Bacterial and ectoparasitic challenges imposed on Cyanistes caeruleus (blue tit) during nesting.

Andrew Devaynes BSc (Hons), PGCertHE

This thesis is submitted to Edge Hill University in partial fulfilment for the degree of Doctor of Philosophy

April 2018

Contents

Abstract

3

Chapter 1

Introduction

5

Chapter 2

Changing bacterial load of Cyanistes caeruleus (blue tit) occupied nest boxes and its potential impact upon breeding success

14

Chapter 3

Bacterial species richness at three stages of the breeding season in Cyanistes caeruleus (blue tit)

28

Chapter 4

Bacteria and ectoparasite presence in Cyanistes caeruleus (blue tit) nests

48

Chapter 5

Addition of vascular plant fragments to Cyanistes caeruleus (blue tit) nests and their protective qualities

70

Chapter 6

Conclusion

98

Acknowledgements

104

References

105

Appendix 1

Full invertebrate species list from Cytochrome c oxidase I primers

124

Appendix 2

Full plant species list from rbcL and ITS Uniplant primers

132

Andrew Devaynes

Presented for the degree of Doctor of Philosophy

Bacterial and ectoparasitic challenges imposed on Cyanistes caeruleus (blue tit) during nesting.

Abstract

Birds face many challenges during a breeding attempt including predation, food availability, pathogens and parasite load. Cyanistes caeruleus (blue tit) prefer human placed nest boxes, however the stable microclimate presented within the nest box exacerbates the challenges posed by ectoparasites and potentially pathogenic bacteria, with reductions in breeding success reported. The addition of green plant material to help control these deleterious effects has been reported within Mediterranean climes but comparable studies have not been undertaken in temperate regions.

This study introduces novel molecular approaches including next generation sequencing methods to assess the nest microbiome. This approach avoids the culturing bias of previous work in this area. Terminal-Restriction Fragment length Polymorphism (T-RFLP) analysis was used to assess bacterial richness progression through the breeding attempt with more traditional methods used to assess bacterial load. DNA barcoding was performed to identify bacteria, ectoparasites and vascular plant fragments present within the nest with vegetation surveys conducted around a subset of nests to assess if any active selection of plant material was occurring.

Bacterial species richness and load were relatively stable between nest build and clutch completion with a significant increase in both post fledging, following the introduction of nestling faeces in the nest and reduced time for nest sanitation. DNA barcoding provided marked increases in the taxonomic knowledge of nest dwelling biota with 169 bacterial taxa, thirteen species of ectoparasite and 154 vascular plant taxa identified.

Although ectoparasites and pathogenic bacteria were detected within the nest no effect was seen upon hatching or fledging success. It is more likely that a reduction in fitness would be observed post fledging. A high proportion of plant material containing volatile compounds was recorded within the nest, however active selection could not be confirmed.

Chapter 1 - Introduction

Birds construct a nest in which to lay their eggs. This nest can take a variety of forms from the barely hollowed mud scrape of Recurvirostra avosetta (avocet) to the ornate construction of Chlamydera nuchalis (great bowerbird). Regardless of their construction the role of the nest is to provide an incubation location for the eggs until hatching and then a safe location until fledging. Like all organisms, a component of reproductive success will be influenced by disease and other biota that the eggs, chicks and adult birds are in contact with within the nest.

Nests are made up of mostly plant material with feathers and mammal hair commonly used for their insulative properties, and as such provide a habitat for other biota to inhabit. This will include invertebrate species, which are well explored, and the development of a nest microbiome, which remains poorly understood, partly due to method limitations. Previous studies have been limited by culturable bias (Shawkey et al. 2003, Goodenough and Stallwood 2010, 2012, Gonzalez-Braojos et al., 2012 Jacob et al.. 2015,et al.) with only between 0.1 – 10% of bacteria able to be cultured under laboratory conditions (Grizard et al.., 2014). Other studies have removed the culturable bias; Benskin et al. (2015) utilising temperature gradient gel electrophoresis (TGGE) to assess wild bird faeces and Lee et al. (2014) and Grizard et al. (2015) using Roche 454 sequencing in assessing the microbiomes of wild bird eggs, however none have been used to assess the nest microbiome.

Next generation sequencing has revolutionised genomic research. It analyses small fragments of DNA in parallel, greatly reducing time and cost whilst increasing accuracy with its increased resolution allowing the analysis of community DNA (Behjati and Tarpey, 2013). An early iteration, Roche 454 sequencing developed in 2007, ceased development in 2013 due to its loss of competitiveness with the introduction of Illumina MiSeq in 2011 bringing improvements in read length, accuracy and cost (Liu et al., 2012). Illumina MiSeq has been utilised across a wide number of community studies including the microbiomes of sea bird plumage (Pearce et al., 2017), ant nests (Lucas et al., 2017) and soil communities (Moroenyane et al., 2018).

Cyanistes caeruleus (blue tit), a small passerine bird in the family Paridae, with a distribution throughout temperate and Mediterranean Europe and western Asia, residing mainly within deciduous woodland is a well-studied species. Their abundance, with an estimated 20-44 million breeding pairs in the UK (IUCN, 2016) and their affinity for occupying nest boxes makes them an ideal study species. Cyanistes caeruleus assess breeding sites and choose a mate over winter with nesting building beginning early April to coincide with the emergence of caterpillars, the key food item for their chicks. Eggs are laid one a day from late April with a clutch size of between six and thirteen the norm dependent upon food availability. Incubation is performed exclusively by the female and lasts fourteen to sixteen days before hatching. The altricial chicks require constant feeding which is predominantly supplied by the male as the female is needed to keep her featherless chicks warm. Chicks spend sixteen to eighteen days in the nest prior to fledging (BTO, 2018).

Previous studies have focused on: mate choice (Kempenaers et al., 1992), feeding behaviour (Naff-Daenzer and Keller, 1999) and factors influencing breeding success (Maicas et al., 2012). However there are no known studies investigating the nest microbiome in relation to breeding success without culturable biases in place. Remarkably, given humans’ close affinity and interest in bird species, bird – microbe research has received little attention outside of commercial poultry. This gap has persisted despite the known deleterious effects of ectoparasitism and pathogenic bacteria on breeding (Tripet and Richner 1999, Ichida et al. 2001, Moreno et al. 2009, Soler et al. 2012).

The preferred use of a nest box, when available, is likely due to the greater protection they offer against adverse weather and predation (Goodenough and Hart, 2011) with birds using natural tree cavities suffering higher losses due to soaking and predation (Wesolowski and Rowinski, 2012). Cyanistes caeruleus are thus classified as secondary cavity nesting species, utilising a cavity that is already present. As opposed to primary cavity nesting species who excavate their own cavity and those species who utilise an open nest (Martin and Li, 1992). However, nest boxes have been shown to harbour significantly higher ectoparasite loads given their favourable microclimate (Dawson 2004, Gwinner and Berger 2005, Moreno et al. 2009, Hebda and Wesolowski, 2012). A stable microclimate within the nest box could also be assumed to favour bacterial growth. Breeding success is subject to high selection pressure and thus determines life history traits (Peralta-Sanchez et al., 2012), thus birds often accept the fitness costs associated with ectoparasites and pathogenic bacteria over the increased chance of breeding failure in natural cavities.

This study will offer a greater understanding of potential factors influencing C. caeruleus breeding success in nest boxes. This in turn may be applied to the wider success of all box-using species, with an estimated 4.7 million nest boxes currently in place in the UK (Hanmer et al., 2017). These are used by a variety of species including Parus major (great tit), Sturnus vulgaris (starling), Ficedula hypoleuca (pied flycatcher), Sitta europaea (nuthatch), Passer domesticus (house sparrow), Passer montanus (tree sparrow) and Phoenicurus phoenicurus (redstart). Nest biome challenges in regards to breeding success of C. caeruleus will be investigated through the following:

Ecology of bacteria in birds’ nests

Bacteria within the nest originate from the nesting material, food items and the adult birds themselves (Mills et al., 1999) as they host a community of bacteria in their plumage. As the breeding season progresses faecal bacteria become more prominent given the reduced time the female bird has for nest sanitation, and thus the protective mucous covering of faecal sacs loses its protective coating (Ibanez-Alamo et al., 2014). Bacteria within the nest microbiome may be symbiotic, playing out beneficial and sometimes essential roles in digestion, nutrient synthesis and protection from pathogen colonisation (Jacob et al., 2014). Conversely, some bacteria are pathogenic with the potential to cause a reduction in fitness or death to an individual (Clayton and Moore, 1997). Certain bacterial species are capable of degrading β-keratin which constitutes more than 90% of feathers (Alt et al., 2015) and are thus collectively termed feather-degrading bacteria (Burtt and Ichida 1999, Shawkey et al. 2005). Any degradation of feathers can reduce fitness by affecting thermoregulation (Ichida et al., 2001) and flight efficiency (Moller et al., 2012) while any alteration to plumage colour can affect feather-based communication (Kilgas et al., 2012). Bacteria transmitted to the egg shell can penetrate through pores and infect the embryo, thus leading to embryo mortality (Soler et al., 2012). Pinowski et al. (1994) found Escherichia coli and Staphylococcus epidermidis present on eggs of Passer domesticus (house sparrow) and Passer montanus (Eurasian tree sparrow) which had failed to hatch. Mills et al. (1999) found that E. coli, Salmonella spp. and Shigella spp. presence in the plumage of Tachycineta bicolor (tree swallow) was positively correlated with wing asymmetry leading to a reduction in flying ability and thus survival.

