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December 2019

Bacterial Diversity of the Gut of Cotinis nitida

Roy Attila Kucuk Clemson University, [email protected]

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Recommended Citation Kucuk, Roy Attila, "Bacterial Diversity of the Gut of Cotinis nitida" (2019). All Theses. 3218. https://tigerprints.clemson.edu/all_theses/3218

This Thesis is brought to you for free and open access by the Theses at TigerPrints. It has been accepted for inclusion in All Theses by an authorized administrator of TigerPrints. For more information, please contact [email protected]. BACTERIAL DIVERSITY OF THE GUT OF Cotinis nitida

A Thesis Presented to the Graduate School of Clemson University

In Partial Fulfillment of the Requirements for the Degree Master of Science Plant and Environmental Science

by Roy Attila Kucuk December 2019

Accepted by: Dr. Michael Caterino, Committee Chair Dr. Peter Adler Dr. Sharon Bewick Dr. Matthew Turnbull ABSTRACT

Adult and larval Holometabolous insects exhibit radically different gut morphologies tied to their differing natural histories. Additionally, like other animals, these organisms frequently show distinctive morphological and physiological partitioning of their digestive systems, and this reflects on resident microbial communities. A review of the literature reveals bacteria have formed various symbioses with holometabolous hosts, differing widely in the context of host-symbiont services and patterns of colonization. The significance of these organisms in shaping host evolution and vice- versa is, at present, unclear, but intriguing in the context of host phylogeny.

Using high throughput 16S amplicon sequencing, the bacterial community of the digestive tract of adults and larvae of the common North American scarab species Cotinis nitida is characterized according to life stage, gut structure, and adult sex. Through statistical analysis of sequence data, I show that the bacterial communities of the digestive system differ significantly between adults and larvae in both taxon richness and relatedness, and that no major differences exist between adult male and adult female beetles in terms of bacterial community. Significant differences are observed between the midgut and hindgut regions in adult beetles. The partitioning between communities of bacteria in the digestive system of larvae is displayed through significant differences in two distinct hindgut regions, the ileum and the expanded paunch., while there is no significant difference between the midgut and ileum portion of the hindgut region in larvae.

ii ACKNOWLEDGMENTS

I thank my advisor Dr. Michael Caterino for providing both professional expertise and steadfast and amiable guidance, and facilitating freedom in my research pursuits. I thank the other members of my committee, Dr. Peter Adler, Dr. Sharon Bewick, and Dr.

Matthew Turnbull for adding novel perspectives on my work that ranged across physiology, morphology, and community ecology, and thus kept its aims and application as holistic as possible. I thank Dr. Barbara Campbell for providing lab space and equipment to carry out some of my methods, but also for providing constant technical advice and encouragement. I thank Lauren O’Connell for providing valuable suggestions and assistance in terms of molecular work and bioinformatics. I thank Matt Green for his assistance in carrying out statistical measures, as well as ready and regular advice. I thank

Dr. Vincent Richards for permitting me to carry out some of my methods in his lab as well as for providing access to his MiSeq machine.

iii TABLE OF CONTENTS

Page

TITLE PAGE ...... i

ABSTRACT ...... ii

ACKNOWLEDGMENTS ...... iii

LIST OF TABLES ...... vi

LIST OF FIGURES ...... vii

CHAPTER

I. BACTERIAL COMMUNITIES OF HOLOMETABOLOUS INSECTS ARE SHAPED BY INTERACTIONS WITH HOST PHYSIOLOGY AND MORPHOLOGY ...... 1

Abstract ...... 1 Introduction ...... 1 Host morphology ...... 3 Host sexual dimorphism ...... 4 Mechanisms of transmission ...... 5 Host nutrition ...... 7 Host development and defense ...... 12 Host behavior ...... 14 Host lineage ...... 16 Concluding remarks ...... 21

II. BACTERIAL DIVERSITY OF THE GUT OF Cotinis nitida ...... 23

Introduction ...... 23 Materials and methods ...... 28 Results ...... 35 Discussion ...... 39 Conclusion ...... 50

APPENDICES ...... 53

iv Table of Contents (Continued) Page

A: Tables ...... 54 B: Figures ...... 55

REFERENCES ...... 73

v LIST OF TABLES

Table Page

A-1 Kruskal-Wallis pairwise comparison of bacterial communities of the gut of C. nitida ...... 54

A-2 Pairwise permanova of bacterial communities of the gut of C. nitida ...... 54

vi LIST OF FIGURES

Figure Page

B-1 The gut of an adult C. nitida ...... 55

B-2 The gut of a larval C. nitida ...... 55

B-3 Detail of the midgut of the larval C. nitida ...... 56

B-4 Detail of the ileum of the larval C. nitida ...... 56

B-5 Detail of the exterior paunch of the larval hindgut C. nitida ...... 57

B-6 Alpha diversity measures of adult and larval C. nitida ...... 58

B-7 Alpha diversity of the gut of two life stages of C. nitida according to the Shannon diversity index and Chao1 ...... 59

B-8 Alpha diversity of the gut regions of C. nitida according to the Shannon diversity index and Chao1 ...... 60

B-9 Beta diversity of the gut of two life stages of C. nitida according to unweighted Unifrac ...... 61

B-10 Beta diversity of the gut regions of C. nitida according to unweighted Unifrac ...... 62

B-11 Beta diversity of the gut of the sexes of C. nitida according to unweighted Unifrac ...... 63

B-12 Beta diversity of the gut of two life stages of Cotinis nitida according to Bray-Curtis dissimilarity ...... 64

B-13 Beta diversity of the gut regions of C. nitida according to Bray-Curtis dissimilarity ...... 65

vii List of Figures (Continued)

Figure Page

B-14 Beta diversity of the gut of the sexes of C. nitida according to Bray-Curtis dissimilarity ...... 66

B-15 Beta diversity of the gut of two life stages of C. nitida according to NMDS (Bray-Curtis dissimilarity) ...... 67

B-16 Beta diversity of the gut of the gut regions of C. nitida according to NMDS (Bray-Curtis dissimilarity) ...... 68

B-17 Relative frequency of bacterial phyla in the gut of C. nitida according to gut region ...... 69

B-18 Relative frequency of bacterial phyla in the gut of C. nitida according to gut region ...... 70

B-19 Relative frequency of Gluconobacter (in purple) in the gut of C. nitida according to gut region ...... 71

B-20 Relative frequency of Pantoea (in purple) in the gut of C. nitida according to gut region ...... 71

B-21 Relative frequency of Methanobrevibacter (in pink and green) in the gut of C. nitida according to gut region ...... 72

viii CHAPTER ONE

DIFFERENTIAL RELIANCE ON OBLIGATE AND FACULTATIVE SYMBIOTIC GUT BACTERIA IN THE HOLOMETABOLA

Abstract

The diversity and ecological variety of Holometabola, partly mediated through their developmental biology, makes them sources for a wide array of dynamic relationships with bacteria. A review of the literature reveals that holometabolous hosts rely primarily on facultative (apparently non-host-dependent) organisms to carry out essential or important roles.

The driving forces behind this relationship can be partly understood through the nature of transmission in Holometabola, including host morphology and reproductive behavior. The predominance of facultative organisms over obligates can also be explained by the various services provided by their symbionts, including nutrition, immune system health, and development. The diversity of Holometabola in the context of bacterial symbiosis can consequently be elucidated through a comparison of obligate (host-dependent) versus facultative partnerships.

Introduction

Many microorganisms colonize animals, which offer a rich supply of resources and a relatively stable habitat. Among the most notable of these relationships are those of bacteria inhabiting the digestive system of their hosts. Some of these microbes are transient commensals

(Zhang et al., 2016, Hammer et al 2017, Zhao et al. 2017), whereas others are capable of colonizing and reproducing in a host and assisting in physiological processes for a portion of its lifespan (Salem et al. 2017). In general, symbiotic gut bacteria may be either obligate organisms

1 that cannot survive outside of their host, or they may be facultative organisms that are apparently capable of living independently in the environment.

The developmental strategy of holometabolous insects and the consequent ecological differential between adults and immatures presents bacterial gut symbionts with a variety of habitats to colonize and exploit, but also presents challenges for bacterial residents, including the potential for metamorphosis-mediated population turnover. Although all insect orders have representatives that possess a microbiome (Kibuchi 2009), the holometabolans show a striking taxonomic diversity that encompasses great ecological variety (and distinctive life stages incumbent on these factors) (Rolff et al. 2019, Truman & Riddiford 2019). Given the occurrences of microbial interaction presented by immense multipartite variation, the biodiversity of holometabolous insects can be seen as a mirror for the biodiversity of communities of bacteria and a particularly broad window into the numerous dynamic relationships between host and prokaryotic symbiont.

Precisely how this biodiversity reflects on bacterial relationships with holometabolous insects and how they are maintained remains a mystery. We can shed light on the processes involved, however, by exploring the particular ways bacteria colonize the gut, transmit between hosts (including different developmental stages), and assist in major physiological needs.

Examining the means by which bacteria colonize the gut and the various manners in which they benefit their host, we see that holometabolous insects rely primarily on facultative organisms compared to inseparable obligates. We can also potentially elucidate the diversity of the

Holometabola through the lens of bacterial partnership, whether or not certain lineages demonstrate more instances of obligate versus facultative symbionts in their guts, how these

2 relationships may have arisen, and how we may use them to predict symbiont diversity across the lineages of Holometabola.

Host morphology

The morphological compartmentalization of the digestive system is a key aspect of its suitability as a habitat for bacterial taxa. Indeed, an insect’s gut shows numerous adaptations to facilitate certain diets, including morphologically distinct and often subdivided foregut, midgut, and hindgut regions. Additionally, further morphological modification has allowed for the settlement and storage of bacteria. This includes specialized diverticula, crypts known as bacteriomes, and wholesale modifications to the regions of the gut.

The security of an obligate relationship with an insect host is often facilitated by localization in the gut in addition to explicit storage in organs called bacteriomes. In early studies of cassidine and eumolpine leaf beetles, for instance, Stammer (1936) made morphological observations about the presence of bacterial symbiotes, which are stored in well-defined

“bacteriome” regions around the foregut-midgut junction. This association with gut morphology was also revealed by the work of Fukumori et al. (2017) in the eumolpine leaf beetle Bromius obscurus. Similarly, in some weevils, a pronounced bacteriome is located on specific portions of the gut, and houses symbiotic bacteria (Buchner 1965, Anbutsu et al. 2017). Such adaptations, while costly, offer a means by which to ensure relative permanence of a bacterial community if only as a region of proliferation—in olive flies obligate bacteria colonize a foregut diverticulum from which they can settle on food material and assist in nutrient provisioning (Ben-Yosef et al.

