend of selected treatments (controls, highest and lowest concentrations) in O. cornuta adult oral toxicity test.

20

Figure 9: Mean normalized food consumption (g/g) for dead and living O. cornuta bees at Day 3. Errors bars

represent standard deviations of the mean normalized food consumption.

Figure 10: Bar graph depicting overview of mean normalized food consumption (g/g) between all treatments

after Day 6, Day 12 and Day 24. Error bars represent standard deviations of the respective mean normalized

food consumption (g/g).

21

Table 4: The significant p-values (<0.05) of all post hoc tests for Day 6 (Nemenyi), Day 12 (Tukey) and Day 24

(Nemenyi).

Day 6 Day 12 Day 24

Mixture 1 Mixture 2 Mixture 1 Mixture 2 Mixture 1 Mixture 2

Control 2 0.0098 0.0104 0.0027

Control 3 0.0027 0.0087

Thiacloprid 2 0.0381

Thiacloprid 3 0.0383

Thiacloprid 4 0.0122 0.008

Thiacloprid 5 0.0325 0.0336 0.0002 0.0164 0.0063

Tebuconazole 5 0.0047

Mixture 4 0.0191 0.0195 0.0002 0.014 0.0079

Mixture 5 0.0455 0.0461 0.0010 0.0486 0.0432

22

4. Discussion

4.1. Results and protocol of adult oral toxicity test

The combination of thiacloprid and tebuconazole affected normalized food consumption in

adult females of O. cornuta throughout the whole experiment (24 days). More specifically,

bees which were exposed to the highest concentrations of the mixture of thiacloprid and

tebuconazole consumed less of the test solution compared to control groups and groups

exposed to the lowest concentrations of thiacloprid, tebuconazole and their mixture after 6,

12 and 24 days. This indicates the presence of a sublethal effect throughout almost the entire

active period of O. cornuta females. Consequently, the combined presence of thiacloprid and

tebuconazole in nectar might cause reduced feeding, which might reduce their own vitality.

Exposure to only thiacloprid and tebuconazole did not affect normalized food consumption,

nor did exposure to thiacloprid, tebuconazole and their combination affect mortality (i.e. no

acute or chronic effects).

In the ecological risk assessment (ERA) framework concerning the placing of plant protection

products (PPPs) on the market in the EU (Regulation (EC) 1107/2009), thiacloprid and

tebuconazole were allowed to be used as pesticides. This allowance was the result of toxicity

testing of the single compounds on several model species. This study confirms this allowance,

in that thiacloprid and tebuconazole alone do not have lethal (mortality) nor sublethal

(reduced normalized food consumption) effects in solitary bees (model species O. cornuta).

However, this regulatory risk assessment framework only focuses on single pesticides,

whereas effects of mixtures of pesticides are not included. This study indicated that the

mixture of thiacloprid and tebuconazole did affect solitary bees, thus pressurizing the

regulatory risk assessment framework to also include toxicity tests on mixtures of pesticides

before allowing them on the market. Especially because it is already known that bees in nature

are exposed to mixtures of pesticides (Mullin et al., 2010).

In this study, O. cornuta mortality was very high during the first three days of exposure in all

treatments, including all control groups (Figure 8), while it was very low for all subsequent

days until day 24. It was shown that normalized food consumption, and not treatment,

affected this mortality (Figure 9). Bees, thus, experienced difficulties in consuming the test

solution from the syringes during the first days of the experiment (even though they had a

two-day habituation period), resulting in mortality. No other factors could seem to cause this

mortality. An identical newly set up toxicity test with only the three control groups during this

study with O. bicornis gave identical results (unpublished results). Only after several days,

bees seemed to learn how to consume the test solution. This contrasts with an identical set

up contact and oral toxicity test with different pesticides on O. bicornis (Eeraerts M.,

unpublished data) where mortality was very low in control groups. The reasons why both O.

cornuta and O. bicornis barely consumed the test solutions during the first days of the

experiment are unknown. It might be that bees were repelled from the chemicals in the test

solutions (Frazier et al., 2014), although this is unlikely because the 50% sugar water control

group also experienced substantial mortality. Another possibility is that bees were not

23

attracted to the syringes or unable to find the food source. This could be explained by the lack

of flower cues as used in the ‘Flower’ and ‘Petal method’ described by Ladurner et al. (2003)

and Hinarejos et al. (2015), respectively. Besides this non-feeding problem, bees were actively

walking around in the artificial cages, and there is no reason to assume that the toxicity test

setup was not sufficient. Therefore, I suggest to repeat this toxicity test with the same setup,

but by using the ‘Petal method’, i.e. constructing a petal of a flower on the tip of the syringe,

although this method is more labor intensive.