However, not all bacteria within the nest microbiome are detrimental. Birds self-preen to condition their feathers through the secretion of uropygial oil. This contains beneficial, symbiotic bacteria, antibacterial compounds (Shawkey et al., 2003) and wax esters believed to be released as a food source for beneficial bacteria within the microbiome (Jacob et al., 2014). These defense mechanisms help to control the load of pathogenic and feather-degrading species (Soler et al. 2008, Martin-Vivaldi et al. 2009). Furthermore, Evans et al. (2016) found that increased exposure to bacteria within the nest increased the adaptive immunity of the chicks, enhancing their bacterial killing ability.

Challenges posed by ectoparasites in birds’ nests

Ectoparasites within the nest are potentially problematic to the female and chicks, not only withdrawing blood thus removing vital resources from the bird but also injecting toxins and disease-causing agents (Dubiec et al., 2013). Consequently nests with high ectoparasites loads are commonly rejected or abandoned (Loye and Carroll, 1998). However, nest site availability is constrained thus some birds have to accept and take on an ectoparasite infested nest (Hanmer et al., 2017). Ectoparasites associated with cavity-nesting birds include fleas, mites, blowfly larvae and black flies (Moreno et al., 2009).

Tripet and Richner (1999) found over 80% of C. caeruleus nests were infested with Ceratophyllus gallinae (hen flea) leading to a reduction in fitness of the female and chicks. P. major nests infested with C. gallinae resulted in a significantly lower fledging success and the chicks that did fledge exhibited reduced body mass, reduced tarsus length, reduced nutritional value and reduced haematocrit levels compared to control nests with fleas removed, thus the number and quality of fledglings was influenced (Richner et al., 1993). Sturnus vulgaris nests infested with Ornithonyssus sylviarum (northern fowl mites) resulted in chicks with reduced survival and innate immune defences, shown through a reduction in haematocrit levels, corticosterone concentrations and reduced bacterial killing ability (Pryor and Casto, 2017). Birds increase nest sanitation in response to a high ectoparasite load but, as with self-preening, this is another time consuming effort in an already energy demanding period and a trade-off against parental effort (Banbura et al., 2001).

Addition of fresh green plant material to nests

Cyanistes caeruleus nest composition broadly comprises a foundation of mosses with dry grass used to construct the nest cup and mammal hair and down feathers used for insulation (Clark and Mason, 1985). However little is known on the full composition of nests of C. caeruleus or similar species with most studies concentrating on broader aspects. These include nest mass in P. major (Lambrechts et al., 2017), determining choice of a few pre-selected plant species in nests of S. vulgaris (Ruiz-Castellano et al., 2018), limiting analysis to only three tree species in F. hypoleuca nests (Briggs and Deeming, 2016) and the use of anthropogenic material in nests of C. caeruleus and P. major. One area that has received greater attention is the continual addition of fresh green vascular plant fragments to the nest with differing hypotheses proposed. Clark and Mason (1985) proposed the nest protection hypothesis, where fresh green plant fragments added to the nests are rich in volatile compounds and control parasitic activity. Gwinner et al. (2000) proposed the drug hypothesis, where the volatile compounds stimulate an immune response, leading to increased haematocrit levels and white blood cell counts in chicks upon fledging. Lambrechts and Dos Santos (2000) proposed the potpourri hypothesis, where the birds maintain a cocktail of strong odours to increase the efficiency of defence against one or more pathogen/parasite species. Finally, Gwinner (1997) proposed the courtship hypothesis, where males bring fresh green plant fragments to the nest during construction to attract a mate, a behaviour that is most commonly observed within S. vulgaris.

Cyanistes caeruleus have demonstrated preference in their choice of nesting material. In a Corsican study only 6-10 plant species from over 200 available in the habitat were found within their nests (Petit et al., 2002) with plant fragments containing volatile compounds found in nests more often than expected, suggesting non-random selection (Pires et al., 2012). The volatiles dissipate over time, hence these fragments are added daily and quickly replaced following experimental removal (Petit et al., 2002).

The addition of such green plant material to nests was shown to be beneficial. Gwinner and Berger (2005) found that green plant material was associated with fewer bacteria and positively correlated with fledging success and body mass. Corsican C. caeruleus mostly preferred Achillea ligustica resulting in a reduction in bacterial richness and density on nestling skin and feathers (Dubiec et al., 2013). Mennerat et al. (2009c) also found the addition of aromatic plants reduced the culturable bacterial richness on nestlings. Soler et al. (2017) found the addition of green plant material to Sturnus unicolor (spotless starling) nests led to an increase in nestling fitness, measured through an increase in telomere length and reduced attrition.

Expanding knowledge through new methods within the field

The use of traditional, culture-based methods in previous studies is only offering a restricted view on the challenges posed to the birds through the nest microbiome. Additionally, the majority of studies have only sampled at a single time point giving no indication on how these challenges progress during a breeding attempt. One study applying a culture-independent technique, Automated Ribosomal Intergenic Spacer Analysis (ARISA), a microbial profiling technique, revealing 180 OTUs from P. major nests (Jacob et al., 2014), a considerable advancement on previous studies, was again limited in only sampling at a single time point. Previous work investigating ectoparasites within nests have used morphological features. While the larger size of ectoparasites compared with bacteria, makes this a valuable method, it can exclude species at low abundance or that have left the nest with the host. Hence DNA approach is likely to be more sensitive.

This study will assess bacterial load and richness at three key stages within the breeding attempt; nest build completion, clutch completion and immediately post fledging. Bacterial richness will be assessed using culture-independent Terminal-Restriction Fragment Length Polymorphisms (T-RFLP), the first use within the field but established in the fields of soil community microbiology (Fry et al. 2016, Gschwendtner et al. 2016), marine microbiology (Tada et al., 2017) and within agriculture (Kaur et al. 2017, Zhu et al. 2017). Nests collected post fledging will be subjected to community DNA extraction and barcoding to identify the plants used in nest construction and bacterial and ectoparasite presence within the nests. This is a proven community assessment technique and will be the first use of next generation sequencing within the field of passerine nest studies.

Study sites

This study will be conducted across six sites; four in Lancashire, one in Gloucestershire and one in Scotland (further detail on site location can be found within chapter 2, Table 1 and Figure 1). An overview of each site and the nest boxes within is provided below.

Ruff Wood – mixed deciduous, broad-leaved woodland covering eight hectares. No previous nest boxes, twenty placed for this study of a uniform wooden construction with internal dimensions 110mm width, 170mm depth and 210mm mid-point height, situated at a height of 3m in an orientation between North and East.

Scutchers Acres – mixed deciduous, broad-leaved and coniferous woodland covering eleven hectares. No previous nest boxes, twenty placed for this study of a uniform wooden construction with internal dimensions 110mm width, 170mm depth and 210mm mid-point height, situated at a height of 2m in an orientation between North and East.

Gorse Hill nature Reserve – mosaic of deciduous, broad-leaved woodland, grassland and arable land covering fifteen hectares. Nest boxes present with monitoring from local volunteer groups, of uniform wooden construction with internal dimensions 110mm width, 170mm depth and 210mm mid-point height, situated at a height of 2m, placed at a random orientation.

Mere Sands Wood Nature Reserve – mixed deciduous, broad-leaved woodland covering 42 hectares. Nest boxes present with monitoring from local volunteer groups, of varying size and wooden construction, situated at a height of 4m, placed at a random orientation. Some nest boxes were in various states of disrepair with occasional dampness observed during sampling.

Scottish Centre for Ecology and the Natural Environment (SCENE) – oak dominated deciduous, broad-leaved woodland covering approximately 100 hectares. 500 nest boxes present, managed by Glasgow University. Nest boxes are cylindrical and of wood-crete construction with an internal diameter 170mm and mid-point height 230mm, situated at a height of 3m to 4m, placed at a random orientation.

Nagshead Nature Reserve – oak dominated deciduous, broad-leaved woodland covering 308 hectares. 400 nest boxes present, managed by RSPB. Nest boxes are of uniform wooden construction with internal dimensions 110mm width, 170mm depth and 210mm mid-point height, situated at a height of 3m, nest boxes utilised in this study were orientated North of East to West.

Although C. caeruleus are listed as a species of least concern with an increasing population trend (IUCN, 2016) it is still ethical to minimise disruption and disturbance during their breeding attempt. Thus swabbing of the nest material was only ever carried out whilst the birds were absent from the nest during nest build and clutch completion and the final nest swabs were only taken post fledging.

Research aims

This thesis aims to enhance current knowledge on the challenges faced by C. caeruleus during their breeding attempt and assess what efforts they make to help address those challenges. Through the use of microbial profiling techniques and next generation sequencing, data absent in the literature through previous methodological biases will be obtained.