2010).

3 Although specialized bacteriomes are often associated with obligate symbionts, more generalized host gut morphology can also be important for maintenance of facultative partners.

This is particularly notable in scarab beetles and their relatives, whose larvae possess a distinctively reduced foregut, greatly enlarged caeca-covered midgut, and an expanded bacteria- rich hindgut paunch for storage of food (Zhang 2018). Interestingly, in these insects, segregation of bacterial communities according to gut region occurs, with each general region of the gut harboring its own distinctive community (Egert et al. 2003, Andert et al., 2010). Similar compartmentalization and specificity of facultative bacteria to particular gut regions is found in crane fly larvae (Klug & Kotarski 1980). In the case of scarabs the bacteria residing in the gut are facultative organisms capable of living outside of their hosts, and in cases of vertical transmission some of these organisms reside in a maternally inoculated food substrate (Shukla et al. 2016). The preference for certain regions of the digestive system by facultative bacteria is not restricted to the hindgut—adult vinegar flies maintain a community of environmentally acquired symbionts of the genus Acetobacter in the foregut (Pais et al. 2018). Thus, while obligate symbionts rely on highly modified structures that specifically suit their proliferation and maintenance, even facultative bacteria can still maintain a foothold in the host according to morphology.

Host sexual dimorphism

Gut morphology can be influenced by host sexual dimorphism (Guillén et al. 2019).

Given the tight relationship between gut morphology and bacterial colonization as well as differences in parental investment it is not surprising that differentiation between sexes plays a

4 role in bacterial symbioses. In the tortoise beetle Cassida rubiginosa, for example, the female possesses a separate community of an obligate gut symbiont near its reproductive tract, whereas the male does not (Salem et al., 2017). Similarly, in weevils harboring the bacteria Nardonella, adult females house the originally gut-associated bacterial community in a similar manner, around the ovarioles and developing oocytes (Anbutsu et al. 2017).

Just as in the case of general morphology, host sexual dimorphism affects both obligate and facultative symbionts. For example, a female-specific bacterial community has been suggested in scarabs, and the guts of female beetles contain taxa that more closely align with microbes that putatively live in the larval brood chamber prior to inoculation (Shukla et al.

2016). Although the closer connection between larval and female gut microbiomes amongst scarabs indicates potential selection pressures associated with maternal transmission, microbiome differentiation among sexes in scarabs has been suggested to play other roles as well. For instance, the bacteria residing in the hindguts of females might produce phenols that likely function as sex pheromones (Hoyt 1971). In some cases sexual dimorphism of the parental microbiome not only determines what organisms get passed on, but also what sorts of key benefits, particularly nutritional and developmental ones, the young insects are receive. Given the routes by which facultative bacteria can be acquired via surrounding substrate, the mechanisms by which hosts evolve to rely on these bacteria may be linked to parent choice

(oviposition site) and parental care (overt provisioning of a fecal inoculate or regurgitant). These differing evolutionary patterns may in turn contribute to host diversification.

Mechanisms of transmission

5 The presence of distinctive larval, pupal and adult life stages, along with a metamorphic period marked by the expulsion of gut contents and a modification of gut morphology from larval to pupal and adult stages, presents a challenge for bacteria in the guts of Holometabola

(Hammer & Moran 2019). Obligate symbionts, relying on endogenous associations with their insect hosts overcome this challenge through close morphological associations. This is prevalent in holometabolans whose adult and larval forms have an identical diet (Hammer & Moran 2019) and thus enjoy similar gut morphology. For example, the thistle tortoise beetle (Chrysomelidae:

Cassidinae) transmits the obligate symbiont Stammera and possesses specialized atria in the reproductive tract of the female beetle that inoculates the eggs with a bacteria-rich cap (Salem et al. 2017). Beyond specialized bacteriomes and sex-specific structures that facilitate bacterial transmission across life stages holometabolans must employ several other mechanisms for acquiring symbionts. The first of these is mere environmental acquisition, which can effectively only involve facultative organisms. Horizontal transmission of bacteria has been shown in various taxa via ingestion of food, as seen in mosquitos (Coon et al., 2015) and the coffee berry borer (Mariño et al., 2018).

Although environmental acquisition of facultative symbionts is common, facultative microbes can also be directly transmitted by parents. In burying beetles (Silphidae), for example, bacteria in the genus Dysgonomonas in conjunction with a similarly abundant fungus species are transmitted to their new larval hosts via hindgut secretions on food substrate (Wang et al. 2017).

Similarly, among scarabs, some dung beetles are capable of providing their larvae with bacterial inoculates in the form of a microorganism-rich brood ball or pedestal (Estes et al. 2013, Shukla et al, 2016). In both of these instances of parental transmission, the utilized taxa are facultative,

6 surviving freely in the substrate on which the larvae feed rather than in a caplet or matrix attached to the egg. The ubiquity of these facultative organisms makes sense in the context of holometabolous insects: they do not require made-to-order morphological adaptations to be stored and transmitted. Indeed, some of them only depend on oviposition substrate.

Host nutrition

The differing diets of insect hosts influence composition of the bacteria residing in their digestive tracts (Yun et al. 2014, Kudo et al. 2019). The holometabolan developmental strategy often corresponds to a marked schism in trophic biology, which in turn affects the types of bacteria that colonize the gut as well as the physiological relationship they form with the host. Moreover, the constant passage of material through the gut makes it a relatively unstable zone of colonization. Combined with the clearing of the gut prior to pupation and subsequent changes in morphology and diet, the gut is a difficult environment for bacteria to colonize, but this constant turnover on both long and short time-scales provides a selective advantage for those that can overcome it.

In general, insects feeding on living plant tissues have less diverse communities of microorganisms (Kolasa et al. 2019), but these communities are varied in their alimentary associations. In the case of nectar-feeding hosts, such as certain adult

Lepidoptera, communities taxonomically correspond to the bacteria in the host’s food source (Phalnikar 2018). Conifer-feeding weevils similarly harbor a unique bacterial community that, though conserved across populations, bears a strong similarity to other arthropods feeding on the same diet, suggesting bacterial preference or association with a

7 particular food source (Berasategui et al. 2016). Bacteria that persist on both the food source and in the holometabolan gut essentially extend their habitat, which can be beneficial both in terms of homeostasis and dispersal. In these cases, however, the relationship between host and microbe is relatively loose, with neither depending on the other for survival.

Insects feeding on more nutritionally difficult plant tissue comprised of complex carbohydrates frequently develop stronger dependencies on their gut microbiomes, for example by outsourcing catabolic labor of digestion to microorganisms. This, in turn, offers numerous niches for bacteria to exploit. Thus, while the task of breaking down food is a physiologically tall order, it furnishes rich habitat for colonists capable of accomplishing it. Bran-feeding house flies (M. domestica), for example, apparently garner a microbiome including Lysinibacillus, Comamonas, Dysgonomonas, Bacteroides and Lactobacillus, which are capable of assisting in digestion, from their food source

(Zhao et al. 2017). Leaf-feeding is linked to bacteria capable of digesting cellulose and xylan (Pinto-Tomás et al. 2007, Anand et al. 2010, Chen et al. 2016, Dantur et al. 2015).

Similarly, pectin and starch-digesting bacteria have been isolated from foliovorous lepidopteran larvae (Anand et al. 2010). Finally, wood-feeding hosts offer habitat for facultative bacterial symbionts, as well as wood-degrading fungi and other eukaryotic organisms. Indeed, xylophagy in the holometabolans has furnished multiple opportunities for bacterial colonization by highly specialized microbes that are necessary for nutritional assistance. Some cerambycid larvae, for instance, house gut bacteria with genes associated with lignocellulose breakdown (Mohammed et al. 2018). Likewise, in the bark

8 and turpentine beetles, which famously rely on fungi to gain access to wood biomass as both adults and larvae, cellulolytic gut bacteria are present (Hu et al. 2013). These bacteria are found in adults and larvae, which have similar diets, and include

Stenotrophomonas maltophilia and Ponticoccus gilvus (Morales-Jiménez et al. 2012). In line with the hypothesis that wood-feeding recruitment of a highly specialized gut microbiome, some taxonomic similarity exists between the bacteria residing in the guts of scarabs and those associated with the obligate eukaryotic symbionts of the wood-feeding termite Mastotermes (Pittmann et al. 2008). In bamboo weevils, lignocellulose digestion is mediated by bacteria as well, with the key assistants being bacteria of the genera

Lactococcus, Dysgonomonas, Serratia, and Enterrococcus (Luo et al. 2019). The efficiency at which these organisms assist their hosts can also grant them access to a regular supply of food source—the seed-eating carabid consumes more food when digestive bacteria are present (Schmid et al. 2014).

In general, bacteria-mediated catabolism of plant fibers in Holometabola is provided by facultative bacteria, rather than obligates. There are, however, exceptions. In a tortoise beetle that produces digestive enzymes capable of processing most plant material except pectin, a bacterial symbiont fills the gap by producing enzymes to take care of the latter in larvae and adults (Salem et al 2017). Similar associations in other chrysomelids and other phytophagous insects as well. Catabolic nutritional services provided by obligates may be explained by the fact that both larvae and adult beetles have the same diet (Engel & Moran 2019)— a relatively uncommon occurrence among certain Holometabola and one that may demand a physiologically bound obligate.

9 In addition to secreting enzymes necessary for digestion and fulfilling the role of an “external stomach” or merely being commensals growing on portions of food substrate, gut-associated bacteria are able to occupy additional niches involved in host nutrition. In herbivorous ants, for example, bacteria aid in nitrogen fixation (Russell et al., 2009). Putatively nitrogen-fixing bacteria have also been found in the deadwood- feeding cerambycid Prionoplus reticularis (Reid et al. 2011). In turpentine beetles and bark beetles, gut bacteria aid in nitrogen fixation as well (Morales-Jiménez et al. 2009,

Morales-Jiménez et al. 2013). In the cerambycid, Anoplophora glabripennis, the microbiome aids in the uptake of essential amino acids (Ayayee et al. 2016). Bacteria may also improve the quality of food substrate in other ways. In leaf-mining caterpillars, cytokinins produced by the insect host’s bacteria enable maintenance of the food substrate for enhanced nutritional value (Kaiser et al. 2010, Body et al. 2013). Just as in the case of enzyme-provisioning bacteria, these organisms are not totally dependent on the host for survival. Host specialization on a particular food source offers opportunities for obligate relationships to form as well, however. This is notably displayed by the presence of Wigglesworthia and Sodalis in the blood-feeding tsetse fly (Wang et al 2013,

Griffith et al., 2018), both of which are dependent on host morphology and physiology. In olive flies (Bactrocera oleae) bacteria of the genus Erwinia perform a similar service, assisting in amino-acid provisioning to supplement the protein-poor honeydew diet of their host. Just as in catabolic assistance, the adult insect requires the symbionts for survival, and such a necessity may select for organisms that can be obligately associated with the host.

