Decoding symbioses in grain pest beetles Ecology, evolutionary history, and specificity of symbiont-mediated nutrient supplementation in Silvanidae and Bostrichidae (Coleoptera) Dissertation Zur Erlangung des Grades Doktor der Naturwissenschaften Am Fachbereich Biologie Der Johannes Gutenberg-Universität Mainz Julian Simon Thilo Kiefer geb. am 18.06.1989 in Freiburg im Breisgau Mainz, 2022 Dekan: Prof. Dr. Eckhard Thines 1. Gutachten: Prof. Dr. Martin Kaltenpoth 2. Gutachten: Prof. Dr. Susanne Foitzik Tag der mündlichen Prüfung: 20.01.2023 „Do. Or do not. There is no try. “ Master Yoda, Star Wars: The Empire Strikes Back Summary Each individual organism interacts with at least one or more other species. When these relationships between two (or more) species are close over the long term, they are called symbioses. Symbiosis is considered a key driver of evolution, as it drives the development of morphological adaptations and new capabilities. A particular form of symbiosis is found between insects, especially beetles, where the symbiosis with bacteria has enabled them to access new habitats and food sources that they would not have had access to without their symbiotic partners. One example of challenging environments are grain storages which are by nature very dry habitats and at the same time the grain only offers a diet poor in nutrients - and yet beetles of multiple families, including Silvanidae and Bostrichidae have independently invaded this habitat. At the same time, beetles from both families are associated with symbionts of the same clade of Flavobacteria, which made it possible for them to spread around the world as pests in grain and wood. In my work, I used various molecular biology methods as well as experimental approaches to study the endosymbiont of the sawtoothed grain beetle Oryzaephilus surinamensis from the family Silvanidae. Via the shikimate pathway, the symbiont Shikimatogenerans silvanidophilus (Bacteria: Bacteroidota) provides precursors for the amino acid tyrosine, which the host requires for its cuticle synthesis. After contact with glyphosate, which specifically inhibits an enzyme of the shikimate pathway, Shikimatogenerans can no longer supply its host with this required nutrient. As a result, the symbiont titer decreases, or the symbiont is completely lost. The same effect is observed when the host has direct access to tyrosine through its diet. In addition to Shikimatogenerans, O. surinamensis is also infected with the Wolbachia strain wSur. Wolbachia is commonly known to manipulate the reproduction of its host. In O. surinamensis, wSur causes cytoplasmic incompatibility that results in the development of less viable embryos derived from spermatozoa of wSur-negative ♂- and oocytes of wSur-positive ♀-beetles. The genome of wSur also contains homologues of a gene responsible for male-killing in various insects. However, in grain storage populations of O. surinamensis, the gene sequence underwent a frameshift mutation. This mutation causes a premature translation stop and the loss of the protein’s functional domain, possibly contributing to the widespread colonisation of grain stores. i My analysis of 29 species from the family Bostrichidae shows that the entire family is consistently associated with Shikimatogenerans bostrichidophilus. In addition, only species of the genera Dinoderinae and Lyctinae are associated with Bostrichicola ureolyticus (Bacteria: Bacteroidota). While Shikimatogenerans provides the host with the precursor to tyrosine via the shikimate pathway, Bostrichicola supports its host by providing additional amino acids as well as recycling nitrogen. A comparison of host and symbiont phylogenies reveals a high degree of co- cladogenesis. One peculiarity stood out in this study: the endosymbiont S. bostrichidophilus of the large grain borer Prostephanus truncatus is divided into three strains in its host. The genes for the shikimate pathway and also the ribosomal proteins are thereby encoded by the three different strains, which complement each other but also exhibit a considerable degree of redundancy in encoded functions. The results of this work highlight the importance of symbioses, in which bacterial symbionts support the biosynthesis of the cuticle in insects by supplying tyrosine. Furthermore, it provides insight into the evolutionary processes of these communities, which have existed for millions of years and have led to mutual dependence between symbiont and host. As a result, contact with substances such as glyphosate could cause lasting damage to the symbiosis and even results in its collapse – an alarming scenario in view of the recent decline in insect populations. ii Zusammenfassung Jeder einzelne Organismus interagiert mit mindestens einer oder mehreren anderen Arten. Wenn diese Beziehungen zwischen zwei (oder mehreren) Arten langfristig eng sind, wird sie als Symbiosen bezeichnet. Die Symbiose gilt als eine der wichtigsten Triebkräfte der Evolution, da sie die Entwicklung morphologische Anpassungen und neuer Fähigkeiten vorantreibt. Eine besondere Form der Symbiose findet sich zwischen Insekten, insbesondere Käfern, wo die Symbiose mit Bakterien ihnen den Zugang zu neuen Lebensräumen und Nahrungsquellen ermöglicht hat, zu denen sie ohne ihre symbiotischen Partner keinen Zugang gehabt hätten. Von Menschen eingerichtete Getreidelager sind von Natur aus sehr trockene Lebensräume und gleichzeitig bietet das Getreide auch nur eine sehr einseitige Nahrungsgrundlage – und dennoch wurden sie unter anderem von Käfern der Familie Silvanidae und Bostrichidae unabhängig voneinander für sich als Habitat erschlossen. Voraussetzung dafür war, dass Käfer beider Familien mit Symbionten aus der gleichen Gruppe der Flavobakterien eine Verbindung eingegangen sind, die es ihnen erst ermöglichte, sich weltweit als Schädlinge in Getreide und Holz zu verbreiten. In meiner Arbeit habe ich mit verschiedenen molekularbiologischen Methoden sowie experimentellen Ansätzen den Endosymbionten des Getreideplattkäfers Oryzaephilus surinamensis aus der Familie der Silvanidae untersucht. Über den Shikimatweg liefert der Symbiont Shikimatogenerans silvanidophilus (Bacteria: Bacteroidota) Vorstufen für die Aminosäure Tyrosin, welche der Wirt für seine Kutikulasynthese benötigt. Nach Kontakt mit Glyphosat, welches gezielt ein Enzym des Shikimatwegs inhibiert, kann Shikimatogenerans seinen Wirt nicht mehr mit diesem benötigten Nährstoff versorgen. Infolgedessen sinkt der Symbiontentiter ab oder der Symbiont geht vollständig verloren. Der gleiche Effekt ist zu beobachten, wenn der Wirt direkten Zugang zu Tyrosin über seine Nahrung hat. Neben Shikimatogenerans ist O. surinamensis auch mit dem Wolbachia-Stamm wSur infiziert. Wolbachia ist allgemein dafür bekannt die Reproduktion seines Wirts zu manipulieren. In O. surinamensis verursacht wSur zytoplasmatische Inkompatibilität, die zur Entwicklung weniger lebensfähiger Embryonen führt, die aus den Spermien von wSur negativen ♂- und Eizellen von wSur positiven ♀-Käfern hervorgegangen sind. Des Weiteren codiert das Genom von wSur für ein Gen, welches für Male-Killing in diversen iii Insekten verantwortlich ist. In Getreidelager-Populationen von O. surinamensis erfuhr die Gensequenz allerdings eine Frameshift-Mutation. Durch diese Mutation bricht die Translation zum Protein frühzeitig ab, dieses verliert seine funktionale Domäne und hat damit möglicherweise zur weit verbreiteten Besiedlung der Getreidelager beigetragen. Meine Analyse von 29 Spezies aus der Familie der Bostrichidae konnte aufzeigen, dass die gesamte Familie durchweg mit S. bostrichidophilus assoziiert ist. Zusätzlich besitzen nur Spezies der Genera Dinoderinae und Lyctinae den zweiten Endosymbionten Bostrichicola ureolyticus (Bacteria: Bacteroidota). Während Shikimatogenerans den Wirt auch hier über den Shikimatweg mit der Vorstufe zu Tyrosin versorgt, unterstützt Bostrichicola zusätzlich seinen Wirt, in dem er weitere Aminosäuren bereitstellen sowie Stickstoff recyceln kann. Eine Gegenüberstellung der Phylogenie von Wirt und Symbiont zeigt zudem ein hohes Maß an Co-Kladogenese. Eine Besonderheit ist bei dieser Untersuchung aufgefallen: der Endosymbiont S. bostrichidophilus des großen Kornbohrers Prostephanus truncatus ist in seinem Wirt in drei Stämme aufgeteilt. Die Gene für den Shikimatweg und auch die ribosomalen Proteine werden dabei von den drei verschiedenen Stämmen kodiert, die sich gegenseitig ergänzen, aber auch ein beträchtliches Maß an Redundanz in den kodierten Funktionen aufweisen. Die Ergebnisse dieser Arbeit unterstreichen die Bedeutung von Symbiosen, bei denen bakterielle Symbionten den Aufbau der Kutikula bei Insekten durch die Versorgung von Tyrosin unterstützten. Des Weiteren gibt sie Einsicht in die evolutionären Prozesse dieser seit Millionen von Jahren bestehende Lebensgemeinschaften, welche zu einer gegenseitigen Abhängigkeit zwischen Symbiont und Wirt geführt hat. Dies hat dazu geführt, dass der Kontakt zu Substanzen wie z.B. Glyphosat die Symbiose nachhaltig schädigen und gar zum Zusammenbruch bringen könnte - mit Hinblick auf den jüngsten Rückgang der Insektenpopulationen ein alarmierendes Szenario. iv Contents List of publications ................................................................................................................................... 1 Chapter 1 General Introduction ............................................................................................................ 3 1.1 Symbioses ...................................................................................................................................... 3 1.2 Symbioses in insects .................................................................................................................... 3 1.3 Ecology of grain pest beetles ...................................................................................................... 6 1.4 Cuticle formation ........................................................................................................................ 7 1.5 Bacteroidota symbionts .............................................................................................................. 9 1.6 Thesis outline ............................................................................................................................. 12 1.7 References ................................................................................................................................... 13 Chapter 2 Inhibition of a nutritional endosymbiont by glyphosate abolishes mutualistic benefit on cuticle synthesis in Oryzaephilus surinamensis ................................................................................. 17 2.1 Abstract ....................................................................................................................................... 18 2.2 Introduction ............................................................................................................................... 19 2.3 Results ........................................................................................................................................ 22 2.3.1 Symbiont genome is extremely reduced and GC poor ............................................... 22 2.3.2 Candidatus Shikimatogenerans silvanidophilus OSUR encodes glycolysis and shikimate pathways .......................................................................................................................... 25 2.3.3 Amino acid titers are influenced by symbiont presence............................................ 27 2.3.4 The symbiont’s shikimate pathway is sensitive to glyphosate and its inhibition results in an aposymbiotic phenotype ........................................................................................... 29 2.4 Discussion .................................................................................................................................. 33 2.5 Material & Methods ................................................................................................................ 37 2.5.1 Insect cultures ................................................................................................................... 37 2.5.2 Elimination of O. surinamensis symbionts .................................................................... 37 2.5.3 Symbiont genome sequencing, assembly, and annotation ....................................... 37 2.5.4 Phylogenetic analyses ...................................................................................................... 38 2.5.5 Comparison of bacteria ................................................................................................... 39 2.5.6 Glyphosate and aromatic amino acid supplementation ............................................ 39 2.5.7 Quantitative PCR .............................................................................................................. 41 2.5.8 Analysis of cuticle traits .................................................................................................. 41 2.5.9 Amino acid extraction .................................................................................................... 42 2.5.10 Derivatisation of amino acids ........................................................................................ 43 2.5.11 Amino acid analysis with gas chromatography-mass spectrometry (GC-MS) ..... 43 2.5.12 Data availability ............................................................................................................... 45 2.6 Data Accessibility Statement ................................................................................................. 46 2.7 Acknowledgments .................................................................................................................... 46 2.8 Contributions ........................................................................................................................... 46 2.9 References .................................................................................................................................. 47 2.10 Supplementary Information ................................................................................................... 52 Chapter 3 Wolbachia causes cytoplasmic incompatibility, but not male-killing in a grain pest beetle ........................................................................................................................................................ 57 3.1 Abstract ...................................................................................................................................... 58 3.2 Introduction .............................................................................................................................. 59 3.3 Material & Methods ................................................................................................................ 62 3.3.1 Insect cultures ................................................................................................................... 62 3.3.2 Elimination of O. surinamensis symbionts .................................................................... 62 3.3.3 Quantitative PCR ............................................................................................................. 62 3.3.4 Fluorescence in situ hybridisation .................................................................................. 63 3.3.5 Symbiont genome sequencing, assembly, and annotation ....................................... 64 3.3.6 Phylogeny and comparative genomics of Wolbachia strains ...................................... 65 3.3.7 Identifying genes important for reproductive manipulation ................................... 65 3.3.8 Bioassays for reproductive manipulation..................................................................... 66 3.3.9 Statistical procedure for qPCR results and differences in hatching rate and sex ration 67 3.4 Results ........................................................................................................................................ 68 3.4.1 Localisation and Infection Dynamics in O. surinamensis ........................................... 68 3.4.2 Genomics and Phylogeny of the Wolbachia strain ...................................................... 69 3.4.3 Analysis of male-killing gene candidates .................................................................... 73 3.4.1 Cytoplasmic incompatibility (CI) ................................................................................ 75 3.5 Discussion .................................................................................................................................. 78 3.6 Acknowledgments .................................................................................................................... 83 3.7 Data Accessibility Statement ................................................................................................. 83 3.8 Benefit-Sharing Statement ..................................................................................................... 83 3.9 Contributions ........................................................................................................................... 83 3.10 References .................................................................................................................................. 84 3.11 Supplementary Information ................................................................................................... 88 Chapter 4 Co-speciation and functional complementarity of dual Bacteroidota symbionts in powderpost beetles (Coleoptera: Bostrichidae) .................................................................................. 91 4.1 Abstract ...................................................................................................................................... 92 4.2 Introduction .............................................................................................................................. 93 4.3 Results ........................................................................................................................................ 96 4.4 Discussion ................................................................................................................................ 104 4.5 Material & Methods .............................................................................................................. 109 4.5.1 Insect collection .............................................................................................................. 109 4.5.2 Symbiont genome sequencing, assembly, and annotation ..................................... 109 4.5.3 Fluorescence in situ hybridisation ................................................................................ 110 4.5.4 Phylogenetic analyses .................................................................................................... 110 4.6 Data Accessibility Statement ............................................................................................... 112 4.7 Acknowledgments .................................................................................................................. 112 4.8 Contributions ......................................................................................................................... 112 4.9 References ................................................................................................................................ 113 4.10 Supplementary Information .................................................................................................. 117 Chapter 5 Loss of genomic cohesion: Symbiont lineage splitting in the large grain borer Prostephanus truncatus (Coleoptera: Bostrichidae) ............................................................................ 123 5.1 Abstract .................................................................................................................................... 124 5.2 Introduction ............................................................................................................................ 125 5.3 Results ...................................................................................................................................... 127 5.3.1 Bostrichidae-symbiont genomes are highly eroded .................................................. 127 5.3.2 Localisation of the three strains of P. truncatus in the bacteriome ......................... 131 5.1 Discussion ................................................................................................................................ 135 5.2 Material & Methods .............................................................................................................. 139 5.2.1 Insect cultures ................................................................................................................. 139 5.2.2 Symbiont genome sequencing, assembly, and annotation ..................................... 139 5.2.3 16S rRNA cloning and phylogenetic analyses .......................................................... 139 5.2.4 Comparative genomics .................................................................................................. 140 5.2.5 Quantitative PCR ........................................................................................................... 140 5.2.6 Fluorescence in situ hybridisation ................................................................................ 141 5.2.7 HCR in situ staining ...................................................................................................... 142 5.3 Data Accessibility Statement ............................................................................................... 145 5.4 Acknowledgments .................................................................................................................. 145 5.5 Contributions ......................................................................................................................... 145 5.6 References ................................................................................................................................ 146 Chapter 6 General Discussion ............................................................................................................ 149 6.1 Ecology of symbioses in Oryzaephilus surinamensis ............................................................. 149 6.2 Symbionts within Bostrichidae ........................................................................................... 151 6.3 Evolution of beetle-associated Flavobacteria ..................................................................... 156 6.4 Grain pest beetles as non-model model organisms ........................................................... 157 6.5 Conclusion .............................................................................................................................. 160 6.6 References ................................................................................................................................ 161 Danksagung .......................................................................................................................................... 165 Erklärung ............................................................................................................................................... 167 Curriculum Vitae ................................................................................................................................. 169 List of publications Kiefer JST, Batsukh S, Bauer E, Hirota B, Weiss B, Wierz JC, Fukatsu T, Kaltenpoth M, Engl T (2021): Inhibition of a nutritional endosymbiont by glyphosate abolishes mutualistic benefit on cuticle synthesis in Oryzaephilus surinamensis. Commun. Biol. 4, 554 (Chapter 2) Kiefer JST, Schmidt G, Krüsemer R, Kaltenpoth M, Engl T (submitted): Wolbachia causes cytoplasmic incompatibility, but not male-killing in a grain pest beetle. (Chapter 3) Kiefer JST, Bauer E, Okude G, Fukatsu T, Kaltenpoth M, Engl T (in preparation): Co-speciation and functional complementarity of dual Bacteroidota symbionts in powderpost beetles (Coleoptera: Bostrichidae). (Chapter 4) Kiefer JST, Bauer E, Kaltenpoth M, Engl T (in preparation): Loss of genomic cohesion: Symbiont lineage splitting in the large grain borer Prostephanus truncatus (Coleoptera: Bostrichidae). (Chapter 5) 1 Chapter 1 General Introduction 1.1 Symbioses No organism on this planet lives in isolation1. Instead, every single organism one can imagine interacts with at least one or multiple other species. If these interactions between two or more species are close over long-term, they are called symbioses2. Symbiosis is a central driver of evolution3,4. In these partnerships, one of the partners can benefit while the other is not affected (commensalism) or suffers (parasitism), or both partners benefit (mutualism)5. If one of the partners provides for example access to additional pathways or metabolites, the partner may gain new abilities, whereas the other may lose some. and access to otherwise unavailable ecological niches6. One of the most prominent examples of symbiosis is the origin of the eukaryotic cell, which arose probably around 2 billion years ago after different prokaryotic lineages fused during an endosymbiotic event7–10. From then until today, many symbioses have evolved, some of which can even be seen with the bare eye (Figure 1). As best known is probably the pollinator-flower- symbiosis, where the flower offers the pollinator nectar and in return, is fertilized through pollen transport by the pollinator11. Another example of two organisms benefiting the other is the symbiotic relationship between a sea anemone and an anemonefish, where the anemone provides the anemonefish with protection and shelter, while the anemonefish provides the anemone nutrients in the form of waste while also defending against potential predators12. Another example is the symbiosis between aphids and ants in which aphids offers honeydew to the ants. In return, the ants protect aphids from natural enemies such as ladybird beetles13,14. However, the majority of symbiotic interactions of eukaryotes occur on a microscopic level with bacteria, fungi, and viruses. 