Thesis outline

Chapter 2 investigates the bacterial load of nest boxes and how it progresses during a breeding attempt, sampling at nest build completion, clutch completion and immediately post fledging. The bacterial load of potentially pathogenic species is also investigated through the use of selective media and any relationship with hatching or fledging success analysed.

Chapter 3 again investigates progression of the microbiome but this time looking at species richness at the different stages. Terminal Restriction Fragment length Polymorphism (T-RFLP) was used to avoid the culturable bias associated in previous studies.

Chapter 4 looks to identify the bacteria and ectoparasites within the nest using DNA barcoding, again removing the culturable bias of previous bacterial work. This data will inform the challenges the birds face within the nest and the likely fitness costs experienced.

Chapter 5 investigates the vascular plant fragments added to the nest whilst determining what role they may play in controlling the bacterial and ectoparasite load within the nest box.

Chapter 6 will set out the overall conclusions of the thesis, describe their implications for birds utilising nest boxes, help inform nest box management and suggest avenues for future research.

Chapter 2 - Changing bacterial load of Cyanistes caeruleus (blue tit) occupied nest boxes and its potential impact upon breeding success

Abstract

Cyanistes caeruleus prefer to use nest boxes to raise their young rather than nests in natural tree cavities. However, nest boxes offer a warm, humid microclimate favourable to rich bacterial communities. This study investigated how the bacterial community developed throughout the breeding season and whether it had any effect on embryo or nestling mortality. Samples were collected across six sites and three breeding seasons at nest build completion, clutch completion and immediately post fledging. Bacterial counts were obtained for each sample, a total count on generic agar whilst also selecting for Staphylococcus spp. and Enterobacter spp., which may indicate pathogenicity to the birds. There was significantly more bacteria (generic, Staphylococcus spp. and Enterobacter spp.) present within the nest box post fledging following nestling feeding and defecating, and reduced time for self-preening and nest sanitation from the adult birds. No positive relationship was found with either embryo mortality, nestling mortality or brood size, however a negative relationship between embryo mortality and generic bacterial count was discovered. Although somewhat unexpected an increase in symbiotic bacteria could offer a greater level of protection.

Introduction

Nesting birds face many potential threats to breeding success, of which predation and adverse weather are major considerations when choosing a nesting site. Breeding success is subject to a high selection pressure and therefore plays an important role in determining the evolution of life history traits of birds, especially behaviour within the nest, including nest site selection, nest material selection and incubation behaviour (Peralta-Sanchez et al., 2012). Most hole-nesting birds prefer to use human placed nest boxes when available as these are sturdier and offer greater protection than natural tree cavities (Goodenough & Hart 2011). Birds using natural tree cavities, suffer higher losses to predation, soaking and predation of adults on the nest (Wesolowski and Rowinski, 2012).

Studies have shown that nest boxes harbour significantly higher loads of ectoparasites than natural nests in tree cavities for C. caeruleus (Moreno et al. 2009, Hebda and Wesolowski 2012), Parus major (Great tit; Hebda and Wesolowski 2012), Sturnus vulgaris (European starling; Gwinner and Berger 2005), Tachycineta bicolor (Tree swallow; Dawson 2004) and Ficedula hypoleuca (Pied flycatcher; Moreno et al. 2009). The issue of high ectoparasite loads in nest boxes (mainly fleas, mites and blowfly larvae) can therefore be deemed to be an association with the nest box itself rather than an individual species. Consequently nest re-use is rare, most birds will construct a new nest for each breeding attempt to reduce exposure of themselves, their eggs and nestlings to ectoparasites and potentially rich bacterial communities (Newton, 1994). However, for cavity nesting passerines nest site availability is the main factor constraining reproduction, consequently old nest cavities and nest boxes may be reused.

In contrast to ectoparasites, bacterial loads of the nest box has received little attention despite the possibility of them being pathogenic agents and the nest box offering favourable conditions for growing bacterial communities given their stable microclimatic conditions (Gonzalez-Braojos et al., 2012a).

Bacteria within the nest originate from the nesting material, food items and the adult birds themselves (Mills et al., 1999) with the adult birds playing host to a community of bacteria in their plumage. These bacteria can be separated into two ecological classes: free-living and attached. Free-living bacteria are labile whilst attached are a much more stable community. Bacteria attached to the plumage can be a problem for the bird itself, as certain species are capable of degrading β-keratin which constitutes more than 90% of feathers (Alt et al., 2015). In relation to nest re-use Gonzalez-Braojos et al. (2012b) found F. hypoleuca nestlings raised in old, reused nests harboured higher bacterial loads on their belly skin than those reared in freshly built nests. Bacteria can also be transmitted on to the egg shell, penetrate through pores and infect the embryo, reducing hatching success (Soler et al., 2012). Increased bacterial abundance transmitted to the egg shell increases the chances of embryo infection. Pinowski et al. (1994) found Passer domesticus (house sparrow) and Passer montanus (tree sparrow) eggs which had failed to hatch were infected with Escherichia coli and Staphylococcus epidermitis. Therefore selection should favour birds who try to limit the bacterial load of their plumage to control potential contamination (Shawkey et al., 2009). One method exercised by the birds to control plumage bacteria is self-preening. During preening uropygial oil is secreted which contains beneficial, symbiotic bacteria which can control the load of pathogenic, feather degrading species (Soler et al. 2008, Martin-Vivaldi et al. 2009). Preening is however a time consuming process and during the breeding season there is a trade off with parental effort (Lucas et al., 2005). This leads to increased free living bacteria abundance within the plumage which are more likely to be transmitted on to the egg (Alt et al., 2015).

Upon hatching, nestlings are essentially sterile but are quickly colonised by bacteria from the nesting material, food items and adult birds (Mills et al., 1999). The colonising bacteria may be commensal or indeed symbiotic, aiding in food digestion and complementing the immune system. However, some bacteria may be pathogenic with the ability to affect the growth and survival of altricial nestlings. Feather degrading bacteria can be transmitted to nestlings, which can lead to problems with thermoregulation (Ichida et al., 2001), flight efficiency (Moller et al., 2012) and any alteration to plumage colour can affect feather based communication (Kilgas et al., 2012). Larger broods equate to higher parental intensity and reduced nest and self-sanitation (Cantarero et al., 2013) which correlates with increased bacterial loads (Gonzalez-Braojos et al. 2012b, 2014). Alt et al. (2015) found female F. hypoleuca with experimentally reduced broods had the lowest number of free living bacteria on their feathers. Conversely, females with the male experimentally removed to assist in feeding the nestlings had the highest. Nestling fecal sacs have a mucous coating encasing the fecal bacteria, however the mucous covering only offers protection for 23 minutes (Ibanez-Alamo et al., 2014) and given the increased workload of the female they can rarely be removed within this time period. In a similar study on S. vulgaris, Lucas et al. (2005) also discovered an increase in free living bacteria however there was no change in the attached bacterial community.

Of the few studies to date to identify bacteria within a nest box, Goodenough and Stallwood discovered 32 species of bacteria (2010) and 28 species (2012) within those of C. caeruleus and P. major. Of the bacteria found, Enterobacter cloacae and Staphylococcus hyicus are negatively associated with fledging success. Berger et al. (2003) identified 12 genera of bacteria in the nests of S. vulgaris, of which Klebsiella sp. and Enterococcus sp. were the most prevalent. Of the bacteria identified, the authors could not determine pathogenicity due to the fact that no bacterial load was calculated. Where a bacterial species may be part of the normal microbial flora and be beneficial, a considerable increase in their numbers can lead to them becoming pathogenic to the host. Studies investigating the bacterial loads of nest boxes have only sampled at a single point whilst nestlings are present in the nest, with the exception of Gonzalez-Braojo et al. (2012a, 2014) sampling at day 7 and 13 post hatching in F. hypoleuca nests and Gwinner and Berger (2005) who sampled day 1, 9 and 14 in S. vulgaris nests. There have been few studies investigating the bacterial load prior to hatching, despite bacteria being closely linked with embryo fatality (Soler et al., 2012). This study aims to address some of these gaps in the literature by investigating the bacterial community development in C. caeruleus nest boxes during the breeding season and whether any impact is made upon the breeding attempt.

Cyanistes caeruleus are a suitable model organism as they readily accept the shelter of a nest box occupying up to two thirds of those available at the study sites (personal observation). There are 20-44 million breeding pairs in Europe (RSPB, 2016) and the species is classified as least concern on the IUCN red list of threatened species with an increasing trend in population (IUCN, 2016). The aim of this study is to discover: (1) bacterial load of the nest box at nest build, once eggs are present and immediately post fledging; (2) any relationship between the bacterial count at clutch completion and embryo mortality; (3) any relationship between the bacterial load immediately post fledging and chick mortality; and (4) any relationship between bacterial load and brood size in C. caeruleus.

Methods

Study sites

The study was performed in the breeding seasons of 2014-2016 across six sites; predominantly in the North West United Kingdom (see Fig 1) though covering a north south range of 500km. Table 1 gives a description of each site.

Table 1: Study site location, description and nest box status

Site

Map reference

Vegetation type

Nest boxes

Ruff Wood

53°33′36.27″N, 002°51′59.43″W

Mixed deciduous woodland.

None present, 20 placed for this study.