Unfortunately, QuEChERS analyses, executed in another master thesis, on the test solutions

(the lowest and highest concentrations of thiacloprid, tebuconazole and the mixture of

thiacloprid and tebuconazole) that were administered at the start of the adult oral toxicity

test on O. cornuta showed substantial deviations from the aimed concentrations (Ostyn,

2017). More specifically, real administered concentrations of the test solution were up to a

factor 100 lower compared to the aimed administered concentrations (Table 5). Potential

causes of these decreased concentrations could be that pesticides were destructed by

acetone or photolytic processes during storage and administration (Gupta et al., 2008).

Despite these deviations, the range of high to low concentrations administered to the bees

was still maintained. This implies that effects found in this study are still applicable but caused

by lower concentrations than the aimed concentrations (Table 2). Therefore, it can be

expected that if this toxicity test was conducted with the aimed concentrations, which are

field realistic concentrations, the effects found in this study might have been more

pronounced. Furthermore, the fact that no effects of thiacloprid, tebuconazole and their

combination on mortality and normalized food consumption were found, must be interpreted

with caution, as actual effects under field realistic concentrations might, in fact, occur.

Table 5 : QuEChERS analyses on several test solutions that were administered at the start of the adult oral toxicity

test on O. cornuta showed substantial deviations from the aimed concentrations, up to a factor 100 lower (real

concentration). <LOD (limit of detection) means that the analyses could not detect thiacloprid in the sample.

Treatment Aimed Concentration Real Concentration

Control 1 50% sugar water 50% sugar water

Control 2 50% sugar water + 2.5% acetone 50% sugar water + 2.5% acetone

Control 3 50% sugar water + 5% acetone 50% sugar water + 5% acetone

Thiacloprid 1 2000 ppb 85 ppb

Thiacloprid 5 50 ppb <LOD

Tebuconazole 1 12500 ppb 4435 ppb

Tebuconazole 5 320 ppb 78 ppb

Mixture 1 2000 + 12500 ppb 111 + 2939 ppb

Mixture 5 50 + 320 ppb 1 + 25 ppb

4.2. Protocol for larval toxicity testing

The first step in contributing to the development of a standardized protocol for larval toxicity

testing is optimizing the methods for larval laboratory rearing. This starts with the choice of

the nest box from which one aims to collect the eggs/larvae. In toxicity testing, larvae should

24

not be older than the first instar larvae, to ascertain that all pollen-feeding larval stages

consume the test solution. Preferably, nest boxes from which it is easy to select and transfer

only eggs and first instar larvae should be used. The wooden nest box used in this study could

only be opened as a whole, which resulted in harvesting numerous larvae that were older

than the first instar stage and, hence, could not be included in the toxicity test. However, a

major advantage of this wooden nest box is that it is opened layer by layer, making transfer

of eggs and larvae with a spatula and/or forceps very convenient. In contrast, individual

cardboard tubes have to be cut open with a sharp razor, which increases the risk of damaging

the eggs/larvae. A wooden nest box with U-shaped holes in the individual layers, from which

layers could individually be removed upon completion of the nesting cavity, seems to be ideal.

This reduces handling impact and secures an easy transfer.

Although a small pre-study indicated that eggs/larvae were very prone to detach from their

provisions even after successful transfer, eggs and larvae in this study very rarely detached

both during handling and the experiment. This was probably due to two major factors: the

use of a layered wooden nest box (easy transfer) and the extra added Biobest pollen layer on

the flat bottom of the individual wells of the 24-well plate. The latter prevented the provision

mass from rolling around in the well, potentially resulting in detachment of the eggs/larvae.

