Callum Richards Research Report Final

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Manipulation experiments highlight the importance of diet as a determinant of bacterial community structure in the gut of the omnivorous cockroach, Pycnoscelus surinamensis. Callum Richards1, Saria Otani1, Aram Mikaelyan2 and Michael Poulsen1 1 Centre for Social Evolution, Section for Ecology and Evolution, Department of Biology, University of Copenhagen,

Universitetsparken 15, Building 3, 2100 Copenhagen East, Denmark.

2 Department of Biogeochemistry, Max Planck Institute for Terrestrial Microbiology, Karl-von-Frisch-Str. 10, 35043,

Marburg, Germany.

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Contents 1. Title Page: Manipulation experiments highlight the importance of diet as a

determinant of bacterial community structure in the gut of the omnivorous cockroach, Pycnoscelus surinamensis.

3.

Abstract Introduction

5. Materials and Methods 9. Results 18. Discussion 21. Acknowledgements 22. References 27. Supplementary Material 30. Failures and Revisions

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Abstract The gut microbiota of cockroaches and termites plays important roles in the symbiotic digestion of dietary components, such as lignocellulose. Diet has been proposed as a primary determinant of community structure within the gut, acting as a selection force to shape the diversity observed within this “bioreactor” and as a key mechanism for the divergence of termite gut microbiota from their omnivorous cockroach relatives. The gut microbiota in most termites supports primarily the breakdown of lignocellulose, one sub-family of higher termites – the fungus-growing termites that grow fungus as their main food source – has become similar in gut microbiota to the ancestral omnivorous cockroaches. In this study, we comparatively analyse community compositions in the guts of experimentally manipulated Pycnoscelus surinamensis cockroaches fed on different dietary proportions of the fungus maintained by fungus-farming termites in order to assess the importance of a fungus diet as a driver of community structure. Using MiSeq amplicon sequencing of the 16S rRNA gene, we compare community structure in cockroaches experimentally exposed to fungus to allow us to identify key bacterial lineages contributing to community structure, suggestive of a potential role in fungus digestion. Analysis of gut microbiotas from 49 gut samples showed a step-wise gradient pattern in community similarity that correlated with an increase in fungal material provided to the cockroaches. Analysis of the taxonomic composition of communities showed that an increase in dietary fungal biomass promoted particular bacterial lineages that were also present within the core microbiota of fungus feeding termites. These results demonstrate the role diet can play in determining gut community composition as a fungal based diet induced the cockroach microbiota to resemble that of a more fungus-growing termite like community. Correspondence: Callum Richards, E-mail: [email protected] Introduction Gut microbes have had a significant impact on animal evolution and play a diverse range of functional roles within their symbiotic hosts (Sachs, Skophammer & Regus, 2011; Sanders et al. 2014). Complex gut microbiotas are found in species ranging from mammals to insects and have crucial roles in digestion, immunity, and development. It is important to gain an understanding of the mechanisms that govern the ecology and evolution of complex microbial communities to gain further insight into the development of these mutualistic (beneficial) symbioses (Heijtz et al., 2011; McFall-Ngai et al., 2013). Research into the microbiology of insect symbionts has increased over recent years with advancements in sequencing technologies that have helped identify the microbes dominating insect guts in Drosophila, honey bees and attine ants (Moran et al., 2012; Scheuring & Yu, 2012; Staubach et al., 2013; Douglas, 2015; Sapountzis et al., 2015). Termite guts are of particular interest as they harbour diverse and unique microbial populations, particularly in their hindguts that act as a major “bioreactor”, as suggested by the low redox potential and the accumulation of hydrogen at this site (Li et al., 2012). Such microbial diversity in their guts has made termites the major decomposers of lignocellulosic biomass in nature (Brune, 2014; Rahman et al., 2015).

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Termites are “eusocial cockroaches” and evolved from an omnivorous cockroach ancestor more than 150 million years ago, accompanied by a specialisation to a wood-feeding lifestyle (Bourguignon et al., 2014; Mikaelyan, Dietrich et al., 2015). The transition from an omnivorous to a wood-feeding life style was enabled by the acquisition of cellulolytic flagellates that can still be observed as predominant members of the enlarged hindguts in primitive “lower” termites and their cockroach sister group, the Cryptocercidae (Ohkuma, 2008; Hongoh, 2011; Brune & Dietrich, 2015). The subsequent loss of gut flagellates in the Termitidae lead to the radiation of the “higher” termites and dietary diversification as this group evolved to feed on a variety of lignocellulosic food sources with the aid of a completely prokaryotic gut microbiota (Dietrich, Köhler & Brune, 2014; Brune & Dietrich, 2015) (Fig. 1). Diet has been suggested as a major driver of bacterial community structure in the guts of higher termites, with major dietary shifts and diversification being associated with compositional changes of the gut microbiota (Otani et al. 2014; Mikaelyan, Dietrich et al., 2015; Poulsen, 2015). Convergence of bacterial community structure would therefore be expected to occur between species that share a dietary specialization, particularly in species with a highly specific diet, such as in the fungus-cultivating Macrotermitinae, where the fungus genus Termitomyces is the main food source for the termites (Leuthold, Badertscher & Imboden, 1988). This symbiosis has lead the termite subfamily to become the most player in plant degradation and nutrient cycling within its ecological range, with members of the Macrotermitinae consuming more than 90% of dry wood litter in tropical ecosystems (Buxton, 1981). Fungus-growing termites have evolved highly organised colonies with optimised division of labour between colony members in order to efficiently degrade plant material with combined efforts from the fungal mutualist and gut bacteria (Leuthold, Badertscher & Imboden, 1988). Older termite workers forage for plant material within the surrounding environment, returning foraged material to the nest were it is consumed by younger workers and inoculated with asexual spores of Termitomyces derived from nodules within the nest (first gut passage) (Badertscher, Gerber & Leuthold, 1982). The digested mixture is then deposited as external sponge-like fungus comb and is the site in which the majority of lignocellulosic material is degraded using fungus-derived cellulases and xylanases (Hyodo et al. 2013; Poulsen et al. 2014). Older workers then consume older parts of the comb after considerable mycelial growth (second gut passage) to finalize the complete breakdown of more simple polysaccharides with termite and microbial derived enzymes (Leuthold, Badertscher & Imboden, 1988; Nobre & Aanen, 2012; Liu et al. 2013; Poulsen, 2015). A shift to a relatively proteinaceous diet in the fungus-growing termites may be responsible for a convergence of community structure between this specialized group and their omnivorous non-eusocial cockroach relatives (Dietrich et al., 2014). Otani et al (2014) sampled guts from nine species of fungus-growing termites and found that the Macrotermitinae are associated with a core set of gut microbiota that are more similar to each other and to cockroach gut communities than to those of other termites. They observed a resurgence of bacterial taxa that prevail in cockroaches within the Macrotermitinae, with a shared predominance of Bacteroidetes and Firmicutes (Dietrich et al., 2014; Otani et al., 2014). These taxa are common in omnivorous animals and may have been promoted by the protein-rich fungal component of the termite diet (Schauer et al., 2012; Dietrich, Köhler & Brune, 2014). This again suggests that the obligate association with Termitomyces has shaped the gut microbiota to be compositionally different to those of other termites (Dietrich et al., 2014; Otani et al., 2014).

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The dense microbial colonisation of the homologous hindgut in the cockroach Shelfordella lateralis and the profile of microbial metabolites (Schauer et al. 2012) suggests that even in cockroaches, the hindgut is the major site for microbial activity. Here we test if a fungal diet can act as a selective force to alter the composition of microbiota in the gut of the litter-feeding cockroach Pycnoscelus surinamensis. Previous diet manipulation studies have indicated the ability of dietary shifts to shape the gut microbiota of omnivorous cockroaches (Huang et al., 2013; Schauer et al., 2014; Pérez-Cobas et al., 2015). By providing fungal material from a pure culture of Termitomyces obtained from a fungus-growing termite colony we attempt to mirror fungus feeding and use MiSeq sequencing of the 16S rRNA gene to compare bacterial community structure between cockroaches fed on increasing dietary proportions of dried Termitomyces pure-culture biomass relative to leaf litter. Materials and Methods Study species Individuals of the litter-feeding cockroach Pycnoscelus surinamensis were derived from a stable lab population housed at the Max Planck Institute for Terrestrial Microbiology, Marburg, Germany. P. surinamensis is a species of burrowing cockroach endemic to the Indomalayan region and is a common plant pest that has colonized New World tropical and sub-tropical regions due to its ability to reproduce quickly via thelytokous parthogenesis; a process that produces functional offspring of exclusively female clones from unfertilized eggs (Roth, 1974; Gade & Parker, 1997; Komatsu et al., 2015). This species is a member of the Blaberidae, a sister family to the combined termite, Cryptocercus, and Blattidae clade (Inward, Beccaloni & Eggleton, 2007). It is therefore well placed to act as a model for termite evolution. The cockroaches were maintained at the University of Copenhagen in climate rooms at 27oC and 50% relative humidity. An initial stock population of 1000 individuals was established and maintained throughout the experimental period in a plastic container (56x39x28cm) containing five to eight centimetres of soil and leaf litter substrate. The cockroaches were fed leaf litter, fruit, and vegetables three times a week and the substrate within the container was replenished weekly until three days before initiation of feeding experiments. Diet experiment After a two-week pre-feeding period, individuals within the holding container were exposed to a control diet of only leaf litter for 72-hours and then juveniles were isolated into subsets of 50 cockroaches within smaller experimental containers (21x17x15cm). Juveniles were chosen to ensure the occurrence of at least two moults and therefore restoration of the gut microbiota during the experimental period (Carrasco et al., 2014). This process was expected to allow the microbiota to change as a consequence of the selection pressure of an altered diet and results in the cockroach appearing white for a