Scutchers Acres

53°35′17.68″N, 002°49′23.79″W

Mixed deciduous and coniferous woodland.

None present, 20 placed for this study.

Gorse Hill nature Reserve

53°33′41.45″N, 002°54′53.46″W

Mosaic of woodland, grassland and arable land.

Present, some monitoring from local volunteer groups.

Mere Sands Wood Nature Reserve

53°38′05.59″N, 002°50′16.06″W

Mixed deciduous woodland.

Present, some monitoring from local volunteer groups.

Scottish Centre for Ecology and the Natural Environment

56°07′13.72″N, 004°35′34.70″W

Oak dominated deciduous woodland.

500 present, managed by Glasgow University.

Nagshead Nature Reserve

51°46′25.59″N, 002°34′20.76″W

Oak dominated deciduous woodland.

400 present, managed by Royal Society for the Protection of Birds.

Figure 1: Geographical representation of study sites; a – Scottish Centre for Ecology and the Natural Environment (SCENE), b – North West sites (Ruff Wood, Scutchers Acres, Gorse Hill Nature Reserve, Mere Sands Wood Nature Reserve) within a 7km radius, c – Nagshead Nature Reserve.

Field sampling

Nest boxes were observed to determine breeding activity and bacterial samples taken under Natural England licence (2014/SCI/0288) at nest build completion, clutch completion and immediately post fledging.

Data were collected on clutch size, number of young to hatch and number of young to fledge. Data from the Scottish Centre for Ecology and the Natural Environment (SCENE) and Nagshead Nature Reserve was obtained from weekly nest box inspections already in place as part of their management. For all other sites clutch size was determined from observation at the clutch completion sampling point of the number of eggs present and whether they were warm and therefore undergoing incubation or cold indicating the clutch had yet to be completed. Hatching and fledging success were calculated once the young had fledged from the observation of any unhatched eggs and deceased nestlings within the nest box. Nesting stages were determined as; nest build completion, nest constructed to either N4 or NL, clutch completion, post swabbing eggs were felt to discover whether warm and thus under incubation indicating the clutch complete or cold indicating an incomplete clutch which were then excluded from analyses, post fledging within two weeks of the chicks leaving the nest.

Sterile cotton tipped swabs (Fisher brand, UK) were pre-moistened in phosphate buffer (pH7.1, 0.2M, Sigma Aldrich Ltd, Dorset, UK) and the nesting material was swabbed for 30 seconds starting in the back left corner of each nest box using a side to side motion until the front right corner had been reached. This was performed by the same person with particular effort to keep this process standardised and remove inter-operational variability. Eggs would be inadvertently swabbed during the clutch completion stage, as would faeces within in the nest at post fledging, given the eggs/birds would be exposed to any bacteria identified this was deemed justifiable and relevant. Swabs were stored in individual 15ml Falcon tubes (VWR, UK) containing 1ml of phosphate buffer (0.2M, pH7.1, Sigam Aldrich Ltd, Dorset, UK). All samples were processed for culturing within 4-6 hours.

Bacterial counts

Bacterial counts of the samples were obtained in the form of CFU/ml (colony forming units) using the Miles and Misra (1938) method. Samples were vortexed for 5 seconds before a series of tenfold dilutions were made to 10-5 using sterile phosphate buffer. Samples were plated onto Nutrient agar (Oxoid CM0309, Fisher Scientific, UK) to achieve an overall count of culturable bacteria, Baird Parker agar (Oxoid CM0275, Fisher Scientific, UK) with the addition of Egg Yolk Tellurite (Oxoid SR0054, Fisher Scientific, UK) to select for Staphylococcus spp. and Eosin Methylene Blue agar (Oxoid CM0069, Fisher Scientific, UK) to select for Enterobacter spp. across the three sample years. Additionally in 2016, MacConkey agar (Oxoid 0007, Fisher Scientific, UK) was utilised to select for Escherichia coli and Soya Flour Mannitol (20g soya flour, 20g mannitol, 20g agar per one litre of tap water, Fisher Scientific, UK) with the addition of 1ml nalidixic acid (25mg/ml, Fisher Scientific, UK) and 1ml nystatin/dimethyl sulfoxide (DMSO) solution (5mg/ml, Fisher Scientific, UK) incubated at 25˚C to determine the presence of any Actinomycetes. Agar plates were divided into six sections and labelled 10-0 to 10-5 before being placed in a drying cabinet for one hour. Three 20µl drops of each dilution were pipetted onto the relevant section of the agar to serve as triplicate repeats and plates were kept upright for 30 minutes allowing the samples to be absorbed before inverting and incubating for 48 (+/-2) hours at 37˚C. Each section was observed for growth with the first dilution showing discrete individual colonies counted and recorded. The mean of the triplicate colony counts within the countable dilution was obtained and CFU/ml calculated.

Statistical analysis

Nonparametric analyses were utilised due to the absence of normal data, even after transformation using log(x+1). Analysis was undertaken using R version 3.2.3 (R Core Development Team; R Studio, 2017). A Freidman test with post-hoc Nemenyi was used to determine any difference in bacterial load at the three sample points. Analysis of embryo mortality and chick mortality was heavily skewed by 0% occurrence in both instances, and although significant correlation was found in some instances, scatter plots revealed there was no clear relationship between bacterial counts and either embryo or chick mortality. The data were therefore converted binomially to give any occurrence of embryo/chick mortality a value of 1 and no occurrence a value of 0. Spearman’s rank correlation followed by binomial regression was then performed to determine any significant relationship. To determine any relationship between bacterial load and brood size, Spearman’s rank correlation followed by Poisson regression analysis was performed.

Results

Generic bacterial counts (CFU/ml) across the nest build and clutch completion stage showed no significant difference (Figure 2). There was a slight decrease in median upon incubation but variation within nests was higher at this stage. Post fledging the bacterial count increased greatly showing a highly significant difference to the previous two stages (p<0.001).

Figure 2: Bacterial CFU/ml in the three stages of nesting in blue tits (build, eggs and fledged; mean ± 0.95 confidence interval). Build-eggs p=0.78, build-fledged p<0.001, eggs-fledged p<0.001, n=434.

Figure 3: Bacterial CFU/ml of Staphylococcal species in the three stages of nesting in blue tits (build, eggs and fledged; mean ± 0.95 confidence interval). Build-eggs p=0.92, build-fledged p<0.001, eggs-fledged p<0.001, n=474.

Bacterial counts for Staphylococcus spp. (Figure 3) showed very little difference between nest build and clutch completion (p=0.92), at these two stages, in 37% of nests no Staphylococcus spp. were detected. Post fledging this dropped to 13% and in significantly higher numbers (p<0.001).

Bacterial counts for Enterobacter spp. (Figure 4) showed a greater difference between nest build and clutch completion however this was not significant (p=0.73). At nest build 63% of nests had no Enterobacter spp. present, reducing to 55% at clutch completion. Post fledging 79% had Enterobacter spp. present and in significantly higher numbers (p<0.001).

Figure 4: Bacterial CFU/ml of Enterobacter species in the three stages of nesting in blue tits (build, eggs and fledged; mean ± 0.95 confidence interval). Build-eggs p=0.73, build-fledged p<0.001, eggs-fledged p<0.001, n=426.

Figure 5: Bacterial CFU/ml of Escherichia coli in the three stages of nesting in blue tits (build, eggs and fledged; mean ± 0.95 confidence interval). Build-eggs p=0.10, build-fledged p<0.001, eggs-fledged p<0.001, n=426.

Bacterial counts for E. coli (Figure 5) reported the least incidence across the utilised agars with only 47% occurrence at nest completion, 18% at clutch completion and 81% upon fledging. There was little change in bacterial counts between nest and clutch completion, however counts upon fledging were significantly higher (P<0.001). Bacterial counts obtained from Soya Flour mannitol agar (Figure 6) showed the highest counts at nest and clutch completion and counts were present in all nests across all stages. Counts between nest and clutch completion differed non-significantly (p=0.52), however they greatly increased upon fledging (p<0.001) Bacterial counts between nests varied greatly across all five agars.

Figure 6: Bacterial CFU/ml (when incubated at 25°C) in the three stages of nesting in blue tits (build, eggs and fledged; mean ± 0.95 confidence interval. Build-eggs p=0.52, build-fledged p<0.001, eggs-fledged p<0.001, n=426.

Generic bacterial count on Nutrient agar and embryo mortality were significantly negatively correlated (rs=-0.25, p<0.001, n=108), with binomial regression also showing significance (z=-2.058, p=0.040, d.f. =1, 106) with 12.75% of embryo mortalities being explained by the model. Neither Staphylococcus spp. counts (rs=0.18, p=0.85, n=111), Enterobacter spp. counts (rs=-0.04, p=0.66, n=111), E. coli counts (rs=-0.15, p=0.40, n=111) nor the generic count gained from Soya Flour Mannitol agar (rs=-0.11, p=0.55, n=111) showed significance with embryo mortality.