However, instead of using Biobest pollen, this problem could also be overcome by using

another kind of well plates or glass vials. In this study, 24- well plates with wells having a

diameter of 15 mm were used. This creates a large surface area which enhances rolling of the

provision mass, as well as potential problems for successful cocoon spinning. Indeed, personal

observations showed that several larvae initiated up to three attempts to spin their cocoon

(Figure 11A), possibly leading to a lack of silk to complete it (Abbott et al., 2008). Therefore,

well-plates with wells or glass vials having a diameter of around 10 mm, more closely

mimicking the natural diameter of nesting cavities, should be used.

To standardize a protocol for larval oral toxicity testing, a major challenge is to administer

equal concentrations of the test solutions to each larva. For this, several aspects need to be

taken into account. First, only eggs and first instar larvae should be used, as these do not yet

feed on the provision. Despite this fact, in this study also second instar larvae were included

to increase the sample size. Second, provision masses should be weighted in order to be able

to administer equal concentrations to eggs/larvae. It is known that the average fresh weight

of female and male provisions in O. cornuta approximates 0.5 g and 0.25 g, respectively (Bosch

and Vicens, 2002). In this study, however, provision masses varied greatly; between 0.1 g and

0.5 g. This was because of the inclusion of different developmental stages (eggs and first and

second instar larvae). Consequently, together with the additional Biobest pollen, the aimed

concentrations of the test solutions are expected to be up to a factor 10 lower, as it was

observed that most larvae consumed their own provisions as well as the additional Biobest

pollen before spinning their cocoon. To account for provision weight differences, test

solutions with different concentrations should be prepared, so finally all eggs/larvae are

exposed to an equal concentration of the test solution. However, this is very labor intensive.

Third, the mode of administration of the test compound. This study opted to inject the test

compound in a longitudinal hole in the provision mass, according to the ‘own pollen’ method

25

described in Abbott et al. (2008). However, according to this method, the test solution might

not diffuse evenly in the provision mass, resulting in topical exposure and/or in larvae

consuming the entire dose in a short period of time. Furthermore, different larvae may be

exposed to the test solution at different developmental stages and background

concentrations of other chemicals in the ‘own pollen’ provision mass cannot be excluded.

Indeed, personal observations indicated that the test solution diffused only locally in the

provision mass and caused different larvae to consume the test solution at different

developmental stages. Also, larvae consumed all the additional Biobest pollen, which was

rendered free of test solutions. This all results in the fact that the different eggs/larvae in this

study were undoubtedly exposed to different concentrations of the test solutions, making

standardized protocols very difficult. Therefore, I opt to retry the ‘new pollen’ method as

described in Abbott et al. (2008). To my knowledge, no other study so far used this ‘new

pollen’ method due to the high observed mortality of eggs/larvae, probably due to

manipulation of the eggs/larvae and the consistency of the pre-blended pollen mixture

(Abbott et al., 2008). However, this method ensures homogenous administering of equal

concentrations of the test solutions to each larva, as each egg/larva is transferred to a pre-

blended mixture of pollen with the test solution. This study showed that mortality both during

transfer of eggs/larvae with their provision mass from the wooden nest box to the well-plates

and during the experiment itself was very low, and that larvae of all developmental stages

rigorously consumed the Biobest honey bee pollen. Consequently, I believe that with the

appropriate kind of forceps and spatulas underneath a stereo microscope, eggs/larvae can be

removed from their provision mass and transferred to the pre-blended pollen mass (See Box

3). This method also eliminates weighing the eggs/larvae with their provision mass to prepare

the individual required concentrations of the test solutions, as well as the extra manipulation

necessary for the weighing step itself.

Box 3: Revision of Abbott et al.’s (2008) ‘new pollen’ method

Place the eggs/larvae with provision on a Petri dish of which the bottom is covered with

Biobest pollen. With a sharp small razor and/or forceps, remove most of the provision from

the egg/larvae without touching the egg/larvae. If during this act the egg/larvae gets

detached, it falls softly on the Biobest pollen. Subsequently, the egg/larvae can be

transferred with a small spatula/spoon into the well with the pre-blended pollen mixture.