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short period after moulting as the exoskeleton loses pigmentation (Fig. 2c) (Korb, Hoffmann & Hartfelder, 2012). Such an appearance was observed and recorded during the experiment to enable the monitoring of the moulting process. Over a one-month treatment period, sub-populations were exposed to one of six diet regimes consisting of 0 to 100% dried fungus material obtained from a Termitomyces culture isolated from Odontotermes sp. Od127 (Otani et al. 2016). Termitomyces was cultured on Potato Dextrose Agar (PDA, 39g/L PDA, 10g/L agar) and incubated at 27°C for at least 96 hrs

Sister group to termites

Both asocial and gregarious

Omnivorous

ICockroaches

Higher Termites

Subsocial

Entirely prokaryotic micobiota

Complex gut structure with differentiated compartments

Feed on variety of dietary componentseg wood, plant litter, dung, humus, soil organic matter

Fungus- growing termites

Fungus feeding - digest plant material via 2 phase gut passage

Tripartite symbiosis with fungal mutualist Termitomyces andthe gut community

Bla

ttodea

Isopte

ra

200 150 100 50 0

MYA

(a)

(b)

(c)

Term

itid

ae

(i)

(iii)

(iv)II

Simple gut structure

Celluloytic flagellates- horizontal transfer via proctodeal trophallaxis

Wood-feeding - capable of digesting lignocellulose

Eusocial

Lower Termites

Other Cockroaches

Blattidae

Cryptocercidae

Mastotermitidae

Stolotermitidae

Hodotermitidae

Archotermopsidae

Kalotermitidae

Serritermitidae

Rhinotermitidae

Macrotermitinae

Sphaerotermitinae

Apicotermitinae

Syntermitinae

Termitinae

Nasutitermitinae

(ii)

Fig. 1(a) Life histories and feeding habits of (i) Cockroaches, (ii) Lower termites, (iii) Higher termites, (iv) Fungus-growing termites. The phylogeny shows the origination of termites within the radiation of the cockroaches (based on Bourguignon et al., 2014). Two key events occurred in the evolution of digestive symbiosis and the termite gut community; Key Event I: acquisition of cellulolytic flagellates by the common ancestor of the termites (Isoptera) and their sister group, the Cryptocercidae, boosting their capacity for lignocellulose digestion to form the “lower” termite gut (ii). Key Event II – the subsequent loss of celluloytic protists in the Termitidae ca. 50 million years ago. This lead to dietary diversification in the higher termites (iii), including specific feeding adaptions such as fungus feeding in the fungus-growing termites (iv) (Brune & Dietrich, 2015). (b) A fungus-growing termite Macrotermes natalensis royal pair (image from Poulsen, 2015). (c) The study species, Pycnoscelus surinamensis, placed in the “other cockroaches” as a member of the Blaberidae, a family of cockroaches that is a sister group to the combined clade of termites, Cryptocercus, and the Blattidae (Inward, Beccaloni & Eggleton, 2007).

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to allow sufficient fungal growth. Contaminated cultures were discarded. Fungal material was harvested from the agar plates by scraping off mycelium taking special care to avoid the medium. Harvested mycelium was dried at 56°C for four hours before being combined with the appropriate dry weight of leaf litter to produce the feed allowance for the treatment sub-populations. Sub-populations were provided with 1.5g of forage material, consisting of one of the following combinations (percentage-by-weight ratios) of dried leaf-litter to fungus material: 100:0, 80:20, 60:40, 40:60, 20:80 or 0:100. Each of six dietary combinations was set-up in triplicate, yielding a total of 18 samples (Fig. 2). Sub-populations were fed twice a week for a one-month treatment period. Uneaten food was removed before new provisioning was made in order to keep the leaf litter: fungus ratios as constant as possible.

Fig. 2 (a-c). Pycnoscelus surinamensis individuals used in the diet experiment: juvenile (a), adult (b) and newly moulted (c). The scale bar represents a length of 30mm and was used to assess and control for the size of selected cockroaches as a guide for their age and therefore likelihood of moulting. Individuals selected for the experiment were approximately 8-10mm in length. (d-f). Images of the treatment boxes with a diet consisting of 0% (d), 60% (e), and 100% (f) Termitomyces, respectively. Survival and behaviour surveys Each subpopulation was surveyed twice a week and their foraging behaviour recorded to establish if the cockroaches consumed the fungal biomass. The number of juveniles, sub-adults and adults were counted for each sub-population at the end of the experiment to compare the well being of cockroaches on different feeding regimes. Age was approximated using the size of the surviving cockroaches within the population.

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Table 1 The number of sequences after filtering of raw reads, the number of identified taxa, the percentage of reads successfully assigned to the phylum, family and genus levels (based on relative abundances) as well as the estimated richness and diversity indices for the bacterial communities (at 3% dissimilarity threshold). Dissections and DNA Extraction Nine cockroaches were randomly picked per sub-population for dissections, and these nine were randomly assigned to one of three technical replicates per sub-population. Given the six feeding regimes with three biological replicates, each of which had three technical replicates, a total of 54 cockroaches were dissected. Because of the labour-intensiveness of the dissections, only 1/6th of all cockroaches included in the experiment could be dissected in one day. Therefore, three sub-populations were randomly picked daily for dissections, while sub-populations yet to be dissected were maintained on their diet regime in order to avoid starvation and to sustain their gut microbiota. Before dissection, cockroaches were subdued by placing individuals on ice for 20 minutes and they were then placed dorsally on a sterile Petri dish. The head was removed and the tergal area opened by coaxial removal of the legs, exposing the body cavity to allow removal of the gut from the anus to the metathorax. The hindgut was then separated from the whole gut under saturation in RNAlater® (Ambion®Thermo Fisher Scientific, Nærum, Denmark). Dissections were carried out under stereomicroscope fine forceps (Wild M3C, Leica Microsystems, Ballerup, Denmark). Guts were stored at -20oC until DNA extraction. The DNeasy blood and Tissue kit (Qiagen, Germany) was used for DNA extractions from the hindgut samples, following the manufacturer’s instructions. Bacterial 16S rRNA PCR amplification and Miseq sequencing The V4 region of the 16S rRNA gene was amplified using the primers v4.SA504 (5’-AATGATACGGCGACCACCGAGATCTACACCTGCGTGTTATGGTAATTGTG-TGCCAGCMGCCGCGGTAA-3’) and v4.SB711 (5’-CAAGCAGAAGACGGCATACGAGATTCAGCGTTAGT-CAGTCAGCCGGACTACHVGGGTWTCTAAT-3’). The V4 region amplification was carried out using a dual indexing sequencing strategy (Kozich et al., 2013), and the PCR

Classification Success (%) Diversity Indices

Leaf litter to fungus ratio

Number of sequences

Number of genus-

level taxa

Number of family-level

taxa Phylum Family Genus Shannon Simpson

100:0 60267 383 243 87.7 44.6 10.4 5.29 0.98

80:20 94796 412 256 90.4 49.0 12.1 5.31 0.98

60:40 78589 379 244 88.9 48.6 17.1 5.26 0.98

40:60 78765 387 245 89.9 46.4 12.0 5.11 0.98

20:80 73661 370 245 89.9 45.8 10.9 5.15 0.98

0:100 84488 406 253 91.1 44.2 12.1 5.24 0.98

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mixture was prepared in 20µl volume containing 18.5µl sterile distilled water, 2µl of each primer (40.0µM), 2µl of 10x AccuPrime PCR buffer II (Life Technologies, USA), 3µl DNA template, and 0.15µl AccuPrime High Fidelity Taq DNA polymerase (Life Technologies, USA). PCR conditions were 95°C for 2 minutes followed by 30 cycles of 95°C for 20 s, 55°C for 15 s, and 72°C for 5 min followed by 72°C for 10 min. Troubleshooting PCR was carried out with 2µl of 1:10 diluted DNA template. More information on the selection of samples for MiSeq sequencing can be found in the supplemental information. Library normalisation was carried out using Life Technologies SequencePrep Normalization plate kit (Life Technologies, USA) following the manufacturer’s instructions. Sample concentration was measured using Kapa Biosystems Library Quantification Kit for Illumina Platforms (Kapa Biosystems, USA) and the size of library amplicons was determined using Agilent Bioanalyser High Sensitivity DNA analysis kit (Invitrogen). The samples were subjected to sequencing on the Illumina MiSeq platform using Miseq Reagent Kit V2 500 cycles (Illumina) (Kozich et al., 2013). Sequence filtering and taxon classification Raw flow grams from sequencing were analysed using the Mothur software (version 1.37.6, Schloss et al., 2009) and the standard operating procedure (SOP) was followed as described at http://www.mothur.org/wiki/MiSeq_SOP (Kozich et al., 2013). The paired end reads were assembled into contigs and subjected to several filtering steps in order to reduce PCR and sequencing errors. High quality sequences were aligned against the SILVA 102 non-redundant database. Operational taxonomic units OTUs were calculated at 3% species level classification and rarefaction curves based on a 97% sequence similarity cut-off were generated using R (R core team, 2013). Analysis of gut community diversity and similarity between different fungal ratios Relative taxa abundances were calculated as the number of sequence reads per taxon for the 54 gut samples. Principal coordinate analysis (PCoA) was performed in R (R Core Team, 2013) based on the Bray-Curtis index to determine community similarity between the 54 gut extracts from the six different fungal treatment ratios. Relative taxa abundances across the different treatment ratios were visualised in a heatmap of the relative abundances of the 20 most common taxa in each of the six treatment diets. Principal Component analysis (PCA) was used to calculate loading values in order to assess the contribution of genus level-taxa to the principal components and identify lineages that contributed most to patterns in community diversity observed within the PCoA analysis (R Core team, 2013). The distribution of the most abundant taxa within the treatment groups was also compared to previously published data on P. surinamensis and fungus-growing termite groups (Otani et al., 2014; Dietrich, Köhler & Brune, 2014; Mikaelyan, Köhler et al., 2015; Otani et al., 2016). Results Mortality and behaviour surveys Behavioural observations indicated that P. surinamensis was able to consume the provided fungal material with frequent occurrences of active feeding observed throughout the experimental period. Cockroaches were observed to drag fungal material down into the soil substrate after a short initial feeding period (Fig 4b), as well as feeding occasionally on material on the soil surface (Fig 4c). Minimal fungal material was