There were no relationship between bacterial counts and chick mortality on either generic or selective agars. Generic count on Nutrient agar (rs=0.07, p=0.50, n=111), Staphylococcus spp. (rs=0.16, p=0.09, n=111), Enterobacter spp. (rs=0.13, p=0.18, n=111), E. coli (rs=-0.15, p=0.40, n=32) nor generic count on Soya Flour Mannitol agar (rs=-0.11, p=0.55, n=32). Brood size had no significant relationship with either the generic bacterial count on Nutrient agar (rs=0.02, p=0.83, n=111), Staphylococcus spp. (rs=0.12, p=0.21, n=111), Enterobacter spp. (rs=0.009, p=0.93, n=111), E. coli (rs=0.15, p=0.40, n=32) nor generic count on Soya Flour Mannitol agar (rs=-0.16, p=0.37, n=32).

Discussion

The main result of this study is the significantly higher number of bacteria in the nests in the fledged state compared to all other stages with minimal difference between nest build completion and clutch completion. A significant negative correlation between generic bacterial count and embryo mortality was also discovered. Although the generic bacterial load showed only a minimal difference between nest build completion and clutch completion, there was a marked increase in the variation of counts between nests upon the initiation of incubation. The bacterial load of Staphylococcus spp. and Enterobacter spp. followed a similar pattern. In the process of laying eggs, bacteria from the female’s cloacal cavity are introduced in to the nest, of those Enterobacter spp. and Coliform spp. were found to be prevalent (Sanders et al., 2005). Upon incubation the temperature of the nest cup and eggs is raised to between 35-38˚C (Nord and Nilsson, 2011), this is closely related to the body temperature of the adult birds and as such the optimum temperature for host associated bacteria. From this it could be expected that the bacterial load would increase significantly compared with the ambient temperature it had been exposed to previously. However this was not the case. Possible explanations could include a lack of nutrients to facilitate growth or swabbing took place early on in the incubation when the bacteria had received insufficient time to benefit from the improved conditions. The significantly higher bacterial loads for generic, Staphylococcus spp. and Enterobacter spp. is associated in the main to the increased activity within the nest. Both parents are continually bringing food leaving little time for preening and nest sanitation, making the removal of faecal sacs before there protective mucous covering deteriorates less achievable The food itself and nestling faeces also offer a constant source of bacteria. Benskin et al. (2015) analysed C. caeruleus faeces and found 55 bacterial OTU’s, although no bacterial load was investigated it illustrates how diverse the bacterial communities within the faeces are.

Embryo mortality showed a significant negative relationship with generic bacterial counts at clutch completion, as bacterial counts increased embryo mortality fell. This is perhaps unexpected. It may be explained by the female removing belly feathers prior to incubation to aid in efficient temperature transfer to the eggs (Nord and Nilsson, 2011). These feathers are then used as lining material in the nest cup. Uropygial oil from the preening process will still be present on these feathers (Peralta-Sanchez et al., 2012) which then provides a community of beneficial bacteria that aid in the control of pathogenic bacteria. These symbiotic bacteria are generally superior competitors compared to their pathogenic counterparts (Brook, 1999) due to co-evolutionary interactions between host and microorganism (Hackstein and van Alen, 1996), bacterial interference through the production of antibiotic substances to impede the establishment of competing bacteria (Riley and Wertz, 2002) or the host modifying its environment for symbiotic bacteria (Hackstein et al., 1996). Although the bacterial load is increased it is to the benefit of the eggs in protecting them from possible pathogens.

No relationship was found between the load of Staphylococcus spp., E. coli or Enterobacter spp. with embryo mortality, despite E. coli and S. epidermitis being associated with embryo mortality within sparrow nests (Pinowski et al., 1994). Pathogenic bacteria present on the egg need to travel through pores in the shell to enable them to infect the embryo. This process is heavily dependent on the presence of water and a major benefit is the increased waterproofing of nest boxes compared with natural cavities. Nest boxes are more prone to condensation, but this is offset by maintaining a warm incubation temperature (D’Alba, 2010). Antimicrobials present within the egg white are a further defence against any bacteria that are able to breach the shell in protecting the embryo.

The data showed no relationship between chick mortality and any of the bacterial counts, supporting the results of Berger et al. (2003). Nestlings are exposed to many potential mortality factors with bacterial load and type likely to be a minor one. Significant factors include; nutrition (Gonzalez-Braojos et al., 2012b) and ectoparasite load (Richner et al., 1993). A more prominent relationship with bacterial load may be a measure of nestling fitness. Benskin et al. (2015) found Bacillus licheniformis was the most prevalent amongst isolates, detected in nearly three quarters of C. caeruleus faecal samples. B. licheniformis can be a feather degrading bacteria which may influence feather condition and coloration (Gunderson, 2008). A reduction in feather quality has a detrimental effect on thermoregulation and flight and any change in coloration can affect mate choice. Feather degrading bacteria are transient within the nest environment with those isolated on the feathers of parents subsequently found in the cloaca of nestlings (Giraudeau et al., 2010).

No relationship was discovered between brood size and bacterial counts, counter to that found by several previous studies (Lucas et al. 2005, Gonzalez-Braojos et al. 2012, 2014, Cantarero et al. 2013, Alt et al. 2015). However these studies concerned plumage bacteria sampled from the feathers of the birds. Of these, attached bacteria are less likely to be found on the nesting material, and although free living plumage bacteria have shown to be transient and could be present, they may have little effect on bacterial load of the nesting material.

In conclusion bacterial counts increase significantly upon the hatching of the eggs, consequently the chicks are exposed to potentially pathogenic bacteria. These have shown to have no detrimental effect on embryo or chick mortality but may influence the general fitness of the nestlings whilst in the nest and upon fledging and initiate an increase in parental effort through compensatory responses such as increased foraging and nest sanitation at an already energy demanding time A longitudinal study measuring both fecundity and survivorship would enable a more thorough examination of the effect the nest bacterial load has upon the adult birds and chicks. Using selective agars gives only an indication of the presence of a species within the selected genus, therefore the pathogenic species may not be present. Although bacterial load gives a useful indication to the pressures the chicks face in their first stage of life, the next step is to investigate bacterial richness at each stage of the breeding process and to identify the bacteria present within the nest box.

Chapter 3 - Bacterial species richness at three stages of the breeding season in Cyanistes caeruleus (blue tit)

Abstract

Blue tits are exposed to a vast array of bacteria throughout their life cycle and are particularly exposed during a breeding attempt. Any pathogenic bacteria within their microbiome can have a detrimental effect on their fitness and that of the nestlings they are raising. This study aims to identify the bacterial species richness that birds of this species are exposed to during three key stages of the breeding cycle: nest build, clutch completion and immediately post fledging. Nests were swabbed at these time points across four deciduous woodland sites in the United Kingdom and genomic DNA extracted prior to T-RFLP analysis. This is the first known instance of this technique being used to assess the nest microbiome and the first culture independent assessment of nest microbiome within this species. This revealed 103 distinct OTUs across all sites and stages with an increase in taxa richness at each stage. There were differences in the microbiomes of each nest across breeding stage and site with evidence suggesting the nest microbiome is largely determined by the local environment.

Introduction

Bacteria form a large part of Earth’s biomass, outnumbering animal and plant cells and have adapted to be able to survive and thrive in all known conditions (Jacob et al., 2014). Symbiotic relationships between animals and bacteria are omnipresent and can play a significant role in animal evolution (van Veelen et al., 2017). Bacteria can be hugely beneficial and in some cases essential in processes such as digestion, nutrient synthesis and protection from pathogen colonisation (Jacob et al., 2014). There is increasing evidence over the importance of host – microbiome relationships to the survival of species (Glasl et al., 2016) with indications that many have co-evolved (Goodrich et al., 2016). Conversely, some bacteria are pathogenic, with the potential to cause a reduction in fitness or death to an individual, as such they impose strong selective pressures on host life-history traits (Clayton and Moore, 1997). It is therefore of upmost importance to obtain knowledge of an individual’s microbiome and the challenges it faces when exposed to new sources of bacteria and any associated pathogenicity (Burtt and Ichida, 1999).

Plumage microbiomes have been the main focus of bird microbiome studies (i.e. Burtt and Ichida 1999, Shawkey et al. 2005, Kilgas et al. 2012) with Kilgas et al. (2012) identifying wild bird’s plumage play host to a vast microbiome with evidence suggesting it influences the host behaviour and life histories. Van Veelen et al. (2017) investigated both plumage and nest microbiomes, finding bird and nest-associated bacteria showed substantial OTU co-occurrences sharing dominant taxonomic groups within Lullula arboea (woodlarks) and Alauda arvensis (skylarks). Similarly, Goodenough et al. (2017) discovered the skin and feather microbiome of individual female Ficedula hypoleuca (pied flycatchers) were closest to their nest microbiome. This convergence is likely due to bi-directional transfer of bacteria between the nest and plumage given the close contact between the female and nest during the breeding attempt. It is thus reasonable to assume that studies relating to only plumage microbiome or nest microbiome offer a fair comparison to each other.