If possible, the pre-blended pollen mixture should weigh approximately 0.5 g and 0.25 g for

female and male O. cornuta eggs/larvae, respectively.

Another advantage of using a pre-blended pollen mixture might be that potential parasites

are excluded from the ‘own’ provision mass of eggs/larvae. This is because in this study it was

observed that the provision of some larvae was parasitized by Cacoxenus indagator.

Cacoxenus indagator larvae also consume on the provision mass, hereby causing starvation

of its host (in this case O. cornuta larvae). Although parasitized, only a very small amount

(maximum three larvae) of Cacoxenus indagator larvae were observed, so O. cornuta larvae

could still develop alongside the Cacoxenus indagator larvae. Figure 11B depicts the

26

Cacoxenus indagator larvae alongside the O. cornuta larvae. The orange-like threads are the

faeces of the Cacoxenus indagator larvae. Nevertheless, whichever method is used for larval

toxicity testing, it is advised to discard larvae which are parasitized by any kind of parasite

from analyses.

Figure 11: A: Wells having a diameter of 15 mm might create potential problems for successful cocoon spinning.

This larva initiated up to three attempts to spin its cocoon, possibly leading to a lack of silk to complete it. B:

Cacoxenus indagator larvae (bottom left) alongside the O. cornuta larvae (top right). The orange-like threads are

the faeces of the Cacoxenus indagator larvae.

27

5. Conclusion and future research

Results of the adult oral toxicity test showed that combined exposure to thiacloprid and

tebuconazole resulted in reduced normalized food consumption (sublethal effect) throughout

the active period of adult O. cornuta females. In contrast, exposure to only thiacloprid and

tebuconazole did not affect normalized food consumption, nor did exposure to thiacloprid,

tebuconazole and its combination affect mortality (i.e. no acute or chronic effects). In the

ecological risk assessment (ERA) framework concerning the placing of plant protection

products (PPPs) on the market in the EU (Regulation (EC) 1107/2009), thiacloprid and

tebuconazole were allowed to be used as pesticides. However, this study indicates the need

for the regulatory risk assessment framework to also include toxicity tests on mixtures of

pesticides before allowing them on the market. I suggest that future studies on toxicity testing

must include mixtures of pesticides to support these results, and test for synergistic and

sublethal effects, as bees are commonly exposed to mixtures of pesticides in nature.

No results of the larval oral toxicity test were obtained as this test was still running at the end

of this study.

The main goal of the ICPPR “non-Apis” working group is to develop standardized testing

methods for bumblebees and solitary bees, as no official standardized test protocols are

available for non-Apis bees. This study explored protocols for both adult and larval oral

toxicity tests on a solitary bee species, Osmia cornuta.

The protocol for adult oral toxicity testing used in this study showed to be practically feasible

(not labor intensive), yielding useful results for toxicity testing. A big issue, however, was that

mortality of bees was very high during the first three days for all treatments (including the

control groups). It was shown that this was probably due to the fact that bees barely

consumed the test solution (including the 50% sugar water control group) from the provided

syringes, and subsequently starved. The reason for this was probably that bees did not find

the food source. Consequently, the protocol used in this study may be optimized by providing

flower cues to the tip of the syringes as described in the ‘Petal method’ (Hinarejos et al., 2015).

Therefore, I suggest to repeat this toxicity test with the same setup, but by using the ‘Petal

method’, to further optimize this testing protocol in adult oral toxicity testing.

The protocol for larval oral toxicity testing used in this study proved to be successful. However,

it also proved that the major problem in designing a standardized protocol for larval oral

toxicity testing includes administration of equal concentrations of the test solutions to each

larva. When the test solution is injected in the provision mass of the eggs/larvae, the test

solution might not diffuse evenly in the provision mass and different larvae may be exposed

to the test solution at different developmental stages. Therefore, I opt to retry the ‘new

pollen’ method as described in Abbott et al. (2008), which ensures homogenous administering

of equal concentrations of the test solutions to each larva. Further research is, however,

necessary to verify the possibility of this method.