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collected after feeding periods during the experimental period and individuals remained highly active in all diet regimes below the 20%:80% regime, further supporting the conclusion that cockroaches were able to consume the fungus material provided. Moulting was observed frequently over the course of the experiment, with depigmented individuals recorded but not quantified across all diet regimes. Activity levels appeared to decrease in diet regimes consisting of 20%:80 and 0:100% fungus material, where individuals moved at slower speeds and were found dormant within the substrate. This did not, however, appear to increase mortality, as the majority of the sub-populations maintained ca. 50 individuals throughout the experiment (Fig 4a). Sub-colonies exposed to a diet regime of 60%:20% leaf litter: fungus did experience an average population reduction of 5 individuals. Statistical analysis however indicated that the total population count in this treatment was not significantly different to the other sub-colonies (F=0.839, DF=11, p=0.56). Illumina MiSeq data from cockroach guts Rarefaction analysis showed insufficient coverage of 5 bacterial communities from the 54 gut samples (Fig. 3). Gut samples ID 8, 26, 42, 46 and 54 (Table S1) were omitted from further downstream analysis of community diversity and taxa abundances due to a low-number of high-quality reads. This filtering however did not result in the loss of any complete biological replicated. 16S rRNA gene sequencing of the remaining 49 cockroach gut samples generated between 60’267 and 94’796 high-quality reads (average±SE: 78’427±4’680) per treatment (Table 1). A total of 5’478 unique OTUs at the 3% cutoff level were identified after filtering and sequence analysis of gut communities. The number of bacterial genera ranges between 370 and 412 (average 390) (Table 1), with cockroaches fed on an 80%:20% leaf litter:fungus regime harbouring the least. Shannon and Simpson diversity indices were similar across all treatments (Table 1).

Num

ber

of O

TUs

Number of reads

ID 42

ID 26ID 46

ID 8ID 54

Fig. 3. Rarefaction curves of sequence depth for the 54 gut samples (R Core Team, 2013). Each curve represents the number of identified OTUs as a function of the number of sequenced reads after filtering. The samples ID 8 (0% fungus, replicate 3, technical replicate 2), ID 26 (40% fungus, replicate 3, technical replicate 2), ID 42 (80% fungus, replicate 3, technical replicate 2), ID46 (100% fungus, replicate 1 technical replicate 1)) and ID 54 (100% fungus, replicate 3, technical replicate 3) (see also Table S1) were omitted from further downstream analysis due to low sequence read count and subsequent poor coverage of bacterial communities in these samples. The remaining 49 samples had sufficient community coverage and were used for analysis of community diversity and taxa abundances.

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Analysis of gut community diversity and similarity Ordination analyses Gut microbiota similarities across the different treatment rations were assessed by Bray-Curtis distances and in PCoA plots (Fig. 5). A clear signal of diet specific community overlapping was observed, where microbiotas from cockroaches fed on diets consisting of similar fungus ratios were more similar to each other than to cockroaches fed on diets of differing fungus contents, resulting in a step-wise gradient pattern in community similarity from 100% to 0% fungus content along the PCO2 axis (Fig.5a). A strong signal of community similarity at the extremes of 20%:80% and 0%:100% leaf litter to fungus ratios was observed as communities from high fungal diets overlap, excluding all but 2 samples from diets with a lower proportion of fungal material. Communities of mid range fungal ratios (60%:40% and 40%:60%) also overlap extensively. Community similarities between cockroaches fed on diets with minimal fungus material (100%:0% and 80%:20%) are more diluted however, with their gut communities sharing some bacterial diversity with mid range fungal diets (60%:40% and 40%:60%) (Fig. 5a).

Fig. 4 (a) Mean ± SE (n=3) number of juvenile (grey), sub-adult (light grey) and total number of cockroaches (dark grey) within each diet regime. No fully-grown adults were observed within the sub-colonies at the end of the experimental period. No significant differences in survival levels were observed between sub-populations fed on the same fungal ratios or between different treatments for the total population size (F=0.839, DF=11, p=0.56). The number of juveniles (F=0.722, DF=11,p=0.64) and sub-adults (F=1.94, DF=11, p=0.17) also remained consistent between sub-populations and treatments. The majority of diet regimes maintained a population size of approximately 50 individuals (intersecting dotted line) or experienced a slight increase in average population size. (b) P. surinamensis sub-adult feeding on Termitomyces and (c) juvenile interacting with Termitomyces material

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Fig. 5. A. PCoA similarity analysis of the 49 gut samples including technical replicates visualized via Bray-Curtis distances across samples. Abundances of the most abundant bacteria averaged across technical and biological replicates are given in Table 1; abundances of the bacteria that based on loading values contribute the most to the separation observed are given in Table S2. B. PCoA similarity analysis of the gut samples visualized via Bray-Curtis distances between 18 gut samples averaged across the technical replicates. Abundances of the bacteria that based on loading values contribute the most to the separation observed are given in Table 3. C. PCoA similarity analysis visualized via Bray-Curtis distances across gut samples from only the treatments receiving 0% or 100% fungus material. Abundances of the bacteria that based on loading values contribute the most to the separation observed are given in Table 4. PCoA visualisation via Bray-Curtis distances between 18 gut samples averaged across the technical replicates indicated a similar step-wise pattern in community similarity as communities from cockroaches fed on high fungal ratios overlap extensively, excluding all fungal ratios below 20%:80% leaf litter to fungus material (Fig 5b). Communities fed on mid range fungus ratios also overlap but shared some similarity to communities from cockroaches fed on a dietary ratio of 80%:20% leaf litter to fungus. Cockroaches fed on no fungus material overlapped together with a sample of 80%:20% leaf litter:fungus material, with one technical replicate of 0% fungus showing a distinct dissimilarity from other communities that were not exposed to fungus material (Fig 5b ). A final PCoA of community similarity from treatments receiving only 0% and 100% fungus material

20:800:100

40:6060:4080:20100:0

PC

O2 (

18%

)

PCO1 (39%)

Leaf litter: Fungus Material (%)A

0.0

PCO1(44%)

PCO2(23

%)

B

0.0 0.4PCO1 (45%)

PCO

2 (

33

%)