10 Not all diets are challenging, and bacteria abound in the guts of Holometabola dealing with more nutritionally balanced food sources as well. Environmentally acquired bacteria that don’t seem to perform any digestive services for their hosts are found in predatory Holometabola. Vespid wasps, for example, show a microbiome partly comprised of the symbionts of their honeybee prey, as shown by Suenami et al. (2019) in the Asian giant hornet and Japanese hornet. The acquisition of a microbiome via predation might be a widespread phenomenon, suggesting a benefit for bacteria capable of colonizing new guts—the destruction of the host is not necessarily the elimination of one’s tenure in a nutrient rich digestive tract. Whether or not diverse communities are regularly established by predation and whether or not they provide any important services to the host is a particularly valuable question when considering a highly biodiverse and widespread predatory groups of generalists.

Bacterial colonization is also shaped by the capacity to thrive in the host digestive system in spite of adult-larval difference in diet and morphology. For example, the wasp

Megastigmus shows a general similarity of bacterial communities between life stages

(Paulson 2014). Some butterflies show a similar absence of community distinctiveness between life stages, with the differing feeding guilds apparently not always influencing general gut communities (Phalnikar et al., 2017). In butterflies, there is a relatively scant association between diet and microbiome with only 4% of the bacterial community difference being influenced by food source in some species (Ravenscraft et al. 2018). In butterfly larvae there is a less marked association between microbiota and digestion, and

11 the resident bacteria of caterpillar guts generally do not correspond to their food substrate

(Minard et al., 2019).

The value of facultative bacteria in assisting in dietary heavy lifting would certainly facilitate relationships in which these organisms are common. We see, however that in many Holometabola, the direct catabolic necessity of such organisms is tenuous at best. These seeming freeloaders offer other services that make their facultative nature a useful asset to insects, and consequently gain access to habitat without sacrificing their own autonomy.

Host development and defense

Bacterial colonization of insect hosts and the evolution of symbioses is not entirely incumbent on the symbiont’s mastery at processing food material via enzyme production or thriving in a given food substrate as a transient commensal. The bacteria of the digestive tract are capable of participating in systems not directly related to alimentation and digestion in their hosts. Mosquitoes, whose aquatic larval stage is spent feeding on detritus or small organisms, require, through ingestion, the presence of a gut microbial community to induce anoxia (via their metabolic activities) to undergo ecdysis (Coon 2014). In the red palm weevil, a core gut community aids the regulation of metabolism (Muhammad et al., 2017). These “core communities” are comprised of environmentally acquired facultative bacterial symbionts, however. An exception to this is seen in many weevil subfamilies, in which the obligate gut- bacteriome-inhabiting bacteria Nardonella synthesize tyrosines that aid in durable exoskeleton

12 construction (Anbutsu et al. 2017). This phenomenon of an obligate bacteria assisting in cuticle development is also shown in the silvanid Oryzaephilus surinamensis (Hirota et al. 2017).

The immune defenses of hosts offer a major challenge for bacteria, making the gut of a living Holometabolous insect a potentially hostile site of colonization. It also, however, provides bacteria with a novel way to form partnerships with their hosts and colonize new habitat. Indeed, the very development of the immune system is enhanced by the bacterial microbiome (Krams et al. 2017, Kwong et al. 2017). This is notable in vinegar flies and honeybees, whose gut bacteria aid in immune system homeostasis (Ryu et al. 2008, Kwong et al. 2017). Facultative bacteria are not the only organisms that assist in these processes: the obligate bacterium Wigglesworthia stimulates normal expression of immunity-related genes and the production of phagocytic hemocytes in tsetse flies

(Weiss et al. 2012). Evidently, even organisms with a seemingly limited functionality from a genomic perspective can still provide novel services for the host.

The holometabolous host’s need to defend itself from pathogens and parasites at various distinctive stages of development offers opportunities for bacteria-mediated aid. Gut bacteria can mount biochemical attacks on pathogens and parasites of their hosts, effectively gaining admission to new habitat and food by acting as bodyguards. The biochemical warfare that these organisms must mount to compete with numerous other microbial competitors makes them ideal for this role, and as in the case of feeding relationships, antimicrobial defense is a domain teeming with facultative bacteria. Flesh-feeding burying beetles rely on specialized hindgut bacteria to inhibit the growth of generalist microbes on the food of their larvae (Shukla et al.

2018). In house flies (Musca domestica), hindgut bacteria of the genus Klebsiella reduce fungal

13 growth on eggs (Lam et al. 2009). This genus has also been found in bacteria associated with the gut-content-lined pupal chamber of stag beetles and is implicated in a similar defensive function

(Miyashita et al. 2015). Adams et al. (2011), considering the chitinase-producing bacteria associated with the fungus-harboring wood wasp Sirex noctilio, hypothesized that these bacteria keep the growth of fungal symbionts in check. In mosquitoes, the infection rate of the

Plasmodium parasite is reduced by the presence of a bacterial microbiome (Dong et al. 2009).

Bacteria in the gut can contribute to suppression of opportunistic pathogens, as is the case with adult mosquitoes harboring Serratia marescens (Wei 2017). In fruit flies (D. melanogaster), a species of Spiroplasma is implicated in defense against nematode parasites (Jaenike et al. 2010).

Facultative bacteria also assist in management and suppression of potentially dangerous toxins, as seen in cowpea beetles exposed to dichlorvos (Akami et al. 2019). The coffee berry borer,

Hypothenemus hampei (Curculionidae), houses bacteria in its gut that assist in the breakdown of the harmful alkaloid caffeine (Ceja-Navarro et al. 2015). Once again, this appears to be an exploitation of a capacity evolved in facultative microbes that are equipped to deal with various environmental stresses otherwise absent from inside insect hosts. Consequently, relationships with facultative organisms potentially dominate certain services by virtue of exaptation—what’s good for the bacteria is incidentally good for the insect host.

Host behavior

The complexity of host behavior offers yet another set of opportunities for environmentally acquired and facultative bacteria. This manipulation can be simple, as in the case of fruit flies (D. melanogaster) whose resident facultative gut bacteria manipulate

14 locomotion via the enzyme xylose isomerase (Schretter et al. 2018). Manipulation can also be more complex, inducing particular habits. The tephritid fruit fly Bactrocera dorsalis, individuals experimentally devoid of a microbiome exhibit radically different feeding behavior from microbiome-possessing controls (Akami et al. 2019). The manipulation of behavior in spite of a need for nutritional assistance is strikingly exhibited in D. melanogaster, in which commensal

Acetobacter pomorum and Lactobacillus sp. prevent their hosts from seeking proteinaceous food sources as a result of low quantities of essential amino acids, while simultaneously protecting their hosts from the low fitness that typically comes with this deprivation (Leitão-Gonçalves et al. 2017). The dearth of obligate gut bacteria in overtly influencing or manipulating host behavior is curious but may be explained by their host dependence. The influence of facultative organisms on behavior is mediated by presence-absence: an ever-present obligate is not going to alter host behavior in this fashion. Whether or not obligate symbionts can also directly influence individual behavior within a host is unknown.

Bacteria demonstrate these behavior-altering capacities in regard to potential hosts via semiochemical production. In scarabs, a behavior-influencing function has been suggested in the form of bacterial metabolic products (phenols) serving as sex pheromones in adults of a melolonthine beetle (Hoyt 1971). In non-holometabolans, such as cockroaches (Wada-Katsumata et al. 2015), who rely on such a system, as well as in holometabolous insects that depend on fungi as pheromone producers (Johnson & Vishniac 1990), the symbionts in question are facultative organisms. It is possible, however, that obligate symbionts could facilitate behavioral manipulation through semiochemical production as well. The ability to manipulate a host even in the fluctuating environment like the gut of a holometabolous insect evidently adds a new avenue

15 of exploitation for bacteria, obligate specialist or commensal. While not all bacterial manipulation of the host from the gut is necessarily an adaptation, it can provide a basis for novel relationships, facilitating new means of acquiring nutrients for a would-be symbiont and moving oneself and future generations to suitable environments.

Host lineage

The relationships holometabolous insects form with bacteria encompass numerous aspects of host biology. The predominance of one general type of symbiont, facultative or obligate, varies greatly within the Holometabola, and elucidating these partnerships on a broader evolutionary scale is difficult. Facultative organisms, by virtue of their capacity to survive outside of the insect host may not exhibit phylogenies that completely mirror that of their hosts.

Consequently, phylogenetic inference about host-symbiont relationship is not as clear as it is in insects with obligate gut symbionts demonstrating co-speciation. Our limited knowledge of the ancestral condition of symbiosis in regard to numerous services provided by bacteria in the gut further clouds our understanding of how readily such symbioses form between insect and symbiont. However, by examining the services provided by gut bacteria in the context of host lineage and keeping the fundamental physiological idiosyncrasies of the Holometabola in mind, we can develop and test hypotheses relating to the presence of one type of bacteria or another in gut.

The Hymenoptera comprise the vast majority of eusocial species in the

Holometabola. Such behavior offers a foothold for microbes, as trophallaxis between colony members enables a given species to maintain a stable colony of gut bacteria

16 (Engel & Moran 2013). Moreover, this behavior may offer bacteria the opportunity to be maintained in a host’s digestive system without relying on morphological and physiological measures that may otherwise be necessary to ensure its transmission to different life stages. Such measures in turn may influence the evolution of the symbiont’s dependence on its host. This is exemplified by the honeybees and other Apidae, in which there is a small core community of gut bacteria (Martinson et al. 2011) dominated by

Snodgrassella and Gilliamella which can exist in culture outside of their hosts (Kwong &

Moran 2013). Among the ants, host-bacteria diversity is underpinned by core gut species, as seen in Tetraponera (van Borm 2002) and neotropical army ants (Łukasik et al. 2017).

Additionally, social Hymenoptera symbionts can assist in the priming of the host immune system against potential pathogens (Kwong et al. 2017) and contribute to development

(Raymann and Moran 2018). Indeed, honeybee and bumblebee health are associated with the presence of a taxonomically limited but highly conserved community of resident bacteria (Raymann and Moran 2018, Koch & Schmid-Hempel 2011). Gut bacteria also show capacity for detoxification and nutrient provisioning in social Hymenoptera (Zheng et al. 2016). Other social Hymenoptera, such as the predatory Vespidae, possess a limited set of core taxa, as is the case of Vespa mandarinia and V. simillima (Suenami et al.,

2019). Given the services that those bacteria provide for their herbivorous hosts, it may be hypothesized that similar benefits are granted to their new residences.

