An Examination of the Characteristic

WHAT’S INSIDE? AN EXAMINATION OF THE CHARACTERISTIC

MICROBIOME OF PRAIRIE GRASSHOPPERS AND KATYDIDS

Thesis

Submitted to

The College of Arts and Sciences of the

UNIVERSITY OF DAYTON

In Partial Fulfillment of the Requirements for

The Degree of

Master of Science in Biology

Melani K Muratore

Dayton, OH

May 2020

WHAT’S INSIDE? AN EXPLORATION OF THE MICROBIOME OF PRAIRIE

GRASSHOPPERS AND KATYDIDS

Name: Muratore, Melani K

APPROVED BY:

Yvonne Sun, Ph.D. Faculty Advisor Assistant Professor Department of Biology

Chelse Prather, Ph.D. Committee Member Associate Professor Chair, Department of Biology

Ryan W. McEwan, Ph.D. Committee Member Associate Professor Department of Biology

Melani K Muratore

2020

iii

ABSTRACT

WHAT’S INSIDE? AN EXPLORATION OF THE MICROBIOME OF PRAIRIE

GRASSHOPPERS AND KATYDIDS

Name: Muratore, Melani K University of Dayton

Advisor: Dr. Yvonne Sun

There are many ways in which microbiomes can influence the health and fitness of their insect hosts. While much research has been done on the microbiomes of economically important species like honeybees, and charismatic groups like butterflies, little work has been done to understand the microbiome of grasshoppers and katydids.

Grasshoppers are an important herbivore in grassland ecosystems and provide important ecosystem services like nutrient cycling. Alternatively, grasshoppers can be a pest organism requiring management and control. In relationship to the total ecosystem, the limitation of abiotic factors like essential nutrients can influence the abundance and fitness of insect herbivores such as grasshoppers. However, the effect of these abiotic factors on grasshopper microbiomes is largely unknown. Using culture-independent, high-throughput 16s and ITS sequencing and statistical analysis, we examined the gut microbiome of six species of grasshoppers (n=60) from the site of a fully-factorial fertilization experiment in a coastal tallgrass prairie ecosystem in order to gain a better understanding of the microbial communities present across the orthopteran order in this ecosystem. We also examined the relationship between environmental nutrients and gut microbes of one orthopteran species, Orchelimum vulgare. Our observations support the

iv hypothesis that there is a “core” group of bacterial families in these grasshopper species and factors such as trophic behaviors and the evolution of the host may contribute to the shifts in prevalence among these core microbial groups. We also found significantly higher Shannon diversity in the gut bacterial communities of those grasshoppers from plots fertilized with added sodium in contrast to plots without sodium. There is significantly lower diversity in gut fungal communities from plots amended with nitrogen and phosphorus, as well as sodium, suggesting that both nitrogen and phosphorus and sodium are limiting nutrients for the fungal community. There was also a strong positive correlation between bacterial and fungal diversity within each sample. These results support the hypothesis that environmental nutrients will alter the characteristic gut microbiome of O. vulgare, though these alterations will be different for bacterial and fungal communities.

ACKNOWLEDGMENTS

I would like to express my appreciation to my advisor, Dr. Yvonne Sun, who first gave me the opportunity to do research in her lab in the Summer of 2017. She has been a supportive mentor, giving me many opportunities to learn and lots of help and direction.

She is someone who is enthusiastic about learning new things and collaborating with other scientists. She has a great love of teaching and a desire to share her knowledge and help her students succeed. She is one of the best instructors that I have had. It is her desire to collaborate and explore new ideas to which I owe my association with Dr.

Chelse Prather. I am grateful for Dr. Prather’s knowledge, patient guidance, and her enthusiasm for her work. I am also grateful for the good advice and kind assistance of Dr.

Ryan McEwan, who served as a member of my committee.

I would also like to express my gratitude to the other members of the Biology department past and present who have offered their help to me over the last several years.

I am very grateful for the other students who have shared their knowledge and friendship with me, especially Amanda Finke, Michaela Woods, and Julia Chapman who all helped with coding. I am grateful to Erica Rhinehart for her help in the lab.

I am most grateful for the support of my family, most of all my husband, Chris.

Ever since I decided to go “back to school,” you have all been behind me, encouraging me to persist.

This research would not have been possible without funding from the University of Dayton Graduate Student Summer Fellowship, a University of Dayton STEM catalyst grant, and a Keck Environmental Fellowship.

TABLE OF CONTENTS

ABSTRACT ...... iv ACKNOWLEDGMENTS ...... vi LIST OF FIGURES ...... viii LIST OF TABLES ...... ix INTRODUCTION ...... 1 CHAPTER 1 THE GUT BACTERIAL COMMUNITIES ACROSS SIX SPECIES FROM A COASTAL TALLGRASS PRAIRIE ...... 8 Abstract ...... 8 Introduction ...... 9 Materials and Methods ...... 11 Results ...... 13 Discussion...... 15 Conclusion ...... 21 Acknowledgements ...... 22 References ...... 22 CHAPTER 2 EFFECTS OF ENVIRONMENTAL NUTRIENTS ON BACTERIAL AND FUNGAL MICROBIOMES IN THE COMMON MEADOW KATYDID, ORCHELIMUM VULGARE ...... 32 Abstract ...... 32 Introduction ...... 33 Methods ...... 36 Results ...... 39 Discussion...... 42 Conclusion ...... 46 References ...... 47

vii

LIST OF FIGURES

Figure 1. Mean relative abundance of bacterial phyla in the gut of six grasshopper species……………………………………………….……….………………………….54

Figure 2. Mean relative abundance of prevalent bacterial families in the gut of six grasshopper species. ……………………………………………..……………………..55

Figure 3. Beta diversity comparison of bacterial communities…………………………56

Figure 4. Comparison of the gut microbiome in male and female grasshoppers……….57

Figure 5. Average relative abundance of microbial phyla in grasshopper samples from four treatments…………………………………………………………………………..58

Figure 6. Average relative abundance of microbial taxa in grasshopper samples from four treatments……………………………………….…………………………….59

Figure 7. Nonmetric multidimensional scaling of grasshopper gut community assemblages at the family level………………………………………………………….60

Figure 8. Analysis of diversity over four treatments………………………………….…61

Figure 9. An analysis of shared groups among the four treatments……………………..62

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LIST OF TABLES

Table 1. Gut bacterial richness and diversity for six grasshopper species……………....63

Table 2. Gut microbiome richness and diversity for four treatments……………………64

INTRODUCTION

Over the last decade, advances in culture-independent methods have led to an explosion in microbiome research. These methods include 16S rRNA gene sequencing which uses variable regions (V3-V4) of the 16S rRNA gene to identify bacterial and archaeal taxa from a variety of samples, such as content inside the gut. The discovery that humans have an intricate symbiosis with their commensal microbes from which they derive health and well-being represented a paradigm shift in the study of human physiology. Microbiomes can include both the friendly and unfriendly bacteria, archaea, fungi, viruses, and protists that gain shelter and a steady source of nutrients from their hosts. Recently uncovered effects of the human microbiome on human health and well- being include alterations to mood and behavior, benefits to digestion and nutrition including the synthesis of vitamins, and support of healthy aging and effective response to disease (1–4).

As research techniques and insight into the human microbiome progress, research into the microbiomes of other animal species has followed in its wake. Many animal gut microbiomes have been sequenced, generating knowledge that has already been put to good use in conservation efforts. Recent research into the microbiome of the eastern black rhino has indicated that there may be specific genera of bacteria that are associated with increased fertility and subsequent successful breeding in these animals (5) .

Microbiome research also facilitated the development of probiotic therapies to more

1 successfully foster whooping crane chicks in captivity. (6, 7). A better understanding of the characteristics and contributions of microbial communities in animals will lead to more effective conservation and management strategies (8–10).