C

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displayed a key separation in community similarity between cockroaches fed on a diet of 0% and 100% fungus material, with communities fed on 100% fungus overlapping more tightly than those fed on 0% in which one sample was clearly distinct from other communities that were not exposed to fungal material (Fig. 5c). Taxonomic Composition of Gut Bacterial Communities Gut community reads were classified using the SILVA database. We were not able to classify the gut sequences using the extended DictDb v .2.3 database due to time and technological restrictions. The most abundant phyla across the six treatments were Bacteroidetes, Firmicutes and Proteobacteria, accounting for 79% of all sequenced reads. A heatmap of the 20 most abundant gut bacteria across the 6 treatments is presented in Table 2 and identifies six different phyla from the 20 gut samples. The top 20 most abundant bacteria represent on average 52% of the the total community abundance across all 49 gut samples. Bacteroidetes, and Firmicutes dominate the top 20 most abundant bacteria (Table 2). Family-level OTU classification of the top 20 most abundant bacteria identified 19 bacterial families from 20 gut communities (Table 2). OTU 1 was not able to be classified despite with an average abundance of 8.8% across all treatments, but the next most abundant bacteria were Enterococcaceae (3.76%) and Lactobacillaceae (3.83%) from the Firmicutes. Synergistaceae (3.43%) and Planctomycetaceae (3.15%) were also in high abundance, with Planctomycetaceae particularly abundant (6.88%) in the 0% fungal treatment cockroaches, dropping steadily to reach a final abundance of 0.29% in cockroaches fed on 100% fungal biomass. Bacteria that were less abundant in the overall top 20 did prove however to be in higher relative abundance in the gut communities of cockroaches fed on higher fungal ratios. For example, Lachnospiraceae increased in abundance in the higher fungal ratios of 80% and 100% with an average abundance of 0.79% across these two treatments, but had an average of only 0.27% across the 4 lower fungal treatments (Table 2). Porphyromonadaceae also experienced an increase in abundance in cockroaches fed on higher fungal ratios, with an average abundance of 0.54% across 0% and 20% treatments, increasing to a 1.05% average abundance across 80% and 100% treatments. Family-level OTU classification success was however low when using the SILVA database at an average success rate of only 46.4% (Table. 1) Genus-level OTU classification identified only six bacterial genera from the 20 gut communities with classification success rate low (average 12.4% (Table1) when referencing the SILVA database. The heatmap revealed that Actinomyces was relatively abundant across all the treatment samples (1.2% average), particularly in the 80% fungal treatment (2.20%). Weisella was also prevalent throughout all treatments, particularly at 20% (2.54%) and 100% (2.68%) fungal treatments. Clostridium_XI (1.54%) and Lactococcus (1.02%) were also highly abundant in the 100% fungal treatment compared to the lower average across all the treatments (0.4% and 0.25%) as they increased with increasing fungal ratios (Table. 2) Taxonomic contributions to community similarity A heatmap of abundances of the 20 bacteria that, based on loading values gained from PCA analysis, contribute most to the pattern observed in Fig. 3a for community similarity is shown in Table S2. Five different phyla contributed most to the stepwise-pattern in community similarity observed within Fig. 3a, with Bacteroidetes, Proteobacteria, Firmicutes and Actinobacteria making the largest contribution (85% of the top 20 bacteria’s loading values) to the pattern (Table S2). Family level-classification identified

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6 different families of bacteria across the 20 gut communities. Porphyromonadaceae, Actinomycetaceae, Microbacteriaceae and Verrucomicrobia in particular contributed to the pattern observed in Fig. 3b, constituting 47% of the top 20’s contribution across these 4 families (as measured from their cumulative loading values) (Table S2). Enterococcaceae also contributes to the pattern and is found to be highly abundant throughout the treatment samples (average abundance of 3.8%). Genus-level OTU classification was low for the top 20 most contributing bacteria. Actinomyces was identified as the genera to contribute most to the pattern, while Parabacteroides was the only other genera identified within the top 20 (Table S2). No other patterns of changes in relative abundance across the different treatment ratios were observed (Table S2). A heatmap of abundances of the 20 bacteria that, based on loading values, contribute most to the pattern observed across averaged technical replicates (Fig. 3b) is shown in Table 3 Phylum-level analysis identified 6 different phyla that contribute most the step-wise gradient pattern observed in Fig. 3b. In particular, Synergistetes, Bacteroidetes, Proteobacteria and Firmicutes contribute heavily to the patterns observed, contributing 87% of the cumulative loading value for these 20 bacteria (Table. 3). At a family level, 10 different phyla were observed across the gut samples. Synergistaceae (3.5% average abundance across technical replicates) and Lachnospiraceae (2.3%) were the top two contributors to the pattern and were also highly abundant across all treatment samples. Families such as Desulfovibrionaceae (0.72%) were on average in lower abundance but increased in abundance in the 100% treatment cockroaches (1.53%) (Table 3). Planctomycetes was again found to decrease in average abundance in cockroaches fed on higher fungal proportions. Genus-level OTU classification identified only 3 genera from the 20 gut communities, identifying Weissella, Paracbacteroides and Actinomyces as contributors to this step- pattern gradient (Table. 3) A heatmap of abundances of the 20 bacteria that contribute most to the pattern of separation observed in Fig. 3c, based on biological replicates from cockroaches fed on only 0% and 100% fungus is shown in Table 4. Phylum-level analysis identified 6 different phyla represented within the 20 gut communities, with Bacteroidetes, Firmicutes, Proteobacteria and Actinobacteria particularly abundant and constituting 83% of the top 20’s contribution to the pattern in community similarity observed (Table.4). At a family-level, OTU classification identified 12 different families of bacteria in the 20 gut communities. Enterococcaceae is again present at high abundances across all treatments (average 2.95%), as well as Lactobacillaceae (4.11%) and Synergistetes (3.25%). Genus level OTU classification indicated 4 different genera that contributed to the step wise pattern. Ruminococcaceae, a genera present at higher abundances within cockroaches fed on 100% (1.18%) fungal diets compared to a 0% fungal diet (0.51%) are shown to contribute to the separation pattern observed between higher and low fungal communities. This pattern in the prevalence of genera that are found in higher fungal treatments is also observed within Clostridium_XI, with an average of 0.1% abundance in cockroaches fed on 0% fungus, rising to 1.5% average abundance in cockroaches fed on only fungus material, and also Parabacteroides that increased from an average abundance of 0.25% in cockroaches fed on 0% fungus, to 1.14% in cockroaches fed on only fungal biomass. This pattern in increasing abundance in the 100% fungus treatments diets is also observed in the family Desulfovibrionaceae, with an average abundance increase of 1.42% from 0% to 100% fungus biomass. A reverse pattern is observed within the Planctomycetes, a family of bacteria that are particularly abundant across cockroaches that were not exposed to fungus (6.93%) compared to those exposed to only fungus (0.29%).

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Table 2. The 20 most abundant taxa in each of the six treatments averaged across technical and biological replicates. The heatmap scale is the percentage of reads assigned to a given taxon out of the total number of the high quality-filtered and classified reads for the treatment sample. 0 1 6 >12

Treatment(%FungusMaterial)

OTU Phylum Family Genus 0 20 40 60 80 100

1 Bacteroidetes(100) Bacteroidetes.UC(100) Bacteroidetes.UC(100) 6.58 6.79 9.39 12.52 9.10 8.66

2 Firmicutes(100) Enterococcaceae(85) Enterococcaceae.UC(71) 2.84 3.33 3.73 4.47 5.17 3.00

3 Firmicutes(100) Lactobacillaceae(100) Lactobacillaceae.UC(98) 3.24 3.94 3.36 2.49 4.96 4.97

4 Synergistetes(100) Synergistaceae(100) Synergistaceae.UC(100) 3.90 3.31 3.79 3.93 3.11 2.54

5 Planctomycetes(100) Planctomycetaceae(100) Planctomycetaceae.UC(100) 6.88 3.86 2.66 3.79 1.44 0.29

6 Firmicutes(100) Lachnospiraceae(100) Lachnospiraceae.UC(100) 1.58 2.50 2.38 2.58 3.10 1.93


8 Actinobacteria(100) Actinomycetaceae(100) Actinomyces(100) 1.16 1.28 1.57 2.04 2.20 1.29

9 Firmicutes(100) Leuconostocaceae(100) Weissella(100) 0.81 2.54 1.58 0.69 1.15 2.68

10 Proteobacteria(100) Betaproteobacteria.UC(100) Betaproteobacteria.UC(100) 3.30 1.64 0.99 1.10 0.38 2.00


12 Bacteroidetes(100) Porphyromonadaceae(100) Porphyromonadaceae.UC(100) 0.56 1.32 1.32 1.38 1.54 1.40

13 Proteobacteria(100) Betaproteobacteria.UC(100) Betaproteobacteria.UC(100) 1.27 1.22 1.14 1.47 0.90 1.89

14 Proteobacteria(100) Proteobacteria.UC(100) Proteobacteria.UC(100) 1.33 1.25 1.23 1.38 1.49 0.22




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18 Firmicutes(100) Clostridiales.UC(100) Clostridiales.UC(100) 1.05 1.07 1.06 1.08 1.07 1.02




22 Bacteroidetes(100) Porphyromonadaceae(100) Tannerella(100) 0.51 0.91 0.84 0.77 0.71 0.55

23 Bacteroidetes(100) Bacteroidales.UC(100) Bacteroidales.UC(100) 0.65 1.00 0.77 0.57 0.68 0.49

24 Bacteria.UC(100) Bacteria.UC(100) Bacteria.UC(100) 0.70 0.74 0.73 0.81 0.55 0.60

25 Proteobacteria(100) Desulfovibrionaceae(79) Desulfovibrionaceae.UC(75) 0.11 0.64 0.58 0.39 0.72 1.52

26 Firmicutes(100) Ruminococcaceae(100) Ruminococcaceae.UC(99) 0.81 0.73 0.65 0.73 0.51 0.52

27 Firmicutes(100) Ruminococcaceae(100) Ruminococcaceae.UC(100) 0.51 0.57 0.46 0.74 0.48 1.19

29 Firmicutes(100) Firmicutes.UC(100) Firmicutes.UC(100) 0.77 0.64 0.42 0.81 0.55 0.55

33 Bacteroidetes(100) Porphyromonadaceae(100) Parabacteroides(97) 0.26 0.32 0.58 0.29 0.31 1.12

34 Firmicutes(100) Lachnospiraceae(100) Lachnospiraceae.UC(100) 0.21 0.36 0.27 0.24 1.14 0.95

35 Bacteria.UC(100) Bacteria.UC(100) Bacteria.UC(100) 1.37 0.59 0.60 0.42 0.49 0.08

36 Verrucomicrobia(100) Verrucomicrobiaceae(100) Verrucomicrobiaceae.UC(100) 0.17 0.33 0.62 0.52 1.15 0.28


46 Firmicutes(100) Peptostreptococcaceae(100) Clostridium.XI(100) 0.01 0.07 0.09 0.23 0.44 1.54

47 Actinobacteria(100) Microbacteriaceae(98) Microbacteriaceae.UC(98) 0.78 0.19 0.21 0.44 0.40 0.76

71 Actinobacteria(100) Actinomycetales.UC(88) Actinomycetales.UC(88) 0.05 0.07 0.11 0.13 0.27 0.97

86 Firmicutes(100) Streptococcaceae(100) Lactococcus(100) 0.01 0.03 0.05 0.06 0.35 1.02

Totalabundanceincommunity 49.0 49.7 51.8 56.0 55.2 53.7

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Table 3. Heatmap of abundances of the 20 bacteria that based on loading values from PCA, contribute the most to the pattern observed in the PCoA in Fig. 3b: the dataset including biological replicates and all six treatments, but where technical replicates were averaged prior to PCoA and loading analyses. The heatmap scale is the percentage of reads assigned to a given taxon out of the total number of the high quality-filtered and classified reads for the treatment sample.