A general remark for future toxicity testing research is that study species must be exposed to

field realistic concentrations and that the exposed test solution must always be double

28

checked to see if aimed administered concentration equals the real administered

concentrations.

Only with a multitude of extra studies concerning (protocols on) toxicity testing of pesticides,

it will be possible to ensure the protection of our bees, our pollinators and ourselves. This

study has brought us a step closer.

29

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Appendix A

Proposed protocol for solitary bee (Osmia spp.) acute oral toxicity test

Silvia Hinarejos1,2

, Xavier Domene1, Jordi Bosch

1

1Centre for Ecological Research and Forestry Applications (CREAF), Autonomous

University of Barcelona, 08193 Cerdanyola del Vallès, Barcelona, Spain 2 Valent U.S.A. Corporation, Valent Technical Center, 6560 Trinity Court, Dublin 94568 CA,

USA

INTRODUCTION This proposal describes a test method designed to assess the acute oral toxicity of pesticides to adult

female solitary bees under laboratory conditions. This is based on the test method developed by

Ladurner et al. (2003, 2005a, 2005b) with improvements according to Hinarejos et al. (in prep.). Osmia populations nest for about 50 days in spring (from March to May depending on the population

origin) (Bosch and Vicens 2005, 2006) during which each female visits about 10 000–25 000 flowers

in ~20 days (Bosch 1994) and consumes about 360–1 540 mg of sugar. During this time exposure the

ingestion of contaminated food may be a significant exposure route for adults. In Osmia adults, exposure

is mainly via ingestion nectar although small amounts of pollen are probably also consumed, especially

by females to complete ovary maturation during pre-nesting period (Richards 1994; Sgolastra 2007).

The acute oral toxicity test is carried out to determine the inherent toxicity of pesticides to solitary bee

adults. The results of this test should be used to define the need for further evaluation. Two mason bees of the genus Osmia (O. cornuta and O. bicornis) are proposed as suitable test species

in the pesticides risk assessment scheme for solitary bees in the European Union (EFSA 2013). O.

cornuta and O. bicornis are very closely related species from the Paleartic region, and share many life

history and behavioural traits. O. cornuta is distributed in central and southern Europe, Turkey and parts

of North Africa and the Middle East. O. bicornis can be found also in northern Europe. Likewise other

species of the genus Osmia, these two species are strictly univoltine and winter as fully-emerged adults

within their cocoons (Bosch and Kemp 2001). CONSIDERATIONS Pesticides can be tested as active ingredients (a.i.) or as formulated products. In order to have bees with a known age, the bees used in this test should be newly emerged adult females

(approx. 24 hours old) A reference item should be used to verify the sensitivity of the bees and the suitability of the test design.

PRINCIPLE OF THE TEST Adult females newly emerged from their cocoon (approx. 24 hours old) are exposed to a range of doses

of the test substance dispersed in 50 % (w/v) during 1 hour. Mortality is recorded daily during at least

35

48 hours and compared with control values. If the mortality rate is increasing between 24 and 48h whilst

control mortality remains at an accepted level, i.e. ≤10%, it is appropriate to extend the duration of the

test to a maximum of 96h. The results are analysed in order to calculate the LD50 at 24h and 48h and, in

case the study is prolonged, at 72h and 96h.

In some cases (e.g. when a test substance is expected to be of low toxicity or when a chemical is poorly

soluble) a limit test may be performed, using 100 μg a.s./bee/day, or the maximum achievable solubility,

whichever is lower. VALIDITY OF THE TEST For the test to be valid, the following criteria apply:

• the average mortality for the controls must not exceed 10 % at the end of the test; • the LD50 of the reference item meets the specified range. As Osmia bee species differ in

size/weight variation in LD50 values can be observed. For O. cornuta and O. bicornis the LD50 of dimethoate, as reference item, should be established by a ring-test in future.