Globally, insects are one of the most abundant forms of animal life. There are over one million species of insects with the majority of insect species still left to be discovered (11). Insects have tremendous economic and ecological significance. They are providers of indispensable ecosystem services like pollination and nutrient cycling, but they also pose threats as destructive pests and vectors of disease. Insect herbivores, grasshoppers included, are integral to grassland ecosystems. Their presence or absence can change the composition of soils, the structure of plant communities, and have cascading effects for other animals that depend on them as a food source (12–14). The order Orthoptera includes grasshoppers, katydids, and crickets. Though Orthopterans are relatively abundant, their microbiomes have not been characterized as extensively as some other, more charismatic insect species. Much of what we do know about insect herbivores comes from studies of Lepidopterans like butterflies, moths, and caterpillars, and Hymenopterans like bees. From these studies, it has been generalized that the insect herbivore microbiome is largely dominated by Proteobacteria, Firmicutes, and

Actinobacteria (15,16). Whether this generalization holds true across all insect herbivores has not been fully investigated.

The connections between the microbiome, the host, and the ecosystem constitute an intricate tripartite relationship. For my thesis research, we have studied the relationship between the grasshopper, its microbiota, and the tallgrass prairie. The tallgrass prairie is a threatened ecosystem which is home to many unique animal and

2 plant species. At one point dominating the North American Great Plains, only 4% of the tallgrass prairie remains today (17). Tallgrass prairies are subjected to a number of disturbances like burning, grazing and fertilization (18). Prairie soils are generally productive and fertile; Their microbial communities are resilient, but they do change with changing soil conditions (19). Fertilization, or changes in nutrient levels, can cause changes to the soil microbiome, including shifts in the constituent microbial community and also shifts in the relative abundances of the constituents. While it is unknown whether changes to the soil nutrient levels in tallgrass prairies will impact the microbiome of the O. vulgare, it is commonly accepted that changes to host nutrients will have an effect on host microbiome. For example, changes in the human microbiome are more positively correlated with the environment than with host genetics (20), plants grown in contaminated versus uncontaminated soils experience significant shifts in dominant microbial groups (21), and insect microbiome can also be determined by the insect host environment, especially host diet (22).

My research includes two main focuses: The first focus is to determine whether there is a characteristic gut microbiome for tallgrass prairie grasshoppers. The second objective is to establish how grasshopper gut microbiomes are affected by changing soil nutrients. We collected six species of orthopteran from a tallgrass prairie, which was the site of a fully-factorial fertilization experiment, and analyzed the microbial communities from the entire gut samples. We expected that grasshoppers from the tallgrass prairie would share characteristic microbiomes similar to other insect herbivores with

Proteobacteria, Firmicutes, and Actinobacteria as the dominant members. Of the six species of grasshopper collected, there is only one strictly herbivorous species (Paroxya

3 atlantica) while the other five species have omnivorous feeding behaviors. It is reasonable to suppose that grasshoppers with different feeding behaviors may have differences in their gut microbiome. We hypothesized that there may be differences between the characteristic gut microbiome of grasshoppers depending on the grasshopper species, causing shifts in the relative abundance of characteristic taxa. The second objective of this work was to determine whether abiotic factors like added soil nutrients affect the compositions of the gut microbiomes. While prior research has established that shifts in soil nutrients impact the microbiomes of the soils themselves (23–25), we hypothesized that based on the connection between grasshoppers and the soil systems, there would also be a significant difference in grasshopper gut microbial community compositions when changes in soil nutrients, like nitrogen, phosphorus, and sodium, were made.

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Disturbances: A Review. Agronomy. 2018 Dec 1;8(12):300.

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CHAPTER 1

THE GUT BACTERIAL COMMUNITIES ACROSS SIX SPECIES FROM A

COASTAL TALLGRASS PRAIRIE

Abstract

Insect microbiomes play an important role in the health and fitness of insect hosts by contributing to nutrient absorption, immune health, and overall ecological fitness. As such, research interests in insect microbiomes have focused on agriculturally and industrially important organisms such as honey bees and termites. Orthopterans, on the other hand, have not been well explored for their resident microbial communities.

Grasshoppers are an integral part of grassland ecosystems and provide important ecosystem services. Conversely, grasshoppers can be an agricultural pest requiring management with broad spectrum pesticides. However, little is known about the microbiomes of grasshoppers and their potential contribution to grasshopper biology.

Here we examine the gut microbiome of six species of grasshoppers (n=60) from a coastal tallgrass prairie ecosystem to gain a better understanding of the microbial communities present across the orthopteran order in this ecosystem. We found that there are bacterial phyla common to all six grasshopper species: Actinobacteria, Proteobacteria,

Firmicutes, and to a lesser degree, Tenericutes. Although the grasshopper species shared a high relative abundance of these groups, there were notable shifts in dominant phyla

8 depending on the grasshopper species. Moreover, measures of alpha diversity revealed a more diverse microbiome in males than females. Our observations support the hypothesis that there is a “core” group of bacterial families in these grasshopper species and factors such as trophic behaviors and the evolution of the host may contribute to the shifts in prevalence among these core microbial groups.

Introduction

The relationships between insect hosts and their microbiomes have wide-ranging effects on the host, and by extension, the ecosystem. For example, termites, wood- feeding cockroaches, and carrion-feeding burying beetles are aided by endosymbionts in nutrient acquisition (1,2). Female red firebugs of the genus Pyrrhocoris smear their eggs with their microbiota in order to ensure the success of their offspring (3). Many lepidopterans also transfer microbes vertically by smearing their microbiota on the eggs after oviposition (4). Other host-microbiome effects include defending the host against exogenous microorganisms that may be pathogenic (5) and influencing host behaviors like aggregation into large groups (6). With the great diversity of insect taxa, our understanding of the extent their microbiomes contribute is still emerging.

Orthopterans, including both short-horned and long-horned grasshoppers, are key herbivores in worldwide grassland ecosystems and are one of the most important insect groups ecologically and economically (7). Grasshoppers vary in their feeding habits.

Some are strict herbivores, including specialist grass- or forb-feeders or generalists that feed on a wide range of plants. There are also omnivorous grasshoppers that are known to opportunistically feed on other insects or carrion. Because of their consumption of large amounts of plant biomass, grasshoppers function as key consumers that can increase the

9 pace of nutrient cycling, serving as a food source for higher trophic level organisms like avian predators (8,9) and affecting both above- and belowground decomposition and soil enzyme activity (10–12). As a result, grasshoppers have been suggested as a key indicator of the level of stability in grassland ecosystems, especially in threatened habitats such as prairies, where the soil fertility inevitably leads to agricultural use (13).

Alternatively, as a key consumer of plant mass, grasshoppers are also associated with

1.25 billion dollars of damage per year in the United States alone, inflicting crop damage and degrading range forage, often necessitating management with pesticides that create additional ecological disturbances (14). Therefore, the ecological and the economic importance of grasshoppers necessitate for a better understanding of their biology— including their microbial communities.

Much work has been done to characterize and understand the function of microbiomes in animal systems, with an increasing range of host models. The lion’s share of research on insect-herbivore microbiomes has focused mostly on model orders like the hymenopterans (including the honeybee), which are highly charismatic and offer obvious ecosystem services (15–17). Although orthopterans are ecologically and economically significant as a group, their microbiome is poorly understood. At the very least, insect herbivores, like orthopterans, likely rely on their microbiomes to extract nutrients from difficult to digest plant tissues (18). Therefore, understanding the microbiomes of live-captured grasshoppers from the field offers an opportunity to better understand the functional roles of an important herbivore host in the broader context of their environment. There is a strong possibility that grasshopper species from the same habitat, with similar natural histories, may share a characteristic or “core” group of

10 microbes in their gut. Recent efforts to understand insect herbivore microbiomes have sampled from frass, the gut in sections, or the gut in its entirety (19–21). In this study, to construct a more complete picture of grasshopper gut bacterial communities, we targeted the microbial community from the entire gut of six different grasshopper species collected from a Texas coastal tallgrass prairie. We identified a core group of common bacterial taxa, including those that contain well-known representatives of plant and animal pathogens, present in all six grasshopper species and detected a significant difference in alpha diversity of bacterial communities present in male versus female grasshoppers.