All organisms have competing energy demands and this is especially true for female birds during the breeding season (Monclus et al., 2017). One energy demand exercised by the birds is to control plumage bacteria by self-preening. During preening, uropygial oil is secreted which contains beneficial, symbiotic bacteria, antibacterial compounds (Shawkey et al., 2003) and wax esters believed to be released as a food source for beneficial bacteria within the microbiome (Jacob et al., 2014), all of which can control the load of pathogenic, feather-degrading species (Soler et al. 2008, Martin-Vivaldi et al. 2009). Preening is however a time-consuming process and during the breeding season there is a trade-off with parental effort (Lucas et al., 2005). Initially this includes nest building, followed by egg incubation and then nest sanitation and collecting food for the chicks once hatched. This activity leads to increased free-living bacteria abundance within the plumage (Alt et al., 2015). Bacteria within the plumage microbiome can be a problem as certain species are capable of degrading β-keratin which constitutes more than 90% of feathers (Alt et al., 2015). Feather degrading bacteria can reduce fitness through thermoregulation problems (Ichida et al., 2001) and lower flight efficiency (Moller et al., 2012) while any alteration to plumage colour can affect feather based communication (Kilgas et al., 2012). Altricial nestlings are prone to such reductions in fitness which can affect their long term survival. However the birds may not be passive recipients of such increase in bacteria. Experimentally altering the bacterial density that Parus major (great tits) were exposed to resulted in an increase in the size of their uropygial gland and quantity of secretions (Jacob et al.., 2014). They do this despite the already increased energy demand of the breeding effort and an increased probability of olfactory detection by predators, illustrating the importance that birds place upon actively influencing their microbiome.

During a breeding attempt nest-building birds are exposed to an ever-changing and highly variable community of bacteria. For instance, P. major were at their most ‘dirty’ during the nest building phase with an increase in microbial density (Kilgas et al., 2012). It is hypothesised that this is a result of increased contact with the ground and nest materials, introducing new and potentially pathogenic bacteria to the bird. Upon laying, bacteria from the cloacal cavity are released on to the nest (Sanders et al., 2005), and what may have been a commensal organism within the gut could have a pathogenic effect on skin or feathers. Following hatching food items are constantly being brought in to the nest as well as the chicks defecating which gives further avenues for bacteria to be introduced (Benskin et al., 2015). We have previously identified C. caeruleus are subjected to an increasing number of bacteria during the breeding attempt (Chapter 2) supporting the introduction of additional bacterial taxa.

Many of the studies which identify the microbiome within the nest are restricted by culturable bias with only 0.1-10% of microbes able to be cultured under lab conditions (Grizard et al., 2014), whilst this is acceptable for comparisons within a study it does not give a true indication of the total bacterial species richness the birds are exposed to. Those using culture-independent techniques are few, with only Jacob et al. (2014) finding 180 OTUs across 52 P. major nests present within the literature. This study was limited as they sampled only once during the breeding attempt so no information was gathered on the development of the nest microbiome over this period. Kilgas et al. (2012) found the density of plumage bacteria changed rapidly during nest building, although here they only assessed bacterial density and not community composition.

This study aims to address this gap in the literature offering an additional non-culture bias comparison and investigating the bacterial species richness of C. caeruleus nests at three time points; nest build, clutch completion and immediately post fledging using Terminal – Restriction Fragment length Polymorphisms (T-RFLP) analysis. This will allow true comparisons on how the bacterial community composition progresses during a breeding attempt. Additionally given that Goodenough et al. (2017) found the nest microbiome to be dependent upon the local environment, samples will be collected across four sites to discover any intra-site difference between nests and inter-site difference between the relative local environments. T-RFLP analysis has been shown to be a robust and reproducible methodology for rapid exploration of microbial community structure with a higher resolution for detecting less abundant species than other microbial profiling techniques (Torok et al., 2008) and used in the fields of; agriculture (Kaur et al. 2017, Zhu et al. 2017), marine biology (Tada et al., 2017) and soil community studies (Fry et al. 2016, Gschwendtner et al. 2016). It does have its vulnerabilities owed to primer choice, template DNA concentration and number of PCR cycles (Zhu et al., 2002) but this is true of all PCR-based technologies. This study will determine how the bacterial community of the nest progresses throughout the breeding attempt whilst utilising a novel method within this field, assessing its efficacy.

Methods

Study sites

The study was performed in the breeding season of 2016 across four sites in England to incorporate local and broader scale geographical differences. Three sites were located in Lancashire, and a fourth site was located within the Forest of Dean, Gloucestershire (refer to Table 1).

Field sampling

Nest boxes were observed to determine breeding activity and bacterial samples taken under Natural England licence (2014/SCI/0288) at nest build completion, clutch completion and immediately post fledging (determined as per chapter 2), subsequently referred to as breeding stage. Given the similarity between nest and plumage microbiomes (Goodenough et al. 2017, van Veelen et al. 2017), nest material was sampled thus causing less distress to the birds whilst still offering a true representation of the microbiome the birds are exposed to. Sterile cotton tipped swabs (Fisher brand, UK) were pre-moistened in phosphate buffer (0.2M, pH7.1, Sigam Aldrich Ltd, Dorset, UK) and the nesting material was swabbed for 30 seconds by the same person starting at the rear left corner with side to side movements until the front right corner of the nest box had been reached, with particular effort to keep this process standardised and remove inter-operational variability. Swabs were stored in individual 15ml Falcon tubes (VWR, UK) containing 1ml of phosphate buffer (0.2M, pH7.1, Sigam Aldrich Ltd, Dorset, UK). All samples were processed for genomic DNA extraction within 4-6 hours.

Table 1: Study site location, description and nest box status

Site

Map reference

Vegetation type

Nest boxes

Nest box orientation

No. of nests sampled

Ruff Wood

53°33′36.27″N, 002°51′59.43″W

Mixed deciduous woodland.

None present, 20 placed for this study.

North to East.

11

Scutchers Acres

53°35′17.68″N, 002°49′23.79″W

Mixed deciduous and coniferous woodland.

None present, 20 placed for this study.

North to East.

10

Mere Sands Wood Nature Reserve

53°38′05.59″N, 002°50′16.06″W

Mixed deciduous woodland.

Present, some monitoring from local volunteer groups.

Random covering all aspects.

10

Nagshead Nature Reserve

51°46′25.59″N, 002°34′20.76″W

Oak dominated deciduous woodland.

400 present, managed by RSPB.

North to East & North to West.

6

Genomic DNA extraction

Falcon tubes containing the sample swabs in phosphate buffer were vortexed for 30 seconds before the swabs were discarded. Total genomic DNA was then extracted from the phosphate buffer using the Bioline ISOLATE II Genomic DNA Kit (Bioline, UK) as per the manufacturer’s protocol. Products were stored in sterile microcentrifuge tubes at -20°C.

PCR amplification for T-RFLP

PCR amplification was performed on total community DNA following the methods of Torok et al. (2008). Primers 27F (AGAGTTTGATCCTGGCTCAG) and 907R (CCGTCAATTCCTTTGAGTTT) were used, which are specific to eubacterial 16S ribosomal rRNA gene sequences. The forward primer of the pair (27F) was labelled with the fluorescent label 6-carboxyfluorescein (6-FAM) (Invitrogen, Paisley, UK) to allow for detection by the 3730 DNA Analyser (Applied Biosystems, CA,USA) during downstream processing of the DNA in Terminal restriction fragment length (T-RFLP) analysis. PCR reactions were performed in 50µl volumes comprising 25µl 2x AmpliTaq Gold® 360 Master Mix (Applied Biosystems, CA, USA), 1µl (20pmol) of each of the primers, 13µl sterile molecular biology grade water (Sigma-Aldrich Ltd, Dorset, UK) and 10µl of template DNA. In negative controls DNA was replaced with sterile molecular biology grade water (Sigma-Aldrich Ltd, Dorset, UK) and genomic Escherichia coli DNA was instead substituted for the positive controls. Ten technical replicates were included throughout the protocol to ensure reproducibility. PCR amplifications were run in a 96 well Thermal Cycler (Prime, Bibby Scientific Ltd., UK) with the amplification conditions; initial denaturation at 95⁰C for 10 minutes followed by 35 cycles of denaturation at 95⁰C for 45 seconds, annealing at 48⁰C for 45 seconds and an extension step at 72⁰C for 1 minute, with a final extension step at 72⁰C for 7 minutes. PCR Products were stored at -20°C.

Visualisation of PCR products by agarose gel electrophoresis

PCR products were visualised using agarose gels (0.7% w/v, Sigma-Aldrich Ltd, Dorset, UK) prepared with 1 x TAE (Sigma-Aldrich Ltd, Dorset, UK) with 2µl ethidium bromide (10mg ml-1) added to 50ml molten agarose prior to pouring. Following PCR, 10µl of PCR product was added to 2µl loading buffer (30 % glycerol (v/v), 2.5g ml-1 bromophenol blue, 2.5g ml-1 xylene cyanol, Sigma-Aldrich Ltd, Dorset, UK). Products were then loaded into agarose gels and were separated by electrophoresis at ~100v for 1 hour. A 1kb ladder (Invitrogen, Paisley, UK) and a 100bp ladder (Invitrogen, Paisley, UK) were run as markers on gels to allow visualised amplicons to be sized. Products were visualised on a UV transilluminator (320nm, Labnet International Inc, NJ, USA) following electrophoresis.