Generally, a taxonomically associated microbiome is greatly represented in social corbiculate bees, whereas variable and transient communities appear to be the norm for other bees and wasps (Engel et al., 2016). Even so, some non-social Hymenoptera exhibit consistent

17 microbial partnerships as well. In members of the parasitoid Pteromalidae, the gut microbiome appears intimately linked to phylogeny (Brucker & Bordenstein 2013). Interspecific transmission between closely related taxa in Nasonia results in greatly decreased fitness compared to intraspecific transmission (van Opstal & Bordenstein 2019). Comparing solitary organisms that are closely related to eusocial or semisocial ones, we may be able to further elucidate the differing methods by which symbiotic gut bacteria evolve from transient organisms.

In both adult and larval Lepidoptera, the bacterial community is relatively unconserved (Hammer et al. 2017, Jones et al. 2019), although general differences in microbiota have been observed across species (Phalnikar et al. 2017, Jones et al. 2019).

While there is distinction on the taxonomic level in adult butterflies, this is generally subordinate to individual differences (Ravenscraft et al. 2018). In a single species of butterfly, neither taxonomy nor ecology showed a strong link to bacterial associations

(Minard et al. 2019). In the case of some taxa, the bacterial community has been tied to the nutritional biology of larvae, however (Anand et al. 2010, Kaiser et al. 2010, Body et al. 2013). The apparent absence of a stable microbiome in some Lepidoptera while other species benefit from bacteria in their diet demonstrates the value of versatility in facultative symbionts—relationships can readily arise to provide particular services, and the mere capacity to harbor such bacteria is useful to the host.

An understanding of the evolutionary context of the relatively symbiont-free

Lepidopteran gut may be enriched by studying the closely related Trichoptera, which display radically different diets and a general physiology that may facilitate symbioses with gut bacteria.

18 Among the Diptera, differing diets and natural histories can be partly explained through a reliance on a variety of facultative organisms. Vinegar flies (Drosophila melanogaster) possess a microbiome comprised of facultative bacteria that can provision nutrients, among other services (Leitão-Gonçalves et al. 2017, Gould et al. 2018), and dominant taxa may vary between study populations (Engel & Moran 2013). Even among species with a conserved community, environmentally acquired symbionts reign, as seen in the cactus specialist Drosophila nigrospiracula (Martinson et al. 2017). The success of calyptrate filth flies in their habitat of decay can be attributed to protective facultative bacteria that keep the environment free of fungi, as seen in the common house fly Musca domestica (Lam et al. 2009). A testament to the value of facultative bacteria can be observed in mosquitos for whom such organisms are essential for molting and pupation

(Coon et al. 2015 and Coon et al. 2016). Even so, obligate symbionts assist in many interesting evolutionary breakthroughs in Holometabola, including, for example, hematophagy and immunity in tsetse flies (Weiss et al. 2012, Wang et al. 2013).

Coleoptera may owe some of their diversity to the exploitation of plants as a food source during at least part of their life cycles (Farrell 1998, Janz et al. 2006), and this may partly be explained by bacterial assistance. The diverse leaf beetles and weevils have obligate bacterial symbionts in their guts (Anbutsu et al. 2017, Salem et al 2017) The services provided by these widespread bacteria are not all overtly nutritional, however, as seen in some weevils whose symbionts synthesize tyrosine for developing larvae (Anbutsu et al. 2017). The presence of two instances of co-speciation and three instances of symbiosis with an obligate symbiont (Toju et al.

2013) in weevils, in addition to obligates performing a similar function in other beetles like the

19 Silvanidae (Hirota et al. 2017) suggests a broader necessity for obligate symbionts among the plant-feeding insects. Further surveying and functional assessment of symbionts in successful phytophagous beetle lineages including the facultative-harboring Buprestidae (Vasanthakumar et al. 2008, Bosorov et al. 2019) and Cerambycidae (Ayayee et al. 2014, Ayayee et al. 2016,

Mohammed et al. 2017) will be necessary to assess the evolution of dependence on obligate bacteria compared to facultatives. The widespread strategy of utilizing facultative bacteria in the

Scarabaeoidea whose larvae feed primarily on living and dead plant tissue (Egert et al., 2003,

Egert et al., 2005, Arias-Cordero et al., 2012, Ceja-Navarro et al. 2014, Miyashita et al. 2015,

Zhang et al. 2018, Chouia et al. 2019) further emphasizes the need to look more precisely at host biology. Patterns based on general feeding guild do little to elucidate bacterial services beyond gross conjecture.

The Carabidae are poorly represented in microbiome research, although they show high gut bacteria diversity within their ranks (Kolasa et al. 2019) and the presence of omnivory in this group demonstrates a link between diet and the advent of facultative symbionts that aid in digestion (Lundgren & Lehman 2010). Whether or not similar trends can be observed in other largely predatory beetle groups is unknown, but additional studies may shed further light on carnivory and high bacterial diversity and whether or not high diversity is not merely an artifact of broad diet. Just as in the case of other beetles as well as Holometabola, further assessment of these organisms must not only consider heightened taxon sampling but also more rigorous texting of organismal function, particularly outside of the context of diet. The absence of entire orders (Mecoptera and

Neuroptera) from extensive functional analysis, as well as large swaths of the phylogeny

20 of more speciose orders precludes our ability to make sense of obligate and facultative bacterial symbionts on a phylogenetic scale.

Concluding remarks

The diversity of holometabolous bacterial taxa, along with difficulties in experimentally manipulating the bacterial community of the gut of holometabolous insects, continues to challenge our ability to develop a well-structured understanding of bacterial services. This struggle is further deepened by the potential of numerous taxa to influence the “performance” of other members of the community, in addition to the various host-specific factors that shape diversity and functionality of bacteria in their guts. While some bacteria have clearly co-evolved with their hosts as demonstrated by phylogeny (Toju et al. 2013), the extent to which bacteria, particularly obligate organisms drive insect host evolution and why some insects favor the use of facultative organisms over obligates is still a mystery. Through continued analysis of these relationships and a steady attempt to contextualize symbiont services and host biology and phylogeny, we can get closer to grasping the dynamics of these oft-appearing relationships.

Future directions in this field should include extensive sampling of host taxa for bacteria, with a greater emphasis on community partitioning based on gut morphology and physiology.

Moreover, such sampling should take into account all life stages of the host. Systematic manipulation of the gut community and isolation of core community members (if present) for more thorough experimentation relating to various host services should follow these approaches.

Additionally, we must consider the community interactions between bacteria as well as other gut-dwelling organisms, including fungi. The broad dichotomy of facultative and symbiotic

21 organisms in Holometabola demonstrates vague patterns than we can easily render clearer with additions to methodology.

22 CHAPTER TWO

BACTERIAL DIVERSITY OF THE GUT OF Cotinis nitida

Introduction

Numerous factors determine which bacteria ultimately dwell on or inside a given host, and what taxa comprise the core community. An important first step in understanding the functionality, interspecific relationships, and ultimate evolutionary significance of the microbiome for anthropocentric application (medicine, agriculture) is to determine the diversity and test for the presence of a “core community” within a host species. That is, knowledge of the presence of certain organisms and of greater patterns relating to their presence enables more precise investigations of certain organisms within the system. This is certainly true for the bacteria of the holometabolous insect digestive system, which is generally complex, structurally diverse, and inter- and intraspecifically variable.

The scarab beetles are a particularly speciose family of the Holometabola whose members show a diversity in their diets. Additionally, like many other holometabolous insects, the diets and feeding strategies of scarab beetles differ between adults and larvae, and this distinction is evidenced in the form of the alimentary canal. Indeed, the digestive system in scarab larvae is different from that of the adults. This species-level diversity as well as the differing diets between adults and larvae and the consequential remodeling of the digestive tract during metamorphosis presents an enormous challenge to our understanding of bacterial diversity and our ability to infer and test function.

23 The putative benefits conferred by the bacterial microbiome of larval scarabs are nutritional—bacteria capable of digesting the celluloses, hemicelluloses, and pectins that comprise plant tissue have been found in select species of Cetoniinae (Cazemier et al.,

1999, Cazemier et al 2003), Melolonthinae (Huang et al., 2012, Handique 2017), (Shukla et al., 2016) and Rutelinae (Chouaia et al. 2019), and in some cases these organisms have been directly tied to digestive faculty (Huang et al. 2010). Likewise, the maternally transmitted community of a dung beetle is associated with enhanced development of larvae (Schwab et al. 2016). In the case of the cetoniine Pachnoda, the bacteria and endogenous physiological conditions of the high pH midgut aid in increasing solubility of food material, “prepping” it for the plant fiber-digesting bacteria located in the hindgut

(Hobbie et al. 2012). Partitioning of bacterial communities is an apparent trend in scarabs, with each general region of the gut harboring its own distinctive community

(Egert et al. 2003, Andert et al., 2010, Chouia et al. 2019) including unique members like

Promicromonospora pachnodae in some species of Pachnoda Cazemier (2003).

Relatives of this bacterial taxon are found in the gut of unrelated scarab taxa in the subfamily Melolonthinae (Pittmann et al. 2008).

The bacterial communities of scarab larvae mirror those of other insects with similar saprophagous diets. Relatedness exists between the bacteria residing in the guts of scarabs and those associated with the obligate eukaryotic symbionts of the wood-feeding termite Mastotermes (Pittmann et al. 2008). Additionally, there are similarities between termites and scarabs at the family level, with both groups possessing fiber-digesting bacterial families like the Christensenellaceae and Ruminococcaceae, as well as genera of

24 Rikenellaceae like Alistipes (Andert et al. 2010, Huang et al., 2013, Chouia et al. 2019,

Schnorr et al. 2019). The morphological analogy and general dietary similarities between these different insects, combined with the presence of similar microbial residents suggests a possible broader similarity between these systems.

There are differences between the bacterial communities of adult and larval scarabs (Arias-Cordero et al., 2012, Shukla et al. 2016), reflecting on the differing habits, as well as varied routes of community transmission. Notably, this is even true among beetles with similar adult and larval diets, such as the dung beetles. In dung beetles, differences not only lie between adults and young, but also between males and females— the latter harbor taxa phylogenetically closer to those found in the larvae than the males do (Shukla et al. 2016).