Materials and Methods

Sample Collection

Grasshoppers were collected using sweep nets (100 sweeps per sampling) from a

Texas coastal tallgrass prairie in 2017 within a large-scale fertilization experiment (22).

The grasshoppers were then shipped on ice to the University of Dayton (Dayton, OH) and stored frozen at -20°C until identification and dissection. Grasshopper species and sex were identified and recorded. We used only grasshoppers that could be easily dissected

(individuals at 4th instar or later in development). Dissection was performed on each grasshopper by removing the entire gut, including contents, from the crop to the hindgut using instruments sterilized in 95% (v/v) lab-grade ethanol between each dissection.

DNA Extraction

Each grasshopper gut sample and its contents were first homogenized into smaller pieces inside sterile microcentrifuge tubes using scissors sterilized by 95% lab-grade ethanol and flaming. DNA was extracted using the Qiagen DNeasy Blood and Tissue Kit

(Qiagen 69504) following the manufacturer’s protocol with some modifications. Briefly, the length of sample digestion time was increased to 60 minutes in a 57°C water bath, followed by mechanical lysis by bead beating in a Fisherbrand Bead Mill 4 Homogenizer

(Fisher Scientific 15-340-152) using 0.5 mm glass beads (Fisher Scientific 15-340-152) for 5 rounds of 60 seconds each at 5 m/s to assure lysis of bacterial cell wall. The bead tubes were then centrifuged for 1 minute at 8,000 rpm to separate lysate from debris. The lysate was placed into a DNeasy collection column for further purification following the manufacturer’s protocol. Following extraction, the concentration of total DNA in each sample was measured by a nanophotometer (Implen, Denville Scientific Inc.). The extracted DNA was stored at -20°C until sequencing.

16S rRNA Sequencing

High throughput DNA sequencing was performed by Zymo Research (Irvine,

CA). DNA from specimens was quantified using a NanoDrop apparatus as well as the

2100 Bioanalyzer system (Agilent) prior to library construction. The sequencing library was prepared by PCR reactions by Zymo Research in real-time PCR machines to control cycles and therefore prevent PCR chimera formation. The V3-V4 region of the 16S rRNA gene was amplified using the Quick-16S Primer Set V3-V4™ (Zymo Research,

Irvine, CA). The primers used included coverage of both bacteria and archaea. The final

PCR products were quantified with qPCR fluorescence readings and pooled together based on equal molarity. The final pooled library was cleaned up with Select-a-Size DNA

Clean & Concentrator™ (Zymo Research, Irvine, CA), then quantified with

TapeStation® and Qubit®. DNA libraries from grasshopper gut samples were sequenced on the Illumina HiSeq2500 platform in “Rapid Run” mode with a v3 reagent kit (600

12 cycles), using 100 bp paired end sequencing, with an average of 10.2 million reads per sample. Samples were collected at various cycles (in 4 cycle intervals) until sufficient amplification had occurred, or at 42 cycles if no amplification occurred (Zymo Research,

Irvine, CA). The sequencing was performed with >10% PhiX spike-in (23–25). DADA2 software, identifying single-nucleotide differences among sequences, was used in concert with the Greengenes database (gg_13_8) to assign bacterial taxa.

Statistical Analysis

Statistical analysis was performed in R (version 3.5.0). After checks for normality indicated non-normal distribution, non-parametric Kruskal-Wallis tests were used for significance calculations. Richness, Shannon diversity, and inverse Simpson diversity calculations were done using the vegan package in R (26). Dominant phyla and families were those that contributed a mean relative abundance of over 1.5% of the sequences for at least one grasshopper species within the taxonomic level being examined. Non-metric multidimensional scaling (NMDS) was performed using the vegan package in R. NMDS plots were used if the model arrived at convergence. The labdsv package in R was used to calculate indicator groups for each grasshopper species using the indval function in R.

Analysis of similarity (ANOSIM) to measure Bray-Curtis dissimilarity, corresponding with the NMDS plots was also performed using the vegan in R.

Results

A total of sixty grasshoppers (Table 1) were sweep-net harvested from the Texas tallgrass coastal prairie, including six different species: Conocephalus fasciatus (n=3),

Conocephalus strictus (n=8), Orchelimum concinnum (n=9), Orchelimum vulgare

(n=32), Paroxya atlantica (n=4), and Scudderia texensis (n=4). The differing numbers of

13 individuals from each species represent the relative abundance of these grasshopper species in the field. While no archaea were detected, a total of 293 bacterial taxa were obtained from the 60 gut samples. Mean species richness for all six grasshopper species was 27.22 (Table 1). Notably, we found that all six grasshopper species overlapped in their dominant bacterial phyla (Fig 1). However, depending on the grasshopper species, there were shifts in prevalence from one phylum to another. The overall composition of the grasshopper gut bacterial community at the phylum level consisted mainly of

Proteobacteria, Actinobacteria, Firmicutes, and to a lesser degree, Tenericutes (Fig 1).

The most prevalent families included: Listeriaceae (n=51 samples),

Lactobacillaceae (n=45), Methylobacteriaceae (n=47), Rhizobiaceae (n=50),

Sphingomonadaceae (n=48), Enterobacteriaceae (n=55), and Pseudomonadaceae (n=59)

(Fig 2). High abundance sequences also included unassigned families from the order

Entomoplasmatales and class Mollicutes. Analysis of similarity at the level of microbial family indicated a high degree of similarity between the microbiomes of the six species of grasshoppers (Bray-Curtis, R = 0.241, p = 0.004), implying a potential “core” group of microbes that compose the gut microbiome of these insects (Fig 3).

The most notable indicator bacterial group for the six grasshopper species analyzed was the phylum Tenericutes, which was present at relatively low levels in most grasshopper species but clearly dominated the microbiome of P. atlantica (Fig 1).

Indicator group analysis generated a strong indicator value (0.7126, p = 0.006) for

Tenericutes in P. atlantica samples. Non-metric multidimensional scaling (NMDS) performed at the family level (Fig. 3) revealed a clear separation between the bacterial communities in P. atlantica and those in other grasshopper species. The NMDS analysis

14 also shows that despite the shifts in phylum and family prevalence for the other species of grasshopper (Fig 1, 2), there is a clear overlap in a “core” or characteristic microbiome.

When males and females of all species were pooled together and compared, there was a significant difference in alpha diversity between males and females with males having higher richness and significantly higher Shannon and inverse Simpson diversity

(Fig 4a, b; non-parametric Kruskal-Wallis test, p = 0.031 and p = 0.038, respectively).

NMDS of the same data showed that although male and female microbiomes overlapped for the most part, female microbiomes were less closely clustered to one another than male microbiomes (Fig 4c). There were strong indicator values for male association with

Listeriaceae (indicator value = 0.6853, p = 0.017) and Staphylococcaceae (indicator value

= 0.5326, p = 0.008). These results highlight a previously unrecognized distinction between male and female grasshopper gut microbiomes across different species.

Discussion

We found that grasshopper guts play host to diverse communities of bacteria. A total of 293 bacterial taxa at the species or strain level were present, with the conservation of Proteobacteria, Actinobacteria, and Firmicutes phyla in all the six grasshopper species.

The highest number of taxa present per gut sample in our study was 65, with an average of 26.82 and a median of 27. Although these six grasshopper species do not have the same gut microbiome diversity as mammals, they do have bacterial families similar to other insects. For example, Lactobacilliaceae and Enterobacteriaceae families were also found in the guts of Mormon crickets, another orthopteran (19). Actinobacteria,

Firmicutes, and Proteobacteria have all been found in locusts, also orthopterans, (27) and in the well-characterized microbiota of honeybees (15,16,28). The six species of

15 grasshoppers studied here also overlapped abundant phyla with neotropical butterflies whose dominant groups included Firmicutes, Proteobacteria, and Tenericutes (29). In contrast, the microbial communities in humans and other mammals tend to be dominated by Bacteroidetes and Firmicutes (30).