DNA purification and quantification

Following visualisation PCR products were purified using the Bioline ISOLATE II PCR and Gel Kit (Bioline, UK) according to the manufacturer’s protocol. Purification was performed in order to remove primers, nucleotides, salts and other impurities from the PCR amplicons prior to endonuclease digestion. DNA concentration was determined to ensure that sufficient DNA within the range suggested by Torok et al. (2008) was included in each digestion reaction. Purified PCR products were quantified using the Nanodrop ND8000 UV-Vis Spectrophotometer (Thermo Scientific, Hampshire, UK) of 1µl of each PCR product.

Endonuclease reaction

Purified PCR products were digested with 2 units of the restriction enzyme MspI (New England Biolabs Inc., Hertfordshire, UK) following the manufacturers guidelines. The volume of DNA required to achieve 1µg was calculated then added to 1µl restriction enzyme, 5µl 10X NEBuffer (New England Biolabs Inc., Hertfordshire, UK) and sterile molecular biology grade water to a total reaction volume of 50µl. Reactions were then incubated at 37⁰C for 15 minutes. Following digestion, enzyme activity was inactivated by incubating at 65⁰C for 15 minutes.

T-RFLP analysis procedure

Endonuclease digestion products were transferred to 96 well plates in volumes of 1.5µl for running on the genetic analyser and an internal size standard was added to each sample. The reaction mixture for the size standard comprised 9µl Hi-Di Formamide (Applied Biosystems, CA, USA) and 0.35µl GeneScan 1200 LIZ Size Standard (Applied Biosystems, CA, USA) for each sample. This was heated at 95°C for 3 minutes, prior to cooling on ice for 5 minutes. Digest fragments were separated and analysed by capillary electrophoresis using the 3730 DNA analyser (Applied Biosystems, CA, USA). The LIZ size standard acted as an internal size marker to allow for the accurate sizing of DNA fragments within each sample. Gene mapper software was then used to manually bin individual T-RFs and assign sizes (in units of base pairs) to each fragment. In order to distinguish true peaks from artefacts and background noise, a baseline threshold of 50 relative fluorescence units (RFU) was applied to T-RF profiles (after Torok et al., 2008), so that peaks below this threshold were excluded from analysis. Furthermore T-RFs sized less than 50 base pairs were removed to exclude potential primer-associated fragments as well as those over the amplicon size of 880 base pairs (Aguilera et al., 2017). T-RFLP profiles remaining were grouped to identify synonymous fragment sizes within 2 base pairs (Torok et al., 2008). The resulting T-RFs (peaks) were then treated as individual OTUs.

Statistical analysis

For each sample the T-RFLP profiles (individual OTUs) were converted in to binary code given (1) for presence and (0) for absence at each site and breeding stage. Given the large number of zeros within the dataset the Hellinger transformation was performed. Total OTUs per sample was calculated, then a Cochran’s Q test with post hoc pairwise McNemar test was used to determine any difference in bacterial (in the form of OTUs) richness at the three breeding stages. Chi squared was utilised to determine whether OTUs were evenly distributed across sites and breeding stages. Non-metric multidimensional scaling (NMDS) was used to explore any difference in bacterial species richness between breeding stages within a site and each breeding stage between sites. PERMANOVA was used to distinguish any significant difference in bacterial species richness in the above treatments. Spearman’s rank correlation co-efficient was used to determine any relationship between bacterial richness and brood size (data from chapter 2). All analyses were carried out using R Studio version 3.1.1 (R Core Development Team; R Studio, 2017).

Results

In total 37 nests were sampled (Scutchers Acres = 10, Mere Sands Nature Reserve = 10, Nagshead Nature Reserve = 6, Ruff Wood = 11) at each breeding stage. Technical replicates revealed 96% similarity. Across the four sites and three breeding stages there were 103 distinct OTUs. Of these two were present in all but one nest and a further five present in over 90% of nests across the three breeding stages. Other OTUs occurred much less frequently with seventeen only present within one nest and a further 28 in fewer than 5% of nests. Within the nest build stage there was 46 distinct OTUs of which two were present in each nest and eighteen showed only one occurrence. Clutch completion breeding stage revealed 72 distinct OTUs with again two present in each nest and a further two in all but one nest, with 26 showing only one occurrence. Post fledging there was 89 distinct OTUs with two present in all but one nest and nineteen showing only one occurrence. Table 2 shows the OTUs within each nest at each breeding stage.

ab

a

b

Figure 5. Bacterial community richness by breeding stage (nest build, clutch completion and post fledging; mean ± 0.95 confidence interval). Build-clutch p=0.086, build-fledged p<0.001, clutch-fledged p=0.052, n=103. Letters ab denote any significant difference between breeding stages.

Table 2: OTUs per nest at each breeding stage

Mean

SD

Max

Nest build

14.4

4.2

23

Clutch completion

18.6

9.1

42

Post fledging

26

12.2

49

Table 3: Number of OTUs present at each site/breeding stage. 2= 19.59, d.f. = 6, p = 0.003

Scutchers Acres

Mere Sands

Nagshead

Ruff Wood

Nest build

28

24

26

42

Clutch completion

39

51

34

54

Post fledging

62

30

70

74

Table 4: Number of unique OTUs per site and breeding stage. Percentage of total OTUs per site/stage in brackets.

Scutchers Acres

Mere Sands

Nagshead

Ruff Wood

Nest build

Clutch completion

2(5.1%)

4(7.8%)

Post fledging

1(1.6%)

5(7.1%)

7(9.5%)

Figure 1 shows the bacterial species richness across the three breeding stages. Species richness increased at each breeding stage with additional and unique OTUs present. The difference between nest build and clutch completion revealed no significance, with clutch completion to post fledging bordering significance at p=0.052. Nest build to post fledging revealed a highly significant difference in the number and presence of OTUs.

Figure 2 illustrates the fifteen most common OTUs are present within each site and breeding stage. Within nest build Nagshead Nature Reserve has fewer OTUs than the other sites, missing OTUs that are common within the breeding stage. Ruff Wood has the greatest OTU richness across nest build and clutch completion whilst Mere Sands Wood has a lower OTU richness post fledging and Nagshead Nature Reserve a lower OTU richness at clutch completion. Figure 3 highlights at Scutchers Acres, Nagshead Nature Reserve and Ruff Wood that OTU richness increases as the breeding stage progresses, conversely at Mere Sands Wood clutch completion has the greatest OTU richness. Overall Ruff Wood shows the greatest OTU richness across the breeding stages. Table 3 shows the number of OTUs present at each site and breeding stage. Ruff Wood has a higher number of OTUs at nest build than the other three sites whilst Mere Sands has fewest post fledging and is the only site to show a decrease in the number of OTUs from clutch completion to post fledging. The Chi squared result illustrates that individual OTUs are not evenly distributed between sites and breeding stages.

1

Nest build

Clutch completion

Post fledging

SA

MS

N

RW

SA

MS

N

RW

SA

MS

N

RW

*

 

 

 

 

 

 

 

 

 

 

 

 

107

 

81-100

 

 

 

 

 

 

 

 

 

 

 

 

107

 

61-80

 

 

 

 

 

 

 

 

 

 

 

 

106

 

41-60

 

 

 

 

 

 

 

 

 

 

 

 

102

 

21-40

 

 

 

 

 

 

 

 

 

 

 

 

99

 

01-20

 

 

 

 

 

 

 

 

 

 

 

 

93

0

 

 

 

 

 

 

 

 

 

 

 

 

85

% OTU abundance within each site

 

 

 

 

 

 

 

 

 

 

 

 

82

 

 

 

 

 

 

 

 

 

 

 

 

77

 

 

 

 

 

 

 

 

 

 

 

 

76

 

 

 

 

 

 

 

 

 

 

 

 

72

 

 

 

 

 

 

 

 

 

 

 

 

68

 

 

 

 

 

 

 

 

 

 

 

 

67

 

 

 

 

 

 

 

 

 

 

 

 

54

 

 

 

 

 

 

 

 

 

 

 

 

51

 

 

 

 

 

 

 

 

 

 

 

46

 

 

 

 

 

 

 

 

42

 

 

 

 

 

 

 

 

 

 

 

 

41

 

 

 

 

 

 

 

 

 

 

 

 

39

 

 

 

 

 

 

 

 

 

 

35

 

 

 

 

 

 

 

26

 

 

 

 

 

 

 

 

26

 

 

 

 

 

 

 

26

 

 

 

 

 

 

 

 

 

 

25

 

 

 

 

 

24

 

 

 

 

 

 

 

 

24

 

 

 

 

 

 

 

 

23

 

 

 

 

23

 

 

 

 

 

 

20

 

 

 

 

 

 

 

 

 

18

 

 

 

 

17

 

 

 

17

 

 

 

 

 

 

 

 

17

 

 

 

 

 

 

 

 

16

 

 

 

 

 

 

16

 

 

 

 

 

16

 

 

 

14

 

 

 

 

 

14

 

 

 

 

 

 

13

 

 

 

 

 

12

 

 

 

 

 

12

 

 

 

 

 

12

 

 

 

 

 

11

 

 

 

 

 

10

 

 

 

 

 

 

10

 

 

 

 

 

10

 

 

 

 

10

 

 

 

 

 

 

9

 

 

 

9

 

 

 

8

 

 

 

 

 

8

 

 

 

 

 

8

 

 

 

7

 

 

 

 

7

 

 

 

 

 

7

 

 

 

 

 

7

 

 

 

 

 

7

 

 

 

6

 

 

 

6

 

 

 

 

 

 

6

 

 

 

 

6

 

 

5

 

 

 

5

 

 

 

 

5

 

 

 

 

5

 

 

 

 

4

 

 

 

4

 

 

4

 

 

4

 

 

 

3

 

 

3

 

 

3

 

 

3

 

 

3

 

 

3

 

 

 

3

 

 

3

 

 

 

3

 

 

3

 

 

2

 

 

2

 

 

2

 

 

2

 

2

 

 

2

 

2

 

 

2

 

2

 

 

2

 

 

2

 

1

 

1

 

1

 

1

 

1

 

1

 

1

 

1

 

1

 

1

 

1

 

1

1

 

1

 

1

 

1

 

1

Figure 2: OTU abundance within a given site and breeding stage. Shading represents percentage abundance within the given site and breeding stage. * Number of nests/ breeding stages each OTU was present within. SA = Scutchers Acres, MS = Mere Sands Wood, N = Nagshead, RW = Ruff Wood.