Thus, the bacterial microbiota of scarabs encapsulates many of the phenomena observed in such communities in the holometabolans as a whole—it is diverse, it varies across “regions” of the digestive system, it varies between life stages and sexes, and it is in some cases vertically transmitted. A valuable approach in understanding microbial community composition and thus differences in bacterial taxa on the community level is next-generation 16S amplicon sequencing. Most bacteria are unculturable with known techniques, and consequently attempts to understand their diversity on a given substrate through culture-based approaches is rife with inaccuracy (Hongoh & Toyoda 2011).

Through high-throughput sequencing of amplicons of the so-called hypervariable regions of the 16S ribosomal RNA gene in bacteria, we are able to get a clearer portrait of what types of bacteria are present (Woo et al. 2008). While this approach provides poor clarity

25 on the species level, particularly with the V3-V4 regions (Bukin et al. 2019) it enables us to answer basic questions about the diversity and community partitioning of bacteria, including those present in the gut of insects. To begin answering more refined questions about bacterial communities we must know what kinds of taxa are present as well as how these organisms are distributed in the gut.

Our present lack of knowledge of the scarab beetle bacterial microbiome, including its diversity, functional capacity in both a physiological and broader ecological context, in combination with the relative ease of bacterial communities via next- generation sequencing makes further exploration a tantalizing prospect. Moreover, such work has implications in regard to numerous broader efforts to utilize novel bacterial taxa for agriculture, antibiotic synthesis, and bioreactor development (Huang et al. 2010). The scarab, Cotinis nitida, is a ubiquitous agricultural pest known to damage ripe fruit as an adult (Hammons et al. 2008). Like other scarabs, it exhibits a radically different adult and larval gut morphology: the adults possess a simplified digestive tract while the latter possesses a gut noticeably subdivided into small foregut, midgut with three crowns of gastric caeca, and a hindgut divided into an ileum and a large baglike paunch. As a larva it is known to cause damage in turf grass (Potter 1991). Little of its microbiome is known, although adult beetles possess yeasts that are acquired only after eclosion and exposure to the environment outside of the pupal chamber. The yeasts of adult beetles apparently produce semiochemicals that enhance aggregation of beetles on their preferred food sources (fresh fruit, including commercial peach and grape crops) (Vishniac &

Johnson 1990).

26 Vishniac and Johnson (1990) and Johnson and Vishniac (1991) revealed through culture-based analysis that the yeast Trichosporon cutaneum is abundant in the gut of adult beetles. Additionally, these eukaryotic symbionts have been shown to play a relatively important role in the behavioral biology of adult beetles in that they produce volatile semiochemicals that serve as aggregation pheromones in adult beetles (Johnson and Vishniac 1991). The larvae of these beetles have largely been neglected in the context of their microbiome. They also possess actively swimming ciliated protists

(personal observation) of uncertain identity.

There is no research on the bacteria of the gut of C. nitida, any of its close relatives, or any scarabs within the large cosmopolitan tribe (Gymnetini) to which it belongs. In addition to enhancing our understanding about an unexplored facet of a particular host system, shedding light on the bacterial community of C. nitida adds to our general understanding of patterns of bacterial diversity in the guts of animals and provides a new basis of comparison from which we may establish a better understanding of the dynamics of that system. It is my aim to test for a core bacterial microbiome, community partitioning based on gut region, and community differences based on life stages in C. nitida through the analysis of 16S high-throughput amplicons.

From the central objective, I have the following hypotheses:

-The adult and larval midgut,and hindgut structures harbor communities of bacterial OTUs significantly different from each other.

-The larval and adult beetles have significantly different bacterial microbiomes

27 -The gut microbiome of C. nitida is sexually dimorphic—males and females have significantly different communities

Materials and methods

Sample collection

Adult beetles were collected after emerging from their pupal cells between June and September of 2018. Third instar larvae were collected from the soil between

December of 2018 and February of 2019. All individuals were collected from Clemson,

South Carolina. Adult C. nitida are distinguishable from all other scarabs in the region by their gross morphology—no other members of the genus or tribe are known from South

Carolina, and the presence of a distinctive hood-like pronotum covering the scutellum distinguishes adults from vaguely similar species like Euphoria fulgida. Cotinis larvae are distinguished from those of other taxa by their size, manner of locomotion, and terminal setae. In total, 12 adults (7 males and 5 females) and 11 third instar larvae of indeterminate sex were sampled.

Dissections

All dissections were performed on individual specimens and glassware was washed and sterilized with 10% bleach between each dissection. Dissections were performed with bleach-washed microscissors and forceps, and bleached glassware. Adult beetles were placed in a -20° freezer for approximately 15 minutes and upon removal they were surface sterilized in 70% ethanol for 1 minute. They were then placed in a glass tray with approximately 20 mL of phosphate buffered saline (PBS). Elytra were

28 removed, as were wings. Microscissors were used to trim around the base of the spiracles and pulled away the tergites, exposing the gut, trachea, and fat body.

The prothorax was separated from the meso- and meta-thorax. The meso and meta-thorax were removed by tearing out the tergum, pulling away any muscle or fat body tissue, and then pulling away the remaining structure delicately by the legs. The prothorax was removed by cutting the pleural region on either side of the pronotum to cut it in half, and both pieces were gently pulled away. To prevent lumen contents from leaking out of the midgut and small delicate foregut, the head was not removed. Using the forceps to clamp the cuticle, I pulled away the respiratory tissue and fat body along with the gut. Forceps were used to pull nervous, circulatory and respiratory tissue, fat body, and Malpighian tubules from the gut. The remainder of the beetle’s abdomen was pulled away from the rectum. The whole gut, including the foregut enclosed by the head, was moved to another tray of approximately 15 mL of PBS.

Microscissors were used to separate the hindgut from the midgut (Figure B-1), and the midgut from the small foregut with the head.

Like adults, larvae were anaesthetized in a -20° freezer 15 minutes prior to dissection and surface sterilized in 70% ethanol prior to being placed in phosphate buffered saline solution. Incisions were made along the entire plural region, below the spiracles. A circum-occipital cut was made, thus leaving a dorsal and ventral portion of the integument, which could be pulled apart and away from the head. Circulatory and respiratory tissue, fat body, and Malpighian tubules were pulled away from the gut.

Remaining cuticle was cut away from the around the anus. The gut, with the head still

29 attached to the foregut, was transferred to a clean glass container filled with phosphate buffered saline solution. The gut was then cut into three sections: the paunch, the ileum, and the midgut (Figure B-2). After dissection, all gut sections were immediately stored in a freezer at -80°. The remains of adults and larvae not processed for extraction were stored in 100% ethanol. Given the partitioning, the adult guts amounted to 24 individual samples (12 midgut samples, and 12 hindgut samples), and the larvae amounted to 33 (11 midgut samples, 11 ileum samples, and 11 paunch samples). The remains of adult and larval specimens not used for extraction were saved as voucher specimens and deposited in the Clemson University Arthropod Collection.

Extraction and test amplification

Bacterial DNA from the gut sections of both adult and larval beetles were extracted using the DNeasy PowerSoil kit (QIAGEN). Given the large volume of liquid harbored by the larval gut sections and the potential to dilute reagents, these were first dehydrated in a vacuum centrifuge for 90 minutes. Both larval and adult gut sections were pulverized with sterile micropestles (Millipore Sigma) prior to extraction.

Extractions were carried out according to the protocol of the manufacturer, including suggested incubation steps. Extracts were stored in a freezer at -20°. Extracts were quantified using a Qubit 3.0 Fluorometer (Life Technologies). Samples with a DNA concentration of above 1 ng/μL were used.

PCRs were prepared for extracts using 16.875 microliters of nuclease-free water,

2.5 microliters of buffer, 2.5 microliters of dNTP mix, 1 microliter of forward and reverse

30 primer for the V4 hypervariable region, and 1 microliter of DNA sample. For larval midgut samples, successful amplification required 6 microliters of samples diluted to

1:100 of their original concentration. The settings used for the PCR were as follows: 94° initial denaturation for 3 minutes, (94° denaturation for 20 seconds, 50° annealing for 15 seconds, 72° extension for 5 minutes) x25 cycles, 72° final extension for 10 minutes, and

4° incubation. These test runs were carried out on a Mastercycler nexus gradient

(Eppendorf). Successful amplification was shown by the presence of a fluorescent band at the 400bp mark (as indicated by a ladder) and absence of primer-dimers. Foreguts of both adult and larval beetles were excluded from sequencing due to their small size, structural simplicity, and inability to yield bacterial DNA for sufficient amplification.

The total number of individual samples suitable for sequencing was 48 (4 larval midgut samples, 11 ileum samples, 10 paunch samples, 12 adult hindgut samples, and 11 adult midgut samples).

Gene amplification, Library prep, Normalization, and sequencing

Amplicon libraries were assembled using PCR to add sample-specific indexes and adapters to individual samples. For this we used 16.25 microliters of nuclease-free water,

4 microliters of buffer, 0.5 microliters of Phusion high fidelity DNA polymerase (New

England BioLabs), and 0.75 microliters of forward and reverse primer, respectively. Test runs were carried out on a C1000 Touch Thermal Cycler (Bio Rad) using Barcoded primers (each primer bearing a unique sequence that would enable the identification of the sample it comes from) for the amplification of the V4 hypervariable region to allow

31 single-step multiplexed sequencing and consequently reduce occurrence of chimeras

(Callahan et al. 2019).

To improve banding on larval midgut, adult midgut, and adult hindgut samples, 6 mL, 4 mL, and 4 mL of diluted (1:100) sample was used for the PCR runs, respectively.

The settings (which were changed to account for differences in reagent, primer, and sample volumes) that yielded successful PCR product, without primer-dimers are as follows: 98° initial denaturation for 3 minutes, (98° denaturation for 20 seconds, 61° annealing for 15 seconds, 72° extension for 5 minutes) x25 cycles, 72° final extension for

10 minutes, and 4° incubation. Negative controls containing primers but no DNA sample were included in the PCR runs to ensure contamination was not occurring. Denaturation cycle time was increased to achieve better banding in adult midgut samples. As larval midgut samples did not yield banding, or resulted in regular primer dimers, KAPA HiFi

DNA polymerase (KAPA Biosystems) was used instead. 6 of the 10 midgut samples did not amplify properly. To improve banding in these midgut samples the denaturation was increased to 30 cycles. All of the remaining sample types successfully amplified. Given the presence of inhibitors in some gut structures, 1:100 dilutions of certain samples

(primarily larval midgut) were performed.