1.2 Symbioses in insects Insects are one of the most successful animal classes on Earth15. They are present in every terrestrial and limnic environment, even in the Arctic and Antarctic16. However, they are not solely responsible for their success. Several hundred million years ago some insect families formed 3 partnerships with another, highly successful kingdom: bacteria1,17. The symbionts provided new capabilities to their hosts, allowing the insects to acquire new abilities and evolve novel life strategies1,18–23. Today, numerous symbioses between an insect host and a bacterial symbiont are known. In some cases, the insect host benefits from secondary metabolites produced by the symbiont24, which provide context-dependent protection against natural enemies. Some examples of defensive symbioses can be found in aphids25, the European beewolf Philanthus triangulum (Hymenoptera: Crabronidae)26, the darkling beetle Lagria villosa (Coleoptera: Tenebrionidae)27, or the Paederus beetles (Coleoptera: Staphylinidae)28,29. Also, the bacterial symbiont can help their insect host with the digestion of plant material by delivering digestive enzymes, e.g. in the tortoise leaf beetles (Chrysomelidae: Cassidinae)30. Figure 1: Examples of symbiosis. Pollinators like the European honeybee Apis mellifera (A) and the blue- chinned sapphire Chlorestes notata (B) fertilize the plant through pollen transport in exchange for food in the form of nectar offered by the flower. (C) Lichens are a symbiotic association of a fungus and a photobiont (algae or cyanobacteria). While the fungus provides a suitable habitat, the partner provides photosynthetically fixed carbon in return. (D) A common remora Remora remora attached to a green sea turtle Chelonia mydas. While remoras use their host as a taxi to travel through the sea and eat food that falls out, they rid the host of parasites in return. (E) A sea anemone and its resident anemonefish Amphiprion ocellaris. While the anemone offers the fish protection and shelter, the fish provides the anemone nutrients in the form of waste while also defending against potential predators in return. 4 The mechanisms for acquiring symbionts or passing them on to the progeny to keep the symbiont in the population are very diverse31. The symbionts have to be acquired de novo from the environment32,33, horizontally from the outside34, or are transmitted vertically from the mother to the offspring. The latter can also be variously shaped, as a layer of bacteria on the eggs27, special egg caplets which contain the symbiont cells35, to cells of the symbiont already present within the developing egg36. When the bacterial symbionts are transmitted vertically from infected mothers to the offspring, the persistence across host generations of the symbiont population relies on the survival and successful reproduction of infected females. This dependency has driven the development of numerous transmission strategies. One of these strategies is to infect the eggs and manipulate the reproduction of the host to increase the number of progeny that can pass the manipulator by avoiding the production of useless males (from the symbiont’s perspective)37. One popular example of a reproductive manipulator is Wolbachia, which represents the most prevalent symbiotic bacterial genus, associated with over 60% of insect species. In infected insects, Wolbachia often manipulates host reproduction to enhance their transmission. The mechanisms of reproductive manipulation caused by Wolbachia include cytoplasmic incompatibility (CI), parthenogenesis, male-killing, or feminisation37,38, and all lead to a higher proportion of female individuals in the population. This increases the fitness of the symbiont that is predominantly transmitted maternally. Thereby, they can rapidly sweep through uninfected populations and then maintain a high prevalence within a population. However, while Wolbachia infection can be detrimental to the species/population level, the individual’s fitness is enhanced as compared to uninfected individuals in the same population39. Furthermore, Wolbachia can even evolve into a mutualist and enhance its host’s fitness by supplementing dietary limited nutrients, such as B- vitamins like riboflavin40–42. The most widespread type of symbiotic interaction is nutritional1. At the origin of the symbiosis, the microorganisms mostly gain the imminent advantage of a stable environment and the provision of nutrients, while the insect host benefits from the more specialized metabolic capabilities of the symbiont - giving both of them a competitive advantage or the ability to expand to a previously inaccessible ecological niche1,19,20. Specifically, insects often benefit from the metabolic capabilities of microorganisms that they lack themselves, for example, the synthesis of essential amino acids and B-vitamins, which affect the host at several biological levels: physiology, ecology, and evolution17,43. This is especially important when the insect host feeds on 5 nutritionally unbalanced diets. For example, the endosymbiont Buchnera aphidicola44 in the plant sap-sucking aphids, and Wigglesworthia45 in the vertebrate blood-sucking tsetse flies deliver limiting essential nutrients to complement the host’s diet. After a certain period, a dependency has evolved between the partners in many symbioses, as a result of which one can no longer survive without the other46. In addition, since they are morphologically and functionally adapted to their respective food source, an insect cannot simply switch the food source, i.e. the host plant30. Figure 2: The sawtoothed grain beetle Oryzaephilus surinamensis on oat flakes. 1.3 Ecology of grain pest beetles Already one century ago, entomologists described symbiotic organs in numerous species of invertebrates, the so-called bacteriome47. This specialised organ is made up mainly of cells called bacteriocytes which house the bacteria intracellularly. Among others, they found bacteriomes in beetle species of various families, several of which are known as grain pests47–49. These beetles appear to benefit from an excess of food but face the challenge of unbalanced diets and low 6 humidity, especially nowadays maintained in storage facilities to control mold growth, but also insect infestation. Several beetle groups nevertheless managed to invade the same ecological niche of stored grain and dried plant products independently. Examples are weevils of the genus Sitophilus, several silvanid beetles including the sawtoothed grain beetle Oryzaephilus surinamensis (Figure 2, Chapter 2) as well as several bostrichid beetles (Figure 3, Chapter 4). The symbiont Nardonella (γ-proteobacteria), which is associated with multiple weevils of the families Curculionidae and Brentidae50 encodes a complete tyrosine biosynthetic pathway while lacking pathways for all other essential and non-essential amino acids and vitamins46. Experimental suppression of Nardonella results in beetles with soft and less melanised cuticles, revealing their role in cuticle sclerotisation and melanisation46,51. Similar effects have been observed in Sitophilus spp. (Curculionidae, Dryophthorinae) and in O. surinamensis (Silvanidae), with both harbouring symbionts that are phylogenetically distinct from Nardonella36,52,53. In addition, several Bostrichidae36 harbour symbionts that are closely related to that of O. surinamensis36, suggesting an involvement in cuticle tanning. In contrast to many other nutritional endosymbionts, the symbionts of Sitophilus and of O. surinamensis can be completely removed by treating the beetles with antibiotics or heat without interrupting the host life cycle36,46,52,54. The loss of a nutritional endosymbiont usually has severe effects on its host, e.g. by arresting development and reproduction and/or causing high mortality46. In contrast, the experimentally symbiont-deprived (aposymbiotic) O. surinamensis beetles were viable, able to reproduce, and could be maintained in stable aposymbiotic populations under laboratory conditions, allowing disentangling the effect of symbiont loss from direct antibiotic treatment36. In comparison to symbiont-containing control beetles, aposymbiotic individuals of O. surinamensis exhibited a thinner and noticeably lighter cuticle, which resulted in a significant fitness decrease due to higher mortality55, especially under desiccation stress in stored grain products, the contemporary predominating habitat of the beetle36. 1.4 Cuticle formation Among the holometabolous insects, beetles (Coleoptera) are the most speciose of all insect orders. Their most outstanding characteristic is that their front pair of wings are hardened into protective wing-cases, called elytrae56. The elytrae guarantee the beetles protection against predators and pathogens and provide a higher desiccation resistance57. Beetles develop their full imaginal cuticle 7 during metamorphosis and the first days as an imago58. The cuticle of insects is primarily composed of chitin fibrils enveloped in protein chains rich in aromatic amino acids57,59–61. In addition, insects modify the native cuticle through the integration of phenolic compounds derived from the aromatic amino acid tyrosine in two processes called tanning (melanisation) and hardening (sclerotisation)61, which lead to a cuticle that is darker, harder, and less permeable to water. However, in many herbivorous diets, nitrogenous compounds including tyrosine and phenylalanine from which tyrosine could be derived by insects, are limited. Thus, the thick cuticle comes with a cost as it requires a very high tyrosine investment during development57,62. Plants, fungi, and bacteria can synthesise precursors of tyrosine via the shikimate pathway63. Since beetles lack this pathway, they have to either ingest the aromatic amino acid via a food source or have it supplied by symbiotic microorganisms. Figure 3: Beetles of the Bostrichidae family. Images of Bostrichus capucinus (A) and Amphicerus bimaculatus (B) were taken by Thomas Hörren in the wild. Autofluorescence microscopy images of Prostephanus truncatus (C), Rhyzopertha dominica (D), and Dinoderus porcellus (E). 8 Like all holometabolous insects, beetles undergo a stepwise development, meaning from the embryo in the egg to the larva, to pupation, and finally to the adult animal. For the animal, this means that the anatomy of internal and external morphological organisations is remodelled fundamentally64. For some insect species, this also means that the individual life stages of an animal feed on different food sources because the morphology of e.g. the mouthparts is also changing65,66. Thus, the larval stage of the butterfly - the caterpillar - feeds on leaves, while the adult butterfly feeds on nectar. Further, in some species, food intake is limited to the larval stage, while the adult animal is only devoted to reproduction and thus to the survival of the species. The morphological reorganisation during metamorphosis requires a high investment in nutrients. As a consequence, young imagines initially have few resources left and have to take them up again first - or have them provided by symbionts. As beetles are not able to feed during metamorphosis, they are reliant on stored nutrients acquired during their larval development for metamorphosis and cuticle formation. In the case of tyrosine, the amount an insect can store is limited, because the amino acid is toxic in high concentrations68. In this situation, harbouring a bacterial endosymbiont that can synthesize tyrosine or its precursors' chorismate or prephenate57,63 in the moment of demand represents a strategy to cope with the storage problem36,46,52,53. 1.5 Bacteroidota symbionts Beetles of the families Silvanidae and Bostrichidae harbour abdominal bacteriomes, housing their bacterial endosymbionts47. The paired bacteriomes are located between the gut, fat body, and reproductive organs, but without direct connection to any of these tissues36,69. Despite the phylogenetic distance of Bostrichidae and Silvanidae of about 105 - 175 Mya87,88, both beetle families are associated with symbionts belonging to the class Flavobacteria in the phylum Bacteroidota36. In addition, they are not the only ones, as phylogenetic analyses of insect-bacteria symbioses have shown in recent years (Figure 4). Accordingly, the Bacteroidota symbionts of the silvanid O. surinamensis and the bostrichids such as the lesser grain borer Rhyzopertha dominica are derived from the same ancestor as Blattabacterium, Uzinura, Walczuchella, and Sulcia muelleri. They diverged about 409 Mya ago36 and evolved different functional specialisations36,69–71. Uzinura (endosymbiont of armoured scales), Walczuchella (endosymbiont of giant scales), Blattabacterium (endosymbiont of roaches and primitive termite), and Sulcia (endosymbiont of numerous sap- 9 feeding hemipterans) are known to supplement their host's diet with essential amino acids and vitamins71–74. In addition, there is evidence for nitrogen recycling in Blattabacterium75–77. As Blattabacterium still retains one of the largest genomes, it can be argued that urea catabolism appears to be an ancestral capability of Bacteroidota symbionts. Figure 4: Phylogenetic placement of intracellular symbionts in silvanid (red) and bostrichid (yellow) beetles within the Bacteroidota, and their close association within the clade of exclusively insect-associated endosymbionts (green), based on 16S rRNA sequences. Node numbers represent local support values. RefSeq assembly accession in square brackets. Since then, the ancestor of Bacteroidota symbionts has been associated with various insect families: scale insects71,72,78, cockroaches and termits75, Auchenorrhyncha e.g. cicadas, leafhoppers, planthoppers and splittlebugs79–81 as well as the silvanid36,52, bostrichid36,82 and nosodendrid beetles69. Bacteroidota symbionts also show a divergent specialisation in different insect orders: Uzinura diaspidicola in armoured scale insects71, Walczuchella monophlebidarum in scale insects72, Blattabacterium spp. in cockroaches, Sulcia muelleri in the Auchenorrhyncha, and now Shikimatogenerans sp. and Bostrichicola sp. in beetles (Chapter 2 and 4). The repeated independent acquisitions of symbionts from this specific clade of Bacteroidota bacteria indicate that these bacteria were once highly capable of infecting and establishing in insects, akin to the α-proteobacterium Wolbachia (Chapter 3) or the γ-proteobacterium Sodalis83,84. This hypothesis is 10 supported by the observation that a basal clade of Bacteroidota endosymbionts is described as male-killing endosymbionts in different ladybird beetles85–87. Hirota et al. (2017) and Engl et al. (2018) provided evidence that the sawtoothed grain beetle O. surinamensis harbours a Bacteroidota endosymbiont which is somehow involved in the beetle’s cuticle biosynthesis. Still, what the symbiont delivers to the host remained unknown. Also, the full metabolic potential beyond supporting cuticle synthesis of the endosymbiont remained hidden. In addition, Okude et al. (2017) and Engl et al. (2018) showed that beetles of the Bostrichidae family are associated with close related Bacteroidota endosymbionts, but the evolutionary origins and ancestral ecological contexts of these symbioses were unknown. Still, it remained elusive how exactly these symbioses between beetles of the Silvanidae and Bostrichidae families and their respective symbiont (or symbionts) function in detail as the metabolic potential of these bacteria was not elucidated. 11 1.6 Thesis outline In this thesis, I aimed to identify the influence of symbiosis on the physiology, ecology, and evolution of grain beetles associated with Bacteroidota symbionts using genome sequencing, fluorescence microscopy, and experimental manipulation. I set out to (i) sequence the genomes and reconstruct the metabolic potential of the Bacteroidota endosymbionts associated with representative species of the families Silvanidae and Bostrichidae, (ii) experimentally verify the function of the endosymbionts in Oryzaephilus surinamensis, and (iii) reconstruct the evolutionary history of the symbiotic association in the species-rich bostrichid family. In Chapter 2, I report the genome of Shikimatogenerans silvanidophilus OSUR, the Bacteroidota endosymbiont of the sawtoothed grain beetle O. surinamensis. Further, as the genome encodes genes of the shikimate pathway, I utilized the herbicide glyphosate, a selective pharmacological inhibitor of the shikimate pathway, to block the beneficial symbiotic contribution to its host’s physiology and assessed the impact of symbiont elimination on host fitness correlates. Doing so, O. surinamensis is introduced as a well-suited system for experimental and detailed work on symbiosis because of the amenability for manipulation of the host-symbiont association. In addition to being associated with the Bacteroides endosymbiont, O. surinamensis is also regularly infected with the reproductive manipulator Wolbachia. Therefore, I characterized this Wolbachia strain and its effect on the beetle’s reproduction in Chapter 3. 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Wierz1, Takema Fukatsu2,3,4, Martin Kaltenpoth1,5,6, Tobias Engl1,5 1Evolutionary Ecology, Institute of Organismic and Molecular Evolution, Johannes Gutenberg-University, Mainz, Germany; 2Bioproduction Research Institute, National Institute of Advanced Industrial Science and Technology, Tsukuba 305-8566, Japan; 3Department of Biological Sciences, Graduate School of Science, University of Tokyo, Tokyo 113-0033, Japan; 4Graduate School of Life and Environmental Sciences, University of Tsukuba, Tsukuba 305- 8571, Japan; 5Research Group Insect Symbiosis, Max-Planck-Institute for Chemical Ecology, Jena, Germany; 6Department of Insect Symbiosis, Max-Planck-Institute for Chemical Ecology, Jena, Germany 17 2.1 Abstract Glyphosate is widely used as a herbicide, but recent studies begin to reveal its detrimental side effects on animals by targeting the shikimate pathway of associated gut microorganisms. However, its impact on nutritional endosymbionts in insects remains poorly understood. Here, we sequenced the tiny, shikimate pathway encoding symbiont genome of the sawtoothed grain beetle Oryzaephilus surinamensis. Decreased titers of the aromatic amino acid tyrosine in symbiont- depleted beetles support the ability to synthesize prephenate as the precursor for host tyrosine synthesis and its importance for cuticle sclerotisation and melanisation. Glyphosate exposure inhibited symbiont establishment during development and abolished the mutualistic benefit on cuticle synthesis in adults, which could be partially rescued by dietary tyrosine supplementation. Furthermore, phylogenetic analyses indicate that the shikimate pathways of many nutritional endosymbionts contain a glyphosate sensitive 5-enolpyruvylshikimate-3-phosphate synthase. These findings highlight the importance of symbiont-mediated tyrosine supplementation for cuticle biosynthesis in insects, but also paint an alarming scenario regarding the use of glyphosate in light of recent declines in insect populations. 18 2.2 Introduction Glyphosate is a widely used, broad-spectrum herbicide that targets the shikimate pathway by inhibition of the 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS)1,2. The shikimate pathway is present in plants, fungi, and bacteria to biosynthesize the aromatic amino acids phenylalanine, tyrosine and tryptophan and folates. As the inhibition of the EPSPS by glyphosate in susceptible plants is lethal3, this herbicide is extensively used in agriculture in combination with genetically modified glyphosate-resistant crops to eliminate competing plants4. Animals, by contrast, are assumed to be unaffected by glyphosate, as they lack the shikimate pathway and meet their demands for aromatic amino acids from external sources5. However, animals do not live in isolation but engage in manifold mutualistic interactions with microorganisms6. While the microorganisms mostly gain the imminent advantage of a stable environment and the provision of basic nutrients7, animals benefit from a more specialized metabolic capabilities of the symbiont, giving both of them a competitive advantage or the ability to expand to a certain niche8–14. Specifically, animals often benefit from metabolic capabilities of microorganisms that they lack themselves, for example, the synthesis of essential amino acids and (B-)vitamins, which is especially important when the insect host feeds on nutritionally unbalanced diets like plant sap or blood14,15. In the same way, semi-essential nutrients, i.e. nutrients like the aromatic amino acid tyrosine that can in principle be derived from the other aromatic amino acids taken up from the diet, but are in certain life stages like the metamorphosis of holometabolic insects still limited, are supplied by many nutritional endosymbionts16–19. Despite several studies emphasizing that glyphosate displays only “minimal toxicity” in animals via off-target activity due to the lack of the shikimate pathway in their genomes1,20,21, recent work demonstrates that glyphosate does have a negative impact on insects, either directly or by inhibiting the EPSPS of animal-associated, mutualistic bacteria, with negative fitness consequences for the host22,23. In blood-feeding tsetse flies, glyphosate interferes with the biosynthesis of folate by the ɣ-proteobacterial symbiont Wigglesworthia glossinidia24. The herbicide also alters the gut microbiota of honeybees that consequently become more susceptible to opportunistic pathogens25,26. Since many obligate insect endosymbionts encode the shikimate pathway and supply aromatic amino acids to their host19,27,28, glyphosate could prove highly detrimental for diverse insect hosts, with potentially severe ecological implications given the widespread nature of these mutualistic interactions in insects29–31. However, the impact of 19 glyphosate on insect-microbe associations remains poorly studied, particularly with respect to obligate nutritional endosymbionts in herbivorous insects. The cuticle of insects is primarily composed of chitin fibrils enveloped in protein chains rich in aromatic amino acids32–34. Additionally, insects modify the native cuticle through the integration of phenolic compounds derived from tyrosine in two processes called melanisation and sclerotisation, which lead to a cuticle that is darker, harder, and less permeable to water35,36. However, in many herbivorous diets nitrogen in general, but also specifically tyrosine as well as phenylalanine and tryptophan from which it could be derived by insects are limited37,38. Among the holometabolous insects, beetles (Coleoptera) are distinguished by an especially strong adult cuticle, including the fully hardened forewing, i.e. the elytrae39. In addition, holometabolous insects develop their full imaginal cuticle during the pupal stage when the insect is undergoing metamorphosis as well during the first days as an imago36. As insects are not able to feed during metamorphosis, they are reliant on stored nutrients acquired during their larval development for metamorphosis and cuticle formation. In the case of tyrosine, the amount an insect can store is limited, because the amino acid is toxic in high concentrations40. In this situation, harbouring an endosymbiont that can synthesize tyrosine or its precursors chorismate or prephenate5,41 in the moment of demand represents a strategy to cope with the storage problem16,19,42,43. We recently described a bacteriome-localised Bacteroidota endosymbiont in the sawtoothed grain beetle Oryzaephilus surinamensis (Coleoptera, Silvanidae)42,43, which is a worldwide distributed pest of cereals and other stored food44. In contrast to many other nutritional endosymbionts, the symbiont of O. surinamensis can be removed by treating the beetle with antibiotics or heat without interrupting the host life cycle42,43. The loss of a nutritional endosymbiont usually has severe effects on its host, e.g. by arresting development and reproduction and/or causing high mortality19. In contrast, the experimentally symbiont-deprived (aposymbiotic) O. surinamensis beetles were viable, able to reproduce, and could be maintained in stable aposymbiotic populations under laboratory conditions, allowing to disentangle the effect of symbiont loss from direct antibiotic treatment42. In comparison to symbiont-containing control beetles, aposymbiotic individuals exhibited a 30% thinner and noticeably lighter cuticle, which resulted in a significant fitness decrease due to higher mortality, especially under desiccation stress in stored grain products, the natural habitat of the beetle42. 20 In this work, we demonstrate (i) that the symbiont genome of O. surinamensis is extremely streamlined, providing precursors for cuticle synthesis via the shikimate pathway, (ii) that exposure to glyphosate compromises the symbiont establishment in O. surinamensis and induces the fitness-relevant cuticular defects of the host in a similar manner as the complete loss of the symbiont, (iii) that these phenotypic effects can be partially rescued by dietary tyrosine supplementation, and (iv) that the shikimate pathways of O. surinamensis as well as many other nutritional endosymbionts contain class I EPSPSs that are predicted to be glyphosate-sensitive. These results experimentally validate the functionality of the shikimate pathway in an obligate endosymbiont and more generally demonstrate the severe impact of glyphosate on organisms that are dependent on bacterial endosymbionts. 