Scutchers

Mere sands

Nagshead

Ruff Wood

NB

CC

PF

NB

CC

PF

NB

CC

PF

NB

CC

PF

*

 

 

 

 

 

 

 

 

 

 

 

 

107

 

81-100

 

 

 

 

 

 

 

 

 

 

 

107

 

61-80

 

 

 

 

 

 

 

 

 

 

 

 

106

 

41-60

 

 

 

 

 

 

 

 

 

 

 

 

102

 

21-40

 

 

 

 

 

 

 

 

 

 

 

 

99

 

01-20

 

 

 

 

 

 

 

 

 

 

 

 

93

0

 

 

 

 

 

 

 

 

 

 

 

 

85

% OTU abundance within each breeding stage

 

 

 

 

 

 

 

 

 

 

 

 

82

 

 

 

 

 

 

 

 

 

 

 

 

77

 

 

 

 

 

 

 

 

 

 

 

 

76

 

 

 

 

 

 

 

 

 

 

 

 

72

 

 

 

 

 

 

 

 

 

 

 

 

68

 

 

 

 

 

 

 

 

 

 

 

 

67

 

 

 

 

 

 

 

 

 

 

 

 

54

 

 

 

 

 

 

 

 

 

 

 

 

51

 

 

 

 

 

 

 

 

 

 

 

46

 

 

 

 

 

 

 

 

42

 

 

 

 

 

 

 

 

 

 

 

 

41

 

 

 

 

 

 

 

 

 

 

 

 

39

 

 

 

 

 

 

 

 

 

 

35

 

 

 

 

 

 

 

26

 

 

 

 

 

 

 

 

26

 

 

 

 

 

 

 

26

 

 

 

 

 

 

 

 

 

 

25

 

 

 

 

 

24

 

 

 

 

 

 

 

 

24

 

 

 

 

 

 

 

 

23

 

 

 

 

23

 

 

 

 

 

 

20

 

 

 

 

 

 

 

 

 

18

 

 

 

 

17

 

 

 

17

 

 

 

 

 

 

 

 

17

 

 

 

 

 

 

 

 

16

 

 

 

 

 

 

16

 

 

 

 

 

16

 

 

 

14

 

 

 

 

 

14

 

 

 

 

 

 

13

 

 

 

 

 

12

 

 

 

 

 

12

 

 

 

 

 

12

 

 

 

 

 

11

 

 

 

 

 

10

 

 

 

 

 

 

10

 

 

 

 

 

10

 

 

 

 

10

 

 

 

 

 

 

9

 

 

 

9

 

 

 

8

 

 

 

 

 

8

 

 

 

 

 

8

 

 

 

7

 

 

 

 

7

 

 

 

 

 

7

 

 

 

 

 

7

 

 

 

 

 

7

 

 

 

6

 

 

 

6

 

 

 

 

 

 

6

 

 

 

 

6

 

 

5

 

 

 

5

 

 

 

5

 

 

 

 

5

 

 

 

 

4

 

 

 

4

 

 

4

 

 

4

 

 

 

3

 

 

3

 

 

3

 

 

3

 

 

3

 

 

3

 

 

 

3

 

 

3

 

 

 

3

 

 

3

 

 

2

 

 

2

 

 

2

 

 

2

 

2

 

 

2

 

2

 

 

2

 

2

 

 

2

 

 

2

 

1

 

1

 

1

 

1

 

1

 

1

 

1

 

1

 

1

 

1

 

1

 

1

1

 

1

 

1

 

1

 

1

Figure 6: OTU abundance within a given breeding stage at each site. Shading represents percentage abundance within the given site and breeding stage. * Number of nests/ breeding stages each OTU was present within. NB = nest build, CC = clutch completion, PF = post fledging.

Table 4 shows that there are 19 unique OTUs across the study although none of these are within the nest build. There are unique OTUs at clutch completion within Scutchers Acres and Mere Sands Wood with the largest number occurring post fledging with seven at Ruff Wood, five at Nagshead Nature Reserve but none at Mere Sands Wood.

The NMDS for Scutchers Acres (Fig. 4) shows the OTUs for nest build breeding stage tightly clustered with a clear separation from those of post fledging, clutch completion shows slight overlap with post fledging and shares some similarity with nest build however are much more dispersed. PERMANOVA reveals there is a significant difference in bacterial composition between the different breeding stages (F2, 27 =3.97, p<0.0001) however the dispersions are not homogenous meaning there is a considerable difference in multivariate spread, therefore either could be the cause for the significant PERMANOVA result. At Mere Sands Nature Reserve there is no clear separation between OTUs at each breeding stage. At Nagshead Nature Reserve there is some separation between nest build and clutch completion and complete separation post fledging, PERMANOVA shows significance (F2,15 =5.12, p<0.0001) and dispersions are homogenous. Similarly Ruff Wood reveals some separation between nest build and clutch completion and complete separation post fledging with PERMANOVA again significant (F2, 29 =6.40, p<0.0001) and dispersion homogenous.

NMDS for breeding stage at each breeding site (Fig.5) reveals some separation between Scutchers Acres and Nagshead Nature Reserve and slight separation between Mere Sands Nature Reserve and Ruff Wood at nest build. At clutch completion there is some separation between Scutchers Acres and Nagshead Nature Reserve, Ruff Wood and Mere Sands Nature Reserve. Post fledging points are much more tightly clustered although there is still separation between Scutchers Acres and Mere Sands Nature Reserve, Scutchers Acres and Nagshead Nature Reserve and between Ruff Wood and Nagshead Nature Reserve. PERMANOVA reveals significance in each of the scenarios; nest build (F3, 33 =2.90, p<0.0001), clutch completion (F3, 33 =2.29, p<0.001) and post fledging (F3, 31 =3.81, p<0.0001) with all dispersions homogenous. Spearman’s rank correlation revealed no relationship between bacterial richness in the nest and brood size (rho = -0.27, p = 0.15).

Figure 7: NMDS to explore bacterial (OTU) richness for each breeding stage at each sample site. SA = Scutchers Acres, MS = Mere Sands Nature Reserve, N = Nagshead Nature Reserve, RW = Ruff Wood. Stress scores; SA = 0.11, MS = 0.09, N = 0.08, RW = 0.15. Stress scores <0.05 = excellent, <0.10 = good, <0.20 = usable (Clark, 1993).

Figure 8: NMDS to explore bacterial (OTU) richness for each sample site at each breeding stage. Stress scores; nest build = 0.14, clutch completion = 0.10, post fledging = 0.11.

Discussion

Across the four sites and three breeding stages 103 distinct OTUs were found. Of these the fifteen most common were present across all sites and breeding stages with the remaining OTUs exhibiting variation amongst sites and breeding stages. Nagshead Nature Reserve had the lowest OTU richness at nest build and clutch completion, Mere Sands Wood the lowest post fledging values, whilst Ruff Wood experienced the greatest richness across all stages. Chi squared analysis revealed that Individual OTUs were not evenly distributed across all sites and breeding stages. The variation in OTUs between sites and breeding stages follows previous studies highlighting that bird associated microbiomes are more dependent upon the local environment than the host and demonstrates the transient nature of the microbiome and thus the exposure to ever changing bacteria the birds are experiencing (van Veelen et al., 2017).

Our study surpasses the bacterial species richness previously associated with C. caeruleus, with Goodenough and Stallwood finding 32 species (2010) and 28 species (2012), although these studies were restricted by culturable bias, they were only carried out at a single site which may explain the reduced numbers. Our study is the first known instance of performing culture-independent analysis of C. caeruleus nest microbiomes however, a greater bacterial richness was found in P. major nests with Jacob et al. (2014) reporting 180 OTUs across 52 nests using a different technique, Automated Ribosomal Intergenic Spacer Analysis (ARISA). OTU richness was an average of 14.4 at nest build, 18.6 at clutch completion and 26 post fledging with a maximum of 23, 42 and 49 respectively highlighting the diverse microbiomes C. caeruleus have to contend with during their breeding attempt. Bacterial species richness increased at e