A sample sheet containing the IDs of samples readied for the MiSeq run was prepared. Each sample ID corresponded to the specific index primer that it was amplified with. Library prep was completed successfully for most sample types. Several midgut samples, which were difficult to amplify in previous runs did not successfully amplify with the barcoded primers despite multiple attempts to improve amplification with the

32 addition of DMSO, BSA, alteration of annealing temperature, different cycle times, and sample dilutions. All samples that did not successfully amplify were omitted from the sequencing run. Consequently, only 4 out of the original 11 larval midgut samples and 11 of the original 10 adult midgut samples were kept.

Normalization was carried out with a SequelPrep Normalization kit

(ThermoFisher) according to the manufacturer’s protocol. After pooling, the samples were quantified. The DNA concentration in the first pool was calculated to be approximately 0.2 ng/μg. The DNA concentration of the second pool was calculated to be approximately 0.4 ng/μg. The sequencing run was shared with amplicons from unrelated samples, which were pooled along with the two normalized pools of larval samples.

Subsequently, a gel was performed on 20 microliters of the pooled sample. Fluorescence indicated banding as well as the absence of primer dimers.

Barcoded amplicons were sequenced on the MiSeq platform using the MS-102-

2003 MiSeq Reagent Kit v2 (500 cycle) (Illumina) according to the protocol of the manufacturer. From this, 251-bp paired-end reads were generated for analysis.

Data analysis

Analysis of genome sequences from C. nitida was carried out in QIIME 2 (Bolyen et al. 2019). Sequences, as paired-end reads, were first joined using deblur. Quality filtering, including chimera removal and other denoising was carried out with deblur

(Amir et al. 2017), and aligned using MAFFT (Katoh et al. 2002) and masked. Fastree

(Price et al. 2010) was used to produce an unrooted tree, which was then rooted with

33 midpoint. Taxonomy was assigned with sequences from the SILVA database (Quast et al.

2013), and a taxon bar plot was created and visualized on QIIME 2 view (Bolyen et al.

2019) (Figure B-18). Sequences were rarefied. To prevent the low frequency midgut samples of adult and larvae from being eliminated, the sampling depth was set at 1,000.

This still excluded 3 adult samples from analysis on QIIME 2. Sequence analysis for alpha and beta diversity were carried out on QIIME 2 and R (Bolyen et al. 2019).

Statistical Analyses

Alpha diversity was used to determine bacterial community differences between adults and larvae, gut regions, and sexes by testing for differences in phylogenetic diversity via Faith’s phylogenetic diversity, abundance via Chao1, and evenness via the

Shannon diversity index. Alpha diversity was calculated using Faith’s Phylogenetic diversity with a Kruskal-Wallis pairwise test. A metadata file containing the individual sample IDs as well as pertinent variables, including specimen life stage (larva or adult), gut type (midgut, hindgut, paunch, ileum), and sex (male or female) was prepared prior to analysis. Additionally, a reads manifest file containing the file path information for the paired-end reads was created. Figures were visualized on QIIME 2 view (Bolyen et al.

2019).

Beta diversity (via unweighted UniFrac) was used to test for community differences between adults and larvae, gut regions, and sexes by examining the taxonomic distinctiveness of bacteria in a given taxon and thus test for a core community shared by one sample type or another. A pairwise PERMANOVA was used to compare differences between the bacterial communities of adult and larval stages, general gut

34 types (midgut and hindgut), and specific gut types (adult midgut, larval ileum, etc.).

Figures for Beta diversity analysis were visualized on QIIME 2 view (Bolyen et al.

2019). PCoA plots were created to demonstrate grouping of samples based on taxonomic similarity of communities. Principle coordinate analysis (PCoA) plots were generated according to Bray-Curtis dissimilarity metrics and unweighted UniFrac phylogenetic distance metrics (Figure B-9, Figure B-10, Figure B-11, Figure B-12, Figure B-13, Figure

B-14) (Lozupone et al. 2011) and visualized on EMPeror (Vázquez-Baeza et al. 2013).

For both alpha and beta diversity, life stage, specific gut type, and general gut type were included as independent variables.

Alpha diversity metrics, including Chao1 and ACE, as well as the Simpson

Diversity index and Shannon diversity index were carried out and visualized on R (Figure

B-6, Figure B-7, Figure B-8). The metadata file, saved in the form of a .tsv file and the results of alpha and beta diversity analysis, saved in the form of .qza files, were imported to Rstudio. These were used to create a phyloseq object which could then be analyzed with the phyloseq package (McMurdie & Holmes 2013) on Rstudio. A feature table of taxa with associated abundances was imported to R and NMDS analyses were carried out and visualized (Figure B-15, Figure B-16). NMDS analysis was executed using the vegan package on R using Pairwise adonis (Martinez Arbizu 2017).

Results

Overview of sequencing results

35 A total of 629,144 reads were obtained from 48 sample. Clustering analysis at

97% similarity yielded a total of 3,634 bacterial and archaeal OTUs. These encompass 27 phyla, 144 families, and 705 species. The prokaryotic communities of both adult and larval C. nitida are dominated by bacteria of the phyla Firmicutes and Proteobacteria.

These phyla did not exceed 30% relative frequency in certain ileum samples, which in turn exhibited a greater abundance of other phyla in the Actinobacteria, Planctomycetes,

Acidobacteria, and Chloroflexi (Figure B-17). Archaea are present in comparatively greater abundance in the larvae where they are primarily found in the paunch region and are dominated by the Methanomethylophilaceae. Methanobrevibacter of the

Methanobrevibacteriacae is also present primarily in larval paunches, and totally absent from adult guts, as well as larval midguts and all but one ileum sample (Figure B-21).

In the comparatively taxon-rich larval gut samples, no OTUs exceeded a relative frequency of 30% (Figure B-18). In larval paunches, no taxa exceeded a relative frequency of 15% (Figure B-18). In the midgut, no taxa with a frequency of higher than

10% are present. In some ileum samples, certain taxa exceeded relative frequencies of above 20%. Taxa of relative frequencies of above 30% were found across adult samples, however. In some adult samples, certain bacterial OTUs dominated the community.

These include a species of Pantoea, which reaches a relative frequency of over 60% in the midgut of one individual (Figure B-20), while a species of Gluconobacter reaches a relative frequency of over 70% in another individual (Figure B-19).

Alpha diversity

36 The gut bacteria of adult and larval beetles showed a marked statistically significant difference. Larval beetles harbored a taxonomically richer community of bacterial and archaeal OTUs than adults, according to Faith’s PD (H: 28.324696, p

<0.001, q<0.001). Additionally, the taxa present in larvae were greatly distinct from those of adults (pseudo-F: 13.222026, p=.001, q=0.001)

Within adults, phylogenetic diversity of the midgut region was significantly lower from that of the hindgut (H: 12.595238, p<0.001, p<0.001) (Table A-1). This distinction in phylogenetic diversity was not observed for the three regions of the larval hindgut that were sampled. The midgut was not significantly more diverse than the ileum (H:

2.454545, p=0.117, q=0.147) or the paunch (H: 1.28, p=0.258, q=0.287) (Table A-1).

Larval paunches were not significantly more diverse than ileums (H:1.115702, p=0.290846) (Table A-1). In adult beetles, males and females did not harbor significant differences in the phylogenetic diversity of their gut bacteria (H: 0.46287 p=0.496, q=0.496) (Table A-1).

Using other Alpha diversity measures including Chao1, ACE, as well as measures of community evenness with the Shannon diversity index, Simpson diversity index and

Fisher’s alpha, plots demonstrated a pattern similar to the one demonstrated by Faith’s

PD (Figure B-6, Figure B-7, Figure B-8). That is, larval midgut, ileum, and paunch samples were more diverse than adults. Moreover, paunch and midgut samples showed consistently high diversity, whereas ileum samples varied.

Beta diversity

37 There was significant taxonomic distinction between the adult midgut and adult hindgut (pseudo-F: 4.436725, p-value=0.001, q-value: 0.001) (Table A-2). There was not a trend of taxonomic distinctiveness observed between the guts of male and female adult beetles (pseudo-F: 0.840, p=0.603, q=0.603) (Table A-2).

All regions of the larval hindgut differed significantly from the regions of the adult gut. The larval midgut did not show significant taxonomic distinction from the larval ileum (pseudo-F: 1.585, p=0.102, q=0.102) (Table A-2). However, the bacteria of the paunch showed a marked taxonomic distinction from the midgut (pseudo-F: 9.316, p=0.003, q=0.0033) and the ileum (pseudo-F: 8.83, p=0.001, q=0.001) (Table A-2).

Unweighted UniFrac PCoA plots showed a distinctively grouped paunch community, as well as a larval midgut community that more closely resembles that of the larval ileum (Figure B-10). However, both models indicated an ileum community that differs. Unweighted UniFrac PCoA plots also show distinctive adult and larval communities (Figure B-9, Figure-B-10) as well as a distinction between adult and larval gut communities (Figure B-10). Bray-Curtis PCoA plots demonstrated a marked separation between adult and larval stages, as well as certain gut regions, namely between the adults and larvae and the hindgut paunch and other regions of the larval gut (Figure

B-12, Figure B-13). Additionally, midgut samples group more closely with hindgut ileum samples from the same individual. No distinction is observed between male and female gut bacterial communities (Figure B-14).

Community distinctiveness was corroborated by NMDS analysis using Bray-

Curtis dissimilarity, which demonstrated a statistically significant difference between the

38 bacterial communities of larvae and those of adults (p=0.001) (Figure B-15). Looking at individual gut sections, there was distinctive partitioning (Figure B-16). Analysis revealed a statistically significant difference between paunches (p=0.01, p=0.01) and larval midguts and ileum. Moreover, there was not a significant difference between ileum and larval midgut samples (p=0.51). Interestingly, there is some overlap between paunch and adult hindgut samples. Moreover, the adult midgut differed from significantly from the adult hindgut (p=0.01).

Discussion

Overview

This study characterized the gut community of adult and larval members of the cetoniine scarab beetle C. nitida. This study revealed that this beetle not only possesses a taxonomically rich community of bacteria, but also that this richness varies markedly between life stages, with larvae possessing far more OTUs in their digestive tract.

Additionally, this study shows that the uniqueness of OTUs differs markedly not only between adult beetles and larvae, but also between certain regions of the adult and larval guts. Such patterns are observed in other animals, including insects, and suggest settlement of bacterial communities in the digestive tract is not merely a consequence of ingestion, but rather that underlying patterns relating to the host system or microbial community determine presence or absence of bacteria. A better understanding of these patterns may be used to make predictions not only about gut communities in other organisms, but also about potential function.