OTU Phylum Family

0

5

10

Genus

0R1

0R2

0R3

20R1

20R1

20R3

40R1

40R2

40R3

60R1

60R2

T60R3

T80R1

T80R2

T80R3

T100R1

T100R2

T100R3

4 Synergistetes(100) Synergistaceae(100) Synergistaceae_UC(100)

6 Firmicutes(100) Lachnospiraceae(100) Lachnospiraceae_UC(100)

15 Bacteroidetes(100) Porphyromonadaceae(100) Porphyromonadaceae_UC(96)

2 Firmicutes(100) Enterococcaceae(85) Enterococcaceae_UC(71)

3 Firmicutes(100) Lactobacillaceae(100) Lactobacillaceae_UC(98)

10 Proteobacteria(100) Betaproteobacteria_UC(100) Betaproteobacteria_UC(100)

14 Proteobacteria(100) Proteobacteria_UC(100) Proteobacteria_UC(100)

8 Actinobacteria(100) Actinomycetaceae(100) Actinomyces(100)

5 Planctomycetes(100) Planctomycetaceae(100) Planctomycetaceae_UC(100)


35 Bacteria_UC(100) Bacteria_unclassified(100) Bacteria_UC(100)

9 Firmicutes(100) Leuconostocaceae(100) Weissella(100)


1 Bacteroidetes(100) Bacteroidetes_UC(100) Bacteroidetes_UC(100)


25 Proteobacteria(100) Desulfovibrionaceae(79) Desulfovibrionaceae_UC(75)

33 Bacteroidetes(100) Porphyromonadaceae(100) Parabacteroides(97)



47 Actinobacteria(100) Microbacteriaceae(98) Microbacteriaceae_UC(98) Totalabundanceincommunity 31.0 40.7 43.6 42.9 30.6 42.3 39.2 43.3 36.9 46.6 39.7 39.5 40.0 41.3 42.3 48.2 40.4 35.6

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Table 4. Heatmap of abundances of the 20 bacteria that based on loading values from PCA, contribute the most to the pattern observed in the PCoA in Fig. 3c: the dataset including biological replicates for only cockroaches exposed on 0% or 100% fungus biomass. Technical replicates were averaged prior to PCoA and loading analyses. The heatmap scale is the percentage of reads assigned to a given taxon out of the total number of the high quality-filtered and classified reads for the treatment sample 0 5 >10

Discussion In this paper we present a comparison of bacterial community diversity within the guts of P. surinamensis cockroaches fed on increasing dietary proportions of fungal material in an attempt to demonstrate the influence that diet can have as a structuring force. By comparing 49 gut samples from individuals fed on diets consisting of 0% to 100% fungal material, we found that communities from cockroaches fed on diets consisting of similar fungal ratios were more similar to each other than to cockroaches fed on diets of differing fungus contents. Ordination analysis revealed a step-wise gradient pattern of community similarity that correlated with an increase in fungal material provided in the manipulated diet regimes (Fig.5a). These results suggest that diet has acted as a selective force over the cockroach gut community. Results also indicate that community similarity is higher in individuals fed on extremes of 80-100% fungus material, while communities of

OTU Phylum

Family Genus

0%R1

0%R2

0%R3

100%R1

100%R2

100%R3


2 Firmicutes(100) Enterococcaceae(85) Enterococcaceae_UC(71)

3 Firmicutes(100) Lactobacillaceae(100) Lactobacillaceae_UC(98)


5 Planctomycetes(100) Planctomycetaceae(100) Planctomycetaceae_UC(100)

4 Synergistetes(100) Synergistaceae(100) Synergistaceae_UC(100)

9 Firmicutes(100) Leuconostocaceae(100) Weissella(100)


33 Bacteroidetes(100) Porphyromonadaceae(100) Parabacteroides(97)


7 Bacteroidetes(100) Bacteroidetes_unclassified(100) Bacteroidetes_UC(100)

35 Bacteria_UC(100) Bacteria_unclassified(100) Bacteria_UC(100)


27 Firmicutes(100) Ruminococcaceae(100) Ruminococcaceae_UC(100)

71 Actinobacteria(100) Actinomycetales_UC(88) Actinomycetales_UC(88)

25 Proteobacteria(100) Desulfovibrionaceae(79) Desulfovibrionaceae_UC(75)

8 Actinobacteria(100) Actinomycetaceae(100) Actinomyces(100)



46 Firmicutes(100) Peptostreptococcaceae(100) Clostridium_XI(100)

Totalabundanceincommunity 28.9 37.8 43.6 45.7 38.9 39.7

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individuals that were exposed to mid and low range ratios of 0%, 20%, 40%, and 60% fungal material show more relaxed community similarities. For example, communities of cockroaches fed on a diet of 80%:20% leaf litter:fungus overlapped with cockroaches fed on a more balanced leaf litter to fungus diet. This suggests that diet has a stronger effect on community structure when the diet is more uniform, i.e., in this experiment when it was almost exclusively fungus material. Further ordination analysis using 18 gut samples averaged across the technical replicates indicated a similar pattern of community structuring, as the same stepwise-gradient pattern in community similarity was observed from 0% to 100% fungus ratios (Fig. 5b). Averaging across technical replicates again resulted in distinct clustering patterns, with high fungal regimes of 20%:80% and 0%:100% leaf litter:fungus material overlapping extensively without interspersing with communities from cockroaches fed on lower amounts of fungus material. Mid range fungal diets also shared high similarity, but again interspersed with some cockroach communities fed on 20% fungus, suggesting overlap in community similarity between these diet regimes. Communities that were not exposed to fungus clustered distinctly from all but one of the other diet samples, sharing similarity only with communities from a cockroach fed on 20% fungus. A final PCoA analysis demonstrated the clear separation in community similarity between cockroaches fed on 0% fungus and 100% fungal biomass (Fig. 3c). These results indicate that the ratio of leaf litter to fungus material provided to cockroaches in manipulated diet regimes plays a role in determining community composition. Below we discuss the associated abundances of the bacteria that may drive the observed patterns in community composition and compare the observed microbial diversity to that of fungus-growing termites. Taxonomic composition of bacterial communities across increasing fungal ratios Gut communities were dominated by Bacteroidetes, Firmicutes and Proteobacteria, but Synergistetes and Planctomycetes were also abundant, with the former being particularly well represented in cockroaches fed on low amounts of fungus material. Former studies have established the predominance of Bacteroidetes and Firmicutes as a common feature in the cockroach gut and represent lineages that are shared amongst omnivorous animals (Schauer, Thompson & Brune, 2012; Dietrich, Köhler & Brune, 2014). Comparisons of 0% fungal treatment samples to that of previously obtained data on P. surinamensis indicates a high abundance of Bacteroidetes, including families such as the Porphyromonadaceae, as well as a prevalence of Firmicutes such as the Lactobacillaceae (Mikaelyan, Köhler et al., 2015). However, unlike our study, the survey of clone libraries of P. surinamensis did not record high abundance of phyla such as Synergistetes and Planctomycetaceae. Differences in the naïve community of P. surinamensis found in our study compared to previous assessments may be due to the use of the SILVA database as reference for taxonomic classification. This database lacks the depth of Dictb and resulted in low classification success rates (Table. 1). To investigate bacterial lineages that may contribute to diet associated patterns in community composition, heatmaps were constructed for the top 20 bacteria that, based on PCA loading values, contributed most to patterns observed under community similarity analysis using PCoAs. A heat map of abundances of the top 20 bacteria that contributed to the clear separation in community similarity between 0% and 100% fungal treatments (Table 4) revealed that Bacteroidetes and Firmicutes dominated both communities, with Enterococcaceae and Lachnospiraceae remaining at high abundance throughout both treatments. Such a result highlights the high degree of community similarity that