21 2.3 Results 2.3.1 Symbiont genome is extremely reduced and GC poor We sequenced the metagenome of O. surinamensis combining short and long-read technologies (Illumina and ONT) into a hybrid assembly to gain first insights into its symbiont’s metabolic capabilities. As expected, we could detect the 16S rRNA sequence of a Bacteroidota bacterium in the assembly that matched PCR-based Sanger sequences from previous studies42,43. In total, 13 contigs were extracted from the metagenomic assembly via taxonomic classification, GC content filtering as well as by manually searching for tRNAs and ribosomal proteins of Bacteroidota bacteria (Figure 1.1, Supplementary File 1). The longest was 74,813 kbp and the shortest 6,352 kbp in length. Together they had a length of 307,680 kbp with an average GC content of 16.2%. The draft genome encoded for 299 genes and had a coverage of 120x with short-read sequences and 61x with long-read sequences. The phylogenetic reconstruction based on the conserved Clusters of Orthologous Group (COG) genes confirmed the placement of the endosymbiont of O. surinamensis in a group of insect- associated Bacteroidota bacteria and specifically the close relationship to Blattabacterium sp. and Sulcia muelleri (S. muelleri) that was already previously reported based on a 16S rRNA gene phylogeny (Supplementary Figure 1)42. The symbiont’s genome encodes for 299 protein-coding sequences, 28 tRNAs and 51 ribosomal proteins (21 SSU and 30 LSU proteins). Based on a set of single-copy marker genes that are assumed to be essential, the O. surinamensis symbiont genome is estimated to be 66.7% complete45. However, this ‘completeness’ measure is based on essential genes in free-living bacteria and known to severely underestimate the completeness of highly eroded genomes of intracellular bacterial symbionts46,47. Concordantly, the estimation falls in the range of other insect-associated Bacteroidota mutualists that exhibit highly eroded, closed genomes (54.20 - 67.38% of S. muelleri PSPU and W. monophlebidarum; Supplementary Table 1 & 2). Despite the remaining gaps in the symbiont genome sequence, the 13 contigs are thus inferred to contain the complete or almost complete set of coding sequences, which is corroborated by the presence of a full complement of tRNAs, tRNA synthetases, and ribosomal proteins to a similar extent as in other Bacteroidota endosymbionts. The contigs are likely also fragments of a single chromosome, as it does not feature duplicated genes, multiple rRNA operons or variable coverage across the different contigs, which are indicative of such fragmentation reported from multiple 22 Figure 1.1: Shikimatogenerans silvanidophilus, the symbiont of O. surinamensis. (a) Fluorescence in situ hybridisation micrograph of a sagittal section of a 5-day-old O. surinamensis pupa stained with CFB563mod-Cy3 (magenta) and DAPI (white). (b) Circular representation of the draft genome of S. silvanidophilus. Single contigs are sorted clockwise by length. The outer grey circles denote coverage with long- and short-reads, respectively, the intermediate circles indicate annotated functional KEGG categories separated by direction of transcription (see legend for depicted categories). The inner grey circle denotes relative GC content and the average GC content of 16.2% by the red line. Below: comparison of the functional gene repertoires of Bacteroidota symbionts in insects. Box colors are based on KEGG’s categories. 23 fragmented chromosomes of Hodgkinia strains of different Magicicada species48–50. As an extremely low GC content (16.2%) and long repeat sequences towards contig ends are known issues for sequencing technologies and PCRs51–54, these features likely prevented successful amplification steps and the in silico assembly of the contigs into a single genome despite the high coverage (120x with short-read sequences and 61x with long-read sequences), especially as the GC content at the ends of most contigs reached even lower values (~ 4% within the last 100 bp). We also compared the presence and arrangement of the genes between Bacteroidota endosymbionts of different host insects. A synteny plot (Supplementary Figure 2) revealed that there is no conserved arrangement of genes between the genomes of related Bacteroidota symbionts. This likely explains futile attempts to assemble the genome by mapping the assembled contigs to Blattabacterium or S. muelleri as reference genomes or to sort contigs into a single scaffold. Figure 1.2. (c) Detailed comparison of the amino acid metabolism gene repertoires between the Shikimatogenerans silvanidophilus genome (pink), other Bacteroidota symbionts (left) and Proteobacteria symbionts (right), some of which are known to exclusively provision tyrosine precursors to their insect host (blue). 24 2.3.2 Candidatus Shikimatogenerans silvanidophilus OSUR encodes glycolysis and shikimate pathways The metabolic repertoire of the O. surinamensis symbiont is highly reduced (Figure 1.2, Supplementary File 1). Apart from general genetic information processing including DNA replication and repair, transcription, and translation, it only encodes an extremely limited set of metabolic pathways including a full glycolysis pathway to process glucose-6-phosphate to erythrose 4-phosphate (E4P) and phosphoenolpyruvate (PEP). In addition, the genome encodes all the genes of the shikimate pathway except a shikimate dehydrogenase (aroE [EC:1.1.1.25]) that utilizes PEP and E4P to produce chorismate, as well as a chorismate mutase to catalyze the conversion of chorismate into prephenate, the precursor of the aromatic amino acids phenylalanine, tryptophan and tyrosine (Figure 2). The lack of aroE is described in other tyrosine- supplementing bacterial symbionts: Cd Carsonella ruddii in Pachypsylla venusta55 and Cd Nardonella EPO in Euscepes postfasciatus19. AroD (3-dehydroquinate dehydratase [EC:4.2.1.10]) and aroE are also often found as a bifunctional enzyme aroDE (3-dehydroquinate dehydratase/shikimate dehydrogenase [EC:4.2.1.10 1.1.1.25])56, possibly complementing the loss of one of the enzymes. Like all other Bacteroidota symbionts, the O. surinamensis symbiont genome encodes the bifunctional aroG/pheA gene (phospho-2-dehydro-3-deoxyheptonate aldolase/chorismate mutase [EC:2.5.1.54 5.4.99.5]). The genomic data revealed no transporter for glucose, so it remains unknown how the symbiont acquires the substrate for glycolysis from the host (Figure 2). Whether the O. surinamensis symbiont can recycle nitrogen, like Blattabacterium sp.57,58 remains unclear, as it encodes the urease α and γ subunits (ureC [EC:3.5.1.5]), but no glutamate dehydrogenase (gdhA) that would allow integrating the resulting ammonium into the amino acid metabolism via glutamate was detected. Instead, it encodes a proline transporter (opuE), proline dehydrogenase [EC:1.5.5.2] and oxidoreductase [EC:1.2.1.88] to import and convert proline into glutamate59,60, which may then be exported to function as an amino group donor for the synthesis of the aromatic amino acids from chorismate/prephenate by the beetle itself19. This alternative pathway for glutamate synthesis was not described in Blattabacterium61, S. muelleri62, Uzinura diaspidicola63 or Walczuchella monophlebidarum64. 25 Figure 2: Metabolism of the symbiont Shikimatogenerans silvanidophilus. (a) Complete, reconstructed metabolism of S. silvanidophilus, as inferred from genomic data. Enzymes and arrows in grey were missing in the genome annotation. Dashed arrows indicate transport processes without annotated transporters, but which are expected to occur based on observed phenotypes. (b) Schematic diagram of the shikimate pathway in S. silvanidophilus. Dashed arrows represent multiple enzymatic steps. Red boxes: enzyme present in the genome. White box: enzymes not annotated in the genome, but reaction probably catalysed by another enzyme. A comparison of the metabolic gene repertoires for amino acid biosynthesis revealed convergent genome erosion between the Oryzaephilus symbiont and ɣ-proteobacterial symbionts known to provision precursors for the host’s cuticle biosynthesis (Figure 1c). We observed a gradual loss of functions across the Bacteroidota insect symbionts, with the O. surinamensis symbiont genome exhibiting the most strongly reduced repertoire of biosynthetic genes. A convergent reduction of metabolic functions was observed in the ɣ-proteobacterial symbionts Westerberhardia and Nardonella sp., whose sole metabolic function appears to be the provisioning of aromatic amino acid precursors that are in high demand for cuticle biosynthesis of their insect host19,27,65. Based on our findings, we propose the name ‘Candidatus Shikimatogenerans silvanidophilus OSUR’ for the endosymbiont of O. surinamensis, henceforth called S. silvanidophilus. The genus name Shikimatogenerans refers to its ability to perform the shikimate pathway. Previous studies have shown that there might be other closely related Bacteroidota bacteria associated with other beetle families42,66,67. Thus, we propose silvanidophilus as species name to indicate that this symbiont is associated with a silvanid beetle. As the same studies also revealed that O. mercator 26 has a similar symbiont we also propose to add OSUR to identify the strain associated with O. surinamensis. 2.3.3 Amino acid titers are influenced by symbiont presence Using the genomic data as a basis, we then asked whether the symbiont-encoded pathways are indeed functional. To this end, we tested if the symbiont presence indeed influenced the titer of aromatic amino acids. This would be expected when the symbiont is delivering prephenate to the host, a precursor that can also be transformed to tyrosine or phenylalanine by the beetles themselves17,27,68. Due to the presence of the proline-transporter and proline-converting enzymes in the symbiont genome, an influence on proline titer and glutamic acid was also expected based on the metabolic reconstruction. After hatching from the egg, the larva of O. surinamensis spends weeks in multiple instars before it pupates and hatches five days later as an imago. During metamorphosis, the biosynthesis of the adult cuticle starts and continues in the first days of the imago36. As tyrosine is used in copious amounts in the cuticle biosynthesis69 amino acid titers might change dynamically. Thus, we expected the major symbiont contribution of tyrosine in the pupal stage and early adulthood19. Accordingly, we found an overall significantly positive influence of the symbiont presence on the titer of tyrosine (FDR corrected GLMs, p = 0.0000858, see Figure 3 and Supplementary Figure 3 and detailed statistic results for all the following values in Supplementary Table 3 and 4). Specifically, late symbiotic pupae had a significantly higher tyrosine titer than aposymbiotic ones (Wilcox rank-sum tests with FDR corrected p-values: p = 0.00078), shortly before adult emergence. By contrast, there was a negative impact of symbiont presence on proline and glutamic acid levels (FDR corrected GLMs, p = 0.0000858), which are interconverted by the symbiont based on the genomic prediction and used to synthesize tyrosine from prephenate. Again, free proline was significantly lower in late symbiotic vs. aposymbiotic pupae (FDR corrected Wilcox rank-sum test, p = 0.00078) and bound proline in early adults (FDR corrected Wilcox rank-sum test, p = 0.00078). Free glutamic acid was significantly lower in early and late symbiotic pupae (FDR corrected Wilcox rank-sum tests, both p = 0.00078, Figure 3) and the elytrae of early symbiotic adults (FDR corrected Wilcox rank-sum test, p = 0.0059). There was no difference in the titer of bound or cuticular tyrosine in the different life stages of the beetle in direct comparison (FDR corrected Wilcox rank-sum test, p > 0.05) and only protein-bound glutamic acid of the full body of early symbiotic beetles was significantly higher than in aposymbiotic ones (FDR corrected Wilcox rank-sum test, p = 0.0059). All other 27 amino acids were not influenced by symbiont presence alone, although we detected several interaction effects (FDR corrected GLMs, p > 0.005; Supplementary Table 3). Figure 3: Comparison of titers of the three amino acid tyrosine, proline and glutamic acid that that were influenced by symbiont presence. Shown are free amino acid titers in the whole body (without elytrae in case of adults) of symbiotic and aposymbiotic O. surinamensis beetles. Red: aposymbiotic beetles, Blue: symbiotic beetles. The data distribution is visualized with violin plots and an additional horizontal line depicting the median. The FDR-corrected unpaired, two-sided Wilcoxon-rank-sum-tests: ns p > 0.05, *0.05 < p < 0.01, **p < 0.01. 28 2.3.4 The symbiont’s shikimate pathway is sensitive to glyphosate and its inhibition results in an aposymbiotic phenotype Next, we investigated the consequences of a pharmacological inhibition of the shikimate pathway on the symbiosis. Therefore, we made use of the pesticide glyphosate, an allosteric inhibitor of the enzyme EPSPS (5- enolpyruvoylshikimate-3-phosphate synthase) which is encoded by aroA1. We experimentally determined the impact of glyphosate exposure and aromatic amino acid supplementation during the entire larval and early adult development on phenotypic parameters that were previously demonstrated to be impacted by the symbiosis, i.e. cuticle thickness and melanisation42,43. As predicted, both the thickness (Kruskal-Wallis χ2 = 27.8525, df = 7, p-value = 0; Figure 4a, Supplementary Table 5) and melanisation (Kruskal-Wallis χ2 = 35.967, df = 7, p-value = 0; Figure 4b, Supplementary Table 5) of the cuticle were influenced by glyphosate exposure and by aromatic amino acid supplementation of the beetle diet. Symbiotic beetles showed a significant reduction of both cuticle traits after inhibition of the shikimate pathway with 1% glyphosate (Dunn’s test: symbiotic vs. symbiotic + glyphosate: p = 0.0038 & p = 0.0049), to the same level as observed in aposymbiotic beetles (Dunn’s test: aposymbiotic vs. symbiotic: p = 0.0002 & p = 0.0008; aposymbiotic vs. symbiotic + glyphosate: p = 0.6039 & p = 0.5360). A lower amount of glyphosate (0.1%) led to intermediate phenotypes that did not differ significantly from either symbiotic or aposymbiotic or 1% glyphosate treated individuals (Dunn’s test: p > 0.05). Importantly, the thinner and less melanized cuticle of both aposymbiotic and glyphosate-exposed (1%) symbiotic beetles could be rescued to an intermediate state by adding aromatic amino acids to the diet, resulting in cuticular traits that differed neither from symbiotic nor aposymbiotic or 1% glyphosate treated beetles (Dunn’s test: p > 0.05). By contrast, aromatic amino acid supplementation did not affect cuticular traits of symbiotic beetles that were not exposed to glyphosate (Dunn’s test: p > 0.05). These findings support the genome-based prediction that S. silvanidophilus supplements precursors for tyrosine biosynthesis via the shikimate pathway. The two plausible scenarios for the impact of glyphosate exposure on symbiont contributions are (i) a reduced chorismate biosynthesis by a stable symbiont population, or (ii) an overall decrease in symbiont titers due to glyphosate exposure. This includes either direct effects trough glyphosate toxicity, but also indirect effects via consequences of an inhibited chorismate biosynthesis like a lack of aromatic amino acids or host feedback mechanisms. Concordant with 29 Figure 4: Effect of glyphosate exposure and aromatic amino acid supplementation on cuticle traits and symbiont titers in symbiotic and aposymbiotic beetles. Cuticle thickness (a), melanisation measured as thorax coloration (b), and symbiont titers (c) of aposymbiotic and symbiotic adults reared on different food compositions. Different letters indicate significant differences between experimental treatments (Dunn’s Test, α ≤ 0.05). 30 the latter hypothesis, we detected significant differences in symbiont titers of one-week-old symbiotic adults across experimental treatments (Kruskal-Wallis χ2 = 59.0049, df = 7, p-value = 0; Figure 5c, Supplementary Table 5). The symbiont titers in symbiotic beetles exposed to high glyphosate concentrations were significantly reduced by 98% (based on medians; Dunn’s test: sym vs. 1% glyphosate: p = 0.00034). Low glyphosate concentrations and supplementation with aromatic amino acids also reduced symbiont titers to an intermediate level that did not differ significantly from either untreated beetles nor those treated with high glyphosate amounts (Dunn’s test: p > 0.05). In combination with low amounts of glyphosate, supplementation of aromatic amino acid reduced the symbiont titers further, leading to a significant difference to untreated beetles (Dunn’s test: AA + 0.1% glyphosate vs untreated: p = 0.028). Finally, we tested whether the EPSPS of other intracellular symbionts besides the one of S. silvanidophilus belong to glyphosate sensitive EPSPS variants by a phylogenetic analysis of their amino acid sequences25. The EPSPSs of all included Bacteroidota and also Proteobacteria symbionts of insects clustered with class I EPSPS and are thus predicted to be sensitive to glyphosate – in contrast to insensitive class II enzymes of several free-living bacteria70 (Figure 5 and Supplementary Figure 4). 31 Figure 5: Phylogenetic classification of EPSPS enzymes. Enzymes from different Bacteroidota and γ-Proteobacteria insect symbionts as well as some free-living bacteria and plants were classified based on FastTree and maximum- likelihood analyses of amino acid sequences of EPSPSs using the Jones–Taylor– Thorton model. Enzymes of plants (green), bacterial symbionts in the honeybee gut (yellow), and obligate intracellular insect symbionts (red) are highlighted. Node values indicate FastTree support values. 32 2.4 Discussion The sawtoothed grain beetle O. surinamensis engages in an intimate association with the Bacteroidota endosymbiont S. silvanidophilus that confers desiccation resistance via enhanced cuticle synthesis42,43. This physiological contribution represents a significant fitness benefit in dry habitats like mass grain storage42. Here, we demonstrated that the genome of S. silvanidophilus experienced a drastic erosion, resulting in a genome of 308 kbp in size and a strongly AT-biased nucleotide composition (16.2% GC), akin to what has been described for other obligate intracellular47,71,72 or extracellular insect symbionts73–76. Moreover, the symbiont genome encodes only a few metabolic pathways, of which the substantial remaining ones are the glycolysis and shikimate pathways. Phosphoenolpyruvate (PEP) and Erythrose-4-phosphate (E4P) - both originating from glycolysis - are used as substrates in the shikimate pathway to synthesize chorismate, which in turn is transformed to prephenate by the symbiont and then converted to the aromatic amino acids phenylalanine, tryptophan and tyrosine by the host, as recently demonstrated in the weevil-Nardonella endosymbiotic system19. Of the seven genes in the shikimate pathway (aroG, aroB, aroD, aroE, aroK, aroA, aroC), the genome only lacks the gene for shikimate dehydrogenase (aroE), which catalyzes the reversible NADPH- linked reduction of 3-dehydroshikimate to shikimate. However, the shikimate pathways of Nardonella EPO, the endosymbiont of the sweetpotato weevil Euscepes postfasciatus (Curculionidae: Cryptorhynchinae) and Carsonella ruddii, the endosymbiont of the gall-forming psyllid Pachypsylla venusta (Aphalaridae: Pachypsyllinae) also lack aroE, but remain functional19,55, suggesting that the function can be taken over by other enzymes, either from the host or the endosymbiont—that have yet to be identified. The genome also encodes for an urease (ureC, Urease α and γ subunits [EC:3.5.1.5]), so S. silvanidophilus may be able to recycle nitrogenous waste products of its host, as has been described for Blattabacterium in cockroaches and Blochmannia in carpenter ants61,64,77. Alternatively, however, this enzyme could be a non-functional remnant in the eroded genome, as no glutamate dehydrogenase (gdhA) synthesizing glutamate from ammonium and 2-oxogluterate was detected and the symbiont genome encodes no other annotated gene that could incorporate the resulting ammonium. Instead, the genome encodes for a proline transporter (opuE) and the genes to convert proline into glutamic acid (EC:1.5.5.2 and EC:1.2.1.88)78,79. Thus, proline as one of the most 33 abundant amino acids in insect hemolymph might be utilized by the symbiont, among others as a source of glutamate80. As the genome encodes no other pathways of potential relevance to the host, we hypothesized that the symbiont’s beneficial impact on host fitness results from the supplementation of the tyrosine precursor prephenate, and possibly, from additional nitrogen recycling. The quantification of amino acid titers throughout insect development in symbiotic and aposymbiotic beetles supports the genome-based predictions of symbiont-mediated tyrosine precursor biosynthesis and proline consumption. Both tyrosine and proline concentrations revealed significant differences between symbiotic and aposymbiotic beetles in late pupae, consistent with proline consumption and tyrosine biosynthesis during metamorphosis, which coincides with the reported maximum symbiont titers in O. surinamensis81. While our results indicate that the symbionts consume proline, the dynamics of proline conversion to glutamic acid remain elusive, as glutamic acid titers were lower in symbiotic than in aposymbiotic beetles across multiple life stages. Possibly, glutamic acid is predominantly utilized by the symbiont itself. Alternatively, as more chorismate is available for tyrosine synthesis in symbiotic than in aposymbiotic beetles, the glutamic acid titers might be constantly depleted, while tyrosine may only accumulate during metamorphosis, and is directly channelled into cuticle biosynthesis in all other life stages, as tyrosine accumulation may be toxic40. A similar phenomenon of increased symbiont titers and intense cuticle synthesis and modification during metamorphosis and early adulthood was described in another grain pest beetle, the weevils S. oryzae16and P. infernalis19, as well as the ant Cardiocondyla obscurior27. Symbiont-mediated contributions to cuticle biosynthesis evolved multiple times convergently in insects, with genomic evidence for tyrosine supplementation by symbionts in the ant genus Cardiocondyla, and both genomic and experimental support in carpenter ants82,83, as well as the weevils P. infernalis19 and S. oryzae16,84. The increasing number of tyrosine-supplementing symbioses provide evidence for this aromatic amino acid being a key nutrient that is limiting for many insects to produce their strongly sclerotized and melanized exoskeleton for protection against desiccation and natural enemies16,18,19,82. While all the previously described symbioses that exclusively provision tyrosine precursors to their hosts involve ɣ-proteobacterial symbionts, S. silvanidophilus belongs to the Bacteroidota, providing an example for functional convergence in 34 genome-eroded symbionts across different bacterial phyla. Such convergence has been previously described for obligate endosymbionts of plant sap-sucking Hemiptera and Coleoptera that provision essential amino acids and/or vitamins to their hosts62,85,86. Interestingly, the Bacteroidota clade containing S. silvanidophilus comprises exclusively insect-associated bacteria, suggesting the possibility that the ancestor was a successful symbiont or pathogen of insects, reminiscent of the widespread reproductive manipulator Wolbachia and the common insect associate Sodalis72,87,88. Symbioses relying on nutritional supplements derived from the shikimate pathway are prone to inhibition of the aroA-encoded EPSPS by the herbicide glyphosate. Glyphosate sensitivity has already been used as an experimental tool to manipulate the obligate symbiont of tsetse flies, Wigglesworthia morsitans: chorismate derived folate (Vitamin B9) biosynthesis by Wigglesworthia was inhibited by glyphosate, resulting in delayed larval development24. Furthermore, glyphosate exposure was found to have a detrimental effect on the honey bee gut microbiota, which translated into increased mortality of the honeybees under pathogen pressure1,25,26,70,89. Concordantly, our results on an intracellular beetle symbiosis show that exposure to agronomically applied90 or previously tested glyphosate levels24,25 decreases symbiont titers and recapitulates cuticular phenotypes of aposymbiotic beetles, indicating that the inhibition of the symbiont’s shikimate pathway results in aromatic amino acid starvation of both host and symbiont. Aromatic amino acid supplementation to glyphosate-exposed beetles partially rescued the host phenotype, but not the symbiont titers, indicating that the host uses the dietary tyrosine preferentially or exclusively for its own supply. Interestingly, aromatic amino acid supplementation to symbiotic beetles suppressed the establishment of the symbiont population in the host to a similar level as glyphosate, suggesting the either the symbionts cannot grow in the absence of self-produced precursors for aromatic amino acids or the possibility that, though speculative, the host may sanction its symbionts when the latter’s nutrient supplementation is no longer needed. While this phenomenon has been documented in some dynamic partnerships of plants and their root-associated mycorrhizae and rhizobia91,92, it seems surprising in an intimate and co-evolved mutualism that may have been expected to show little potential for conflict between the partners72. Conceivably, however, symbiont titers may be regulated by the host based on tyrosine or L-DOPA concentrations during 35 normal host development, resulting in symbiont suppression in times of high tyrosine availability. The use of glyphosate in agriculture is currently heavily debated, based on increasing evidence for its detrimental impact on animals due to symbiont depletion25,26,89,93 or inhibition of cuticle melanisation22. Our findings on the glyphosate susceptibility of a beetle via its prephenate- supplementing endosymbiont exacerbate these concerns, particularly when considering our predictions based on an EPSPS phylogeny that many insects harbour endosymbionts susceptible to glyphosate. Whether insects whose symbionts provision tyrosine or its precursors among other amino acids or vitamins28,65,74,94 equally suffer from glyphosate exposure needs to be experimentally tested and will sharpen our predictions. However, the widespread occurrence of nutritional endosymbionts relying on shikimate pathway-derived nutrients paints an alarming picture and suggests that glyphosate application holds a tremendous risk of severe ecological impacts. Especially in light of recent declines in the number and diversity of insects56,95,96 and its impact on higher trophic levels97–101, the use of herbicides with potential side-effects on animals or their associated microorganisms should be carefully reconsidered. 