DESCRIPTION OF THE METHOD Collection of the bees Newly emerged from cocoons (approx. 24 hours) non-mated, meconium-free, adult female bees should

be used for the test. All bees should be of the same subspecies, obtained from adequately fed, healthy,

and as far as possible disease and chemical-free populations with known history and physiological

status. The optimal temperature condition during development and the period of adult emergence

depends on the species and the origin of the population used in the test. In the autumn, after ~ 20-30

days from adult eclosion (monitored by x-rays or dissecting cocoons (Bosch and Kemp 2004)), the bees

are cooled for wintering (15 days at 14 °C + 150-180 days at 3–4 °C). In spring, wintering females

within their cocoons are selected based on cocoon size and incubated (2 hours at 12 °C + 22 hours at

22-24 °C). It is recommended that female cocoons have the same average size, so overly small and

overly large cocoons should be excluded. Cocoons should be weighed (without fecal particules) and

then transferred to a clean and well-ventilated flight cage (e.g. insect rearing tents of 60 x 60 x 60 cm)

to allow to them fully emerge and deposit the meconium. The temperature should be maintained at 22-

24 °C during acclimatisation. Meconium-free females should be used within 24 hours after emergence.

If any males are present, they will emerge earlier than the females and should be removed from the cage

to prevent mating. Handling conditions Handling procedures, including treatment and observations may be conducted under (day) light. Preparation of bees On the day of test start the bees can be collected from the flight cage and randomly allocated into the

individual feeding test cages. Anesthetics are normally not needed for collecting bees, but if used,

chilling for 30 minutes at 4-5°C is recommended. The bees are deprived of food prior to treatment so

that all bees are equal in terms of their gut contents at the start of the test. Moribund bees should be

rejected and replaced by healthy bees before starting the test.

Individual feeding test cages In the oral toxicity test for Apis mellifera (OECD TG 213 1998) a common feeder is provided to a group

36

of workers assuming that, through trophallaxis, all individuals will receive similar doses of test solution.

However, this does not applies to most non-Apis bees in general, and Osmia in particular, because they

do not perform trophallaxis. Therefore an individual feeding method is required. Individual test cages should be easy to clean and well-ventilated. Any appropriate material can be used,

e.g. cups made of plastic or waxed cardboard covered with a transparent perforated lid. The size of test

cages should be appropriate to provide adequate space (e.g. cups of ca. 12 cm diameter and 6 cm height

is sufficient for 10 bees). Feeding solutions The feeding solutions for the control, test and reference item treatments are prepared with 500 g/l (50

% w/v) sucrose solution in water. Preparation of the stock and treated feeding solutions Test item should be measured using a balance, if density is available or if the test item is solid. A stock

solution of test item is prepared or the test item can be directly mixed with 50 % (w/v) sucrose solution

(treated feeding solution). In case of good water solubility deionized water is used as the solvent. For

substances of low water solubility acetone can be used as a solvent. The concentration of solvent used

depends on the solubility of the test item and it should be the same for all concentrations tested. An

acetone concentration of 5 % in the final feeding solution should not be exceeded. Depending on the

stability of the test item in the solution, the stock solution can be prepared only once for the whole test

period and stored tightly closed under cool conditions in the dark (refrigerator, ca. 6±2°C). If the test

item is assumed to degrade quickly in the aqueous or acetone solution, the stock solution has to be

prepared freshly every day or of adequate time intervals. The final feeding solution is prepared from the stock solution or dilution of higher concentrated

solutions with 50% (w/v) aqueous sugar solution. The final solution should be prepared freshly every

day. Where acetone is used as solvent, two control groups will be necessary, i.e. one with pure 50% (w/v)

aqueous sugar solution and one with 50% (w/v) aqueous sugar solution containing the same content of

acetone as in the test item group. PROCEDURE Test and control groups The number of doses and replicates tested should meet the statistical requirements for determination of

LD50 with 95% confidence limits at the end of the test period. Normally, five doses in a geometric series,

with a factor not exceeding 2.2, and covering the range for LD50, are required for the test. However, the

dilution factor and the number of concentrations for dosage have to be determined in relation to the

slope of the toxicity curve (dose versus mortality) and with consideration taken to the statistical method

which is chosen for analysis of the results. A range-finding test enables the choice of the appropriate

concentrations for dosage.