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omnivorous cockroaches share with their descendants, the fungus-growing termites (Dietrich, Köhler & Brune, 2014; Otani et al., 2014). This heatmap also displayed abundance shifts of several bacterial lineages that are scarce in cockroaches fed on 0% fungus but increase in abundance in communities of cockroaches fed on only fungal biomass, a pattern mirrored in analysis of the top 20 most abundant bacteria across the 49 gut samples (Tables 2) and in the heatmap displaying the abundances of bacteria that contributed most to the pattern observed across averaged technical replicates (Table 3). At the family level, Desulfovibrionaceae and Ruminococcaceae increase in average abundance from the 0% to 100% treatment. At the genus-level, genera such as Clostridium_XI and Parabacteroides increase with an increase in fungus ratio. Lineages that appear to contribute to separation between the two extremes in diet are however in relatively low abundance compared to other bacteria that are instead dominant throughout all treatments, such as Enterococcaceae and Lactobacillaceae (Table 2). This pattern suggests that it is small changes in the abundance of less dominant bacteria within the cockroach gut community that are contributing to the separation observed between these two extremes of diet. This suggests that a switch to a fungal diet may be promoting specific lineages that are in low abundance in the cockroach gut, but may have the capacity to breakdown fungal material as cockroaches are exposed to a more proteinaceous diet (Hyodo et al. 2003; Deitrich et al. 2014). Community comparison to the gut communities of fungus-growing termites Fungus-growing termites have a shared core of gut bacteria that is consistently associated in relatively high abundance across the diversity of fungus-growing species (Otani et al., 2014). This core is dominated by taxa from the Bacteroidetes and Firmicutes, as well as phyla including Proteobacteria, Spirochetes, Synergistetes, Planctomycetes and Actinobacteria. Previous studies have established the strong similarity reflected between this core set of microbiota and that of the termite’s omnivorous ancestors, the cockroaches, and the gut community of P. surinamensis within this study is also dominated by taxa from the Bacteroidetes, Firmicutes and Planctomycetes (Dietrich et al., 2014). In order to conclude whether diet manipulations were able to shift the cockroach gut community to that of a fungus-growing termite community, it is important to access whether lineages that contributed to the stepwise change in community similarity from 0% to 100% fungus in this study are also found in fungus-growing termite gut community. Analysis of the bacteria that contributed most to the separation between cockroaches fed on 0% and 100% cockroaches revealed shifts in the abundance of a number of taxa that were in low abundance in the overall cockroach gut community. For example, Desulfovibrionaceae experiences an average increase in abundance of 1.42% from the 0% to 100% treatment and Ruminococcaceae also increased in abundance in the 100% treatment cockroaches. These bacteria constitute a major part of the fungus-growing termite microbiota, with Otani et al. (2014) indicating that high abundances of Desulfovibrio 3 (3.3% average abundance across 9 termite species) and gut cluster 1 in the Ruminococcaceae (4.3%) help to drive the pattern of community similarity observed between the fungus-growing termite core and cockroach gut community (Otani et al. 2014). Other bacteria that also increase in abundance as cockroaches are fed on a completely fungal diet include the genus level taxon Clostridium_XI and family-level Porphyromonadaceae. These bacteria are also found in lower abundances within the

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fungus-growing termite core and suggests that rarer lineages found within P. surinamensis are promoted by a fungal diet as the cockroach gut become more fungus-growing termite-like (Otani et al., 2014). Conclusions This study demonstrates the role that dietary shifts can have in determining gut community composition in the cockroach P. surinamensis. In particular, an increase in dietary fungal biomass is able to promote bacterial lineages that are otherwise found in low abundance within the cockroach gut, such as members of the Desulfovibrio, Ruminococcaceae and Porphyromonadaceae. The shared presence of these taxa in the fungus-growing termite core suggests a shift in the cockroach community to that of a more fungus-growing termite-like community, demonstrating the role diet may have played in driving the community similarity observed between the fungus-growing termites and their omnivorous ancestors. However, the main bacteria that constitute the P. surinamensis gut community, such as Enterococcaceae and Lactobacillaceae, remained in high abundance across all fungal ratios. These taxa are not found in the fungus-growing termite core and suggests that, like other omnivorous cockroaches, P. surinamensis harbours a highly dynamic core of gut bacteria that is maintained even after fundamental dietary shifts (Schauer & Thompson et al. 2014; Pérez-Cobas et al. 2015). Among the dominant bacteria that displayed high abundance across dietary treatments were Synergistetes and Planctomycetes. These bacteria were recorded in only very low abundance in previous studies of this cockroach species (Mikaelyan & Köhler et al. 2015). Their high relative abundance within this study could be due to the relatively low classification success achieved under the SILVA database. In order to further clarify the ability of diet to structure the gut communities of P. surinamensis, alternative reference databases such as Dictb (generated from the SILVA database with additional termite and cockroach gut 16S rRNA sequences (Mikaelyan & Köhler et al.2015)) must be used to accurately determine the identity of key community members within the cockroach gut. A more in depth taxonomic analysis will be needed to clarify the ability of the dominant community to remain resilient against dietary shifts or alternatively indicate that it is strong individual variation that is able to mask taxonomic responses to dietary shifts (Schauer & Thompson et al. 2014). Further diet manipulation experiments are required to clarify the role diet can play in structuring gut bacterial communities. For example, gradient manipulations of fibre-based diets may help identify lineages involved in fibre degradation (Mikaelyan & Strassert et al., 2014). Future studies should also utilise published metagenomic data on the fungus-growing termites to help identify potential complementary roles of major bacterial lineages that increase in abundance in cockroaches fed on increasing ratios of fungal biomass (Liu et al., 2013; Poulsen et al., 2014). Acknowledgements

We thank Harriette Carrington from the Microbial Systems molecular biology lab at the University of Michigan Medical School for laboratory assistance in the MiSeq library preparation, as well as Sylvia Mathiasen and Panagiotis Sapountzis for laboratory assistance. We thank Imperial College London’s biology with research abroad course coordinator Dave Hartley for comments on previous drafts of the manuscript and guidance throughout the project. The work was funded by the Erasmus student exchange program as part of Imperial College London’s Biology with Research Abroad degree to

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CR, a PhD stipend to SO jointly funded by the Department of Biology, University of Copenhagen and the Danish National Research Foundation Centre of Excellence Centre for Social Evolution (DNRF57) to Jacobus J. Boomsma, and a Villum Kann Rasmussen Young Investigator Fellowship (VKR10101) to MP.

Data Deposition

MiSeq data will be deposited in a public repository such as GenBank once this manuscript is being prepared for publication in a peer-reviewed journal.

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Mikaelyan, A., Thompson, C. L., Hofer, M. J. & Brune, A. (2015) The deterministic assembly of complex bacterial communities in germ-free cockroach guts. American Society of Microbiology. Available from: doi: 10.1128/AEM.03700-15.

Moran, N. A., Hansen, A. K., Powell, E. J. & Sabree, Z. L. (2012) Distinctive gut microbiota of Honey Bees assessed using deep sampling from individual worker bees. PLOS One. 7 (4), e36393. Available from: doi: 10.1371/journal.pone.0036393.

Ni, J. & Tokuda, G. (2013) Lignocellulose-degrading enzymes from termites and their symbiotic microbiota Biotechnology Advances. 31 (6), 838-850. Available from: doi: 10.1016/j.biotechadv.2013.04.005.

Nobre, T. & Aanen, D. (2012) Fungiculture or Termite Husbandry? The Ruminant Hypothesis. Insects. 3 (4), 307-323. Available from: doi: 10.3390/insects3010307.

Ohkuma, M. (2008) Symbioses of flagellates and prokaryotes in the gut of lower termites. Trends in Microbiology. 16 (7), 345-352. Available from: doi: http://dx.doi.org/10.1016/j.tim.2008.04.004.

Otani, S., Hansen, L. H., Sørensen, S. J. & Poulsen, M. (2016) Bacterial communities in termite fungus combs are comprised of consistent gut deposits and contributions from the environment. Microbial Ecology. 71.1, 207-220. Available from: doi: 10.1007/s00248-015-0692-6.

Otani, S., Mikaelyan, A., Nobre, T. & Hansen, L. H. et al. (2014) Identifying the core microbial community in the gut of fungus-growing termites. Molecular Ecology. 23, 4631-4644. Available from: doi: 10.1111/mec.12874.

Pérez-Cobas, A. E., Maiques, E., Angelova, A., Carrasco, P., Moya, A. & Latorre, A. (2015) Diet shapes the gut microbiota of the omnivorous cockroach Blattella germanica. FEMS Microbiology Ecology. 91 (4), fiv022. Available from: doi: 10.1093/femsec/fiv022.

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Poulsen, M. (2015) Towards an integrated understanding of the consequences of fungus domestication on the fungus growing termite gut microbiota. Environmental Microbiology 81, 6577-6588. Available from: doi: 10.1128/AEM.02637-15.

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Rahman, N., Parks, D. H., Willner, D. L., Engelbrektson, A. L., Goffredi, S. K., Warnecke, F., Scheffrahn, R. H. & Hugenholtz, P. (2015) A molecular survey of Australian and North American termite genera indicates that vertical inheritance is the primary force shaping termite gut microbiomes. Microbiome. 3 (5), 1-16. Available from: doi: 10.1186/s40168-015-0067-8.

Roth, L. M. (1974) Reproductive potential of bisexual Pycnoscelus indicus and clones of its parthenogenetic relative, Pycnoscelus surinamensis. Annals of the Entomological Society of America. 67 (2), 215-223. Available from: doi: http://dx.doi.org/10.1093/aesa/67.2.215.

Sachs, J. L., Skophammer, R. G. & Regus, J. U. (2011) Evolutionary transitions in bacterial symbiosis. Proceedings of the National Academy of Sciences of the United States of America 108, 10800-10807. Available from: doi: 10.1073/pnas.1100304108.