The composition of bacterial communities associated with a host organism can be attributed to both an organism’s ecology and evolution (31,32). Differences in the bacterial community structure among different organisms, therefore, may depend on host species behavior, modes of bacterial community transmission, timing in the lifecycle of the host, evolution, or changes to the environment (33,34). By analyzing the gut microbiome of 6 different grasshopper species, we have a novel opportunity to examine the correlation between the bacterial communities and the ecology and evolution of their host organisms. The distinctive gut microbiome in P. atlantica in comparison to other five grasshopper species can be rationalized by the organism’s phylogenetic separation as well as its different dietary preferences. Although commonly referred to as long-horned grasshoppers, C. fasciatus, C. strictus, O. concinnum, and O. vulgare, are all meadow katydids from the subfamily Conecephalinae. S. texensis is also a katydid of the subfamily Phaneropterinae, a closely related group to the Conecephalinae (35). P. atlantica, on the other hand, is in the subfamily Cyrtacanthacridinae, a short-horned grasshopper. While long-horned grasshoppers look very similar to true grasshoppers, they are more closely related to crickets. Also, the dietary tendency for katydids is omnivory, while short-horned grasshoppers, like P. atlantica, tend to eat almost exclusively plant matter. The structure of the tooth mandible in P. atlantica suggests that this species specializes on forbs, but a tendency to eat grasses has also been experimentally verified

(36). Therefore, the distinctive bacterial communities associated with P. atlantica, in comparison to the other five species, can be supported by the phylogenetic relationship or the dietary preferences. A wider investigation into the gut microbiome in additional grasshopper species in the future could help establish whether there are evolutionary associations between the phylogeny of different grasshopper species and their microbial symbionts or are there simply differences in dietary preferences that shape the grasshopper microbiome.

It is interesting to note that the shift in bacterial prevalence for P. atlantica is towards the phylum Tenericutes, which has a high indicator value when in association with P. atlantica. Tenericutes are a phylum of bacteria with small genomes and no cell wall (37). They are known to have symbiotic relationships with arthropods as well as other animals and plants [37–40]. Within the phylum Tenericutes is the class Mollicutes, which includes the order Entomoplasmatales that houses many species which have widely reported associations with insects, including spiroplasmas (38,39). Although the bacterial taxa associated with the Mollicutes did not resolve to specific families in our sequence analysis, multiple unique, unclassified Entomoplasmatales family reads were present, indicating the possibility of spiroplasmas in the grasshopper gut. It remains to be seen whether there is a significant host-microbe relationship for P. atlantica and a specific spiroplasma species. In some cases, spiroplasmas are harmful, disease-causing pathogens, and in other cases, vertically-transferred symbionts (38). The absence of spiroplasmas in pea-aphids affects host fitness by decreasing offspring production and survival (39). Other known Entomoplasmatales-insect associations include Dabulus maidis leafhoppers, Heliconius butterflies, New World army ants, and Drosophila hydei,

17 which have been shown to have nonpathogenic entomoplasmatales in their gut that may be linked to increased host survival and fitness (38-43). Finally, in mammals, mice fed a high-fat diet have decreased numbers of Tenericutes in their gut, suggesting that there is a possible connection between diet in mammals and levels of Tenericutes (44). At this point, the relationship of P. atlantica with Entomoplasmales in particular, or Tenericutes in general, is unknown. It is also not known whether the relationship is evolutionary or dietary in origin. Both rationales are possible given both P. atlantica’s phylogenetic distance from the other grasshoppers in this study and its difference in trophic ecology

(45).

Male grasshoppers in our study showed a significantly higher alpha diversity (Fig

4a, b). Because the prevalence of the main phyla does not shift between male and female

(Fig 4c), it is possible that the differences between male and female grasshopper gut microbiota could be attributed to rare taxa. This is true for some species of butterflies, who despite having no significant differences in alpha diversity between males and females, still have different rare taxa (43). However, given that the overlap between male and female grasshopper microbiome is not complete, it is also possible that there are subtle shifts in main taxa groups that account for the difference in alpha diversity between males and females. Functionally, it is unclear what drives the sexual dimorphism in the grasshopper microbiome, but potential roles have been reported in other animals.

In mice, the gut microbiome is integral to the sexual dimorphism in the regulation of hormones for growth and sexual maturity, as well as improving disease resistance

(46,47). Female Drosophila melanogaster mating patterns are correlated with male microbiota so that females mate preferentially with males with intact microbiota (48). In

18 the moth, Spodoptera littoralis, adult female microbiomes contain a higher number of genes for energy metabolism than male microbiomes (5). Future studies that further dissect potential sexual dimorphism in the insect gut microbiome will likely reveal additional functional relationships between insects and their resident microbes.

All the prevalent families found in the grasshopper guts (Fig 2) appear to have members with well-known associations with soils and plants that may contribute to their associations with grasshoppers. Understandably, the presence of these taxa, as established by DNA sequencing, is not a direct indication of functions or permanent residency.

Nevertheless, it is of interest to note that some of these bacterial families may have a functional role to play in the gut, while others may be plant or insect pathogens for which the grasshopper may serve as a reservoir or vector (4,49). For example,

Enterobacteriaceae family was found in almost every gut sample analyzed. Insect symbionts from this family have been found to counteract plant defenses and insecticides

(50). Erwinia, which has been identified as a probable symbiont of western flower thrips

(51), was also present in our samples. Another family with relatively high abundance was

Lactobacilliaceae, which has been reported extensively in honeybees. The microbiome composition in honey bees appear to correlate with behavioral tasks such as foraging, caring for young, or food processing—with Lactobacillus mellis present at a significantly higher level in bees performing nursing or processing food than bees performing foraging tasks (52,53). The bee-associated Lactobacillus species (Family Lactobacillaceae) are known to be phylogenetically and metabolically distinct from human-associated

Lactobacillus species (54). Similarly, Bifidobacterium (Family Bifidobacteriaceae) associated with humans and monkeys also possess higher levels of glycosyl hydrolases

19 than those from insects or other animals and environment (54). These observations, while supporting the metabolic roles of gut microbiome in host behavior and nutrient acquisition, also highlight the need to better understand the evolutionary history behind the association of animals, from grasshoppers to mammals, with these common bacterial families. It is also worth noting the presence of potential plant and animal pathogens within the grasshopper gut microbiome. This fact raises questions on the role of grasshoppers in disease transmission. Families Pseudomonadaceae and

Xanthomonadaceae both have members that are highly critical plant pathogens (55).

Family Methylobacteriaceae, present in 75 percent of grasshoppers sampled, includes

Methylobacterium species that are typically found in soil and water that may be transiently associated with grasshoppers. However, despite its typical free-living lifestyle, one species, Methylobacterium mesophilicum, has a demonstrated ability to stably colonize Catharanthus roseus, or Madagascar periwinkle, through the transmission by the insect Bucephalogonia xanthophis (Hemiptera: Cicadellidae) (56). Most known plant pathogens are transmitted by Hemipteran insects (57); however, if grasshoppers prove to be carriers of plant pathogens, even transiently, they may be able to contribute to disease transmission and undermine efforts to protect crops by sustaining the pathogens in the environment.

The relatively high prevalence of the family Listeriaceae, especially among male grasshoppers (indicator value = 0.6853, p = 0.017), observed in this study has not been routinely acknowledged in any insects. The family Listeriaceae includes two genera:

Listeria and Brochothrix (58), of which several species of Listeria are known human and animal pathogens with others containing pathogenicity-associated genes (59). Therefore,

20 for the interest of food safety, the presence of Listeria in insects, especially potentially edible insects, has been an ongoing investigation. In numerous other surveys of insects, including bush crickets (Ruspolia diffferens), lesser mealworms (Alphitobius diaperinus), tropical house crickets (Gryllodes sigillatus), field crickets (Gryllus bimaculatus), superworms (Zophobas atratus), and cockroaches, no Listeria has been found (60–64).