39 While mirroring trends observed in other insects, the differences in taxon richness and community partitioning based on gut region and life stage in C. nitida is emblematic of the scarab gut microbiome. However, the lack of significant difference in taxon richness between midgut and hindgut regions of larvae differs from that of other scarabs, which either have a more taxon-rich hindgut (Andert et al. 2010) or midgut (Arias-

Cordero et al. 2012). Additionally, the absence of a significant difference between male and female beetles is markedly different from other scarab beetles, notably the dung beetles whose female gut communities share more bacterial taxa with the larvae than those of the males (Shukla et al. 2016). Parental care has been demonstrated in at least one cetoniine, Dicronocephalus wallichii (Kojima & Lin 2018). However, a microbial element to the rearing of offspring has not yet been demonstrated in Dicronocephalus or in flower chafers like Cotinis whose females, once fertilized, oviposit in suitable substrate and do not provide a specially crafted food source inoculated with their own gut bacteria.

Given the presence of unshared taxa in larvae and adults, it can be inferred than many of these organisms are environmentally acquired. It is likely that the substrate chosen for larval alimentation also contains bacteria, archaea, and possibly eukaryotes like fungi that assist in the digestive processes. In terms of adult acquisition, the colonization of various bacteria may not be unlike that of fungal colonization (Vishniac & Johnson 1990), in which the “clean” adult garners symbionts through environmental exposure (ingestion of food substrate, contact with mates, crawling and flying around in various environments).

Whether or not the shared taxa are a consequence of vertical transmission cannot be inferred from the present data. Additional tests are needed to determine the role parental

40 bacterial microbiome has on that of larvae and whether or not sexual interactions facilitate significant transmission.

The crossover between taxa found in the larval ileum and midgut may be indicative of human error in regard to dissection approach and gut morphology—the junction between the midgut and ileum is quite large and permissive toward “spillover” even when forceps are used to prevent gut contents from transferring. Although the gut is filled with large pieces of plant material that does not move readily, lumen fluid can easily shift from one portion of the gut to another during the process of dissection. On the other hand, the wide passage of the anterior ileum and the physiological similarities between this region and the anterior midgut may naturally facilitate a community crossover. The absence of a significant presence of shared organisms in the paunch, which was more morphologically delimited from the other two segments and thus less prone to disturbance during the dissection process further bolsters this hypothetical cause.

It is also possible that the ileum simply shares taxa by virtue of a number of physiological and morphological conditions of the beetle gut, including but not limited to condition of ingested substrate, gut pH, oxygen level, host digestive enzymes, and host immune response. Additionally, presence or absence of certain taxa may be influenced by the microbial community itself,

The poor clustering of ileum samples suggests a less consistent community of organisms compared to that of the paunch. The hypothesis that more than dissection error is at play is supported by the clustering of ileum samples with midguts from the same individual, compared to ileums from other individuals. This less stringent grouping of

41 communities may indicate a far more transient community in the midgut and ileum.

Additionally, it is possible that hindgut paunch physiology and morphology favors particular taxa over others. Oxygen content can influence a community, for example

(Chouaia et al. 2019). Regarding its morphological uniqueness, the paunch is separated from the ileum by a greatly constricted length of hindgut (Figure B-4). It is also internally lined with papillae, the bases of which can be observed through the translucent colon.

(Figure B-5).

Additional factors which were not considered for the present study include age of the sampled organisms. The gut communities of some scarabs change across instars

(Alonso-Pernas et al. 2017, Chouaia et al. 2019, and the provenance of taxa that appear to be present across one region or another may be influenced by shifts in diet and physiology. This may even be true for adults, and while metamorphosis ceases after the imago stage, feeding behavior as well as certain physiological shifts incumbent on maturation may alter the community profile of bacteria.

The distinctiveness of bacterial communities according to alpha and beta diversity measures is not unfounded, and other insects including scarab beetles exhibit such patterns of bacterial diversity. This is also the case for other animals (Donaldson et al.

2016). There is also some taxonomic similarity on the genus and family level between the bacteria of the gut C. nitida and the digestive systems of animals with similar diets as well as pronounced compartmentalization of the digestive system (e.g. termites, ruminant mammals) (Huang et al. 2010). Whether or not this relatedness reflects on functional similarity between these organisms in the guts of their respective hosts is tenuous,

42 although the identification of such similarities provides a basis for future hypothesis testing.

Bacteria of the larval ileum and midgut

Strikingly, the differential abundance between bacteria in the larval ileums occurs at the level of phylum. This sharp distinction at such a basal taxonomic level shows that a diverse array of organisms are capable of colonizing the midgut and ileum.

Unfortunately, the absence of corresponding midgut samples for the remainder of ileum samples prevented a determination of whether this trend of midgut-ileum similarity was present elsewhere. A larger sample size of midguts and ileums would be crucial to lending further support to this.

In some scarabs, reduction of ferric compounds is implicated in the midgut—a process mediated by a species of Bacillus in one cetoniine (Hobbie et al. 2012). In C. nitida a species of Bacillus appeared in relatively great abundance in the midgut and ileum. It is possible a similar iron-reducing function is being performed at the efforts of this organism. The presence of relatives of other iron-reducing bacteria in the mid- and hindgut ileums of larvae point toward this hypothetical function. For example, members of the genus Geobacter known for the capacity to reduce iron (Mahadevan et al. 2006) are present. Growth assay and experimental manipulation would be necessary to offer support for this hypothesis. Bacteria of the genus Turicibacter, in C. nitida found in some ileum and paunch samples have also been reported from the guts of another cetoniine and melolonthine beetles (Egert et al. 2003, Egert et al. 2005, Arias-Cordero et al. 2012). In

43 the midgut of one scarab, a species of Turicibacter produces lactate (Arias-Cordero et al.

2012). That this organism was found in the ileum of only some of the samples further confounds hypotheses about potential function through the comparison of similar habitat alone.

The presence of Rhizobiales bacteria in the midgut and hindgut ileum regions correlates with the beetle’s diet. These organisms, and the Rhizobiales as a whole are associated with soil environments (Garrido-Oter et al. 2018) and frequently occur as nitrogen-fixing root symbionts of legumes. Whether the specific organisms found in the gut maintain this faculty in their Cotinis hosts, as is the case in some other insects

(Morales-Jiménez et al. 2009, Russell et al. 2009) is unknown, however.

The taxa found in the gut of Cotinis nitida do not merely resemble those found in related insects and their habitats. The Planctomycetes, a phylum abundant in the ileum and midgut has affinities with the digestive system of termites (Köhler et al. 2008). It is possible that the functional role provided by these organisms, if they do indeed provide a service is similar to that proposed by similar bacteria in termites—degradation of microbial polymers (Richards et al. 2017).

Distinctive bacteria of the larval paunch

The hindgut of scarabs is where the bulk of plant fiber digestion occurs, through the machinations of a robust community of microroganisms including bacteria (Huang et al. 2010). The bacteria found localized in the paunch at reasonable abundance follow a

44 trend observed in other “hindgut fermenters” of the insect world, including both

Holometabola like the various scarabs and Hemimetabola like termites. Strikingly, many of the organisms associated with the paunches of other scarabs and termites are also found in Cotinis. The genus Alistipes, in the family Rikenellaceae of Bacteroidetes is also known from the guts of those groups (Andert et al. 2010, Schnorr et al. 2019). This is also the case for OTUs belonging to Ruminococcaceae and Christensenellaceae (Huang et al., 2013,

Chouia et al. 2019, Schnorr et al. 2019). Tannerella of the C. nitida paunch is also found in termite guts (Makonde et al. 2015) This highlights the differing strategies in which these organisms are transmitted—in termites both juveniles and adults possess these communities and they are inoculated via trophallaxis. Scarab adults apparently do not harbor these particular taxa, and thus they are likely environmentally acquired and settle in the hindgut paunch. Further work must examine the soil community of the larval habitat to determine specific presence. The hypothetical functional role of these organisms is not limited to insects. The strain Ruminococcaceae UCG-010 of the larval paunch has been reported from deer guts (Li et al. 2017).

Some Rhizobiales appear exclusively in the hindgut paunch, as in the case of an

OTU assigned to the genus Bradyrhizobium, which harbors nitrogen-fixing members

(Bünger et al. 2018). This partitioning may be explained by this organism’s role in nitrogen fixation in the soil. Indeed, nitrogen-fixing bacteria are known from the paunches of other scarabs, and Cotinis may be similarly benefitting from nitrogen fixers.

Paunch-wide organisms related to nitrogen fixers also include a member of the genus

Diplosphaera (Wertz et al. 2012).

45 Despite the apparent partitioning of bacterial communities, varied abundances, and uniqueness of taxa in the paunch, specialization is not necessarily the sole determinant of the condition. Indeed, organisms related to those generally found in the guts of other animals are present. These include, but are not limited to

Christensenellaceae, Lachnoclostridium, Ruminococcaceae, and an OTU of the “Termite planctomycete cluster.” That bacteria are capable of capitalizing on such a varied number of animal habitats is intriguing in a number of ways. Firstly, it demonstrates the well- known versatility of bacteria in their capacity to settle and take advantage of highly varied energy sources. Additionally, however, it raises the question of strain. While these organisms are indeed related, how much do the shared bacteria of humans, cattle, termites, and cockroaches actually resemble each other in situ? Finer taxonomic analysis is necessary to breach this question, in addition to environmental manipulation and cross inoculation.

The hindgut paunch of other scarabs is an anaerobic environment (Huang et al.

2010). The most abundant taxa found in this region in C. nitida are related to organisms that are obligately anaerobic. Even among less abundant paunch taxa we observe putatively obligate anaerobes like a member of the genus Pelospora, which contains species that are known fermenters of glutarate (Matthies et al. 2000). With one exception

(an adult male), Archaea were only found in the hindgut paunch of larvae. The most abundant of these is a species of Methanobrevibacter—a methanogenic organism and obligately anaerobic (Enzmann et al. 2018). The scarab paunch along with that of other insects with similar modifications is considered to be a largely anaerobic environment

46 (Arias-Cordero et al. 2012), a condition further suggested in Cotinis. In addition to oxygen levels and pH, other chemical conditions may be at play in the hindgut paunch, but this analysis does not permit more than cursory speculation.

Bacteria of the midgut and hindgut of adult C. nitida

The diet of adult beetles may explain the lack of a consistent gut community, although it leaves a lot of speculation about midgut-hindgut distinctions. The dearth of diversity of bacterial communities in adult beetles and the high abundance of select taxa may also be explained by diet. The most abundant taxa found across adults are related to organisms known to be common in the guts of other animals as well as environmental samples (Dubin & Pamer 2017). Enterococcus, for example, is a common facultatively anaerobic generalist taxon with representatives known from the digestive systems of other animals, including humans (Dubin & Parner 2017). That the hindgut communities differ from the midguts in terms of richness is intriguing considering the great abundance of some taxa in adults. That the community profile can shift so rapidly in a morphologically less compartmentalized gut suggests various physiological conditions or interspecific competition may be at play.