Sanders, J., Powell, S., Kronauer, D., Vasconcelos, H., Frederickson, E. & Pierce, N. E. (2014) Stability and phylogenetic correlation in gut microbiota: lessons from ants and apes. Molecular Ecology. 23 (6), 1268-1283. Available from: doi: 10.1111/mec.12611.

Sapountzis, P., Zhukova, M., Hansen, L. H., Sørensen, S. J. & Schiøtt, B., J.J. (2015) Acromyrmex leaf-cutting ants have simple gut microbiota with nitrogen-fixing potential. Applie and Environmental Microbiology. 81 (16), 5527-5537. Available from: doi: 10.1128/AEM.00961-15.

Schauer, C., Thompson, C., Brune, A. & Korb, J. (2014) Pyrotag Sequencing of the Gut Microbiota of the Cockroach Shelfordella lateralis Reveals a Highly Dynamic Core but Only Limited Effects of Diet on Community Structure Plos One. 9 (1), e85861. Available from: doi: 10.1371/journal.pone.0085861.

Schauer, C., Thompson, C. L. & Brune, A. (2012) The bacterial community in the gut of the Cockroach Shelfordella lateralis reflects the close evolutionary relatedness of cockroaches and termites Applied and Environmental Micobiology. 78 (8), 2758-2767. Available from: doi: 10.1128/AEM.07788-11.

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Schloss, P. D., Westcott, S. L., Ryabin, T. & Hall, J. R., et al. (2009) Introducing mothur: open-source, platform-independent, community-supported software for describing and comparing microbial communities Applied and Environmental Microbiology. 75 (23), 7537-7541. Available from: doi: 10.1128/AEM.01541-09.

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Supplementary Material Table S1 Gut Samples selected for amplification and MiSeq Sequencing Sample ID Leaf Litter: Fungus (%) Biological Replicate Technical Replicate Yield Dilution

1 100:0 1 1 1st NONE

2 100:0 1 2 1st NONE

3 100:0 1 3 1st NONE

4 100:0 2 1 1st NONE

5 100:0 2 2 2nd NONE

6 100:0 2 3 1st NONE

7 100:0 3 1 2nd NONE

8 100:0 3 2 2nd NONE

9 100:0 3 3 2nd NONE

10 80:20 1 1 1st NONE

11 80:20 1 2 1st NONE

12 80:20 1 3 1st NONE

13 80:20 2 1 1st NONE

14 80:20 2 2 2nd NONE

15 80:20 2 3 1st NONE

16 80:20 3 1 2nd 01:10

17 80:20 3 2 1st 01:10

18 80:20 3 3 1st 01:10

19 60:40 1 1 1st NONE

20 60:40 1 2 1st NONE

21 60:40 1 3 1st NONE

22 60:40 2 1 1st NONE

23 60:40 2 2 2nd NONE

24 60:40 2 3 2nd NONE

25 60:40 3 1 2nd NONE

26 60:40 3 2 2nd NONE

27 60:40 3 3 1st NONE

28 40:60 1 1 1st NONE

29 40:60 1 2 1st NONE

30 40:60 1 3 1st NONE

31 40:60 2 1 1st NONE

32 40:60 2 2 1st NONE

33 40:60 2 3 1st NONE

34 40:60 3 1 1st NONE

35 40:60 3 2 1st NONE

36 40:60 3 3 1st NONE

37 20:80 1 1 1st NONE

38 20:80 1 2 1st NONE

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39 20:80 1 3 1st NONE

40 20:80 2 1 1st NONE

41 20:80 2 2 1st NONE

42 20:80 2 3 1st NONE

43 20:80 3 1 1st NONE

44 20:80 3 2 1st NONE

45 20:80 3 3 1st NONE

46 0:100 1 1 1st NONE

47 0:100 1 2 1st NONE

48 0:100 1 3 2nd NONE

49 0:100 2 1 2nd 01:10

50 0:100 2 2 2nd NONE

51 0:100 2 3 2nd NONE

52 0:100 3 1 2nd NONE

53 0:100 3 2 1st 01:50

54 0:100 3 3 2nd NONE

Table S1. Target PCR products were visualized by agarose gel electrophoresis before submission to MiSeq. 1st yield samples that were unable to be visualized clearly on a gel were run again using the 2nd yield elution samples. Samples that still failed to display a significant banding pattern were diluted in order to counter any impurities present in the sample. DNA template samples were diluted to 1/10 and 1/50 of their original concentration with the addition of sterile distilled water and run using the same PCR conditions and visualised on an agarose gel. Samples that were then clearly visible on an agarose gel and therefore contained quantifiable DNA were submitted for MiSeq

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Table S2 Heatmap of abundances of the 20 bacteria that based on loading values from PCA, contribute the most to the pattern observed in the PCoA in Fig. 3a: the full dataset including both technical and biological replicates and all six treatments. -alsosenttoDrDaveHartleyasattachment,asfilewastoolargetopositionintoworddocument*

OTU Phylum Family Genus8 Actinobacteria(100) Actinomycetaceae(100) Actinomyces(100) 0.72 0.41 0.05 0.27 0.31 1.57 0.73 4.28 1.13 0.35 0.69 0.84 1.49 1.83 3.14 0.98 1.75 2.87 0.86 2.14 2.84 1.87 1.17 0.51 1.01 4.44 2.33 1.60 2.74 1.70 1.62 1.92 2.05 1.06 3.28 0.63 2.20 2.53 2.31 2.96 2.25 1.35 0.88 2.91 1.78 1.12 0.77 1.01 0.50

15 Bacteroidetes(100) Porphyromonadaceae(100) Porphyromonadaceae_unclassified(96) 0.25 0.36 0.59 0.25 0.20 2.06 0.22 0.29 0.41 0.94 0.57 0.41 0.50 0.47 0.90 1.00 0.25 2.51 1.20 1.00 1.33 0.44 1.61 1.66 1.22 1.06 0.95 0.99 0.84 1.37 0.84 2.12 1.57 1.37 0.54 1.17 2.44 1.87 1.44 1.98 1.14 1.90 1.43 1.04 2.51 3.02 2.33 1.42 1.0347 Actinobacteria(100) Microbacteriaceae(98) Microbacteriaceae_unclassified(98) 0.64 0.42 0.75 2.11 1.28 0.02 1.17 0.03 0.02 0.36 0.35 0.01 0.00 0.02 0.00 0.20 0.55 0.00 0.00 0.07 0.06 0.66 0.14 0.33 0.02 0.02 0.64 0.14 0.99 0.71 1.18 0.11 0.10 0.06 0.67 0.73 0.64 0.06 0.08 0.82 0.06 0.47 2.79 0.38 0.64 0.31 0.26 0.38 0.4236 Verrucomicrobia(100) Verrucomicrobiaceae(100) Verrucomicrobiaceae_unclassified(100) 0.24 0.04 0.08 0.00 0.00 0.31 0.09 0.53 0.63 0.04 0.08 0.05 0.44 0.59 0.02 0.14 1.00 0.74 0.19 0.74 0.21 1.12 0.33 0.16 1.02 1.36 0.62 0.15 0.47 0.32 0.32 0.92 0.11 0.86 1.18 0.18 0.89 0.15 1.32 2.86 2.25 0.60 0.36 0.26 0.19 0.46 0.18 0.38 0.1612 Bacteroidetes(100) Porphyromonadaceae(100) Porphyromonadaceae_unclassified(100) 0.17 0.44 0.54 0.47 0.49 1.02 0.90 0.37 0.61 0.84 0.64 1.88 0.77 1.16 1.22 3.60 0.95 0.97 1.01 1.29 0.76 0.03 0.81 3.42 1.39 0.54 0.53 1.20 1.23 2.29 1.95 1.00 1.82 1.31 0.43 2.94 2.20 2.51 1.14 0.54 1.02 1.78 1.24 1.10 2.04 1.98 1.71 1.64 0.7032 Proteobacteria(100) Campylobacterales_unclassified(100) Campylobacterales_unclassified(100) 1.97 0.61 1.55 0.55 0.14 0.31 0.77 0.59 0.75 1.43 1.41 0.24 0.87 0.28 0.24 1.27 0.21 0.19 0.36 0.12 0.39 0.61 0.25 0.60 0.60 0.22 0.59 0.58 0.95 0.27 0.63 0.25 1.03 0.72 0.54 0.61 0.86 0.54 0.46 0.24 0.15 0.12 0.21 0.15 0.49 1.03 0.64 0.22 0.3814 Proteobacteria(100) Proteobacteria_unclassified(100) Proteobacteria_unclassified(100) 1.95 1.36 1.07 0.02 0.81 2.34 1.72 1.21 0.16 2.33 0.20 1.83 1.76 0.33 0.87 1.36 1.35 1.77 1.93 1.60 0.31 1.74 0.25 1.30 0.14 1.29 0.16 1.77 0.24 1.72 1.13 1.42 2.60 1.39 0.63 2.83 2.12 1.25 1.16 0.16 2.91 1.38 0.40 1.23 0.14 1.93 0.43 0.25 0.1320 Proteobacteria(100) Proteobacteria_unclassified(100) Proteobacteria_unclassified(100) 1.46 0.81 0.59 0.40 0.13 0.57 0.42 0.25 1.42 0.59 0.91 2.33 0.30 0.26 0.61 0.65 0.27 0.19 1.57 1.14 0.55 0.84 0.75 0.67 1.22 1.64 1.24 0.48 0.80 1.24 0.75 0.73 1.36 0.57 0.55 0.89 0.65 1.47 1.37 0.42 1.17 1.04 1.38 0.74 1.07 0.09 0.71 1.05 0.2611 Bacteroidetes(100) Bacteroidetes_unclassified(100) Bacteroidetes_unclassified(100) 2.57 1.85 1.42 2.03 1.42 0.93 1.82 1.29 1.49 1.72 2.05 1.08 1.25 2.69 1.76 1.83 0.84 3.10 1.45 1.00 0.47 2.58 0.67 0.95 1.23 1.10 1.58 1.58 1.79 2.12 1.42 1.92 1.51 1.11 1.75 2.15 1.20 1.00 1.26 1.03 1.32 2.03 0.92 0.27 0.64 0.84 0.85 0.85 0.5533 Bacteroidetes(100) Porphyromonadaceae(100) Parabacteroides(97) 0.33 0.28 0.11 0.07 0.29 0.60 0.17 0.23 0.54 0.24 0.24 0.36 0.13 0.93 0.22 0.10 0.32 0.24 0.26 0.55 1.01 0.15 0.47 1.32 0.51 0.39 0.07 0.17 0.32 0.20 0.35 0.60 0.53 0.08 0.05 0.55 0.35 0.27 0.48 0.25 0.05 0.35 0.28 0.85 3.59 1.30 2.68 0.43 0.2539 Bacteroidetes(100) Porphyromonadaceae(95) Porphyromonadaceae_unclassified(95) 0.06 0.07 0.19 0.20 0.02 0.13 0.02 0.02 0.47 0.48 0.23 0.35 0.65 0.30 0.31 0.28 0.07 0.26 0.03 0.13 0.13 0.11 0.33 1.02 0.78 0.17 0.24 0.01 0.15 0.09 0.05 0.12 1.02 0.57 0.00 0.55 0.27 2.26 2.52 0.36 0.26 0.67 0.67 2.06 0.39 0.87 2.01 0.16 0.1313 Proteobacteria(100) Betaproteobacteria_unclassified(100) Betaproteobacteria_unclassified(100) 1.64 1.21 1.63 1.02 1.46 1.44 1.31 0.82 0.61 1.41 0.62 1.41 1.67 0.99 1.15 2.57 0.31 1.29 1.88 1.58 1.14 0.32 2.78 0.65 1.26 1.42 1.34 1.75 2.24 0.96 1.87 1.59 0.52 1.89 0.99 1.61 1.28 1.14 0.80 0.42 0.98 0.20 3.32 1.17 0.96 0.91 1.07 3.22 1.9210 Proteobacteria(100) Betaproteobacteria_unclassified(100) Betaproteobacteria_unclassified(100) 3.11 4.72 4.68 3.52 5.09 0.35 3.11 2.10 0.13 0.66 0.44 2.37 4.46 1.08 1.45 1.64 2.07 1.18 1.56 2.57 0.23 1.20 1.34 0.10 0.45 0.32 1.88 1.02 0.45 0.11 1.41 1.27 2.45 0.66 0.05 0.68 0.46 0.23 0.68 0.47 0.08 0.16 3.07 2.82 0.74 1.17 0.47 0.71 3.63