3

37

In case of unknown toxicity, a preliminary range finding test can be used to derive appropriate

concentrations in the final test. For a dose-response test a minimum of thirty replicates (bees) should be used per test dose groups and

control. If acetone is used an additional solvent control with thirty replicates (bees) have to be included. Limit tests should be performed with fifty replicates (larvae) for the control and the test item treatment

and with at least thirty replicates for the reference item treatment. Reference item A reference item should be included in the test. The preferred toxic item is dimethoate (active ingredient

or formulated product). A minimum of one dose of the reference item, which leads to an expected

mortality of ≥ 50% at the end of the test period, should be used to prove the sensitivity of the bees and

the reliability of the test system. The expected dose response of dimethoate, as reference item for O.

cornuta and O. bicornis should be established by an inter-laboratory ring-test in future. Meanwhile, the

reported oral LD50 (48 h) values for O. cornuta of 0.32 µg a.s./bee (Hinarejos et al., in prep.) and O.

lignaria of 0.26 µg a.s./bee (Ladurner et al. 2005b), can be used as references. Exposure (dose feeding method) In the oral toxicity test for Apis mellifera (OECD TG 213 1998) the test solution is provided directly in

a solution feeder (syringe, test tube, etc) because honey bees recognize it as food. Ladurner studies

(Ladurner et al. 2003, 2005a, 2005b) showed that Osmia bees do not readily feed out of these devices.

The same studies propose a method (“flower method”) whereby most Osmia bees would feed within

minutes of exposure. This method has been simplified as “petal method” (Hinarejos et al., in prep.)

while maintaining similar feeding success ratios. In the “petal method”, the test solution is pipetted into a tiny plastic ampoule (inside diameter 2 mm,

outside diameter 3 mm, height 5 mm). This ampoule can be built with a pipette tip (0.1-10 µl) sealed at

the bottom. Sealing may be accomplished by approaching the tip to a heat source. The ampule is attached

to a petal and both are inserted into the base of floral foam (3 cm x 1.5 cm x 1 cm). Each Osmia females should be provided with 10 μL of test solution using an individual feeder with the

“petal method”. Bees are individually housed in the feeding test cages and kept in an incubator at 22 ±

2°C and 60 ± 20 % relative humidity under natural or artificial light. Bees are exposed to the feeding solutions over a period of 1 hour. The amount of feeding solution

consumed can be determined by weighing the feeders before and after feeding. Alternatively, only bees

that have consumed 100% of the 10 μL of test solution are considered. Post-exposure Housing cages After single exposure to the test solution, groups of bees, e.g. three sets of 10 bees per dose, are

transferred to housing cages. To transfer bees to the housing cages chilling at 4-5º C during 30 minutes

is recommend.

The housing cages are provided with an artificial feeder containing a sucrose solution 50% w/v as food.

Food should be provided ad libitum. The material of the holding cages should be similar to the individual

38

feeding cages and their size should be appropriate to provide adequate space. Plastic or cardboard cups

of ca. 12 cm diameter and 6 cm height are sufficient for 10 bees. To avoid confinement side effects, e.g.

bees stacking up, it is recommended to introduce a wire mesh in form of a small bridge (Hinarejos et al,

in prep). The artificial feeder can consist of a 5-mL Eppendorf vial, with a soaked cigarette filter inserted

through the lid of the vial, or a 1 or 2.5 ml syringe with the cut out, both with a glued petal as visual and

olfactory cues (Hinarejos et al., in prep). The used glue should have shown to be harmless to bees. Test conditions Bees should be held in an experimental room or incubator at: T = 22± 2 °C, RH = 60 ± 10 %, under

natural light or under an artificial 12:12 photoperiod. Temperature and humidity should be recorded

continuously with appropriate equipment. Duration The duration of the test is 48 h after the test solution has been replaced with sucrose solution alone. If

mortality continues to rise by more than 10 per cent after the first 24 h, the test duration should be

extended to a maximum of 96 h provided that control mortality does not exceed 10 per cent. Observations Mortality is recorded at 4 h after start of the test (initial feeding) and thereafter at 24h and 48h (i.e. after