Not all taxa found in adult guts are found across a wide range of host taxa. Two species of Gilliamella, one of which was highly abundant in the guts of adults is particularly fascinating, as it is a genus commonly found in the pollen and nectar-feeding honeybees and bumblebees, comprising their core microbiome. Unlike these organisms, however, the genus is not found consistently across adults. This may be indicative of its non-social transmission in Cotinis. The presence of a second species of Gilliamella,

47 highly abundant in one individual but of relatively low abundance or absent in others gives credence to this hypothesis of irregular transmission as well. That is, without a pool of colony members to maintain this taxon as a core OTU, its presence is determined incidentally as a result of individual feeding habits.

Crossover taxa in the guts of larvae and adults

Certain taxa are shared by both adult and larval C. nitida. Some taxa are found both in the paunches of larvae and the hindguts of adults. These include some of the same species of Ruminococcaceae, and the RsK70 Termite group of the Deltaproteobacteria,

Disulfovibrio, certain Dysgonomonadaceae, Mucispirillum, and Rhodocyclaceae. The most intriguing of these crossover organisms are those with associations with necessary catabolic function. For example, some strains of Ruminococcaceae are found in the hindgut paunch exclusively, but not across all samples and at a much lower abundance

(e.g. Papillibacter and Ruminococcaceae UCG-014).

It is possible that these more dominant organisms are better at thriving in the hindgut paunch, better at excluding taxa with similar catabolic capabilities, or their colonization was incidental, and these select taxa were merely able to get a foothold first.

That certain groups like the Ruminococcaceae were found in both larval and adult hindguts may be more indicative of the ubiquity of such organisms, their versatility in colonizing varied habitats, and the specific biochemical and morphological qualities shared by larval and adult hindguts rather than transmission from adults to their young.

48 An OTU of the family Acidaminococcaceae curiously appears in both the hindguts and midguts of adults and the paunches of larvae in abundance but has only a scant presence in the midgut and ileums of larvae. This is an organism closely associated with cellulolytic bacteria in ruminants, particularly members of the family

Ruminococcaceae—a group that is also present in adult hind and midguts and larval paunches, but not in larval midguts or ileums. Members of this family are succinate consumers and propionate producers (Mulder et al. 2017). This putative function based on taxonomy is supported by a similar occurrence in the gut of the cetoniine Pachnoda marginata which yields low propionate production in its midgut, but a high level of propionate production in its hindgut (Lemke et al. 2003).

Additional questions

The dearth of quality sequences garnered from the midgut raises important questions not only about the community but about the ability to standardize approaches for sampling the microbiome. This study did not experiment thoroughly with different extraction and amplification procedures, but my observations suggest that a number of factors may have mitigated either successful bacterial and archaeal DNA extraction or subsequent PCR. The former includes insufficient denaturing or pulverizing of gut tissue, and the latter includes PCR inhibitors produced endogenously by the beetle or resulting from its diet and insufficient denaturation of gut tissue harboring bacteria. The small sample size and small sample range of beetles in this study may confound its findings as well. The question of whether or not patterns observed here vary across the range of C. nitida should be considered in the future.

49 The potential for external contamination of samples is present during all parts of specimen preparation, and despite precautions, some of the bacteria found in samples may be an artifact of sample processing. The addition of negative controls consisting of samples from the processing environment, including the glassware, dissection tools, and dissection fluid (PBS) in sequencing would shed some light on potential contaminants.

Moreover, the potential of contamination via other parts of the host system, including organs closely associated with the digestive tract like the Malpighian tubules and trachea, could be addressed with sequencing the bacteria communities of these organs. A comparison between whole specimen microbiome and that of dissected guts may also provide a useful means by which to determine the efficacy of the dissection approach for isolating specific bacterial communities.

The general lack of taxa delimited to genus is a reminder of the need to characterize bacterial communities and describe bacterial taxa. Many questions arise from the present survey that cannot be soundly addressed with the garnered data. Further comparison of the OTUs of Cotinis across a wider sampling range is necessary to determine the consistency of the “core taxa” found in the small, highly localized sample size used in this study. Further analyses of strain and prevalence across larger populations are necessary to test many of the hypotheses emerging from this survey.

Conclusion

The concept of the holobiont can greatly enhance our understanding of insect physiology, ecology, and evolution. The sheer diversity of insects and the resulting

50 differences in physiological systems and ecological function, in addition to individual variance, effectively ensure that precise relationship between presence and function of gut bacteria cannot be elucidated from phylogenetic relationship alone. Just as in humans, our knowledge of bacterial relationships with insects can begin with a non-culture-based approach—a broad view of an ecosystem—that spans multiple individuals and life stages, eventually moving on to different populations, and related taxa.

For the diverse scarab beetles, the green June beetle, C. nitida, is one of many interesting taxa to begin this exercise of connecting the dots. The taxon richness of larvae and adults is markedly different, with the latter harboring a more diverse bacterial community overall. Additionally, partitioning of bacterial colonies does occur between life stages, although not entirely so in specific regions of the digestive system. Rather, the larval paunch harbors several bacterial and archaeal taxa that are distinctive to it whereas the midgut and ileum are generally similar. The distinctiveness of the paunch bacterial community is implicated a potential functional role by the family- and genus-level relatedness of its members to known plant fiber degraders, including those found in insects with similar diets as well as gut morphology and physiology, and the role of the paunch as a source of bacteria-mediated plant-fiber breakdown in other scarab species.

Partitioning in terms of richness and taxonomic distinctiveness is also present in adults. A

“core microbiome” is suggested in larvae, but “core taxa” are only found in certain regions of the gut. The presence of taxa associated with certain diets and gut morphology and physiology in other insects points toward potential microbial assistance manifest through a diverse consortium of taxa.

51 Possessing ecologically distinctive life stages, relatively few close relatives within its genus, and a wide geographic distribution, C. nitida permits future studies in community variation in the context of symbiotic services and ecological partitioning of symbiotes. Further efforts to culture and experimentally alter the bacterial community and its scarab host will aid in our understanding of its myriad of potential services as well as yield fertile ground for future application.

52 APPENDICES

53 Appendix A

Tables

Table A-1: Kruskal-Wallis pairwise comparison of bacterial communities of the gut of C. nitida

Table A-2: Pairwise Permanova of bacterial communities of the gut of C. nitida

54 Appendix B

Figures

Figure B-1: The gut of an adult C. nitida, head to the right.

Figure B-2: The gut of a larval C. nitida, head to the right.

55 Figure B-3: Detail of the midgut of the larval C. nitida. The anterior portion of the gut midgut is facing the right.

Figure B-4: Detail of the ileum of the larval C. nitida. The ileum is located between the row of fingerlike gastric caeca and the greatly expanded hindgut paunch to the bottom left.

56 Figure B-5: Detail of the exterior paunch of the larval hindgut C. nitida, showing the off-white bases of papillae.

57 Figure B-6: Alpha diversity measures of adult and larval C. nitida according to gut region. Larval gut samples demonstrate higher abundance of observed OTUs (Observed, Chao1, ACE), and greater evenness (Shannon, Simpson) than adults. Additionally, higher alpha diversity is represented among larval gut regions compared to adult gut regions based on Fisher’s alpha.

58 Figure B-7: Alpha diversity of the gut of two life stages of C. nitida according to the Shannon diversity index and Chao1. Larvae demonstrate greater evenness (Shannon) and abundance of taxa (Chao1) than adults.

59 Figure B-8: Alpha diversity of the gut regions of C. nitida according to the Shannon diversity index and Chao1. The larval midgut and larvae paunch display greater evenness (Shannon) and abundance of taxa (Chao1) compared to the larval ileum and adult gut regions.

60 Figure B-9: Beta diversity of the gut of two life stages of C. nitida according to unweighted UniFrac. The larval and adult communities are taxonomically distinct from one another, demonstrating a prominent clustering of sample sites (adult and larval gut regions) based on life stage.

61 Figure B-10: Beta diversity of the gut regions of C. nitida according to unweighted UniFrac. The larval and adult communities are taxonomically distinct from one another and demonstrate a prominent clustering of sample sites (adult and larval gut sections) based on gut region.

62 Figure B-11: Beta diversity of the gut of the sexes of C. nitida according to unweighted UniFrac. The larval and adult communities are taxonomically distinct from one another, but the adult beetles (the only individuals that were sexed) show no distinctive clustering based on sex.

63 Figure B-12: Beta diversity of the gut of two life stages of C. nitida according to Bray-Curtis dissimilarity. The larval and adult communities are taxonomically distinct from one another, demonstrating a prominent clustering of sample sites (adult and larval gut regions) based on life stage.

64 Figure B-13: Beta diversity of the gut regions of C. nitida according to Bray-Curtis dissimilarity. The larval and adult communities are taxonomically distinct from one another and demonstrate a prominent clustering of sample sites (adult and larval gut sections) based on gut region.

65 Figure B-14: Beta diversity of the gut of the sexes of C. nitida according to Bray-Curtis dissimilarity. The larval and adult communities are taxonomically distinct from one another, but the adult beetles (the only individuals that were sexed) show no distinctive clustering based on sex.

66 Figure B-15: Beta diversity of the gut of two life stages of C. nitida according to NMDS (Bray-Curtis dissimilarity). The larval (Lmidgut, ileum, Paunch) and adult (Amidgut, Hindgut) communities are taxonomically distinct from one another and demonstrate a prominent clustering of sample sites (adult and larval gut sections) based on sex.

67 Figure B-16: Beta diversity of the gut of two life stages of C. nitida according to NMDS (Bray-Curtis dissimilarity). The larval (Lmidgut, ileum, Paunch) and adult (Amidgut, Hindgut) communities are taxonomically distinct from one another and demonstrate a prominent clustering of sample sites (adult and larval gut sections) based on gut region.

68 Figure B-17: Relative frequency of bacterial phyla in the gut of C. nitida according to gut region. The order of the key corresponds to order of taxon bars in each column—top to bottom).

69 Figure B-18: Relative frequency of bacterial OTUs in the gut of C. nitida according to gut region. Each colored bar per column corresponds to a different OUT for each sample.

70 Figure B-19: Relative frequency of Gluconobacter (in purple) in the gut of C. nitida according to gut region.

Figure B-20: Relative frequency of Pantoea (in purple) in the gut of C. nitida according to gut region.

71 Figure B-21: Relative frequency of Methanobrevibacter (in pink and green) in the gut of C. nitida according to gut region.

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