This absence of Listeria in insects, despite the prevalence of Listeria in natural environments (65), might result from Listeria acting as an insect pathogen. Multiple insects, including greater wax moth (Galleria mellonella), 2-spotted cricket (Gryllus bimaculatus), house fly (Musca domestica), and common fruit fly (Drosophila melanogaster), have all been suggested as host models to study Listeria infections (65–

68). Curiously, Listeria has been cultured from ants collected from Pamplemousses,

Mauritius, as well as from citrus black flies (Aleurocanthus woglumi) (69,70), suggesting that some insects, perhaps grasshoppers included, are capable of establishing a symbiotic relationship with Listeria species. Whether grasshoppers from a coastal prairie system serve as an environmental reservoir for potential human pathogens remains to be determined. However, there might be an association between Listeria and certain insects that contribute to the ubiquity of Listeria in the environment.

Conclusion

In this study, we demonstrated the conservation of three phylum, Actinobacteria,

Proteobacteria, and Firmicutes across six different species of grasshoppers. A fourth bacterial phylum, Tenericutes, is present in four of the six grasshopper species and dominates the microbiome of the short-horned grasshopper herbivore, P. atlantica. This core microbiome overlaps with the microbiome found in many other insect herbivores.

Differences in the bacterial community compositions in the six species of grasshoppers may be influenced by evolutionary mechanisms or dietary preferences. While there are consistent bacterial groups across all six species, the function of these host microbiomes and their influence on the ecosystem remains an open question. A deeper understanding of these important host-microbiome interactions will provide insight into both host ecology and bacterial functions that may ultimately lead to better conservation of these animals and their habitats and more safe and effective pest control measures.

Acknowledgements

The authors would like to acknowledge Dr. Julia Chapman, Amanda Finke, and

Michaela Wood for assistance with statistical analysis, Dr. Angela Laws for help with field work, and Dr. Sumant Grover for spectrophotometry assistance.

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CHAPTER 2

EFFECTS OF ENVIRONMENTAL NUTRIENTS ON BACTERIAL AND FUNGAL

MICROBIOMES IN THE COMMON MEADOW KATYDID, ORCHELIMUM

VULGARE

Abstract

Microbiomes consist of bacteria, fungi, and viruses that can act at mutualists to influence the health and fitness of their insect hosts, however, the factors that determine the constituents and diversity of these microbial communities is poorly understood. The limitation of essential nutrients can influence the host abundance. However, the effect of abiotic factors like environmental nutrients on most insect microbiomes is yet unknown.

Here we examine the relationship between environmental nutrients and the gut microbial communities in individuals (n=13) of the orthopteran species Orchelimum vulgare collected from the site of a fully-factorial fertilization experiment. We found significantly higher Shannon diversity for the gut bacterial communities in grasshoppers from plots fertilized with sodium compared to those from plots without sodium. In contrast, diversity decreased in the gut fungal communities of grasshoppers collected from plots amended with nitrogen and phosphorus or sodium. Nevertheless, there was also a strong positive correlation between the gut bacterial and gut fungal community diversity within each sample. Upon further analysis, indicator groups for sodium-treated plots included a

32 number of salt-tolerant bacterial and fungal taxa. Together these results highlight a potential connection between the bacterial and fungal communities inside grasshoppers and environmental nutrients conditions experienced by the host. While it is unclear whether the insect gut microbiome is experiencing similar nutrient conditions as its host, the gut microbial constituents are responsive to changes in environmental nutrient levels.

Taking into account abiotic factors like soil nutrients along with characteristic gut microbial groups is necessary for better understanding and conservation of this important insect herbivore.

Introduction

Recently, the rise in the research of insect microbiomes has been contributing to a better understanding of the ecology of insect hosts. Both resident and transient microbes can affect the health and fitness of their insect hosts including (1) improved nutrition for the host (2,3), increased host reproductive success (4), host protection from environmental pathogens and pesticides (3), and changes to host behaviors (5).

Conversely, the insect host’s environment and behaviors like feeding and social roles can be reflected in changes in the host microbiome (6–9). In the case of orthopterans, we have demonstrated that orthopterans share a characteristic microbiome dominated by

Proteobacteria, Firmicutes, and Actinobacteria. However, how changes to the environmental conditions of the host will affect these host microbiomes remains to be determined.

In general, the addition of nitrogen and phosphorus to the soil will result in increases in plant biomass, but declines in plant richness and diversity (10–12). The success of fertilizing treatments like nitrogen and phosphorus in increasing plant growth is explained in part by Leibig’s law of the minimum which proposes that plant biomass will be limited by the essential nutrient in shortest supply. In other words, if a nutrient limiting plant biomass is added to the soil, plant biomass will increase (13). Once there is no longer limitation of an essential nutrient like nitrogen, fast-growing, nitrophilic plant species can increase in abundance to the exclusion of other slower growing species (14).

Consequential impacts of nitrogen and phosphorus fertilization on organisms at higher trophic levels are also evident. For example, laboratory feeding trials found that grasshoppers experienced both nitrogen and phosphorus limitation (15) and a field study found that fertilization of grasses with ammonium nitrate increased the abundance of grasshoppers (16).

In addition to the evidence that nutrients like nitrogen and phosphorus impact insect host abundance, there are also indications that sodium may be a limiting nutrient for insect herbivores like O. vulgare (17). Sodium is a nutrient present in small amounts in plant tissues. While typically considered non-essential for most plants (18), sodium is essential for animal physiology, controlling the osmolarity of body fluids (19), and regulating of growth and reproduction. Evolutionarily speaking, sodium ion channels appeared even before the nervous system itself and are crucial for its proper function

(20). Animals, including insects, have been known to satisfy their needs for sodium through activities outside of the normal feeding behaviors (21,22). Therefore, with limited sodium present in their food source, grassland plant consumers like O. vulgare,

34 likely experience sodium limitation and may exhibit sodium-seeking behaviors (23,24).

Insect abundance and diversity has been shown to increase in areas where sodium has been added to the soil. Other grasshoppers collected from the same fully-factorial fertilization experiment showed an increase in abundance when they came from plots with nitrogen and phosphorus and sodium added, suggesting colimitation (17). Sodium can be added to the soil in the form of chemical treatments like ice-melting road salts or can be carried into an area by ocean winds and deposited in the soil where it has subsequent ecosystem impacts (25,26).

Insect herbivores, play pivotal roles in many ecosystems, including the grassland ecosystems of North America where O. vulgare is commonly found (27,28). O. vulgare is a key consumer of plant biomass as well as a key food source for predators thus contributing to nutrient cycling and plant community composition in grasslands (29).

Though primarily feeding on plants, O. vulgare has been observed feeding on carcasses of insects and other small animals (30,31). It is a common orthopteran species of both economic and ecological significance. Our earlier work has identified a “core” bacterial community in the gut of O. vulgare. However, how the microbiome of O. vulgare responds to environmental nutrient conditions has not been determined.

In this study we aim to better understand the impact of environmental nutrient conditions on the gut microbiome of O. vulgare by analyzing the bacterial and fungal communities of the gut in individuals from large plots in a tallgrass prairie. These plots had been amended with two different two-level fertilizer treatments including added nitrogen and phosphorus, added sodium, as well as ambient nitrogen and phosphorus, and ambient sodium. If the gut microbiomes responded to these nutrient treatments, we would

35 expect to see a significant decrease in microbial diversity in individuals collected from added nitrogen and phosphorus plots. If sodium is a limiting nutrient for the gut community, we would also expect to see a similar pattern of decreased gut microbe diversity in added sodium plots. Here, we did not see the same pattern with added sodium plots.