given dose). If a prolonged observation period is required, further assessments should be made at 24

hours intervals, up to a maximum of 96h, provided that the control mortality does not exceed 10 per

cent. All abnormal behavioural effects observed during the testing period should be recorded. LIMIT TEST In some cases (e.g. when a test substance is expected to be of low toxicity or when a chemical is poorly

soluble) a limit test may be performed using a high dose (100 μg a.s./bee/day), or the maximum

achievable solubility, whichever is lower. The same procedure as above should be used, including the test group, the relevant controls, and the use

of the reference item. If mortality occurs, a full study should be conducted. If sublethal effects are

observed, these should be recorded. DATA AND REPORTING Data The data should be summarized in tabular form, showing for all test groups, the number of bees used,

mortality and number of bees with adverse behaviour at each observation time. Data on mortality at the

end of the test are analyzed by appropriate statistical methods (e.g. probit analyses) (Finney 1971) in

order to calculate the median lethal dose (LD50) values with 95 % confidence limits. Correction for

control mortality could be made according to the formula of Abbot (1925). Where treated diet is not

completely consumed, the dose of test substance consumed per bee should be determined. LD50 value

should be expressed in µg of test substance/bee:

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Test report The test report must include the following information: Test and reference item: • Details of the test and reference item (chemical identification data, purity, identity and

concentration of active substances);

Test bee species: • Details on the test species (scientific name, subspecies, age, incubation and collection method,

and information on the population used such as rearing methods, health status, pre-treatment

conditions, etc.); Test conditions: • Conditions during incubation, acclimatization (if applicable) and test period; • Description of the test cages (type, material, size); • Cocoon average weight (without fecal particules) and estimated adult weight according to Bosch

and Vicens (2002); • Method of the preparation of the stock solution and the feeding solutions; • Test design (number of treatment groups (test item, reference item, controls, number of replicate,

number of bees per cage); • Date of the start and the end of the test; Results: • Feeding success rate: percentage of bees that consumed the test solution within 1 h; • Mortality at each observation time for all treatments tested; • Uptake of feeding solution at each observation time for all treatments tested; • LD50 values with 95 % confidence limits for the test item and reference item at the each

recommended observation time;

• Description of all statistical procedures used in the study; • Any other biological effects observed e.g. sub-lethal effects, anti-feeding effects; • Deviations from the guideline and any other relevant information.

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Appendix B

Treatment Start Day 1 Day 2 Day 3 Day 6 Day 9 Day 12 Day 15 Day 18 Day 21 Day 24

Control 1 20 18 15 14 12 12 12 12 12 12 11 Control 2 20 18 17 15 12 12 12 11 11 10 10 Control 3 20 17 13 11 11 11 11 11 10 9 9

Thiacloprid 1 16 15 10 8 8 8 8 8 8 8 8 Thiacloprid 2 18 17 16 15 15 15 15 15 15 15 15 Thiacloprid 3 17 16 14 13 13 13 13 13 11 11 11 Thiacloprid 4 19 17 13 10 10 10 10 9 9 9 9 Thiacloprid 5 17 15 14 14 13 12 12 12 11 9 9

Tebuconazole 1 18 17 16 12 9 8 8 8 8 8 8 Tebuconazole 2 19 16 12 7 6 6 6 6 6 6 6 Tebuconazole 3 17 16 11 10 10 10 10 10 10 10 10 Tebuconazole 4 16 13 8 7 7 7 7 7 7 6 6 Tebuconazole 5 18 15 13 13 10 10 10 9 9 8 8

Mixture 1 18 17 14 13 12 12 12 12 12 12 12 Mixture 2 15 12 11 11 10 9 9 9 8 8 8 Mixture 3 18 17 14 13 12 12 12 12 12 12 12 Mixture 4 18 17 14 11 9 9 9 9 9 9 9 Mixture 5 18 16 13 10 10 10 9 9 9 7 7

Table B1: Osmia cornuta mortality data throughout the adult oral toxicity test. Numbers represent the number

of living bees.

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