36 2.5 Material & Methods 2.5.1 Insect cultures The initial Oryzaephilus surinamensis culture (strain JKI) was obtained from the Julius-Kühn- Institute/Federal Research Centre for Cultivated Plants (Berlin, Germany) in 2014 and kept in culture since then. Continuous symbiotic and aposymbiotic (see below) O. surinamensis cultures were maintained in 1.8-L plastic containers, filled with 50 g oat flakes, at 28°C, 60% relative humidity and a day and night cycles of 16 to 8 hours. Another O. surinamensis population (strain OsNFRI) was obtained from the National Food Research Institute (Tsukuba, Japan) and used for genome sequencing. 2.5.2 Elimination of O. surinamensis symbionts An O. surinamensis sub-population was treated for twelve weeks with tetracycline to eliminate their symbionts and then kept for several generations on a normal diet to exclude direct effects of tetracyclin on the host physiology42. Before the following experiments the aposymbiotic status of this beetle sub-population was confirmed. Therefore, 10 female adult beetles were individually separated in a single jar with oat flakes to lay eggs, as were symbiotic beetles in parallel populations. After four weeks, the adult generation was removed before their offspring finished metamorphosis and DNA of these females extracted and the symbiont titer was analyzed by quantitative PCR (see below; 42). 2.5.3 Symbiont genome sequencing, assembly, and annotation Total DNA was isolated from 20 pooled adult abdomina (without wings) of O. surinamensis JKI using the Epicentre MasterPureTM Complete DNA and RNA Purification Kit (Illumina Inc., Madison, WI, USA) including RNase digestion. Short-read library preparation and sequencing was performed at the Max-Planck-Genome-centre Cologne, Germany (SRR12881563 - SRR12881566) on a HiSeq2500 Sequencing System (Illumina Inc., Madison, WI, USA). Two further libraries library were created from O. surinamensis strain OsNFRI. For the first library the DNA was extracted by QIAamp DNA Mini Kit (Qiagen, Germany) from 210 bacteriomes dissected from 60 adults. The library was prepared using the Nextera XT DNA Library Preparation Kit (Illumina Inc., Madison, WI, USA) and sequenced on a MiSeq (Illumina Inc., Madison, WI, USA) of AIST (Japan). For the second library the DNA was extracted by QIAamp DNA Micro Kit (Qiagen, Germany) from 24 bacteriomes dissected from 6 adults (uncounted). 37 The library was prepared using the Nextera DNA Library Preparation Kit (Illumina Inc., Madison, WI, USA) and sequenced on a NovaSeq 6000 (Illumina Inc., Madison, WI, USA) of Novagen (China). Adaptor and quality trimming was performed with Trimmomatic102. In addition, we used two publicly available metagenome libraries of O. surinamensis (SRR5279855 and SRR6426882). Long-read sequencing (SRR12881567 - SRR12881568) was performed on a MinION Mk1B Sequencing System (Oxford Nanopore Technologies (ONT), Oxford, UK). Upon receipt of flowcells, and again immediately prior to sequencing, the number of pores on flowcells was measured using the MinKNOW software (v18.12.9 and 19.05.0, ONT, Oxford, UK). Flowcells were replaced into their packaging, sealed with parafilm and tape, and stored at 4°C until use. Library preparation was performed with the Ligation Sequencing Kit (SQK-LSK109, ONT, Oxford, UK) and completed libraries were loaded on a flowcell (FLO-MIN106D, ONT, Oxford, UK) following the manufacturer’s instructions. Quality-controlled long reads were mapped using a custom-made kraken2 database containing the publicly available genomes of Bacteroidota bacteria103,104 to filter beetle-associated sequences using the supercomputer Mogon of the Johannes Gutenberg-University (Mainz, Germany). Hybrid assembly of MinION and Illumina reads was performed using SPAdes (v3.13.0) with the default settings105. This resulted in ~70,000 contigs that were then binned using BusyBee Web106, screened for GC content and taxonomic identity to Bacteroidota bacteria, and additionally checked manually for tRNAs and ribosomal proteins of Bacteroidota bacteria. In total, 13 contigs were extracted, which were then automatically annotated with RAST107 using the app Annotate Microbial Assembly (RAST_SDK v0.1.1) on KBase108. The annotated contigs were plotted using CIRCOS109 (v0.69-6) for the visualisation of gene locations, GC content and coverage. Additionally, the completeness of the obtained genome was assessed with the app Assess Genome Quality with CheckM - v1.0.18 in KBase45. 2.5.4 Phylogenetic analyses A phylogenetic tree for placement of the intracellular symbiont of O. surinamensis within the Bacteroidota was reconstructed using the KBase app Insert Set of Genomes Into Species Tree v2.1.10 (SpeciesTreeBuilder v0.0.12) based on the FastTree2 algorithm110, including 49 highly conserved Clusters of Orthologous Groups (COG) genes111. 38 A phylogenetic tree of the aroA gene (which codes for the EPSPS enzyme in the shikimate pathway) from the symbiont of O. surinamensis to predict its sensitivity to glyphosate was performed according to Motta et al.25. Manually selected aroA sequences from plants, gut bacteria as well as several intracellular insect symbionts were obtained from Uniprot (UniProt Consortium 2019), translated and aligned using MUSCLE112 (v3.8.425) implemented in Geneious Prime 2019 (v2019.1.3, https://www.geneious.com). Phylogenetic reconstruction was performed with FastTree110 (v2.1.12) and PhyML113 (v2.2.4) implemented in Geneious Prime 2019 (v2019.1.3, https://www.geneious.com) using the Jones-Taylor-Thorton model with 20 rate categories and an optimized Gamma20 likelyhood (FastTree) and 1000 bootstrap replicates (PhyML). The obtained trees were visualized using FigTree (v1.4.4, http://tree.bio.ed.ac.uk/software/figtree/). 2.5.5 Comparison of bacteria Previously published Bacteroidota genomes were re-annotated with RAST107,114 in KBase108 to compare the bacteria and to estimate the genome-wide nucleotide sequence divergence level. Therefore, we identified single-copy orthologs in each genome pair using OrthoMCL115 (v2.0) in KBase. KEGG categories were then assessed via GhostKOALA116 (v2.2) of each gene’s amino acid sequence. Heatmaps were visualized using the ‘ComplexHeatmap´ package in RStudio (V1.1.463 with R V3.6.3). CIRCOS109 (v0.69-6) was used to link orthologous genes. Genomes of Bacteroidota bacteria and other bacteria described as cuticle supplementing symbionts were compared in KBase108 in more detail. Therefore, all genomes were re-annotated with RAST107 and used to classify all annotated genes according to the SEED Subsystem117 using the app View Function Profile for Genomes (v1.4.0, SEED Functional Group: Amino Acids and Derivatives). The resulting raw count of genes with annotation was visualized as a heatmap using the function ‘heatmap.2’ in the ‘ggplot´ package in RStudio (V1.1.463 with R V3.6.3). 2.5.6 Glyphosate and aromatic amino acid supplementation Eight treatments were prepared to assess the supplementation of chorismate by the symbiont to the host (Table 2). For each treatment, jars were filled with 5 g finely ground oat flakes and different combinations of aromatic amino acids (1 w/w% of each L-tyrosine, L-tryptophan and L-phenylalanine; Sigma-Aldrich, Germany) and glyphosate (0.1 w/w% or 1 w/w%; Sigma- Aldrich, Germany). 39 Table 2: Experimental treatments for assessing impact of symbiont elimination, glyphosate exposure, and dietary aromatic amino acid supplementation on cuticle traits and symbiont titers in O. surinamensis. SYM APO 1% AROMATIC 0.1% 1% AMINO ACIDS GLYPHOSATE GLYPHOSATE A x B x x C x x D x x E x x x F x x x G x H x x The glyphosate concentration of 0.1% and 1% (or 0.059 and 0.0059 mmol/g) was chosen based on the experiment by Snyder and Rio24 on tsetse flies. The 10 and 20 mM glyphosate added to the tsetse flies bloodmeal correspond to 0.1595 and 0.319% w/w glyphosate based on a blood density of 1.06 g/cm³. Helander et al report 250 mg per 48  L of soil to be equivalent to the maximum recommended amount of glyphosate for agronomical applications which translates to 0.0004% based on a fertile soil density of 1.3 g/cm³90. Food with the different supplements was weighed, mixed with distilled water, and dried overnight at 50°C in an incubator. Afterwards, the dry material was ground and filled into the jars, before 50 adult apo- or symbiotic O. surinamensis with undefined sex were added to each jar to lay eggs. After four weeks of incubation at standard conditions mentioned above, the adult beetles were removed, and the jars checked daily for adult offspring. Freshly hatched beetles were isolated in a 48-well plate with the corresponding manipulated diet and developed for seven days until cuticle biosynthesis is largely completed in symbiotic beetles43. Then the beetles were stored at -80°C for DNA extraction or fixated in 4% paraformaldehyde in PBS for histological analysis118. 40 2.5.7 Quantitative PCR DNA of 8 to 10 adult beetle abdomina (without wings) per treatment group (symbiotic and aposymbiotic parent generations and diet supplementation treatments), respectively, was isolated using the Epicentre MasterPureTM Complete DNA and RNA Purification Kit following the manufacturer’s instruction (Illumina Inc., USA) to confirm infection status and evaluate the impact of glyphosate and amino acid addition on symbiont titer. Bacterial 16S rRNA copies were quantified via quantitative PCR (qPCR) from single adult abdomina of O. surinamensis. DNA was dissolved in 30 µL low TE buffer (1:10 dilution of 1x TE buffer: 10 mM Tris-HCl + 1 mM EDTA). qPCRs were carried out in 25 µl reactions using EvaGreen (Solis BioDyne, Estonia), including 0.5 µM of each primer and 1 µl template DNA. All reagents were mixed, vortexed and centrifugated in 0.1-mL reaction tubes (Biozym, 711200). To amplify a symbiont specific 16S rRNA gene fragment the forward OsurSym_fwd2 (5’-GGCAACTCTGAACTAGCTACGC-3‘) and reverse mod.CFB563_rev (5’-GCACCCTTTAAACCCAAT-3’) primers were used42. qPCR was carried out in a Rotor-Gene Q thermal cycler (Qiagen, Germany). Standard curves with defined copy numbers of the 16S rRNA gene were created by amplifying the fragment first, followed by purification and determination of the DNA concentration via NanoDrop1000 (Peqlab, Germany). After determination of the DNA concentration, a standard containing 1010 copies/µL was generated and 1:10 serial dilutions down to 101 copies/µL. 1 µL of each standard was included in a qPCR reaction to standardize all measurements. Influence of amino acids and glyphosate on the symbiont titer of the adult beetles was analyzed using Dunn’s test from the package ‘FSA‘ in RStudio (V 1.1.463 with R V3.6.3) with two-sided, for multiple testing corrected post-hoc tests using the Benjamini-Hochberg method119. Plots were visualized using ‘ggplot2’. 2.5.8 Analysis of cuticle traits First, we determined melanisation single-blinded via the inverse of digital red color values of 8 to 10 beetles from each treatment group to evaluate the impact of glyphosate, aromatic amino acids and symbiont elimination on cuticle formation. Photographs were taken with Zeiss StereoDiscovery V8 dissection stereoscope (Zeiss, Germany) under identical conditions using ZENCore software (Zeiss, Germany). Average red values were measured within a circular area covering the thorax with Natsumushi120 and transformed into inverse red values16,42. 41 Further, we measured cuticle thickness single-blinded of 8 to 10 adult beetles per treatment group, which were fixated in 4% paraformaldehyde in PBS. These beetles were embedded in epoxy resin (Epon_812 substitute; Sigma-Aldrich, Germany), and 1 µm cross-sections of the thorax next to the second pair of legs were cut on a microtome (Leica RM2245, Wetzlar, Germany) with a diamond blade and mounted on silanised glass slides with Histokitt (Roth, Germany). Images to measure cuticle diameter were taken with an AxioImager Z2 (Zeiss, Germany) at 200x magnification and differential interference contrast. Mean cuticle diameter was measured at one randomly chosen dorsal, ventral and lateral position, respectively, with the ZEN 2 Blue software distance tool (v2.0.0.0, Zeiss, Germany). Influence of aromatic amino acids and glyphosate on the thickness and melanisation of the adult beetles was analysed by Dunn’s test from the package ‘FSA‘ in RStudio (V1.1.463 with R V3.6.3) with two-sided, for multiple testing corrected post-hoc tests using the Benjamini-Hochberg method119. Plots were visualized using ‘ggplot2’. 2.5.9 Amino acid extraction For both aposymbiotic and symbiotic O. surinamensis beetles, eight individuals for each of five different developmental stages were analysed: 5th instar larvae (of random age), one-day-old pupae, five-day-old pupae, one-day-old adults, and seven-day-old adults. Aged individuals were obtained by separating 5th instar larvae individually into a 48-well plate coated with Fluon (AGC Chemicals, UK) to avoid beetles to escape, filled with three oat flakes for provision, and daily monitoring for pupation or emergence of adults. Once they reached the desired age, individuals were frozen until further analysis. Adults had their elytrae removed to analyse them separately and both were dried overnight at 50°C. The following extraction, derivatisation, and analysis procedure of amino acids (AAs) were adapted from Perez-Palacios et al.121. For the extraction of free AA in larvae, pupae, and elytra- free adults, three stainless steel beads and 500 µL 0.1 M HCl (Merck, Germany) were added to each insect sample and homogenized in a mixer mill MM200 (Retsch, Germany) at 30 Hz for 6 min. Subsequently, samples were centrifuged at 5,000 rpm for 15 minutes at 4°C using a CT15RE microcentrifuge (Himac, Japan) and the supernatant transferred to glass vials. Pellets were kept for extraction of bound AA (see below), supernatants containing free AA were mixed with 300 µL Acetonitril (Carl-Roth, Germany) for deproteinisation and 5 µL L-norleucine (2.5 µmol/mL; 42 Sigma, Germany), which was used as an internal standard. Following centrifugation at 10,000 rpm for 3 min, samples were transferred to new vials and dried in a Savant DNA 110 SpeedVac Concentrator (Thermo Fisher Scientific, Germany). For the extraction of protein-bound AAs, the aforementioned pellets of the remaining body were resuspended in 500 µL 0.1 M HCl and transferred to separate glass vials. Similarly, dissected elytra were homogenized as described above and transferred to separate glass vials. After these samples were dried, 12 open vials (Kimble, USA) were placed in a larger hydrolysis vial (Kimble, USA) and a mixture of 187.5 µL HPLC grade water (Carl-Roth, Germany), 62.5 µL saturated phenol solution (84 g/L, Carl-Roth, Germany) and 250 µL 12 M HCl was added for gas-phase derivatisation to the bottom of the hydrolysis vial. To ensure oxygen-free atmosphere the hydrolysis vial was three times evacuated and aerated with argon and afterwards tightly sealed. The hydrolysis was performed for 24 hours at 110°C. After equilibration to room temperature, 200 µL HCl were added to the single sample vials, the supernatant transferred to a new GC vial and the samples dried again for 10 minutes in a Speedvac evaporator (Thermo Scientific, Germany). 2.5.10 Derivatisation of amino acids Derivatisation is needed to analyse amino acid via gas chromatography-mass spectrometry (GC- MS). N-tert-butyldimethylsilyl-N-methyltriflouroacetamid (MTBSTFA, Sigma-Aldrich, Germany) was utilized to silylate amino acids121: 50 µL MTBSTFA and 50 µL acetonitrile were added to all dried extracts of bound and free AA, followed by derivatisation at 100°C for 1 hour. Also, a mixture of 17 AA (each 25 µmol/L: L-alanine, L-arginine, L-aspartic acid, L-glutamic acid, glycine, L-histidine, L-isoleucine, L-leucine, L-lysine, L-methionine, L-phenylalanine, L-proline, L-serine, L-threonine, L-tyrosine, L-valine; and 12.5 µmol/L L- cysteine, Sigma-Aldrich, Germany) was prepared and derivatised, also including L-norleucine as an internal standard to identify single amino acid derivatives. 1 µL was used for subsequent analysis. 2.5.11 Amino acid analysis with gas chromatography-mass spectrometry (GC-MS) GC-MS analysis was performed with a Varian 240MS ion trap mass spectrometer coupled to a Varian 450 gas chromatograph (Agilent, USA), using external ionisation for all analyses and splitless injection with 280°C injector temperature. Initial temperature of the GC oven was 100°C for 2 min, followed by steady increase by 25°C per minute up to 300°C and final isothermal hold 43 for 5 min. The carrier gas helium had a constant flow of 1 mL/min through a DB-5ms column (30 m x 2.25 mm ID, 0.25 µm film Agilent, USA). Electron impact spectra were recorded with the ion source at 160°C and ion trap at 90°C and analyzed using the Varian MS Workstation Version 6.9.3. AA derivatives were identified using the fragmentation patterns and retention times according to Pérez-Palacios et al.121 and the external standards. Quantification was carried out with two to four fragment ions to increase signal to noise ratio (L-alanine m/z = 158 & 232 & 260 & 428; L-arginine m/z = 199 & 442; L-aspartic acid m/z = 302 & 316 & 390 & 418; L- cystine m/z = 58 & 341 & 442; L-glutamic acid m/z = 272 & 286 & 359 & 432; glycine m/z = 189 & 218 & 246; L-histidine m/z = 196 & 280.5 & 338.5 & 440.5, L-isoleucine m/z = 200 & 274 & 302; L-leucine m/z = 200 & 274 & 302; L-lysine m/z = 198 & 272 & 300; L-methionine m/z = 218 & 292 & 320; L-norleucine m/z = 200 & 274 & 302; L-phenylalanine m/z = 234 & 392 & 308 & 336; L-proline m/z = 184 & 258 & 286; L-serine m/z = 288 & 362 & 390; L- threonine m/z = 303 & 376 & 404; L-tyrosine m/z = 302 & 364 & 438 & 466; L-valine m/z = 186 & 260 & 288). The single amino acid measurements were first normalized by the internal standard to account for deviations in amount or concentration during the analysis procedure and further normalized by the total amount of all amino acids to account for differences in body size between life stages but also animals with and without symbionts. The single normalized amino acid measurements were transformed to approximate Gaussian distribution. The transformation was chosen using the powerTransform command from the package ‘car’ (alanin: negative square root, arginine: cubic root, aspartic acid: decadic logarithm, glutamic acid: square root, glycine: cubic root, isoleucine: decadic logarithm, leucine: decadic logarithm, lysine: square root, ornithine: square root, phenylalanine: square root, proline: decadic logarithm, serine: cubic root, threonine: decadic logarithm, tyrosine: square root, valine: cubic root; the amino acids cystine, histidine and methionine were not detected). Symbiont influence on the titer of the single amino acids was analysed with separate generalized linear models with life stage (larva, two pupa stages, two adult stages), amino acid source (free amino acids in body, protein-bound amino acids in body, total amino acids in elytrae) and symbiont presence/absence as factors allowing for interactions between all of them using the command glm from the ‘stats’ package in R Studio (V1.1.463 with R V3.6.3). P-values were 44 corrected for multiple testing following the classical Bonferroni122. In case of significant symbiont influence, we conducted pairwise post-hoc tests between symbiotic and aposymbiotic samples for each life stage/amino acid source using unpaired Wilcoxon rank-sum tests including Bonferroni correction122 to identify the specific life stage/amino acid source of symbiont influence. Plots were visualized using ‘ggplot2’. 2.5.12 Data availability Sequencing libraries and the assembles genome of the Oryzaephilus surinamensis symbiont (proposed Candidatus Shikimatogenerans silvanidophilus OSUR) were uploaded to the DNA Databank of Japan (accession numbers DRA010986 and DRA010987), the NCBI Sequence Read Archive (accession numbers SRR12881563 - SRR12881566) and Genbank (JADFUB000000000). Raw data of quantitative cuticle measurements are available at the data repository of the Max Planck Society ‘Edmond’123. 45 2.6 Data Accessibility Statement Sequencing libraries and the assembles genome of the Oryzaephilus surinamensis symbiont (proposed Candidatus Shikimatogenerans silvanidophilus OSUR) were uploaded to the DNA Databank of Japan (accession numbers DRA010986 and DRA010987), the NCBI Sequence Read Archive (accession numbers SRR12881563–SRR12881566) and GenBank (JADFUB000000000). Raw data of quantitative cuticle measurements are available at the data repository of the Max Planck Society ‘Edmond’123. 2.7 Acknowledgments The authors thank Dagmar Klebsch and Rebekka Janke for valuable technical assistance, Cornel Adler for the original provisioning of an O. surinamensis culture, John McCutcheon and Piotr Lukasik for their initial input on our genome sequencing approach, the Johannes Gutenberg University Mainz for computation time granted on the supercomputer Mogon, Christian Meesters’ administrative assistance on Mogon and Minoru Moriyama’s support for genome sequencing. The authors further acknowledge the financial support of the Johannes Gutenberg University Mainz (intramural funding to T.E.), a Consolidator Grant of the European Research Council (ERC CoG 819585 “SYMBeetle” to M.K.), the Japan Science and Technology Agency ERATO Grant (JPMJER1803 and JPMJER1902 to T.F.), the Japan Society for the Promotion of Science KAKENHI Grant (20J13769 to B.H.), and the Max-Planck-Society (to T.E., B.W., and M.K.). T.E. also acknowledges the stimulating International Symbiosis Society meeting 2018 in Oregon, especially inspiring presentations by Rita Rio and Joel Sachs. 2.8 Contributions T.E. and M.K. conceived the study. J.S.T.K., E.B., B.H. and T.E. sequenced and assembled the symbiont genome. J.S.T.K. and E.B. annotated the genomes and performed symbiont genomic analysis and J.S.T.K. and T.E. performed phylogenetic analyses of the symbionts. S.B., J.C.W., and T.E. performed amino acid analysis. 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The completeness of the single genomes was calculated with CheckM, which is using collocated sets of genes that are ubiquitous and single-copy within a phylogenetic lineage45. Note that this often strongly underestimates true genome completeness for obligate endosymbionts, because they tend to lose many core genes due to genome erosion. Supplementary Table 2. Aminoacyl tRNA synthetases and tRNAs encoded by the S. silvanidophilus genome in comparison with other Bacteroidota insect symbiont genomes. Three letter entries indicate the presence of the tRNA anticodons, and highlighted cells in blue indicate the presence of the corresponding aminoacyl tRNA synthetases. 52 Supplementary Figure 1. Phylogenetic tree for the placement of the intracellular symbiont S. silvanidophilus in O. surinamensis within the Bacteroidota, based on a defined list of 49 orthologous genes. The phylogeny was reconstructed using the KBase app Insert Set of Genomes Into Species Tree v2.2.0, based on FastTree2 algorithm. Node numbers represent local support values. RefSeq assembly accession in square brackets. 53 Supplementary Figure 2. Comparison of the functional gene repertoires of the endosymbiont and other Bacteroidota symbionts in insects. Colour: orthologs between S. silvanidophilus and another genome. Grey: Pairwise orthologs between the other Bacteroidota insect symbionts. 54 Supplementary Figure 3. Comparison of titers of the three amino acid tyrosine, proline, and glutamic acid that that were influenced by symbiont presence. Shown are (a) bound amino acid titers in the whole body (without elytrae in case of adults) and (b) combined free and bound titers in adult elytrae of symbiotic and aposymbiotic O. surinamensis beetles. Red: aposymbiotic beetles, Blue: symbiotic beetles. FDR corrected unpaired, two-sided Wilcox-rank-sum-tests: ns p > 0.05, * 0.05 < p < 0.01, ** p < 0.01. 55 Supplementary Figure 4. Phylogenetic classification of EPSPS enzymes from different Bacteroidota and γ-Proteobacteria insect symbionts based on PhyML and Maximum Likelihood analyses of amino acid sequences of EPSPSs using the Jones-Taylor-Thorton model and 1000 bootstrap replicates. Enzymes of plants (green), bacterial symbionts in the honeybee gut (yellow) and obligate intracellular insect symbionts (red) are highlighted. Node values indicate bootstrap values. 