2 Firmicutes(100) Enterococcaceae(85) Enterococcaceae_unclassified(71) 1.56 0.73 0.94 1.41 1.87 4.23 3.93 6.37 3.06 2.97 5.43 2.20 1.22 2.43 4.77 2.00 6.00 6.21 2.02 3.17 6.21 4.87 2.98 1.76 3.29 6.63 4.31 4.48 5.95 4.24 3.32 3.98 4.70 3.79 7.60 3.06 6.29 4.57 4.35 6.07 6.77 3.64 2.34 3.82 3.34 2.62 2.88 3.28 2.6535 Bacteria_unclassified(100)Bacteria_unclassified(100) Bacteria_unclassified(100) 2.59 2.52 2.25 0.87 0.38 0.97 0.94 0.79 2.12 0.74 0.51 0.26 0.17 0.43 0.70 0.51 0.30 0.46 0.73 0.32 0.58 1.31 0.14 0.38 0.31 0.34 0.43 0.45 0.30 0.32 0.37 0.59 0.69 0.29 0.31 0.49 0.11 0.29 0.90 0.89 0.25 0.39 0.04 0.09 0.03 0.11 0.10 0.13 0.04

6 Firmicutes(100) Lachnospiraceae(100) Lachnospiraceae_unclassified(100) 0.86 0.71 0.27 0.13 0.63 2.03 0.91 5.81 1.76 0.93 1.98 2.05 1.96 2.06 5.53 2.13 4.61 4.04 1.58 2.43 2.22 3.30 1.67 1.29 2.19 3.51 2.79 3.72 0.71 2.49 2.05 2.21 2.85 2.25 5.31 1.90 2.80 3.43 3.31 2.65 3.35 1.85 1.45 2.37 3.33 2.04 1.87 1.75 1.3016 Bacteroidetes(100) Bacteroidetes_unclassified(100) Bacteroidetes_unclassified(100) 0.25 0.25 0.21 0.15 0.27 0.97 0.19 0.16 0.65 0.70 1.40 1.61 0.74 1.21 0.73 1.14 0.69 1.16 1.16 1.24 2.01 0.30 1.28 1.63 1.82 0.95 2.75 0.96 1.88 1.04 0.49 1.35 2.07 0.86 0.64 0.98 1.08 1.41 1.81 1.33 1.34 1.67 1.66 1.12 1.11 1.86 2.88 1.06 1.4617 Proteobacteria(100) Proteobacteria_unclassified(100) Proteobacteria_unclassified(100) 0.84 0.50 1.69 1.57 0.74 1.46 0.56 0.63 1.22 1.16 0.41 1.43 1.53 1.35 0.82 1.03 0.16 0.96 1.53 0.66 2.00 0.67 1.86 1.84 1.12 0.52 0.74 1.70 1.12 1.81 1.48 1.21 1.25 0.96 0.36 1.97 1.64 1.35 0.53 1.04 0.63 1.60 0.56 1.40 0.85 0.72 1.10 1.33 0.67

7 Bacteroidetes(100) Bacteroidetes_unclassified(100) Bacteroidetes_unclassified(100) 1.68 1.68 1.53 1.61 1.57 1.28 2.55 2.82 1.06 1.40 1.22 1.66 1.34 1.39 1.78 2.54 0.74 2.13 2.09 1.26 0.76 2.25 1.42 2.39 2.04 0.84 1.54 2.55 1.94 2.17 1.42 1.25 1.23 2.60 1.82 2.18 2.75 1.87 2.22 0.89 0.68 1.72 0.90 0.90 2.02 1.94 1.43 2.34 2.1034 Firmicutes(100) Lachnospiraceae(100) Lachnospiraceae_unclassified(100) 0.00 0.00 0.00 0.00 0.04 1.17 0.07 0.41 0.36 0.61 0.99 0.06 0.21 0.11 0.18 0.10 0.50 0.38 0.22 0.18 0.24 0.12 0.83 0.06 0.60 0.28 0.38 0.29 0.43 0.20 0.16 0.34 0.09 0.09 1.63 0.39 1.22 0.60 0.69 2.14 2.12 0.94 0.60 0.98 0.83 2.10 1.08 0.89 0.48

Total abundance in community 22.91 18.97 20.12 16.64 17.11 23.74 21.60 28.98 18.61 19.92 20.37 22.44 21.44 19.92 26.39 25.05 22.95 30.62 21.63 23.19 23.46 24.48 21.10 22.05 22.21 27.03 25.12 25.60 25.52 25.38 22.80 24.92 29.53 22.50 28.34 26.49 31.43 28.81 28.85 27.52 28.78 23.84 24.49 25.65 26.69 26.43 25.46 22.50 18.76

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Failures,frustrationsandrevisionsTermitomycesculturingandprovisiontocockroaches.ThemassculturingofthefungusTermitomyceshasprovidedasignificantpracticalchallengetothisstudy.Determiningtheproportionsoffungustoinitiallyfeedtherespectivecockroachsubpopulationsishardtocontrolfor,withfungusoftenprovingdifficulttoharvestfrompreparedPDAmediainatimeefficientmanner.Determiningthehealthofthecockroachesandtheviabilityofthefungusasaneffectivedietcomponentisalsochallenging,withcockroachesfedonan100%fungaldietfindingithardtosurviveonthisrestrictivediet.Decidingwhethertofeedcockroachesfresh,dryorinactivatedfungusisalsodifficult.ApilotstudymayberequiredinthefuturetodeterminewhichstateofTermitomycessuitscockroachconsumption.Theuniquenessofthisapproachtosuchadietmanipulationexperimenthasbroughtmanychallenges,withonlyaverylimitedsourceofbackgroundliteraturetoprovideinsighttosuchdietmanipulations.Thisapproachtoprovidecockroacheswithaoncelivingdietarycomponent,ratherthanusingasimplenutrientsupplementsuchasaproteinpowder,appearstobeunique(Schauer,Thompson&Brune,2013;Pérez-Cobasetal.,2015).

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