Methods

Sample Collection

We determined how nutrients affected the microbiomes of O. vulgare by collecting individuals from a large-scale fertilization experiment at the University of

Houston’s Coastal Center near Houston, Texas. Many orthopteran species have been documented in this prairie, of which O. vulgare is one of the most common (17). The prairie topography is generally flat and experimental plots were separated by at most a 2 cm gradient in elevation. This experiment was part of a larger experiment looking at the effect of different nutrients on orthopteran communities. We collected individuals via sweep netting from plots of a fully factorial design manipulating nitrogen and phosphorus

(N and P together at two levels, ambient and fertilized) and sodium (Na at two different levels, ambient or fertilized) with eight replicates in each treatment (n=2 levels of NP x 2 different levels of Na x 8 replicates = 32 experimental plots). The prairie was divided into large plots (30 x 30 m2) which were subsequently treated with both macro and micronutrient fertilizers. Fertilizers were applied in March of 2016 and 2017 before the beginning of the growing season. Nitrogen and phosphorus, and sodium were added at rates of 10g/m2 to bring the top 10 cm of soil to approximately 30% higher than ambient levels (14,16). We collected O. vulgare individuals, one of the most common

36 orthopterans at this field site, from as many replicates as possible via sweep-netting during one-day of June 2017. In total we found 13 O. vulgare individuals: 3 individuals were from plots with no added nutrients (None), 4 were from plots treated only with sodium (Na), 3 were from plots with added nitrogen and phosphorus (NP), and 3 were from plots treated with both added nitrogen and phosphorus and sodium (NP x Na). The insect samples were shipped on ice to the University of Dayton (Dayton, OH) and stored frozen at -20°C until dissection. Only O. vulgare individuals that could be easily dissected (individuals at 4th instar or later in development) were included in this study.

Dissection consisted in removal of the entire gut, including contents, from the crop to the hindgut. Instruments were sterilized in 95% (v/v) lab-grade ethanol between each dissection. Gut samples were stored at -20oC until DNA extraction.

DNA extraction

Each frozen O. vulgare gut sample with its contents was homogenized into smaller pieces inside sterile microcentrifuge tubes using scissors sterilized by 95% lab- grade ethanol and flaming before further processing. DNA was extracted using the

Qiagen DNeasy Blood and Tissue Kit (Qiagen 69504.) Manufacturer’s protocol was adhered to with some modifications. Modifications included increasing the length of sample digestion time to 60 minutes in a 57°C water bath, as well as introducing a mechanical lysis step by bead beating in a Fisherbrand Bead Mill 4 Homogenizer (Fisher

Scientific 15-340-152) using 0.5 mm glass beads (Fisher Scientific 15-340-152) for 5 rounds of 60 seconds each at 5 m/s. This step was included to assure lysis of bacterial cell wall and release of DNA. The bead tubes were then centrifuged for 1 minute at 8,000 rpm to assure separation of lysate from debris. The lysate was aliquoted into a DNeasy

37 collection column for purification of the DNA following the manufacturer’s protocol.

The concentration of total DNA in each sample was measured by a nanophotometer

(Implen, Denville Scientific Inc.). The extracted DNA was stored at -20°C until sequencing.

16s rRNA sequencing

High throughput DNA sequencing was performed by Zymo Research (Irvine,

CA). DNA from the O. vulgare gut samples was quantified prior to library construction using both a NanoDrop apparatus and also 2100 Bioanalyzer system (Agilent). The sequencing library was constructed by PCR reactions performed in real-time PCR machines to control cycles which prevented PCR chimera formation. Bacterial and archaeal sequencing focused on the V3-V4 region of the 16S rRNA gene which was amplified using the Quick-16S Primer Set V3-V4™ (Zymo Research, Irvine, CA).

Fungal sequencing focused on the ITS2 region using the ZymoBIOMICS Services ITS2

Primer Set. (Zymo Research, Irvine CA). The final PCR products were quantified with qPCR fluorescence readings and, based on equal molarity, subsequently pooled. The final library was cleaned with Select-a-Size DNA Clean & Concentrator™ (Zymo Research,

Irvine, CA), followed up by quantification with TapeStation® and Qubit®. DNA libraries from grasshopper gut samples were sequenced on the Illumina HiSeq2500 platform in “Rapid Run” mode with a v3 reagent kit (600 cycles).100 bp paired end sequencing was used with an average of 10.2 million reads per sample. Samples were collected at various cycles (in 4 cycle intervals) until sufficient amplification had occurred. If no amplification occurred, samples were collected at 42 cycles (Zymo

Research, Irvine, CA). The sequencing was performed with >10% PhiX spike-in [23–25].

DADA2 software. Single-nucleotide differences were identified among sequences and were used in concert with the Greengenes (gg_13_8) database to assign bacterial taxa.

Data Analysis

Statistical analysis was performed in R (version 3.5.0). Diversity calculations included Shannon and Inverse Simpson diversity as well as richness. Calculations of diversity indices were performed using the vegan package in R. To test for differences between treatments, we used ANOVA with 2 independent variables (NP or Na) at 2 levels (ambient and added) To look at effect sizes, we calculated Cohen’s d using the cohen.d function in the effsize package in R. Regression comparing bacterial and fungal diversity was carried out in base R. Non-metric multidimensional scaling (NMDS) was performed using the vegan package in R. NMDS plots were used if the model arrived at convergence. Analysis of similarity (ANOSIM) to measure Bray-Curtis dissimilarity, corresponding with the NMDS plots was also performed using R. Indicator values were calculated using the indval function in the labdsv package in R. All indicator values specified had a p value of less than 0.05.

Results

A total of 270 species-level operational taxonomic units or OTUs were detected in the 13 samples (Table 2). No archaea were detected. Of the 270 OTUs, 171 were bacterial with a mean bacterial richness of 31.6 (±3.9). There was overlap between the bacterial phyla present in all treatments (None, NP, NP x Na, and Na), including

Proteobacteria, which were predominant among all treatment groups (Fig 5a), Firmicutes, and Actinobacteria. One of the most notable shifts among bacterial phyla was the predominance of Tenericutes in the NP treatment group. There were eleven bacterial

39 families present at an average relative abundance of 2% or higher in all treatment groups

(Fig 6a) with the most abundant families being: Enterobacteriaceae, Lactobacillaceae,

Listeriaceae, Methylobacteriaceae, Pseudomonadaceae, and Rhizobacteriaceae.

Uncategorized family OTUs from the order Entomoplasmatales were present in four out of the 13 samples, representing three of the four possible treatment groups. They comprise a large proportion of the NP treatment group families. They are not relatively abundant in either the Na or NP x Na treatments. Of the bacterial families present in the gut bacterial communities, five are clear indicators of added Na communities, including

Streptococcaceae, Propionibacteriaceae, Phyllobacteriaceae, Listeriaceae, and

Corynebacteriaceae (indicator values respectively: 0.9979, 0.9801, 0.9627, 0.8745,

0.7094). Only one of these, Listeriaceae, is found at high relative abundance. In contrast, only one family, Sphingomonadaceae, was an indicator of added NP (indicator value,

0.7986).

The other 99 OTUs were fungal, with a mean fungal richness of 25.4 (±3.3). The two most dominant fungal phyla in all four treatments were Ascomycota and

Basidiomycota, though a relatively high number of fungal OTUs in the NP treatment group were unassigned (Fig 5b). Fungal data had a higher number of specified OTUs at the order level of taxonomic classification (Fig 6b). Again, there are a high number of unspecified fungal OTUs for the NP treatment group even at order level. There were six predominant fungal orders that appeared in every treatment group: Capnodiales,

Dothideales, Pleosporales, Sporidiobolales, Tremallales, and Ustilaginales. An unspecified species of Cladosporium was indicative of added Na gut communities ( indicator value = 0.7806).