56 Chapter 3 Wolbachia causes cytoplasmic incompatibility, but not male-killing in a grain pest beetle Molecular Ecology, submitted Julian Simon Thilo Kiefer1, Gerrit Schmidt1, Ronja Krüsemer2, Martin Kaltenpoth1,2, Tobias Engl1,2 1Department of Evolutionary Ecology, Institute of Organismic and Molecular Evolution, Johannes Gutenberg- University, Mainz, Germany; 2Department of Insect Symbiosis, Max-Planck-Institute for Chemical Ecology, Jena, Germany 57 3.1 Abstract The endosymbiotic Wolbachia is one of the most common intracellular bacteria known in arthropods and nematodes. Its ability for reproductive manipulation can cause unequal inheritance to male and female offspring, allowing the manipulator to spread, but potentially also impact evolutionary dynamics of infected hosts. Estimated to be present in up to 66% of insect species, little is known about the phenotypic impact of Wolbachia within the order Coleoptera. Here, we describe the reproductive manipulation by the Wolbachia strain wSur harboured by the sawtoothed grain beetle Oryzaephilus surinamensis (Coleoptera, Silvanidae), through a combination of genomics approaches and bioassays. The Wolbachia strain wSur belongs to supergroup B that contains well-described reproductive manipulators of insects and encodes a pair of cytoplasmic incompatibility factor (cif) genes, as well as multiple homologues of the WO- mediated killing (wmk) gene. A phylogenetic comparison with wmk homologues of wMel of Drosophila melanogaster identified 18 wmk copies in wSur, including one that is closely related to the wMel male-killing homologue. However, further analysis of this particular wmk gene revealed an eight-nucleotide deletion leading to a stop-codon and subsequent reading frame shift mid- sequence, likely rendering it non-functional. Concordantly, utilizing a Wolbachia-deprived O. surinamensis population and controlled mating pairs of wSur infected and non-infected partners, we found no experimental evidence for male-killing. However, a significant ~50% reduction of hatching rates in hybrid crosses of uninfected females with infected males indicates that wSur is causing cytoplasmic incompatibility. Thus, Wolbachia also represents an important determinant of host fitness in Coleoptera. 58 3.2 Introduction Symbiotic bacteria influence the ecology and evolution of animals in various ways1,2. Insects harbour an especially high abundancy and diversity of microbial associations that span the entire range from parasitism to mutualism3. While some symbionts exhibit a very strict phenotype, others incur context-dependent impacts along the parasite-mutualist continuum4,5. Thus, already a single symbiont can incur context-dependent fitness benefits or costs6. However, a large proportion of insects are also infected by reproductive manipulators7 and many are co-infected with both. In consequence, their ecology and evolution can be driven by multiple symbionts with possibly different selective interests. The bacterium Wolbachia (α-Proteobacteria) is one of the most common intracellular bacteria known in arthropods and nematodes8. They are predominantly parasitic and transmitted maternally between host generations, but horizontal transmission occurs occasionally. Wolbachia employ several distinct strategies to maximize their transmission by influencing the germ line of their host. Thereby, they can rapidly sweep through uninfected populations and then maintain a high prevalence within a population. These mechanisms include cytoplasmic incompatibility (CI), parthenogenesis, male-killing or feminisation8. While CI leads directly to a higher proportion of infected individuals, the other mechanisms lead to a higher proportion of female individuals in the population. This in turn increases the fitness of Wolbachia which is predominantly transmitted maternally9. However, Wolbachia infection does not necessarily result in reproductive manipulation with negative fitness consequences for the host10. Furthermore, Wolbachia can even evolve into a mutualist and enhance its host’s fitness by supplementing dietary limited nutrients, such as B-vitamins like riboflavin11–13. CI and male-killing are the predominant strategies of reproductive manipulation in insects14,15. CI generally refers to factors localised in the cytoplasm of sperm and eggs that render them incompatible with each other, resulting in inviable embryos16–18. Wolbachia causes CI by expressing a “killing” factor in the male sperm. In eggs of uninfected females, this modification leads to non- viable embryos, whereas in infected females a “rescue” factor reverses this modification so that the zygote can develop normally19. While unidirectional CI occurs when infected males mate with uninfected females resulting in fertilized but unviable eggs, bidirectional CI occurs when two individuals are infected by different, yet incompatible Wolbachia strains8. Recently, two 59 cytoplasmic incompatibility factor genes (cifA and cifB) have been identified as key factors in CI- inducing Wolbachia strains20. The pair of CI-inducing genes were not found in the chromosomal Wolbachia genes, but in the integrated eukaryotic association module of phageWO20. A two-by- one genetic model has been suggested, specifying that while both cifA and cifB induce CI, only cifA is able to rescue the CI phenotype when transgenically expressed in the host’s ovaries16,21. The other widespread phenotype of Wolbachia inducing reproductive manipulation is male-killing. During embryogenesis, the development of the male embryo is disturbed by Wolbachia, leading to embryonic lethality8. In consequence, the fitness of infected sister embryos is enhanced by higher allocation of resources during ovogenesis and reduced intraspecific competition during juvenile development and adult life22,23. The gene WO-mediated killing (wmk) of the Wolbachia strain wMel of the fruit fly Drosophila melanogaster, has been identified to recapitulate this male-killing phenotype when transgenically expressed in D. melanogaster flies24. So far, wmk homologues were found in all Wolbachia strains associated with male-killing, surprisingly also localised within the eukaryotic association module of phageWO, only a few genes upstream from the CI-inducing genes cifA and cifB24. There are at least five homologues of the wmk gene encoded in the genome of wMel and the function of many of these remains enigmatic as only the transgenic expression of the original wmk gene, but not other homologues caused male-killing15. Wolbachia strains causing CI and male-killing phenotypes are well studied within the insect orders of Diptera and Hymenoptera, e.g. the fruit fly Drosophila melanogaster15, the southern house mosquito Culex quinquefasciatus25 and the parasitoid wasp Nasonia vitripennis26,27. Although beetles infected with Wolbachia were repeatedly reported in the last years, little is known about the functional consequences of Wolbachia infections within the order Coleoptera14,28–32. The sawtoothed grain beetle Oryzaephilus surinamensis (Coleoptera, Silvanidae) is a worldwide distributed pest of cereals and other stored food33. It is associated with the bacteriome-localised Bacteroidota endosymbiont Candidatus Shikimatogenerans silvanidophilus OSUR (called Shikimatogenerans silvanidophilus from here on)34–37. The endosymbiont S. silvanidophilus provides aromatic amino acid precursors for cuticle synthesis of the host via the shikimate pathway36. In addition, O. surinamensis is commonly infected with Wolbachia28,38. Sharaf et al. (2010) identified a higher Wolbachia infection rate in feral populations of O. surinamensis compared to adapted silo 60 populations, but also a strong female bias among adults emerging under lab conditions suggesting active reproductive manipulation by these Wolbachia strains. Elucidating Wolbachia’s capabilities of reproductive manipulation in the sawtoothed grain beetle O. surinamensis is therefore relevant in understanding the biology of this agricultural pest as well as a symbiotic model insect. In this work, we localised Wolbachia in the O. surinamensis JKI strain and quantified its growth dynamics across developmental stages. A phylogenetic analysis and functional prediction of the associated Wolbachia genome revealed it to be a member of the supergroup B, presumably capable of CI as it encodes homologues of the cytoplasmic incompatibility factor genes cifA und cifB. However, the strain is likely incapable of inducing male-killing due to a single nucleotide insertion in the identified male-killing gene wmk creating a stop codon as well as a subsequent reading frame-shift. Finally, we experimentally tested the predicted phenotype of reproductive manipulation – unidirectional CI and no male-killing – using mating assays of beetles with manipulated infection status, where we were able to verify the phenotype of reproductive manipulation via unidirectional CI. 61 3.3 Material & Methods 3.3.1 Insect cultures The initial Oryzaephilus surinamensis culture (strain JKI) was obtained from the Julius-Kühn- Institute / Federal Research Centre for Cultivated Plants (Berlin, Germany) in 2014 and kept in culture since then. Continuous symbiotic and aposymbiotic (with aposymbiotic we refer in this manuscript to beetles without both Wolbachia and S. silvanidophilus symbionts) O. surinamensis cultures (see below) were maintained in 1.8-L plastic containers, filled with 50 g oat flakes, at 28°C, 60% relative humidity and a day and night cycle of 16 / 8 hours. 3.3.2 Elimination of O. surinamensis symbionts An O. surinamensis sub-population was treated for 12 weeks with tetracycline (150 mg/5 g oat flakes, see for details Engl et al. (2018)) to eliminate both of their symbionts (S. silvanidophilus and wSur) and then kept for several generations on a normal diet to exclude direct effects of tetracycline on the host physiology. A control group was established in parallel with all steps except the addition of tetracycline to account for any unforeseen effects of the handling, population bottlenecks, etc. The apo-/symbiotic status regarding both symbionts of these beetle sub-populations was confirmed before each following experiment. Therefore, female adult beetles were individually separated into single jars with oat flakes to lay eggs. After 4 weeks, the adult generation was removed before their offspring finished metamorphosis, DNA of these parent females was extracted, and the symbiont titer was analysed by quantitative PCR (see below). 3.3.3 Quantitative PCR Absolute titers of wSur and S. silvanidophilus during host development and after different treatments from previous publications34,36,39 were determined via quantitative PCR (qPCR) amplifying respective single copy 16S rRNA gene fragments. DNA was extracted from individual beetles using the Epicentre MasterPureTM Complete DNA and RNA Purification Kit (Lucigen, Middleton, WI, USA) and dissolved in 30 µL low TE buffer (1 mM Tris–HCl + 0.1 mM EDTA). qPCRs were carried out in 25 µl reactions using EvaGreen (Solis BioDyne, Tartu, Estonia), including 0.5 µM of each primer and 1 µl template DNA. All reagents were mixed, vortexed, and centrifugated in 0.1-mL reaction tubes (Biozym, Hessisch Oldendorf, Germany). The Wolbachia- specific 16S rRNA gene fragment of O. surinamensis was amplified with the primers Wolb_16S_qPCR_fwd (5’-TTGCTATTAGATGAGCCTATATTAG-3‘) and 62 Wolb_16s_qPCR_rev (5’-GTGTGGCTGATCATCCTCT-3’)29, and the 16S rRNA of S. silvanidophilus OSUR was amplified with the primers OsurSym_fwd2 (5’-GGCAACTCTGAACTAGCTACGC-3‘) and mod. CFB563_rev (5’-GCACCCTTTAAACCCAAT-3’)34,36. qPCR was carried out on a Rotor-Gene Q thermal cycler (Qiagen, Hilden, Germany). The initial temperature was 95°C for 12 minutes, followed by 60 cycles of 95°C for 40 seconds followed by 20 seconds at 60°C. A melting curve analysis was used to assess the specificity of the qPCR reaction by a gradual increase of temperature from 60 to 95°C, with 0.25 K per second. The qPCR results were analysed using the Rotor Gene Q Software (Qiagen, Hilden, Germany). Standard curves with defined copy numbers of the 16S rRNA gene were created by amplifying the fragment first via PCR using the previously mentioned primers, followed by purification via innuPREP PCRpure (Analytik Jena GmbH, Jena, Germany) and determination of the DNA concentration via NanoDrop1000 (Peqlab, Erlangen, Germany). After the determination of the DNA concentration, a standard containing 1010 copies/µL was generated and 1:10 serial dilutions down to 101 copies/µL were prepared. 1 µL of each standard was included in a qPCR reaction to standardize all measurements. 3.3.4 Fluorescence in situ hybridisation Wolbachia was localised in O. surinamensis tissues by fluorescence in situ hybridisation (FISH) on semi-thin sections of adult beetles. Therefore, five-day-old pupae and maximum two-week-old adult beetles were fixated in tertiary butanol (80%; Roth, Karlsruhe, Germany), paraformaldehyde (37 - 40%; Roth, Karlsruhe, Germany) and glacial acetic acid (Sigma-Aldrich, Germany) in proportions 6:3:1 for 2 hours, followed by post-fixation in alcoholic formaldehyde (paraformaldehyde (37 – 40%) and tertiary butanol (80% in proportion 1:2). After dehydration, the specimens were embedded in Technovit 8100 (Kulzer, Germany)41 and cut into 8 µm sagittal sections using a Leica HistoCore AUTOCUT R microtome (Leica, Wetzlar, Germany) equipped with glass knives. The obtained sections were mounted on silanised glass slides. Each slide was covered with 100 µL of hybridisation mix, consisting of hybridisation buffer (0.9 M NaCl, 0.02 M Tris/HCl pH 8.0, 0.01% SDS; Roth, Germany) and 0.5 µM of the modified Bacteroidota probe CFB563 (5’-GCACCCTTTAAACCCAAT-3‘)26,31 or the Eubacteria probe EUB338 (5’-GCTGCCTCCCGTAGGAGT-3’)32 labelled with Cy3, as well as the two Wolbachia 63 specific probes Wolb_W2 (5’-CTTCTGTGAGTACCGTCATTATC-3‘)33 and Wolbachia-Wol3 (5’-TCCTCTATCCTCTTTCAATC-3‘)34 labelled with Cy5. DAPI (0.5 µg/mL) was included as a general counterstain for DNA. Slides were covered with glass cover slips and incubated in a humid chamber at 50°C overnight. After washing and incubating them for two hours at 50°C in wash buffer (0.1 M NaCl, 0.02 M Tris/HCl, 5 mM EDTA, 0.01% SDS), they were washed in deionized water for 20 minutes and mounted with Vectashield (Vector Laboratories, Burlingame, CA, USA). The sections were either observed under a Zeiss AxioImager Z2 with Apotome.2 (Zeiss, Jena, Germany) illuminated by a SOLA Light Engine (Lumencor, Beaverton, OR, USA), or a Leica THUNDER imager Cell Culture 3D (Leica, Wetzlar, Germany). Images obtained on the Leica microscope were processed with the instant and small volume computational clearing algorithm using standard settings in the Leica Application Suite X software (Leica, Wetzlar, Germany). 3.3.5 Symbiont genome sequencing, assembly, and annotation Total DNA was isolated from 20 pooled adult abdomina (without wings) of O. surinamensis JKI using the Epicentre MasterPureTM Complete DNA and RNA Purification Kit (Illumina Inc., Madison, WI, USA) including RNase digestion. Short-read library preparation and sequencing were performed at the Max Planck Genome Centre in Cologne, Germany (SRR12881563– SRR12881566) on a HiSeq2500 Sequencing System (Illumina Inc., Madison, WI, USA). Adaptor and quality trimming was performed with Trimmomatic35. In addition, we used two publicly available metagenome libraries of O. surinamensis (SRR5279855 and SRR6426882). Long-read sequencing (SRR12881567–SRR12881568) was performed on a MinION Mk1B Sequencing System (Oxford Nanopore Technologies (ONT), Oxford, UK). Upon receipt of flowcells, and again immediately before sequencing, the number of pores on flowcells was measured using the MinKNOW software (v18.12.9 and 19.05.0, ONT, Oxford, UK). Flowcells were replaced into their packaging, sealed with parafilm and tape, and stored at 4°C until use. Library preparation was performed with the Ligation Sequencing Kit (SQK-LSK109, ONT, Oxford, UK), and completed libraries were loaded on a flowcell (FLO-MIN106D, ONT, Oxford, UK) following the manufacturer’s instructions. Hybrid assembly of MinION and Illumina reads was performed using SPAdes36 (v3.13.0) with the default settings. The resulting contigs were then binned using BusyBee Web37 and screened 64 for taxonomic identity to α-proteobacteria. The single resulting circular contig was extracted, which was then automatically annotated with RAST38 using the app Annotate Microbial Assembly (RAST_SDK v0.1.1) on KBase39. The annotated contig was curated manually and plotted using CIRCOS40 (v0.69-6) for the visualisation of gene locations, GC content, and coverage. Additionally, the completeness of the obtained genome was assessed with the app Assess Genome Quality with CheckM—v1.0.18 in KBase39. 3.3.6 Phylogeny and comparative genomics of Wolbachia strains A phylogenetic tree for placement of the Wolbachia strain of O. surinamensis was reconstructed using the KBase app Insert Set of Genomes Into Species Tree v2.1.10 (SpeciesTreeBuilder v0.0.12) based on the FastTree2 algorithm41, including 49 highly conserved Clusters of Orthologous Groups (COG) genes. Therefore, 74 additional publicly available and published genomes of Wolbachia endosymbionts were obtained from NCBI (https://www.ncbi.nlm.nih.gov/assembly). The resulting tree was visualized using FigTree (v1.4.4, http://tree.bio.ed.ac.uk/software/figtree/). 3.3.7 Identifying genes important for reproductive manipulation The obtained genome was manually searched for wmk, cifA, and cifB genes. For the wmk gene, CDSs annotated as “Transcriptional regulator” were extracted and identified as wmk homologue by a BLASTn search of NCBI’s nucleotide collection (nr/nt). To identify the potentially functional wmk gene, the nucleotide sequence of all 18 wmk homologues of wSur and five phenotypically described wmk homologues of wMel (WD0255, WD0508, WD0622, WD0623, WD0626 (wmk))7 were aligned using MUSCLE42 in Geneious Prime 2019 (v2019.1.3; https://www.geneious.com). Phylogenetic reconstruction of the nucleotide alignment was performed with the MrBayes-plugin43 of Geneious Prime using the HKY85 substitution model and invgamma rate variation as recommended by jModelTest 2.1.10 v2016030344. The analysis ran for 1,100,000 generations, with a burn-in of 100,000 generations and trees sampled every 200 generations until the likelihood values stabilized. Protein domains were identified and annotate by running the protein sequences from the NCBI database through SMART (Simple Modular Architecture Research Tool; http://smart.embl-heidelberg.de/). Additionally, the annotated genome of wSur was manually checked for cif genes. Due to the incomplete annotation, the cif genes were first identified by whole-genome alignment to the 65 genome of wPip and translation alignment with the annotated genes of wNo (WNO_RS01055/ WNO_RS01050) and wMeg (CAI20_01650/ CAI20_01645). To identify whether the cif-genes belonged to the same type, we performed a phylogenetic analysis following Lindsey et al. (2018) and Ün et al. (2021). Briefly, the nucleotide sequences were translation-guided aligned as implemented in Geneious Prime 2019 (v2019.1.3; https://www.geneious.com). Phylogenetic reconstruction of the alignment was performed with the MrBayes-plugin43 of Geneious Prime using the GTR substitution model and gamma rate variation as predicted by jModelTest 2.1.10 v2016030344 using the same parameters as above. According to Ün et al. (2018), potential protein domains of the Cif genes were searched using HHpred’s version 3.2.0 web server (https://toolkit.tuebingen.mpg.de/tools/hhpred)45 with default parameters and the following databases: SCOPe70 version 2.07, COG/KOG version 1.0, Pfam-A version 32.0, and SMART version 6.046. The seven phage WO regions in the wSur genome were compared and visualized using Clinker47. 3.3.8 Bioassays for reproductive manipulation By mating experiments with differentially infected individuals, we tested whether Wolbachia wSur is causing reproductive manipulation in O. surinamensis. To ensure the virginity of the female and male individuals and prevent unwanted crossbreeding, pupae, and 5th instar larvae of aposymbiotic (Wolbachia & Shikimatogenerans uninfected) and symbiotic (Wolbachia & Shikimatogenerans infected) O. surinamensis were isolated into 24-well TC plates (Sarstedt AG & Co., Germany), closed with Adhesive Foil (Kisker-Biotech™, Germany) with several needles punctures to allow for air exchange and maintained under general rearing conditions (see above). The isolated individuals were observed until hatching, and the sex of the individual insect was determined by the presence (males) or absence (females) of spikes on the third femur48. Males and females were combined into mating pairs at an age of 7 - 10 days. In total, 30 mating pairs were prepared, ten for each group: The first group consisted of mating pairs where both partners, female, and male, were aposymbiotic, whereas the second group was made up of crossings with two symbiotic partners. The third group contained symbiotic males of O. surinamensis paired with aposymbiotic females. The mating pairs were given one microspatula scoop of ground oat previously filtered through a 0.6 mm sieve. To prevent the specimens from escaping the setup, the edge of each individual well was coated with Polytetrafluoroethylene 60 wt% dispersion in H2O (PTFE-dispersion; Sigma-Aldrich, Germany). For the first two weeks of the experiment all 66 pairs were left undisturbed. In the following six-week period, the number of laid eggs and hatched larvae were counted twice weekly, and the adults were placed one well further down in the 24 well-plate. In addition, we quantified the sex ratio of 100 randomly picked individuals in both symbiotic and aposymbiotic stock cultures to test for a bias induced by male-killing. 3.3.9 Statistical procedure for qPCR results and differences in hatching rate and sex ration Influence of glyphosate and tetracycline on the symbiont titer of the adult beetles (Supplement Figure 2) was analysed using Dunn’s test from the package ‘FSA‘ in RStudio (R V4.1.1) with two- sided post hoc tests corrected for multiple testing using the Benjamini–Hochberg method49,50. Compact letter display (CLD)51 was generated with the package ‘rcompanion’52. Comparison between hatching rates (Figure 7) was performed with Wilcoxon rank sum tests including correction for false discovery rates (FDRs) by repeated testing following the Benjamini–Hochberg procedure 50, implemented in the R package ‘stats’. Plots were visualized using ‘ggplot2’53. Sex ratio in beetle cultures was analysed using a manually calculated χ2 test of homogeneity. 67 3.4 Results 3.4.1 Localisation and Infection Dynamics in O. surinamensis qPCR quantification of Wolbachia titers in 106 control samples of multiple experiments indicated a wSur prevalence of 100% within laboratory cultures of the O. surinamensis JKI strain. Based on fluorescence in situ hybridisation, Wolbachia is localised throughout the entire body of O. surinamensis (Figure 1 A). Wolbachia-induced CI has been linked to sperm modification during spermatogenesis64, but Wolbachia must also be present in the female reproductive tissues for successful transmission. A close inspection of the reproductive organs of female and male O. surinamensis confirmed a high abundance of Wolbachia in both testes and ovaries (Figure 1 B and C). Further, we compared infection titers of the two bacterial endosymbionts in O. surinamensis during all life stages of O. surinamensis via qPCR (Figure 2). The population of Wolbachia reached its maximum already in the pupa during early metamorphosis (early pupa: 5.9 × 106 median copies; late pupae: 3.9 × 106 median copies; Figure 2, left), while we observed a peak of S. silvanidophilus only within the first week after metamorphosis (male 6.7 × 107 median copies and female 6.7 × 107 median copies; Figure 2, right). After our findings on S. silvanidophilus conferring enhanced cuticle synthesis and higher fitness under biotic and abiotic stresses34,36,39, we assessed whether Wolbachia could have contributed to the previously reported cuticular phenotypes in O. surinamensis. Therefore, we also quantified wSur titers (in addition to S. silvanidophilus) in O. surinamensis samples from different previous treatments (Supplement Figure 2)36. While strict tetracycline treatment eliminated both S. silvanidophilus34,36 and wSur (Kruskal–Wallis χ2 = 52.605, df = 7, p-value = 0.000000004437, Dunn’s test: p < 0.05; Supplement Figure 2), resulting in dual aposymbiotic (from here on aposymbiotic) beetles, the herbicide glyphosate had a differential effect: S. silvanidophilus was drastically reduced, yet still present in low amounts while wSur was not negatively affected (Kruskal–Wallis χ2 = 52.605, df = 7, p-value = 0.000000004437, but Dunn’s test: p > 0.05; see Supplementary Table 1 for pairwise comparisons; Supplement Figure 2). 68 Figure 1: Fluorescence in situ hybridisation micrographs of Wolbachia (green) and Shikimatogenerans (magenta) in sagittal sections (A) a 5-day-old O. surinamensis pupa stained with a Bacteroidota specific probe highlighting S. silvanidophilus (CFB563mod-Cy3, magenta), and in the gonads of (B) an adult female and (C) an adult male stained with a Eubacteria specific probe highlighting S. silvanidophilus (EUB338-Cy3, magenta), Wolbachia specific probes (Wol-W3-Cy5 and Wolb-2-Cy5, green) and DAPI targeting DNA in general (white). b=bacteriomes, c=cuticle, o=ovariole, t=testes, sv=seminal vesicle. Fig. 1A was originally published without the Wolbachia channel in Kiefer et al., (2021). 