Alpha diversity was assessed in terms of taxa richness, Shannon Index and

Inverse Simpson Index (Table 2). Shannon diversity of bacterial taxa assessed at the species level (Fig 7a) was significantly higher for gut samples from added Na plots (p =

0.014) with a large treatment effect size (Cohen’s d = 1.04), and with some, but not significant, interaction between added Na and added NP groups (p = 0.08). Shannon diversity of fungal species (Fig 7b) was significantly lower for gut samples from the added Na added plots (p = 0.035) and also for added NP (0.006), with a large negative treatment effect (Cohen’s d = -1.12 and -1.28 respectively). Notable, for fungal community diversity, there was significant interaction between added Na and added NP

(p = 0.004). More specifically, interaction between NP and Na in the NP x Na treatment group rescued the decrease in diversity present in the NP treatment group to the same level as Na alone. Regression analysis (Fig 7c) of bacterial species diversity versus fungal species diversity showed a strong positive correlation between the two indices (R2 =

0.75).

Beta diversity was assessed using non-metric multidimensional scaling (NMDS) performed at the species level for both bacteria (Fig 8a) and for fungi (Fig 8b). There appeared to be an overlap between the characteristic community of the control group with no added nutrients (None) and the two added Na groups (Na and NP x Na). The characteristic communities of the NP only treatments did not have as much overlap with the characteristic communities of the other three treatments (ANOSIM, R = 0.299). This pattern was observed for fungal communities as well. (ANOSIM, R = 0.335). Beta diversity was further characterized by assessment of shared species level OTUs between treatment groups, revealing that 60% of bacterial species level OTUs and 51.5% of

41 fungal species level OTUs were unique to each treatment group (Fig 9). Conversely, only

5.8% of bacterial OTUs and 18% of all fungal OTUs were shared amongst all four treatment groups. In bacterial treatment groups with added sodium, the numbers of unique bacterial OTUs were higher overall, with 46.5% (NP x Na) and 40.7% (Na). The

NP treatment group samples yielded far fewer unique bacterial groups with only 10.8% unique bacterial OTUs. This contrasted with fungal OTUs which with had the highest number of unique OTUs in the treatment that received no added nutrients (39.4%).

Discussion

The results from this field study indicate that environmental nutrients can have a significant effect on host microbiome. While gut bacterial community diversity and gut fungal community diversity throughout all treatments had a strong positive correlation, there are differences between the effects of added NP and added Na on community composition. As expected, in the case of limitation by NP, the gut fungal microbiome community diversity decreases with addition of NP. This pattern of decreased diversity is also observed in gut fungal communities from plots with added Na, indicating that gut fungal microbiomes in O. vulgare may be limited by both NP and Na. In contrast, while gut bacterial community samples from plots with added NP were less diverse overall and still trend downward, the effect was not significant. Furthermore, the gut bacterial community diversity actually increased in hosts collected from added Na plots. This indicates that although diversity may be depressed in the bacterial community when NP is added, bacterial community diversity is not limited by Na. Instead, the bacterial communities from Na added plots are experiencing effects that are not the direct result of nutrient limitation. Also, in the NP x Na plots, the level of diversity is much higher than

42 in the plots with NP alone, suggesting that the effect of added Na is overriding the effect of NP.

As mentioned previously, adding atmospheric nitrogen to the soil is known to decrease the diversity of plants, as well as animals in the affected area (14,16). Added NP has also be shown to contribute both directly and indirectly to a decrease in microbial richness and shifts in microbial communities in fertilized soils (30,31). Other research also indicates that there are also shifts in community functional traits in added NP soil microbial communities that could affect nutrient cycling (32). The decreases in diversity in our added NP communities demonstrates that NP can limit host fungal communities and, to a lesser extent, host bacterial communities. However, whether or not this is a direct or indirect relationship is difficult to determine. Soil bacterial and fungal communities response directly to added NP with increased growth and shifts in community composition (33,34). In our study, the only indicator group for added NP was

Sphingomonadaceae, a common soil and plant associate. We also observed shifts in the bacterial community from relatively high abundances of Proteobacteria and Firmicutes to a predominance of Tenericutes. These shifts may be the direct result of added NP in the gut microbiome or the indirect result of shifting plant and soil microbiomes. Also, changes in plant community structure due to added NP that result in changes to insect feeding behaviors might also trigger changes in host gut microbiome.

The significant decrease in diversity observed in the added Na fungal communities is evidence that Na may be also be a limiting nutrient for gut fungal communities. However, increased diversity was detected in bacterial communities suggesting that these bacterial communities are not experiencing Na limitation, but some

43 other effect of added Na. Increased salinity does effect both microbial and fungal community composition and can lead to increases in richness and diversity in some soil communities, though salinity that is too high has also been shown to decrease fungal diversity in estuary soils (35,36). Salinity has also been directly linked to changes in fitness and abundance of soybean aphids and oviposition choices in tiger beetles (37,38).

Therefore, it is possible that the changes to community composition are both a direct effect of added Na, perhaps as a stressor to the microbial communities that selects for more halophilic bacteria and fungus, and an indirect effect of changing insect behaviors.

There are a number of unique taxa observed in added Na gut bacterial communities.

While these unique groups may be the result of changes to plant communities or feeding behaviors, they may also indicate stress associated with increased amounts of sodium.

Our added Na communities consist of a number of indicator groups. One of these indicator groups for added Na is Corynebacterium. This genus has been associated with increased soil salinity (45). Phyllobacteriaceae and Streptococcaceae were also indicator groups for added Na and are bacterial families previously associated with high saline soil microbiomes (46,47). Listeraceae and Propionibacteriaceae, also both indicator groups for added Na, are also families with species that are capable of growth in relatively high concentrations of NaCl (48,49). These indicator groups may represent bacterial community restructuring due to the stress of increased salinity. Again, whether this community restructuring is happening in the gut, in the soil, or on plants is difficult to tell.

Our previous work characterizing the O. vulgare microbiome has indicated that

O. vulgare has characteristic bacterial community associations which may depend on diet

44 or other environmental factors (39). While almost nothing is known about the mode of transmission for bacterial communities in O. vulgare, it can be assumed that some of these resident bacteria come from diet, environmental contact and feeding behaviors

(40,41). A large study with over 200 different insect species has indicated that gut bacterial communities among omnivorous insects are more diverse in general than gut bacterial communities in insects which are solely herbivorous or carnivorous (6). This has considerable implications for the omnivorous O. vulgare. It follows that a shift in host feeding behaviors, perhaps from fewer plants to more scavenging behaviors or vice versa, may correlate with a shift in the host microbiome. This does not necessarily mean that the host microbes are delivered solely by the plants or animals that these insects feed on. It is possible that many of the microbes in the grasshopper gut community may come directly or indirectly from the soil and be reflective of those microbial communities.

Changes to the soil microbiome may also result in changes to insect host microbiome; In feeding trials, caterpillars who fed on intact dandelion plants had a microbiome more closely linked in composition to the soil microbiome than the phytobiome of the dandelion (42). While prior research indicates that both soil and plant microbiomes can be affected by changes in environmental nutrients like NP and Na (33,36), it is also possible that the changes to gut microbiome are the direct result of changing nutrient levels inside the grasshopper gut. (43,44).

Historically, research into the microbiome of insects has been biased towards bacterial community characterization, rather than characterization of the microbiome as a whole. This one-kingdom approach to learning about insect microbiomes has inevitably lead to gaps in understanding (48). Also, the characterization of the host-fungal

45 microbiome relationship is largely informed by the studies of insect fungal pathogens and insects as vectors for plant pathogens. Characterization of the fungal microbiome of insects like O. vulgare helps us to begin to gain a more comprehensive perception of these host-fungal communities. The O. vulgare gut samples in this study yielded 99 species level OTUs. However, taxonomic resolution is still relatively low for fungal communities, and a number of these OTUs were unspecified or uncategorized. This in part due to the lack of well-established databases of ITS sequences, and in part due to the convolutions of fungal taxonomy (49,50). In our samples, there were a higher number of shared fungal OTUs than unique fungal OTUs. Some of these shared groups, like the order Capnodiales, have established associations with insects like scale and aphids (51).

Other common taxa included the genera Cryptococcus and Hannaella, have been previously associated with plants (52). The one unique group distinctive as an indicator of added Na gut fungal communities were members of the genus Cladosporium, an osmotolerant mold (53).

Conclusion

Our results support the hypothesis that nutrient limitation can affect the host microbiome both directly and indirectly. We demonstrate significant limitation by NP and Na in the fungal microbiome of O. vulgare. The O. vulgare gut bacterial microbiome is also affected by added Na, however, the result of added Na is an increase in diversity that suggests the community is altered by other factors, perhaps as a result of stresses to the microbial community or changes in feeding behaviors of the host. Here we identified salt-tolerant genera Corynebacterium and Cladosporium, as well as families like

Streptococcaceae, Propionibacteriaceae, Phyllobacteriaceae, and Listeriaceae that may be an indication of insect microbiomes exposed directly or indirectly to increases in sodium.

Here we have demonstrated that abiotic factors, in this case, environmental shifts in soil nutrients like NP and Na, can alter the microbiome of O. vulgare. Our study has some inherent limitations given the small sample size for each treatment group, the artifacts of field study and collection, and the gaps in knowledge about insect microbiome structure and function. It is important to be conservative in the broader conclusions that we draw from this dataset. However, while these results may be considered preliminary in nature, they are nonetheless compelling. In the future, a better understanding of characteristic insect microbiomes under the influence of changing ecosystem conditions will help us to identify disruptions that may represent a threat to hosts like O. vulgare, or ecosystems like the tallgrass prairie.

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Figure 1: Mean relative abundance of bacterial phyla in the gut of six grasshopper species. Bacterial phyla, predicted from the amplified V3-V4 region of the bacterial 16s rRNA gene, were shown if they were present at >1.5% relative abundance in each grasshopper species. Phyla that were present at <1.5% relative abundance in each grasshopper species are grouped together as “Other.”

Figure 2: Mean relative abundance of prevalent bacterial families in the gut of six grasshopper species. The abbreviations for grasshopper species that appear on the x axis are as follows: cf, C. fasciatus; cs, C. strictus; oc, O. concinnum; ov, O. vulgare; pa, P. atlantica; and st, S. texensis. Bacterial families with mean relative abundance <1.5% across each grasshopper species are grouped together as “Other.”

Figure 3: Beta diversity comparison of bacterial communities. Nonmetric multidimensional scaling (stress = 0.084) of grasshopper gut community assemblages at the family level (n=60) was plotted using the vegan package in R. Dispersion ellipses are placed for each grasshopper species. Bray-Curtis dissimilarity was well-preserved in two dimensions. Analysis of Similarity (ANOSIM) was performed with a Bray-Curtis dissimilarity measure, and showed an overall significant difference in gut microbiota among grasshopper species (R = 0.241, p = 0.004).

Figure 4: Comparison of the gut microbiome in male and female grasshoppers. Shannon diversity (a) and inverse Simpson diversity (b) indices for each female or male individuals were calculated with the vegan package in R. Nonmetric multidimensional scaling with dispersion ellipses placed for male and female pooled species (c) was plotted with the vegan package in R. NMDS was constructed using Bray-Curtis dissimilarity. Analysis of Similarity (ANOSIM) was performed and did not show a significant difference in the characteristic gut microbiota of males and females (R = 0.004, p = 0.352).

Figure 5: Average relative abundance of microbial phyla in grasshopper samples from four treatments. The four treatments include: no added nutrients or “None” (n=3), added sodium or “Na” (n=4), added nitrogen and phosphorus or “NP” (n=3), and nitrogen and phosphorus added with sodium or “NP x Na” (n=3). The “Other” category is comprised of groups that do not appear in all four treatments and comprise less than 2% of reads. (a) Relative abundance of bacterial phyla. (b) Relative abundance of fungal phyla.

Figure 6: Average relative abundance of microbial taxa in grasshopper samples from four treatments. The four treatments include: no added nutrients or “None” (n=3), added sodium or “Na” (n=4), added nitrogen and phosphorus or “NP” (n=3), and nitrogen and phosphorus added with sodium or “NP x Na” (n=3). In all The “Other” category is comprised of groups that do not appear in all four treatments and comprise less than 2% of OTUs. The only exception is the Entomoplasmatales families included which were not present in the nitrogen and phosphorus with sodium treatment, but were present in the other three treatments. (a) Relative abundance of bacteria families. (b) Relative abundance of fungal orders.

Figure 7: Analysis of diversity over four treatments. (a) Bacterial species Shannon diversity for each treatment was calculated with the vegan package in R. Two-way ANOVA in base R was used to compare levels of diversity between treatments and test for significant difference. Treatment effect size, or Cohen’s d, as calculated in R with the effsize package for Na compared to control was large (1.04). Effect size for NP was smaller (-0.32). (b) Fungal species Shannon diversity for each treatment was also calculated in a similar manner. Cohen’s d effect size for Na as compared to control was large (-1.12), as was NP compared to control (-1.28). (c) Regression analysis performed in R indicates a significant positive correlation between bacterial Shannon diversity and Fungal Shannon diversity among all 13 samples (R2 = 0.75).

Figure 8: Nonmetric multidimensional scaling of grasshopper gut community assemblages at the family level (n=13) was plotted using the vegan package in R. Dispersion ellipses are placed for each treatment group. Bray-Curtis dissimilarity was well-preserved in two dimensions. (a) Bacterial characterization. Analysis of Similarity (ANOSIM) was performed with a Bray-Curtis dissimilarity measure and showed an overall significant difference in bacterial families among treatments (R = 0.299). (b) Fungal characterization. ANOSIM was performed with a Bray-Curtis dissimilarity measure and showed an overall significant difference in fungal families among treatments (R = 0.335).

Figure 9: An analysis of shared groups among the four treatments. (a) A Venn diagram of bacterial species richness. Total number of bacterial species OTUs = 171. (b) A Venn diagram of fungal species richness. Total number of fungal species OTUs = 99.

Table 1: Gut bacterial richness and diversity for six grasshopper species

Mean Mean Mean Species Sex n= Species Shannon Inverse Richness Diversity Simpson All 60 27.22 1.95 5.18 All samples Female 30 25.47 1.61 4.18 Male 30 28.50 2.10 6.37 All 3 19.67 2.23 10.52 Conecephalus Female 1 10.00 0.88 1.81 faciatus Male 2 24.50 2.90 14.88 All 8 23.88 1.19 5.46 Conecephalus Female 3 16.00 1.00 2.12 strictus Male 5 28.60 2.31 7.46 All 9 26.44 1.65 4.44 Orchelimum Female 4 24.40 1.51 4.69 concinnum Male 5 29.00 1.83 4.14 All 32 30.28 2.00 5.15 Orchelimum vulgare Female 16 31.13 1.86 4.64 Male 16 29.44 2.13 6.19 All 4 21.50 1.06 2.35 Paroxya atlantica Female 1 10.00 0.16 1.05 Male 3 25.33 1.36 2.78 All 4 24.33 1.95 5.18 Scudderia texensis Female 4 24.33 1.95 5.18 Male* - - - -

Table 2. Gut microbiome richness and diversity for four treatments

Bacterial Fungal Inverse Bacterial Inverse Fungal Treatment Bacterial Simpson Shannon Fungal Simpson Shannon group Sample Richness Index Index Richness Index Index None #1-36 40 10.35 2.81 42 10.58 2.94 #2-40 40 3.18 1.74 50 6.5 2.61 #3-44 17 6.15 2.23 22 5.63 2.04 NP #4-37 21 4.19 2.12 24 2.12 1.39 #5-43 19 2.13 1.18 21 1.36 0.75 #6-46 7 1.17 0.39 7 1.01 0.04 NP x Na #7-38 30 8.32 2.68 25 3.78 1.88 #8-42 37 14.59 3.03 16 9.45 2.44 #9-45 65 3.58 2.44 42 4.97 2.32 Na #10-39 26 11.17 2.76 11 6.71 2.12 #11-41 35 4.84 2.26 19 8.23 2.41 #12-47 30 6.3 2.4 26 5.14 2.04 #13-65 41 12.81 2.92 25 5.28 2.23