3.4.2 Genomics and Phylogeny of the Wolbachia strain We previously sequenced the metagenome of O. surinamensis combining short and long-read technologies (Illumina and ONT) into a hybrid assembly. Besides the Bacteroidota endosymbiont S. silvanidophilus36 we also extracted the full genome of a Wolbachia strain in a single, circular contig in the assembly (Figure 3). The circular genome is 1,728,764 bp in length with an average GC content of 34.1% and a coverage of 186× with short-read sequences and 94x with long-read sequences. The phylogenetic reconstruction based on 49 conserved Clusters of Orthologous Group (COG) genes classified wSur as a member of supergroup B, closely related to 69 the Wolbachia endosymbiont wEcas of the common brassy ringlet Erebia cassioides, but also within a clade with wPip of the southern house mosquito Culex quinquefasciatus and wVitB of the parasitoid wasp Nasonia vitripennis (Figure 4). Figure 2: Symbiont titers in different life stages of O. surinamensis from the JKI stock line. Titers of Wolbachia wSur (left) and S. silvanidophilus (right) were measured as 16S rDNA copies by PCR in single individuals. Juvenile life stages (eggs, larvae, and pupae) contained mixed sexes, adults were separated by sex. Larvae stages 1 to 5 (L); one-day and five-day-old pupae (P1 and P5); female adults 1, 4, and 12 weeks (W1-12) and male adults 1, 4 and 12 weeks (M1-12) after metamorphosis. The data distribution is visualized with violin plots and an additional horizontal line depicting the median. The scales of the vertical axes are logarithmic. Filled circles represent specific target amplification, and empty circles off- target amplification during late qPCR cycles, identified by melting curve analysis. The genome of the Wolbachia strain wSur of O. surinamensis encoded for 1688 protein-coding sequences, 34 tRNAs and 50 ribosomal proteins (20 SSU and 30 LSU proteins, Table 1). Besides general genetic information processing including DNA replication and repair, transcription, and translation, the genome also contained a full glycolysis pathway to process glucose-6-phosphate to erythrose 4-phosphate (E4P) and phosphoenolpyruvate (PEP). Further, it contained a full riboflavin pathway and the pathways to synthesize the amino acids lysine, glutamine, threonine, glycine, and serine, but no single gene of the shikimate pathway to synthesize aromatic amino acids, explaining its insensitivity to glyphosate65–67. 70 Figure 3: Top: Circular representation of the genome of Wolbachia wSur. The outer blue circles denote coverage with short- and long-reads, respectively (dark blue: Illumina, light blue: ONT), and the intermediate circles indicate ORFs with KEGG functional annotations separated by the direction of transcription (see legend for depicted categories). The inner grey circle denotes relative GC content and the average GC content of 34.1% is indicated by the red line. Phage WO modules are highlighted in black. Bottom: Comparison of the prophage WO modules in wSur and the well-studied strains of Wolbachia wMel, wInn, and wVitB. Prophage WO gene regions containing wmk, wmk-like homologues, and CI genes cifA and cifB are listed by Wolbachia strain in bold and then the corresponding prophage module. At least one wmk homologue is associated with each Wolbachia-inducing male-killing strain. 71 Figure 4: Phylogenetic relationship of wSur and other Wolbachia strains deposited in sequence databases. The phylogeny was reconstructed based on a defined set of 49 orthologous genes using the KBase app Insert Set of Genomes Into Species Tree v2.2.048, based on the FastTree2 algorithm50. Node numbers represent local support values. RefSeq assembly accession numbers are given in square brackets. The supergroups are colour-coded and indicated on the right93–96. Wolbachia endosymbionts of O. surinamensis (wSur, highlighted in red font) belongs to supergroup B. Wolbachia strain genomes highlighted in bold font were utilized for subsequent phylogenetic analyses of wmk (Figure 5) and cif genes (Figure 7). Table 1: Genomic characteristics of Wolbachia wSur in comparison to other strains. 72 3.4.3 Analysis of male-killing gene candidates The genome of wSur contained seven regions with phage WO-associated genes (WOSurA – WOSurG) in total, each with two to three homologues of the wmk gene (Figure 3). Overall, the genome encodes for 18 wmk homologues which were numbered from wmk1 to wmk18. As these copies may share the ability to induce male-killing, we compared these wmk homologues of wSur with the functionally described wmk homologues in the wMel strain as other known male-killing strains. The phylogenetic analysis identified homologues wmk1 and wmk12 in the phage region WOSurB as the most likely candidates to confer male-killing due to their high sequence similarity with the functional homologue wmk in wMel (for wmk12), as well as wInn and wBor (for wmk1; Figure 5, top). A closer inspection of the coding sequence revealed that wmk12 experienced an eight-nucleotide deletion that resulted in a stop codon and subsequent shift of the reading frame which led to the loss of the second XRE-family HTH DNA-binding region (Figure 5, bottom). We also screened different sequence read archives from an Israeli grain storage and two feral (field) populations of O. surinamensis individuals for wmk1/wmk12-like homologues. We found all individuals from feral populations to encode complete wmk12-like homologues clustering together in an own clade, while all individuals from the grain storage facility contained the deletion & frame-shift mutation and clustered with the wmk12 gene from wSur JKI (Supplement Figure 1). In addition, we only found wmk1-like homologues in individuals from the grain storage population, but in no individual from the feral populations. We tested for the symbiont-mediated male-killing phenotype in the JKI strain of O. surinamensis by quantifying the sex ratio in symbiotic and aposymbiotic beetle cultures. We found a uniform frequency of both sexes in both cultures (SYM 50W+50M, APO 52W+48M; χ2 test of homogeneity: χ2 = 0.080, p-value = 0.888). In addition, the male-killing phenotype should also result in a reduced hatching rate of around 50% in mating pairs with symbiotic females and males in comparison to mating pairs with aposymbiotic individuals. However, we did not observe such differences (BH corrected Wilcoxon rank sum test, p-value = 0.84; Figure 6, right). 73 Figure 5: Top: Bayesian phylogeny of wMel and wSur wmk homologues based on a nucleotide alignment. Consensus support values are shown at the branches. Bottom: Schematic of wMel and wSur wmk native nucleotide sequences. The blue tick marks indicate non-synonymous nucleotide substitutions. The red tick mark indicates eight-nucleotide deletion resulting in frame-shift mutation with a stop-codon at the deletion site. The two loci (helix-turn-helix (HTH) protein domain) of wmk are highlighted in grey. The hatched area indicates the region of wmk12 that is predicted to be not translated based on the stop codon (red tick). 74 3.4.1 Cytoplasmic incompatibility (CI) Single homologues of both previously identified CI factor genes cifA and cifB were encoded in the Wolbachia prophage region WOSurB. Bayesian phylogenetic inference identified both cifA and cifB as type II following the classification scheme of Lindsey et al. (2018) (Figure 7). The cifA gene found in wSur was closely related to those found in the Wolbachia strain wRi of the fruit fly Drosophila simulans and wSuzi of the spotted wing drosophila Drosophila suzukii, while cifB did not cluster closely with any previously described genes from other Wolbachia strains. Although the cifA gene showed no homology to known domains (Figure 7, bottom), putative domains (PD- (D/E)XK nuclease/DpnII-MboI) were found in the cifB gene. Figure 6: Influence of wSur on embryo development of O. surinamensis. Number of laid eggs (left) and hatching rate (right) in the three mating groups. The data distribution is visualized with violin plots and an additional horizontal line depicting the median. A filled sex sign indicates a symbiotic wildtype specimen, meaning infected with wSur and S. silvanidophilus, whereas an empty sign indicates these specimens as aposymbiotic (regarding both symbionts). n is the number of O. surinamensis mating pairs. Statistical significance between the groups is based on Benjamini-Hochberg corrected Wilcoxon rank sum tests (ns = not significant, * = p < 0.05, *** = p < 0.005). We tested the ability of Wolbachia infection to cause cytoplasmatic incompatibility by mating experiments with differential wSur infection. First, the impact of Wolbachia infection on the number of laid eggs was determined. As expected, infection with wSur had no effect on the number of laid eggs (Kruskal Wallis test: χ2 = 0.29, df = 2, p-value = 0.86; Figure 6, left). 75 Following further development, we observed overall differences between the three groups’ hatching rates (Kruskal Wallis test: χ2 = 10.85, df = 2, p-value = 0.004397; Figure 6, right). While the hatching rate between the control groups did not differ (aposymbiotic females and males, as well as symbiotic females and males: BH corrected Wilcoxon rank sum test: p-value = 0.84; Figure 6, right), the hatching rate in the CI cross with aposymbiotic females and symbiotic males right was reduced by 43 – 47% in comparison to both control groups (BH corrected Wilcoxon rank sum test, p-value = 0.04 and 0.0018; Figure 6). 76 Figure 7: Phylogeny and domain structure of cif genes. Top: Bayesian phylogenies based on a nucleotide alignment of cifA (top) and cifB (middle) genes. Consensus support values are shown at the branches. Coloured shapes around branches designate monophyletic “types”. Bottom: Domain structure for the cif genes of wSur. The two loci (PD-(D/E)XK nuclease/DpnII-MboI protein domain) of cifB are shown and indicated with orange bars. 77 3.5 Discussion The sawtoothed grain beetle O. surinamensis harbours not only the nutritional Bacteroidota endosymbiont S. silvanidophilus but is also infected by a pervasive Wolbachia strain. A phylogenetic analysis of the wSur core genome classified it as a member of supergroup B. Wolbachia strains of supergroup B together with supergroup A primarily infect arthropod hosts and are generally capable of reproductive manipulation, particularly by causing male-killing and CI8. The genome of the Wolbachia strain wSur of O. surinamensis encodes for 18 wmk homologues, all of which contain two helix-turn-helix (HTH) DNA-binding domains that are important for their function as a transcriptional regulator. The genomic distribution of some of the wmk homologues inside the phageWO is comparable to homologues found in other Wolbachia strains like wMel of Drosophila melanogaster which were proposed as candidate genes responsible for the induction of Wolbachia’s male-killing phenotype15,24. A phylogenetic comparison of their nucleotide sequences with previously characterized wmk homologues of wMel and wmk homologues of known male-killing strains predicted the homologue wmk1 and wmk12 of wSur as the most likely candidates to cause male-killing. However, further analysis revealed an eight-nucleotide deletion leading to a stop-codon mid- sequence and subsequent reading frame-shift of wmk12. The loss of half of the encoded protein and one of the two HTH DNA-binding domains presumably abolishes its ability to interfere with transcriptional regulation and the male-killing phenotype of wmk12 and wSur, while wmk1 could still be functional. We found no indication of symbiont-mediated male-killing in the context of the JKI strain as no differences in hatching rate could be observed between symbiotic and aposymbiotic (free of both symbionts) mating pairs. Further, JKI stock cultures of symbiotic and aposymbiotic beetles both exhibited homogenous distributions of both sexes. However, Sharaf et al. (2010) compared a feral population of O. surinamensis from the field with a population adapted to a grain storage facility. They observed a strong female bias and reduced larval survival among offspring from these feral populations emerging under lab conditions in contrast to a balanced sex ratio and higher larval survival in the population collected from the grain storage facility. They also observed incomplete Wolbachia infection in both populations: an 84% infection rate in the feral and 66% in the storage population. In combination, these data suggest active sex ratio 78 distortion in the feral populations, likely by Wolbachia, but not in the population adapted to grain storage. Individuals from the same collection site were used for genome sequencing by Hong et al. (2020) with a focus on host genomes. Our analysis of Wolbachia encoded wmk1 & wmk12-like homologues in these sequences read archives revealed the absence of wmk1 together with intact wmk12 in the two feral populations. In contrast, individuals of the Israeli population collected from the grain storage facility did encode wmk1 homologues as well as the truncated wmk12 version. Thus, we hypothesise that the intact wmk12 represents the ancestral state that mediates wSur male-killing in feral populations in Israel, while wSur from populations adapted to grain storage facilities acquired a gene duplication of wmk12 in the WOSurB region (=wmk12) and a deletion in the original wmk12 gene as well as a loss of the male-killing phenotype at least in this host genetic background. These changes probably occurred recently, possibly in the process of invasion and adaption to storage grains within co-adapted hosts that evolved probably under isolation from feral populations and facilitated by repeated strong population bottlenecks during the invasion of novel stored grain facilities or batches, as well as a relaxed selection pressure on wSur in completely infected hos populations. However, whether the changes in the wmk12 gene are causative for the loss of the male-killing phenotype remains elusive. Addition factors like host evolution or resistance to male-killing effectors could also play a role in the loss of male-killing68. Multiple homologues of wmk have been described in other Wolbachia strains, although all except one did not induce male-killing when transgenically expressed in Drosophila melanogaster15. As of right now, the function of the additional wmk homologues in wSur, as well as wMel, remains unknown. wSur and other strains might be multipotent and capable of inducing male-killing under specific conditions, or when infecting another host, like the Wolbachia strain wRec inducing CI in its main host Drosophila recens but causing male-killing when transferred to the closely related species Drosophila subiquinaria69. In addition, the Wolbachia strains might serve completely different functions to manipulate the host beyond reproductive manipulation, e.g. by affecting pheromone biosynthesis, perception, or behaviour70–73. The wmk12/1 duplication in wSur at least suggests that it is beneficial to retain a functional wmk gene, although possibly in a different context. Additional experiments, utilizing both feral and storage-adapted O. surinamensis 79 populations with hybrid crosses, or transgenic expression of different wmk genes in aposymbiotic hosts might help to shed light on their function. CI induced by Wolbachia occurs when the sperm of infected males is expressing the cif genes which lead to infertile embryos in uninfected females, while in infected females the rescue factor cifA can reverse this effect16–18. The genome of wSur encodes homologues for both CI-inducing genes cifA and cifB in one of the phage WO regions. cifA and cifB gene products are classified based on the similarity of their expressed amnio acid sequence as type I to type V20,74,75. The CI phenotype was demonstrated in cif genes of type I, II and IV20. Our analysis classified the cifA of wSur as a type II homologue, while cifB clustered in between type I and II homologues. Our experimental data indicate wSur to be a reproductive manipulator by causing unidirectional CI to its host. Crossing Wolbachia-infected males with uninfected females resulted in a hatching rate that was reduced by 45% compared to crossings between infected males and females or uninfected males and females, respectively. Findings in Drosophila simulans showed strong induction of CI leading to a hatching rate reduction of up to 95%76, while data from Drosophila melanogaster showed weak induction of CI resulting in a hatching rate reduced by 15 – 30%77, depending on environmental conditions78 as well as individual life history79. As we were so far not able to manipulate S. silvanidophilus and Wolbachia presence in O. surinamensis individually, symbiont-mediated phenotypes must be considered with great care in dual symbiont-depleted experiments. However, with the addition of genomic and ecological information, we confidently attribute the here reported CI to Wolbachia. While S. silvanidophilus presence mirrored Wolbachia in the present experiments on CI and male-killing, we have no indication for the presence of known CI factors encoded in the highly reduced S. silvanidophilus genome36, while wSur clearly contains homologues of both so far identified cytoplasmatic incompatibility factors. Thus, the Wolbachia strain wSur is likely able to influence its fitness by increasing its transmission in partially infected populations, which is reflected by its high, observed prevalence in lab conditions. Whether Wolbachia influences O. surinamensis beyond reproductive manipulation remains unclear. Previously reported cuticle supplementation of O. surinamensis is probably solely caused by S. silvanidophilus, because only the Bacteroidota endosymbiont contains the ability to synthesize aromatic amino acids precursors via the shikimate pathway to support the host’s cuticle synthesis, while wSur and Wolbachia, in general, lack the entire pathway (Kiefer at al., 2021 and this study). 80 Further, cuticle deficiencies (reduced thickness and melanisation) were not only reported in dual aposymbiotic individuals after strict tetracycline treatment (deficient of both S. silvanidophilus and wSur), but also after glyphosate treatment, which only reduced S. silvanidophilus, but not wSur titers (Supplement Figure 2)36. Thus, S. silvanidophilus is solely responsible for supplementation of cuticle synthesis as well as ecological consequences in terms of elevated resistance to abiotic desiccation stress, pathogen and predation pressure80, but also costs of symbiont infection on reproduction39. Certain Wolbachia strains were previously reported to supplement the hosts’ diet with limited nutrients (esp. B-Vitamins)11 or provide pathogen defence81. The Wolbachia strain wSur of O. surinamensis also encodes pathways to synthesize the amino acids lysine, glutamine, threonine, glycine, and serine as well as the vitamin riboflavin. While riboflavin does not seem to be limited on cereal-based diets82, lysine is83. It remains unclear whether Wolbachia might synthesize lysine for its benefit, or also contributes to its host’s metabolism. Similarly, it is unclear whether Wolbachia infection inflicts additional costs beyond unidirectional CI which is only relevant in populations with incomplete Wolbachia infection84–86. In combination with our previous work on the Bacteroidota symbiont Shikimatogenerans silvanidophilus34,36,39,80, we demonstrate that the O. surinamensis harbours two notable symbionts. Both impact the host’s physiology, ecology and thereby also its and each other’s evolution. Based on the high prevalence of both nutritional symbionts87,88 and reproductive manipulators7,29 in coleopteran and insects in general, dual infections with both types of symbionts are not uncommon and probably even underestimated31,89,90. However, currently both symbioses are usually only studied by experimental approaches in isolation, or from a descriptive perspective on the prevalence and genomic potential of both symbionts. Thereby, we miss out on potential higher levels of ecological interactions of both types of symbioses, mediated either via the host’s physiology or even directly between different symbionts. Future work should thus try to integrate multipartite, symbiotic relationships. Available tools include selective removal or inhibition of individual symbionts, e.g. by targeting specific, obligate biosynthetic pathways of symbionts. The here utilized glyphosate, inhibiting the shikimate pathway65, but e.g. also inhibitors of the Diaminopimelate pathway responsible for synthesizing lysine are prominent agents suggested for the manipulation of specific biosynthetic capabilities 81 or organisms encoding them91. Alternatively, expression of target symbiont genes in suitable host systems is a powerful tool to address gene function in insect symbionts that are elusive to genetic manipulation themselves15,17,24. Finally, the example of O. surinamensis highlights again the importance to identify systems with interesting combinations of symbionts and certain amenability for experimental manipulation and observation to understand more complex eco- evolutionary dynamics of multipartite symbioses. 82 3.6 Acknowledgments We thank Dagmar Klebsch and Benjamin Weiss for technical assistance with insect maintenance and histology, Eugen Bauer for his support on genome analysis, the Max Planck-Genome Centre Cologne (http://mpgc.mpg.de/home) for performing library preparation and Illumina sequencing, the Johannes Gutenberg-University Mainz for computation time granted on the supercomputer ‘MOGON’, and Christian Meesters for administrative assistance on ‘MOGON’. R.K., M.K. and T.E. acknowledge funding from the Max Planck Society, and further financial support of the Johannes Gutenberg-University Mainz (intramural funding to T.E.), and a Consolidator Grant of the European Research Council (ERC CoG 819585 “SYMBeetle” to M.K.). 3.7 Data Accessibility Statement Genetic data: Raw sequence reads are deposited in the SRA (SRR12881563–SRR12881566; SRR12881567–SRR12881568; BioProject PRJNA670819). The annotated wSur genome is available on GenBank (CP092526). Bioassay data is available on the data repository of the Max- Planck-Society Edmond92. 3.8 Benefit-Sharing Statement All specimens utilized in this work were obtained from a long-standing laboratory culture (pre- 2014). 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Evol. 23, 437–449 (2006). 87 3.11 Supplementary Information Supplement Figure 1: Phylogeny of wmk homologues including wmk1 and wmk12-like homologues from O. surinamensis sequencing libraries from different collection sites in Israel, including two feral field populations (SFS, SRR-52779830 – SRR5279837 and NFS, SRR5279838 – SRR5279845) and a storage facilities population (S, SRR5279846 - SRR5279853). wmk1-like homologues only occurred in wSur from O. surinamensis JKI and Israeli storage populations (S). wmk12-like homologues from storage populations (S) clustered strictly with wSur wmk12 and exhibited the same frame shift mutation as wmk12 in wSur from the O. surinamensis JKI population, while the respective homologues clustered separately and do encode a complete protein analogous to wmk from wMel. Node numbers represent posterior probabilities of Bayesian analyses. Phylogenetic reconstruction was done by Bayesian inference applying a GTR+G+I model using MrBayes (v3.2.7). The analysis ran for 200,000 generations with a “Burnin” of 25% and tree sampling every 1,000 generations. RefSeq assembly accession in square brackets. 88 Supplement Figure 2: Titer of wSur in O. surinamensis adults reared on different food compositions. The data distribution is visualized with violin plots and an additional horizontal line depicting the median. Filled circles represent specific target amplification, and empty circles off-target amplification during late qPCR cycles, identified by melting curve analysis. Different letters indicate significant differences between experimental treatments (Dunn’s Test, α ≤ 0.05). 89 Supplementary Table 1: Results of Dunn’s Test assessing the impact of glyphosate exposure on wSur titer in one-week-old O. surinamensis. Significant results (P.adj < 0.05) are highlighted in bold. Apo = aposymbiotic, Sym = symbiotic, 1% AA = addition of 1% (w/w) of each aromatic amino acid (tyrosine, phenylalanine, tryptophan), 0.1 / 1% G = addition of 1% (w/w) glyphosate Comparison Z p Adjusted p Apo - Apo + 1% AA 1.17E-01 9.07E-01 9.07E-01 Apo - Sym -3.85E+00 1.16E-04 5.41E-04 Apo + 1% AA - Sym -3.97E+00 7.13E-05 3.99E-04 Apo - Sym + 0.1% G -3.17E+00 1.52E-03 4.24E-03 Apo + 1% AA - Sym + 0.1% G -3.29E+00 1.01E-03 3.13E-03 Sym - Sym + 0.1% G 6.83E-01 4.95E-01 6.29E-01 Apo - Sym + 1% G -4.51E+00 6.53E-06 4.57E-05 Apo + 1% AA - Sym + 1% G -4.63E+00 3.73E-06 3.49E-05 Sym - Sym + 1% G -6.54E-01 5.13E-01 6.25E-01 Sym + 0.1% G - Sym + 1% G -1.34E+00 1.81E-01 2.99E-01 Apo - Sym + 1% AA -3.52E+00 4.39E-04 1.54E-03 Apo + 1% AA - Sym + 1% AA -3.63E+00 2.84E-04 1.14E-03 Sym - Sym + 1% AA 2.36E-01 8.13E-01 8.43E-01 Sym + 0.1% G - Sym + 1% AA -4.28E-01 6.68E-01 7.20E-01 Sym + 1% G - Sym + 1% AA 8.73E-01 3.83E-01 5.10E-01 Apo - Sym + 1% AA + 0.1% G -2.55E+00 1.09E-02 2.53E-02 Apo + 1% AA - Sym + 1% AA + 0.1% G -2.66E+00 7.72E-03 1.96E-02 Sym - Sym + 1% AA + 0.1% G 1.31E+00 1.91E-01 2.97E-01 Sym + 0.1% G - Sym + 1% AA + 0.1% G 6.25E-01 5.32E-01 6.21E-01 Sym + 1% G - Sym + 1% AA + 0.1% G 1.96E+00 4.98E-02 9.96E-02 Sym + 1% AA - Sym + 1% AA + 0.1% G 1.04E+00 3.00E-01 4.20E-01 Apo - Sym + 1% AA + 1% G -5.06E+00 4.09E-07 5.72E-06 Apo + 1% AA - Sym + 1% AA + 1% G -5.18E+00 2.20E-07 6.15E-06 Sym - Sym + 1% AA + 1% G -1.21E+00 2.26E-01 3.33E-01 Sym + 0.1% G - Sym + 1% AA + 1% G -1.89E+00 5.83E-02 1.09E-01 Sym + 1% G - Sym + 1% AA + 1% G -5.56E-01 5.78E-01 6.47E-01 Sym + 1% AA - Sym + 1% AA + 1% G -1.41E+00 1.57E-01 2.75E-01 Sym + 1% AA + 0.1% G - Sym + 1% AA + 1% G -2.52E+00 1.18E-02 2.54E-02 90 Chapter 4 Co-speciation and functional complementarity of dual Bacteroidota symbionts in powderpost beetles (Coleoptera: Bostrichidae) Julian Simon Thilo Kiefer1, Eugen Bauer1, Genta Okude2,3, Takema Fukatsu2,3,4, Martin Kaltenpoth1,5, Tobias Engl1,5 1Department of Evolutionary Ecology, Institute of Organismic and Molecular Evolution, Johannes Gutenberg- University, Mainz, Germany; 2Bioproduction Research Institute, National Institute of Advanced Industrial Science and Technology, Tsukuba 305-8566, Japan; 3Department of Biological Sciences, Graduate School of Science, University of Tokyo, Tokyo 113-0033, Japan; 4Graduate School of Life and Environmental Sciences, University of Tsukuba, Tsukuba 305-8571, Japan; 5Department of Insect Symbiosis, Max-Planck-Institute for Chemical Ecology, Jena, Germany 91 4.1 Abstract Insects frequently engage in stable, nutritional symbioses with bacteria and exhibit co- evolutionary processes that result in co-adapted metabolisms and co-speciation. Further, multiple insects are associated with two or more endosymbionts with complementary biosynthetic pathways to synthesize amino acids and/or vitamins. Here, we report the first case of dual co- obligate symbiosis with closely related endosymbionts of the same family in beetle hosts. Beetles of the family Bostrichidae harbour consistently the aromatic amino acid supplementing Bacteroidota bacterium Shikimatogenerans bostrichidophilus. Further, a sub-clade is associated with the co-obligate symbiont Bostrichicola ureolyticus which probably complements the function of Shikimatogenerans by recycling urea and provisioning the essential amino acid lysine, providing additional benefits on nitrogen- and/or lysine-poor diets. Both symbionts represent ancient associations within the Bostrichidae. While Bostrichicola was repeatedly lost, Shikimatogenerans was retained throughout the family and exhibits a perfect pattern of co-speciation. 92 4.2 Introduction Many insects are associated with microbial partners in host-beneficial symbioses1–4. Supplying the symbiont with general nutrients in a stable niche in exchange for essential nutrients allows the host to thrive on challenging, often nutritionally imbalanced diets, like plant sap, wood, or vertebrate blood5. A stable symbiotic association can lead to co-evolutionary dynamics, including co-adaptation and speciation6,7. Strong population bottlenecks during symbiont transmission, genetic drift, and host-level selection result in such scenarios in rapid genomic changes which leads to metabolic specialisation of the nutritional symbiont8. The outcome is drastically reduced symbiont genomes encoding besides the obligate pathways to sustain their metabolism under extensive host provisioning only for biosynthetic pathways to supplement absolutely necessary nutrients to the host metabolism9. Symbionts can also be lost or replaced when they are no longer needed or capable of sufficiently supporting their host’s metabolism – as genome erosion can lead to reduced efficiency in the symbionts10–12. It is also possible that symbionts are not replaced, but only part of their function is complemented by novel microbial partners. This has led to e.g. dual symbioses in which essential amino acids are still synthesized by the original symbiont, but vitamins are synthesized by a secondary symbiont13–15 or pathways for the essential amino acids are split up between two symbionts16–18. Several insects benefit from symbionts that mainly or even uniquely provide aromatic amino acid precursors for cuticle synthesis, which can be limited in the diet and/or in high demand during metamorphosis. This has been found to be important for cuticle melanisation and sclerotisation, as all of the cuticular crosslinking agents are derived from the aromatic amino acid tyrosine19,20. Shortly after metamorphosis, when the cuticle of the adult beetle has fully formed, the symbiont contribution and the population is at their peak – which was demonstrated in the sawtoothed grain beetle Oryzaephilus surinamensis21 (Chapter 3) and the grain weevil Sitophilus oryzae22. Tyrosine-supplementing symbionts can be bacteria in the gut like in turtle ants of the genus Cephalotes23,24 or the bean weevil Callosobruchus maculatus25, but also bacteriome-localised symbionts like the Candidatus Westeberhardia cardiocondylae (Enterobacteriaceae) in the tramp ant Cardiocondyla obscurior26, the γ-proteobacterial symbionts Candidatus Nardonella and Candidatus Sodalis in weevil species22,27, as well as the Bacteroidota endosymbiont Candidatus Shikimatogenerans silvanidophilus in O. surinamensis (Chapter 2). For simplicity’s sake we will skip the Candidatus nomenclature from here on for all symbionts with sequenced genomes. 93 Tyrosine-supplementing symbioses provide evidence for this aromatic amino acid being a key nutrient for many insects to produce their strongly sclerotised and melanised exoskeleton, thereby increasing desiccation resistance and protection from predators and pathogens - coincidentally also susceptibility for herbicides targeting the shikimate pathway22,27–30. Based on this, it has been hypothesized that beetles inhabiting ecological niches with low ambient humidity or high antagonist pressure may have been predisposed towards evolving symbiotic associations with tyrosine-supplementing symbionts28,31. Concordantly, O. surinamensis and – to a lesser extent – Sitophilus have been able to invade dry grain storage facilities, with the help of their symbionts32,33. While several tyrosine-supplementing symbionts are known, several additional beetle families have recently been described to harbour related symbionts for which the functions are not yet known31,34,35. In addition, the evolutionary histories of the symbioses remain largely unknown. A beetle family with associated Bacteroidota symbionts closely related to the endosymbiont Shikimatogenerans silvanidophilus of O. surinamensis are the powderpost beetles (Coleoptera: Bostrichidae)31,35. The family Bostrichidae (Latreille, 1802) is between 17036 and 15537 Mya in age and consists of nine subfamilies (Liu & Schönitzer 2011) with varying ecology: Apatinae (Jacquelin du Val, 1861), Bostrichinae (Latreille, 1802), Dinoderinae (C. G. Thomson, 1863), Dysidinae (Lesne, 1921), Endecatimonae (LeConte, 1861), Euderinae (Lesne, 1934), Lyctinae (Billberg, 1820), Pylcvaoninae (Lesne, 1896) and Psoinae (Blanchard, 1851). While all Bostrichidae are phytophagous and bore into twigs, branches or trunks of dead or dying trees, in which their offspring develop, the most well-known species are pests of wood products or stored foods38,39. The close relation of the Bacteroidota endosymbionts with Blattabacterium spp., Sulcia muelleri, but especially S. silvanidophilus and a similar ecological niche of the hosts as several of the beetle species profiting from cuticle supplementation suggest a similar symbiont function. Thus, to understand the functional contribution and evolution of the symbiosis within the Bostrichidae, we collected 29 beetle species across the family and performed metagenome sequencing. Based on multiple closed or draft symbiont genomes and multiple mitochondrial and nuclear host genes, we reconstructed the molecular phylogenies of both partners. Thereby we demonstrate (i) that most bostrichids are associated with a symbiont whose genome is highly degraded, retaining only the shikimate pathway for tyrosine precursor provisioning. (ii) Only beetles of the genera Lyctus and Dinoderus were associated with a second Bacteroidota symbiont. 94 (iii) This second co-obligate symbiont also exhibited a highly eroded genome but encodes complementary functions of nitrogen recycling, and lysine biosynthesis. Finally, (iv) host and symbiont phylogenies exhibited a high degree of co-cladogenesis, suggesting an ancient association that early resulted in obligate dependencies and co-diversification. 95 4.3 Results We collected and sequenced the metagenomes of 28 beetle species of the family Bostrichidae (Supplement Tables 1 and 2). In addition, we supplemented our dataset with three publicly available datasets from NCBI (Apatides fortis [FJ613421], Sinoxylon sp. SIN01 [JX412742], and Xylobiops basilaris [SRR2083737]). The resulting 31 species covered five subfamilies and nine tribes within the Bostrichidae. For thirteen of the 31 species, we were able to assemble the full and circularized genome of the Bacteroidota endosymbiont Shikimatogenerans bostrichidophilus, with the longest symbiont genome being 200,377 bp and the shortest 172,971 bp in length, and an average GC content of 15.1% (Supplement Table 2). For ten additional species, we assembled draft genomes of Shikimatogenerans based on multiple contigs extracted from the metagenome assemblies via taxonomic classification, GC content filtering, as well as by manually searching for tRNAs and ribosomal protein genes as well as enzymes of the shikimate pathway of Bacteroidota bacteria. For one species (Dinoderus bifoveolatus) we only obtained the 16S rRNA sequence of the endosymbiont. In the remaining species we sequenced, we were not able to detect any sequence of Bacteroidota bacteria in the metagenome data. In addition, we extracted the 16S rRNA and aroA gene sequences of the Bacteroidota symbiont from a publicly available transcriptomic dataset of Xylobiops basilaris (SRR2083737). As expected from a previous study31, we found the genome of a second Bacteroidota endosymbiont, which we named Bostrichicola ureolyticus (see below), in some species of the subfamilies Lyctinae and Dinoderinae. However, we only detected this co-obligate symbiont within the Lyctus and Dinoderus species, but not the other members of the Lyctinae (Trogoxylon impressum) and Dinoderinae (Rhyzopertha dominica). The genomes of Bostrichicola were on average 337,500 bp in length and had an average GC content of 22.4%. The metagenomic datasets were used to reconstruct the phylogeny of the host species (Figure 1 left and Supplement Figure 1). We constructed two phylogenies of the host, one based on the assembled mitochondrial genomes (Figure 1 left), and the other one on 22 aligned, and concatenated Benchmarking Universal Single-Copy Ortholog (BUSCO)40 genes found across all species (Supplement Figure 1). Both phylogenies gave very similar results, revealing two main clades of the Bostrichidae beetles, separating the Lyctinae and Dinoderinae from the Euderinae, Apatinae and Bostrichinae. However, they differ in the placement of Micrapate scabrata, within 96 the Sinoxylonini as a sister clade to the Xyloperthini (BUSCO genes) versus as an outgroup to the before mentioned clades (based on the mitochondrial genomes; Supplement Figure 1). Similarly, the genome assemblies were used to generate a whole genome-based phylogeny for the endosymbionts (Figure 1 right). This phylogenetic reconstruction based on 350 conserved genes confirmed the monophyly of the Shikimatogenerans endosymbionts of Bostrichidae beetles and their close relationship to other insect-associated Bacteroidota bacteria, specifically to Blattabacterium spp. and Sulcia muelleri, which had been previously reported based on 16S rRNA phylogenies31,34,35. The Bostrichicola symbionts in the Lyctus as well as Dinoderus species clustered in a distinct, more basally branching, also monophyletic clade. The two clades of Bostrichidae endosymbionts were separated by the Bacteroidota symbiont Shikimatogenerans silvanidophilus OSUR of the sawtoothed grain beetle Oryzaephilus surinamensis (Silvanidae) as well as the clade of Sulcia muelleri endosymbionts of the Auchenorrhyncha (Hemiptera). A second phylogeny of the endosymbionts based on the 16S rRNA sequences that allowed us to include more taxa revealed a highly similar distribution within the Bacteroidota (Supplement Figure 2). The main difference between both phylogenies (Supplement Figure 2) was the placement of the endosymbiont of Calopertha truncatula, which forms an outgroup to endosymbionts of Sinoxylonini and of Xyloperthini in the 16S rRNA-based phylogeny, while in the whole genome phylogeny it groups within the endosymbionts of Sinoxylonini as a sister clade to endosymbionts of Xyloperthini. A comparison of the endosymbiont phylogeny based on the 350 conserved genes and the host mitochondrial phylogeny showed a high degree of co-cladogenesis (Figure 1). We chose this comparison as we were able to retrieve a higher number of full mitochondrial genomes than BUSCO gene sets. However, Micrapate scabrata was an outgroup for the Xyloperthini and Sinoxylonini while its endosymbiont grouped within the Sinoxylonini as a sister clade to the Xyloperthini, rather than reflecting the host evolutionary background observed in the BUSCO gene phylogeny (Supplement Figure 2). The comparison between the full continuous genomes shows a high degree of synteny within, but not between, the two symbiont genera (Figure 3). On the following page, Figure 1: Comparison of phylogenetic relationships between Bostrichidae beetle hosts and their Bacteroidota endosymbionts. Left: Bayesian phylogeny of 30 Bostrichidae beetle species inferred from concatenated nucleotide alignment of 13 mitochondrial genes. Right: Bayesian phylogeny of 28 Bacteroidota symbionts of Bostrichidae beetles inferred from concatenated nucleotide alignment of 350 genes. Node numbers represent posterior probabilities of Bayesian analyses. Host-symbiont associations are highlighted by connecting trapezoids between the phylogenies. 97 98 Figure 2: (A) Reconstructed metabolism of the two Dinoderus porcellus endosymbionts Shikimatogenerans bostrichidophilus DPOR and Bostrichicola ureolyticus DPOR, inferred from genomic data. Enzymes and arrows in grey were missing in the genome annotation. Dashed arrows indicate transport processes without annotated transporters. (B) Comparison of the functional gene repertoires of Bacteroidota symbionts in Bostrichidae beetles. Coloured boxes indicate the presence, and white boxes the absence of genes in the symbiont genomes. Box colours are based on KEGG’s categories (see legend for depicted categories). 99 As expected, the metabolic repertoires of both Bostrichidae endosymbionts were highly reduced and showed clear signs of genome erosion. Both endosymbionts retained genes involved in the cellular core processes of general genetic information processing including DNA replication and repair, transcription, and translation (Supplement Figure 3). In addition, Shikimatogenerans that were present across the entire family encoded all the genes of the shikimate pathway except a shikimate dehydrogenase (aroE [EC:1.1.1.25]) (Figure 2). Also, these genomes encoded the bifunctional aroG/pheA gene (phospho-2-dehydro-3-deoxyheptonate aldolase/chorismate mutase [EC:2.5.1.54 5.4.99.5]), capable of catalysing the Claisen rearrangement of chorismate to prephenate and the decarboxylation/dehydration of prephenate to phenylpyruvate41. The genome of Bostrichicola encoded both urease α and γ subunits (ureC [EC:3.5.1.5]) to recycle nitrogen, as well as a glutamate dehydrogenase (gdhA [EC:1.4.1.4]) that allows integration of the resulting ammonium into the amino acid metabolism via glutamate (Figure 2 A). In addition, they encoded for aspartate aminotransferase (aspB [EC:2.6.1.14]) to transfer the amino group from glutamate to oxaloacetate, as well as an almost complete diaminopimelate pathway to synthesize the essential amino acid lysine from aspartate. They also retained a methionine synthase to convert L-homoserine to L-methionine and can synthesise menaquinone, but we did not find a single gene of the shikimate pathway to synthesize aromatic amino acids. However, the genomes encoded for a complete fatty acid and peptidoglycan biosynthesis, albeit other cell envelope components apparently cannot be synthesized. The genomic data revealed no transporters, so it remains unknown how the symbionts exchange metabolites with the host and with each other (Figure 2 A). In addition, genes encoding signal transduction, cell surface structures, and motility were absent (Figure 2 B). Further, we also compared the set of encoded genes which are not encoded in all genomes (Supplement Figure 4). For Shikimatogenerans, it is particularly noticeable that mutL (DNA mismatch repair protein) is still present in the Dinoderinae+Lyctinae symbionts but has already been lost in the Euderinae+Apatinae+Bostrichinae. For Bostrichicola, all genes for peptidoglycan biosynthesis (murA, murB, murC, murD, murE, murF, murG, mraY and mrcA) are still encoded in the Dinoderinae, whereas in the Lyctinae L. brunneus lost murC, murD and murG, while L. cavicollis has lost all of them. 100 Figure 3: Gene order comparison between S. bostrichidophilus and B. ureolyticus genomes that could be assembled into full continuous genomes, showing a high degree of synteny within, but not between, the two symbiont genera. Grey shades show the percentage of identity between homologous proteins from different genomes (based on amino acid sequences). The phylogenetic tree on the left is based on the symbiont phylogeny displayed in Figure 1. Based on the close phylogenetic relationship to the Shikimatogenerans silvanidophilus OSUR and the presence of the shikimate pathway in the highly eroded genome, we propose the name ‘Shikimatogenerans bostrichidophilus’ for the endosymbiont of Bostrichidae beetles, henceforth called S. bostrichidophilus. The genus name Shikimatogenerans refers to its ability to perform the shikimate pathway. Previous studies have shown that closely related Bacteroidota bacteria are also associated with other beetle families such as the Silvanidae and the Nosodendridae31,42. Thus, we propose bostrichidophilus as a species epithet to indicate that this symbiont clade is associated with beetles of the family Bostrichidae. As all the symbionts encode highly similar genomes, we propose to add a four-letter abbreviation of the host species to identify the strain (first letter of the host genus and first three letters of the host species epithet), e.g. 101 Shikimatogenerans bostrichidophilus RDOM for the endosymbiont of Rhyzopertha dominica. For the second co-obligate endosymbiont found in Bostrichidae beetles of the subfamily Dinoderinae and Lyctinae, we propose the name ‘Bostrichicola ureolyticus’. Its genus name refers to its association with bostrichid beetles, while ureolyticus refers to its metabolic potential to recycle nitrogen from urea as inferred from the genomic data. In analogy to Shikimatogenerans, we propose to add a four- letter abbreviation of the host species to identify the strains, e.g. Bostrichicola ureolyticus LBRU for the Bostrichicola endosymbiont of Lyctus brunneus. Based on rRNA fluorescence in situ hybridisation with eight of the 32 species, the bacterial symbionts were localised intracellularly in bacteriomes in the abdomen of the host (Figure 4). The bacteriomes are located between the gut, fat body and reproductive organs, but without direct connection to any of these tissues. Bostrichid beetles of all subfamilies harboured one paired bacteriome with symbionts stained by a probe specific for members of the Shikimatogenerans symbiont clade30,31 (Figure 4 a, e, f, g and h). In addition, species of the genera Dinoderus and Lyctus contained a pair of bacteriomes stained by a probe specific to the second symbiont B. ureolyticus31 (Figure 4 b, c and d). In the species containing both Shikimatogenerans and Bostrichicola, the bacteriomes were close to each other, sometimes with direct physical contact yet distinct ultrastructure (Figure 4 b). 102 Figure 4: Fluorescence in situ hybridisation micrographs of Shikimatogenerans bostrichidophilus and Bostrichicola ureolyticus in sections of (a) Trogoxylon impressum, (b) Lyctus cavicollis, (c) Dinoderus minutus, (d) Dinoderus porcellus, (e) Rhyzopertha dominica, (f) Prostephanus truncatus, (g) Xyloperthella picea and (h) Sinoxylon anale. Sections are stained with a S. bostrichidophilus specific probe (magenta), a B. ureolyticus specific probe (yellow), and DAPI targeting DNA in general (white). Scale bar representing 20 µm. 103 4.4 Discussion In this study, we characterized the intracellular bacterial symbionts across 29 species of the beetle family Bostrichidae and assessed their functional potential and co-speciation with their hosts based on comparative genomics. We used the metagenomes to reconstruct the first molecular phylogeny of these so-called auger or powderpost beetles (Coleoptera: Bostrichidae), following the morphological phylogeny by Liu et al (2011). Based on shotgun metagenomics datasets, we extracted beetle nuclear and mitochondrial gene sequences and reconstructed two phylogenies of the Bostrichidae, one based on 13 mitochondrial genes (Figure 1 left), and the other one on 22 BUSCO genes found across all species (Supplement Figure 1). Both phylogenies were generally well supported, highly congruent, and separated the Bostrichidae into two main clades: The Lyctinae and Dinoderinae grouped together, as did the Euderinae, Apatinae and Bostrichinae. The main difference was the placement of Micrapate scabrata. In the host phylogeny based on BUSCO genes, Micrapate scabrata clustered within the tribe Sinoxylonini as a sister clade to the Xyloperthini (BUSCO genes). In contrast, in the mitochondrial phylogeny, it was an outgroup to both tribes (Supplement Figure 1). Our study serves as the first, solid molecular evaluation of the relationships within this family that had been repeatedly reclassified43 and only been addressed with a well-sampled phylogeny based on morphological characteristics by Lui & Schönitzer in 2011. Overall, we arrived at similar conclusions with one major exception: Liu & Schönitzer placed the Euderiinae with a single monotypic genus as a basal branch of the Bostrichidae and suggested to even placing them in a separate family. In our analyses, Euderia squamosa always clustered within the Bostrichidae in a separate branch between the Lyctinae/Dinoderinae and Apatinae/Bostrichinae. The Lyctinae and Dinoderinae are in both studies closely related, with the Psoinae and Polycaoninae interspersed in Liu & Schönitzer but lacking in our work, as we were not able to obtain any specimens. The Apatinae and Bostrichinae form here two sister groups within a monophyletic clade. The Sinoxylini and Xyloperthini are monophyletic groups within the Bostrichinae, while the Bostrichini represents a polyphyletic group39, regardless of whether using the mitochondrial or BUSCO gene phylogeny. 104 The presence of endosymbionts in several species of Bostrichidae beetles known as grain and wood pests was already described by Gambetta (1927)44, Mansour (1934)45, Koch (1936)46, and Buchner (1954)47. Okude et al. (2017) and Engl et al. (2018) identified them as members of the insect-associated Bacteroidota (Flavobacteriaceae) clade31,35. In this study, we were able to detect Shikimatogenerans bostrichidophilus in almost all examined Bostrichidae species. For the few species where we could not detect any symbiont (Dinapate wrightii, Amphicerus bicaudus, Heterobostrychus aequalis and Sinoxylon japanicum), three scenarios are possible. First, it is known from several symbiotic insects that the endosymbiont is degraded or lost in male beetles sometime after metamorphosis48. In cases where nutrient supplementation by the symbionts is only needed during larval development and/or metamorphosis, males can benefit from killing their symbionts and recycling symbiotic organs, given that they do not transmit the symbionts to their offspring21,22. Thus, individual beetle specimens in our study may have been males, resulting in false negatives in our symbiont screenings. Second, in some species, aposymbiotic individuals and even populations occur in the field, due to elevated sensitivity of the symbionts to environmental stressors like heat46,49 or certain contemporary agrochemicals. Third, Shikimatogenerans may have been lost within these species. As we had only single specimens available for the species in which we failed to detect symbionts, we cannot confidently reject any of these hypotheses. However, based on their similar ecological niches, we assume the entire family of Bostrichidae to harbour S. bostrichidophilus originating from a single acquisition event. In contrast, the co- symbiont B. ureolyticus was only present within the genera Lyctus and Dinoderus. In these cases, it is unlikely that we missed the co-symbiont in other species of these two subfamilies, T. impressum and R. dominica. We had multiple individuals and life stages of R. dominica available as well as four specimens of T. impressum and never found any indication for a second bacteriome-localised symbiont, neither within our genomic datasets nor during FISH. Concordantly, previous studies31,35 did not report on a second symbiont in R. dominica, while it could already be discerned based on morphologically differentiated bacteriomes in Lyctus and Dinoderus species31,46. Thus, B. ureolyticus was likely acquired by either an ancestor of all Bostrichid beetles, or only the ancestor of the Lyctinae and Dinoderinae, and repeatedly lost. Phylogenetic reconstructions based on either the symbiont 16S rRNA gene or 350 genes resulted in highly congruent phylogenies that only differed in some of the deeper splits, which were better 105 supported in the multi-gene phylogeny than the 16S rRNA phylogeny. The endosymbiont and the host phylogeny showed a high degree of co-cladogenesis, mirroring each other’s branching patterns almost perfectly. The major disagreement was again the clustering of Micrapate scabrata, respectively its symbiont. M. scabrata itself was an outgroup for the Xyloperthini and Sinoxylonini while its endosymbiont groups within the Sinoxylonini symbionts as a sister clade to the Xyloperthini. Both endosymbionts are characterized by extremely small, heavily eroded and A+T-biased genomes with very limited biosynthetic capabilities50. The genomes of S. bostrichidophilus encoded for the shikimate pathway to synthesize precursors of aromatic amino acids. Of the seven canonical genes in the shikimate pathway (aroG, aroB, aroD, aroE, aroK, aroA, aroC), the S. bostrichidophilus genomes only lack the gene for shikimate dehydrogenase (aroE), which catalyses the reversible reduction of 3-dehydroshikimate to shikimate. However, the shikimate pathways of Nardonella EPO, the endosymbiont of the sweetpotato weevil Euscepes postfasciatus (Curculionidae: Cryptorhynchinae)27,51 and Carsonella ruddii, the endosymbiont of the gall- forming psyllid Pachypsylla venusta (Aphalaridae: Pachypsyllinae)52, as well as the closely related S. silvanidophilus OSUR in the sawtoothed grain beetle O. surinamensis also lack aroE, but remain functional30, suggesting that the function of aroE is likely taken over by other enzymes, either host or endosymbiont. S. bostrichidophilus transformers phosphoenolpyruvate (PEP) and erythrose- 4-phosphate (E4P) to prephenate/chorismate via the shikimate pathway53,54, which is then converted by the host to the aromatic amino acid tyrosine55. As all of the cuticular crosslinking agents are derived from the tyrosine19,20, it plays a key role in melanisation and sclerotisation, which is what gives the cuticular its protective and desiccation-resistant abilities31,56. As the contribution of aromatic amino acids by Shikimatogenerans, Nardonella and Sodalis30,51,57 is especially important as precursors for cross-linking chitin and proteins in the context of cuticle synthesis in beetles, the contribution of B. ureolyticus might be relevant in the same context. The genome of B. ureolyticus encodes the gene for urea recycling and the diaminopimelate pathway to synthesize lysine. In addition, B. ureolyticus retained partial pathways to convert intermediates of the lysine biosynthesis into methionine, folate and menaquinone biosynthesis5,58 as well as certain components of the cell envelope biosynthesis (fatty acids and peptidoglycan). Insights from other symbionts indicate that even incompletely retained biosynthetic pathways are often functional 106 and beneficial for the host17,27,30,52,59. Multiple insects are associated with two or more endosymbionts with complementary biosynthetic pathways to synthesize amino acids and/or vitamins15,17,58,60–62, but so far not with such closely related endosymbionts of the same family. Nitrogen recycling is well documented within Bacteroidota endosymbionts63–65 and can be an important benefit for insects developing in nitrogen-poor sapwood38,39,66–68. The recycling of urea as a source of amino groups might be important for the formation of tyrosine, but also other amino acids and amino acid-derived components like N-acetyl-glucosamine, the monomer of chitin24. However, why B. ureolyticus retained a lysine biosynthesis pathway is less clear. Lysine is an important component of cuticular proteins as its ε-amino group represents an anchor point for cross linking69. Grain diets are specifically limited in lysine70, which could be relevant for the stored product pest beetles of the genus Dinoderus, but also other species of the Dinoderinae subfamily. Thus, the specific role of B. ureolyticus and why it was only retained in certain genera remains speculative. A profound understanding of why certain genera of Bostrichidae might benefit from such provisioning and thus retain Bostrichicola, while others do not, is currently hampered by the scarcity of information on the ecology and physiology of most Bostrichidae43. Based on our phylogenetic analyses, Shikimatogenerans and Bostrichicola are derived from the same ancestor as Blattabacterium spp., Walczuchella monophlebidarum and Uzinura diaspidicola, but then diverged and evolved different functional specialisations30,31,42,71. Interestingly, S. silvanidophilus OSUR30 – the sister taxon of S. bostrichidophilus – retained the shikimate pathway as well as one urease subunit putatively involved in nitrogen recycling30 - and there is evidence for nitrogen recycling in Blattabacterium63,65,72 and Walczuchella64, so urea catabolism seems to be a widespread benefit provided by Bacteroidota bacteria. Within the Bostrichoidae the Bostrichidae family split from the Ptinidae family between 17036 and 15537 Mya ago in the Jurassic period. S. bostrichidophilus started to radiate at the same estimated time, around 274-158 Mya ago31. The ancestor of the Bostrichidae beetle family must have acquired the Shikimatogenerans symbiont before that point in time, indicating that tyrosine- provisioning was likely an important benefit and remained important for the wood- and grain- feeding species. Bostrichicola was likely acquired 100 Mya ago31, in the ancestor of the Dinoderinae and Lyctinae (plus possibly Psoinae and Polycaoninae), and then lost in Rhyzopertha and Trogoxylon. Alternatively, the related bacteria have been acquired twice independently, i.e. in 107 Dinoderinae and Lyctinae, respectively. The repeated independent acquisitions of these specific clades of Bacteroidota symbionts indicate that these bacteria were once specialized in infecting insects, akin to Wolbachia (Chapter 3) or Sodalis73,74. This hypothesis is supported, as a basal clade of Bacteroidota endosymbionts is described as male-killing endosymbionts in different ladybug beetles75–77. Whether this represents an uptake of both symbionts within the ancestor of the Bostrichidae, or an even older association remains enigmatic, as closely related families within the Bostrichoidea (Dermestidae, Ptinidae)37 are not known to harbour nutritional Bacteroidota or other nutritional, bacterial endosymbionts, but in some cases (Anobiinae) yeast-like endosymbionts78. In this context it is worth mentioning that Bacteroidota endosymbionts of this clade were in the last years described in multiple beetle families, including the Silvanidae31,34 and Nosodendridae42, indicating either a widespread ancestral association or high potency for infection and establishment in beetles. 108 4.5 Material & Methods 4.5.1 Insect collection Specimens of 29 species were collected or provided by experts in the field from Germany, the Czech Republic, Yemen, the United Arabic Emirates, the United States of America, Japan, and New Zealand (Supplement Table 1). In addition, three publicly available data sets of bostrichid beetles were retrieved from NCBI (SRR2083737, FJ613421 and JX412742). 4.5.2 Symbiont genome sequencing, assembly, and annotation Total DNA was isolated using the Epicentre MasterPureTM Complete DNA and RNA Purification Kit (Illumina Inc., Madison, WI, USA) including RNase digestion, or the QIAGEN Genomic- tip kit using 20/G columns (Qiagen, Hilden, Germany). Short-read library preparation and sequencing were performed at the Max-Planck-Genome-Centre Cologne, Germany (SRR19201352 - SRR19201388) on a HiSeq3000 Sequencing System (Illumina Inc., Madison, WI, USA), at CeGaT on a HiSeq2500 Sequencing System (Tübingen, Germany). Adaptor and quality trimming was performed with Trimmomatic79. Long-read sequencing (SRR19201386, SRR19201357 and SRR19201352) was performed on a MinION Mk1B Sequencing System (Oxford Nanopore Technologies (ONT), Oxford, UK). Upon receipt of flowcells, and again immediately before sequencing, the number of pores on flowcells was measured using the MinKNOW software (v18.12.9 and 19.05.0, ONT, Oxford, UK). Flowcells were replaced into their packaging, sealed with parafilm and tape, and stored at 4°C until use. Library preparation was performed with the Ligation Sequencing Kit (SQK-LSK109, ONT, Oxford, UK) and completed libraries were loaded on a flowcell (FLO-MIN106D, ONT, Oxford, UK) following the manufacturer’s instructions. PacBio long-read sequencing of D. porcellus was performed at the Max-Planck-Genome-Centre Cologne, Germany (SRR19201385) on a Sequel II (PacBio, Menlo Park, CA, USA). Quality-controlled long reads were taxonomy-filtered using a custom-made kraken2 database80,81 containing the publicly available genomes of Bacteroidota bacteria to extract beetle-associated Bacteroidota sequences using the supercomputer Mogon of the Johannes Gutenberg-University (Mainz, Germany). Assembly of Illumina reads and additional hybrid assemblies with long-read libraries were performed using SPAdes (v3.15.0)82. The resulting contigs were binned using BusyBee Web83, and screened for GC content and taxonomic identity to Bacteroidota bacteria. 109 The extracted contigs were de novo assembled in Geneious Prime 2019 (v2019.1.3, https://www.geneious.com). The resulting contigs were then automatically annotated with PROKKA84 using the app Annotate Assembly and Re-annotate Genomes (v1.14.5) on KBase85. Synteny analysis of complete endosymbiont genomes was performed using Clinker86. 4.5.3 Fluorescence in situ hybridisation Endosymbionts of bostrichid beetles were localised by fluorescence in situ hybridisation (FISH) on semi-thin sections of adult beetles, targeting the 16S rRNA sequence. Adult beetles were fixated in tertiary butanol (80%; Roth, Karlsruhe, Germany), paraformaldehyde (37 - 40%; Roth, Karlsruhe, Germany) and glacial acetic acid (Sigma-Aldrich, Germany) in proportions 6:3:1 for 2 hours, followed by post-fixation in alcoholic formaldehyde (paraformaldehyde (37-40%) and tertiary butanol (80% in proportion 1:2). After dehydration, the specimens were embedded in Technovit 8100 (Kulzer, Germany)100 and cut into 8 µm sagittal sections using a Leica HistoCore AUTOCUT R microtome (Leica, Wetzlar, Germany) equipped with glass knives. The obtained sections were mounted on silanised glass slides. Each slide was covered with 100 µL of hybridisation mix, consisting of hybridisation buffer (0.9 M NaCl, 0.02 M Tris/HCl pH 8.0, 0.01% SDS; Roth, Germany) and 0.5 µM of the Shikimatogenerans bostrichidophilus specific probe (5’-CTTCCTACACGCGAAATAG-3’; Engl et al. 2018) labelled with Cy5, as well as the Bostrichicola ureolyticus specific probe (5’-TACTCGATGGCAATTAACAAC-3’; Engl et al. 2018) labelled with Cy3. DAPI (0.5 µg/mL) was included as a general counterstain for DNA. Slides were covered with glass cover slips and incubated in a humid chamber at 50°C overnight. After washing and incubating them for 20 minutes at 50°C in wash buffer (0.1 M NaCl, 0.02 M Tris/HCl, 5 mM EDTA, 0.01% SDS), they were washed in deionized water for 20 minutes, dried and mounted with Vectashield (Vector Laboratories, Burlingame, CA, USA). The sections were observed under a Zeiss AxioImager.Z2 equipped with an Apotome.2 (Zeiss, Jena, Germany) and illuminated by a SOLA Light Engine (Lumencor, Beaverton, OR, USA). 4.5.4 Phylogenetic analyses We generated phylogenetic trees based on the metagenome data generated from our bostrichid taxa (SRR19201352 - SRR19201388) as well as three Bostrichidae from NCBI (SRR2083737, FJ613421 and JX412742). 110 A phylogenetic tree of the mitochondrial genes of the hosts was reconstructed by assembling the mitochondrial genome using NOVOPlasty87 and MitoZ88 and afterwards annotating them with Mitos89 (http://mitos.bioinf.uni-leipzig.de/index.py). Subsequently, 13 mitochondrial genes were translated and aligned using MUSCLE90 (v3.8.425) as implemented in Geneious Prime 2019 (v2019.1.3, https://www.geneious.com). Additionally, we generated a second (codon- based) nucleotide alignment based on Benchmarking Universal Single-Copy Orthologs (BUSCO) using a custom pipeline40 to extract the genes from the metagenome datasets. According to Shin et al. (2021)91, BUSCO analysis was performed for each dataset using the insecta_odb10 database (1,658 genes) to extract BUSCO genes that were found across all species. The corresponding nucleotide sequences were then extracted and aligned in MAFFT92 with --auto and default options. Gaps in the resulting alignment were then trimmed from the alignment using trimAl93 (v1.2), accepting 5% gaps for each position. Afterwards, the aligned nucleotide sequences for each taxon were concatenated. For the phylogenetic placement of the intracellular symbionts of bostrichid beetles, the encoded gene sequences were extracted from the genomes, aligned based on the nucleotide sequence, and concatenated in Geneious Prime 2019 (v2019.1.3, https://www.geneious.com). Additionally, beetle symbiont 16S rRNA sequences were aligned to representative Bacteroidota 16S rRNA sequences obtained from the NCBI database, using the SILVA algorithm31,94,95. Since complete genomes were not available for some of the species, the 16S rRNA alignment allowed us to incorporate a larger number of species in the phylogenetic analysis, albeit at a lower resolution due to the limited amount of information contained in this single gene. Phylogenetic reconstructions for all alignments were done by Bayesian inference applying a GTR+G+I model using MrBayes96–99 (v3.2.7). The analysis ran for 10,000,000 generations with a “Burnin” of 25% and tree sampling every 1,000 generations. We confirmed that the standard deviation of split frequencies converged to < 0.01. The obtained trees were visualized using FigTree (v1.4.4, http://tree.bio.ed.ac.uk/software/figtree/). 111 4.6 Data Accessibility Statement Sequencing libraries and the assembles genome of the bostrichid symbionts (proposed Shimatogenerans bostrichidophilus and Bostrichicola ureolyticus) were uploaded to the NCBI Sequence Read Archive (see Supplement Table 1 for accession numbers) and GenBank (see Supplement Table 2 for accession numbers). 4.7 Acknowledgments We thank Philipp-Martin Bauer, Hans-Georg Folz, Cornel Adler, Rudy Plarre, Rich Leschen, Michael Eifler, and Miguel Diaz for providing Bostrichid specimens. We thank Benjamin Weiss for technical assistance in histology, Bruno Hüttel and the Max Planck-Genome-Centre Cologne (http://mpgc.mpipz.mpg.de/home/) for performing library preparation and sequencing of most samples in this study, Yu Okamura for help with the BUSCO pipeline as well as the Johannes Gutenberg-University Mainz for computation time granted on the supercomputer ‘MOGON’, and Christian Meesters for administrative assistance on ‘MOGON’. M.K. and T.E. acknowledge funding from the Max Planck Society, and further financial support of the Johannes Gutenberg-University Mainz (intramural funding to T.E.), as well as a Consolidator Grant of the European Research Council (ERC CoG 819585 “SYMBeetle” to M.K.). 4.8 Contributions J.S.T.K. and T.E. designed the project, and J.S.T.K., E.B., G.O. and T.E. sequenced and assembled the symbiont genome. J.S.T.K. and E.B. annotated the genomes and performed symbiont genomic analysis and J.S.T.K. and T.E. performed phylogenetic analyses. 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Supplement Table 2: General features of the symbiont genomes based on annotations with PROKKA. LSU: Large subunit ribosomal protein, SSU: Small subunit ribosomal proteins. 117 Supplement Figure 1: Comparison of symbiont phylogenies based on the 16S rRNA gene alone (left), and 350 genes conserved across at least two genomes (right). Node numbers represent posterior probabilities of Bayesian analyses. 118 Supplement Figure 2: Comparison of host phylogenies based on 22 BUSCO genes (left) and 13 mitochondrial genes (right). Node numbers represent posterior probabilities of Bayesian analyses. 119 Supplement Figure 3: Relative abundance of Clusters of Orthologous Groups (COG). Annotated functional categories (A-Z) and relative proportion of the encoded genes represented as a heatmap are indicated on the right-hand side. 120 Supplement Figure 4: Comparison of differentially encoded genes in genomes of Bostrichicola ureolyticus (left) and Shikimatogenerans bostrichidophilus (right). Genomes were annotated with PROKKA. Hypothetical proteins were removed. Filled: present, white: missing 121 Chapter 5 Loss of genomic cohesion: Symbiont lineage splitting in the large grain borer Prostephanus truncatus (Coleoptera: Bostrichidae) Julian Simon Thilo Kiefer1, Eugen Bauer1, Martin Kaltenpoth1,2, Tobias Engl1,2 1Department of Evolutionary Ecology, Institute of Organismic and Molecular Evolution, Johannes Gutenberg- University, Mainz, Germany; 2Department of Insect Symbiosis, Max-Planck-Institute for Chemical Ecology, Jena, Germany 123 5.1 Abstract The endosymbionts Shikimatogenerans bostrichidophilus (Bacteroidota), associated with beetles of the Bostrichidae family, supply their host's diet solely with prephenate, the precursor for the aromatic amino acid tyrosine. Within the larger grain borer Prostephanus truncatus, the genome of the associated endosymbiont has diverged into three strains. To better understand the lineage splitting of S. bostrichidophilus PTRU we utilized different microscopy approaches to verify the genomic predictions by localising the individual genes and genomes in the bacteriome. The genomes of the three strains are very similar, encoding for a core set of genes involved in processing genetic information. However, genes for the shikimate pathway and the ribosomes are encoded by the three different strains, complementing each other, suggesting that these genes or their products have to be shared between symbiont cells. As this phenomenon of lineage splitting did not add any functionality to the symbiosis, it is likely a non-adaptive result of ongoing genome erosion. 124 5.2 Introduction Similar to organelles, intracellular symbionts depend on the host for their existence. A phenomenon that can be observed very often in endosymbiotic bacteria is genome reduction1,2. In this process, the genome undergoes a process in which the encoded genes are undergoing inactivation and complete loss, leading to a genome with a stable set of necessary encoded functions3. The erosion of a genome is thought to be driven by unselective processes like genetic drift and mutation accumulation1 as well as by selection at the symbiont and host level4,5. The result of this process is a genome with the minimum set of functions, typically encoding a few genes outside of the core processes of replication, transcription, translation, and nutrient provisioning1. Ongoing genome reduction puts the symbiont under tremendous evolutionary pressure as purifying selection acts on the symbiont genes that are important for functions that the host needs6,7. Further, endosymbionts can also be lost, replaced, or complemented when they are no longer needed or incapable of sufficiently supporting their host’s metabolism. In several cases, this has led to dual symbioses in which the essential amino acids are still synthesised by the original symbiont, but vitamins are synthesised by a secondary symbiont, or certain pathways being split between the symbionts8,9. One example is the glassy-winged sharpshooter Homalodisca vitripennis (Hemiptera: Cicadellidae), where the β-proteobacterial symbiont Baumannia retains pathways for vitamins needed by the host, while the Bacteroidota symbiont Sulcia muelleri retains genes for the production of most essential amino acids, resulting in metabolic complementarity8. Beetles of the family Bostrichidae associate with two Bacteroidota endosymbionts, i.e. Shikimatogenerans bostrichidophilus and Bostrichicola ureolyticus (Chapter 3). However, the lesser grain borer Rhyzopertha dominica and the large grain borer Prostephanus truncatus in the subfamily Dinoderinae have lost B. ureolyticus and retain only S. bostrichidophilus10,11 (Chapter 3). Both R. dominica and P. truncatus are found in a similar ecological niche as the sawtoothed grain beetle Oryzaephilus surinamensis with its closely related endosymbiont Shikimatogenerans silvanidophilus (Chapter 2). Shikimatogenerans symbionts exhibit highly reduced genomes specialised in supplementing their hosts diet with nutrients via the shikimate pathway crucial for a healthy cuticle synthesis. However, in contrast to S. silvanidophilus, S. bostrichidophilus even lost the 125 glycolysis pathway to produce precursors for the shikimate pathway. Additionally, early studies by Koch and Huger suggest a higher degree of integration and host dependency, as it is not possible to eliminate the endosymbiont of R. dominica and P. truncatus12,13, while aposymbiotic O. surinamensis can be readily obtained by antibiotic treatment or exposure to elevated temperatures10,13, and symbiont-free cultures can be maintained for years10. In this study, we used whole-genome sequencing and fluorescence microscopy to report an unusual phenomenon in Dinoderinae beetles, in which the endosymbiont S. bostrichidophilus PTRU of P. truncatus has diverged into three strains within its host. The genes for the shikimate pathway and the ribosomes are encoded by the three different strains, complementing each other but also showing a considerable level of redundancy in encoded functions. Lineages splitting apparently did not add any functionality to the symbiosis—the three symbiont genomes of the S. bostrichidophilus PTRU strains in P. truncatus perform together exactly the same functions as the single genome of S. bostrichidophilus RDOM in R. dominica, so this phenomenon is likely a non- adaptive result of ongoing genome erosion. 126 5.3 Results 5.3.1 Bostrichidae-symbiont genomes are highly eroded The metagenomes of the lesser grain borer Rhyzopertha dominica and the large grain borer Prostephanus truncatus were sequenced using short-read (Illumina) technology to gain deeper insights into symbiont metabolic capabilities. For R. dominica, we could confirm the 16S rRNA sequences of a Bacteroidota bacterium that matched PCR-based Sanger sequences from previous studies10. The de novo assembled genome of the symbiont of R. dominica is 173 kbp in length with an average GC content of 13.3% (Figure 1). The genome encodes for 169 genes and had a coverage of 1,751x. Like the genome of S. silvanidophilus OSUR (Chapter 2), the genome of S. bostrichidophilus RDOM is highly reduced, with most of the encoded genes dedicated to genetic information processing including DNA replication and repair, transcription, and translation. Additionally, it encodes all genes of the shikimate pathway except a shikimate dehydrogenase (aroE [EC:1.1.1.25]). Also, the genome encodes the bifunctional aroG/pheA gene (phospho-2- dehydro-3-deoxyheptonate aldolase/chorismate mutase [EC:2.5.1.54 5.4.99.5]), capable of catalysing the Claisen rearrangement of chorismate to prephenate and the decarboxylation/dehydration of prephenate to phenylpyruvate14. In contrast, the metagenome of P. truncatus revealed sequences of more than one Bacteroidota endosymbiont. We were able to assemble genomes of three highly similar, yet distinct endosymbiont strains (henceforth called PTRU strains A, B, and C; Figure 1). All three of them share a highly similar 16S rRNA sequence, but their genomes diversify in length and encoded genes (pairwise identity of aligned PTRU strain 16S rRNA sequences: A + B = 98.8%, B + C = 98.5%, A + C = 98.7%, A + B + C = 98.7%). The genome of PTRU strain A is 157 kbp in length with a GC content of 15.4%, encoding for 136 genes with an average coverage of 704x. PTRU strain B is 152 kbp in length, has a GC content of 15.4% and the genome encodes for 124 genes with an average coverage of 655x. Finally, the genome of PTRU strain C is 160 kbp in length, has a GC content of 15.3% and is encoding for 139 genes with an average coverage of 792x. The genomes of the three Shikimatogenerans bostrichidophilus PTRU strains together show a very similar gene composition as the genome of S. bostrichidophilus RDOM, but some of the genes are only present in one or two of the three strains (Figure 2, Supplementary Figure 1). In consequence, the genes of the shikimate pathway are distributed over the three strains, whereas only one of the 127 six genes (aroF) is present in all three strains (Figure 2). For strain A, the gene 5- enolpyruvylshikimate-3-phosphate synthase (aroA [EC:2.5.1.19]) is unique, while the other four genes of the pathway (aroC, aroD, aroK and aroQ) are unique to strain C. Hence, the shikimate pathway is only complete when at least strain A and strain C complement each other, while strain B is dispensable. Figure 1: Circular representation of the genomes of Shikimatogenerans bostrichidophilus RDOM of Rhyzopertha dominica and the three PTRU strains in Prostephanus truncatus. The outer grey circles denote coverage with short reads, and the intermediate circles indicate annotated functional KEGG categories separated by direction of transcription (see legend for depicted categories). The inner grey circle denotes relative GC content and the average GC content in per cent is indicated by the red line. 128 Figure 2: (A) Comparison of the functional gene repertoires of the endosymbionts. Grey: Genes present in RDOM and also in the three PTRU strains, red: shikimate genes, blue: small subunit ribosomal proteins, orange: large subunit ribosomal proteins. (B) Comparison of shikimate pathway genes and ribosomal proteins present in genomes. Filled: present, white: missing. (C) Comparison of all differentially encoded genes in genomes of RDOM and PTRU strains. Genomes were annotated with PROKKA15. Hypothetical proteins were removed. Filled: present, white: missing. 129 By PCR amplification, cloning and sequencing of the 16S rRNA gene, SNPs could indeed be confirmed between the individual 16S rRNA sequences of the three strains, which makes it possible to reconstruct a 16S rRNA-based phylogeny (Figure 3 A). However, these differences between the three sequences are not detectable in a pooled bacterial PCR Sanger sequencing approach, explaining why we previously missed the strain differentiation10. By combining the cloned 16S rRNA sequences with the shotgun short-read assembly, we were able to assign the three 16S types to the respective strains. The phylogenetic reconstruction based on the 16S rRNA sequences confirmed the placement of the Bacteroidota endosymbionts of R. dominica and P. truncatus as sister species in a group of insect-associated bacteria, closely related to Shikimatogenerans silvanidophilus OSUR (S. silvanidophilus), Blattabacterium sp. and Sulcia muelleri (S. muelleri) (Figure 3 A and Chapter 4). We also used the shikimate pathway gene aroF to reconstruct a phylogeny as it represents the only gene of the pathway being present in all three strains (pairwise identity of aligned nucleotide aroF sequences: A + B = 96.2%, A + C = 95.5%, B + C = 95.1%, A + B + C = 95.5%; pairwise identity aligned amino acid aroF sequences: A + B = 96.3%, A + C = 94.6%, B + C = 94.9%, A + B + C = 95.4%; Figure 3 B). Besides the shikimate pathway, also the ribosomal proteins (rProteins) of the large and the small subunit are distributed across the three strains (Figure 2). We investigated if all three strains of the endosymbiont in P. truncatus are present in the same ratio during development. Therefore, we used a gene of interest (GOI) encoded by the strains to measure the titer in all life stages of the host: First, the sequence of aroA encoded in the genome of PTRU strain A, second the sequence of aroF encoded in the genome of PTRU strain B and third the sequence of aroC encoded in the genome of PTRU strain C. With the three GOIs, we detected constant ratios in symbiont titers across development, reaching their maximum in the adult life stage. Interestingly, titers of strain A were consistently higher than those of strain B (approximately 2.4-fold on average), and titers of strain B were approximately 11.2-fold higher than those of strain C. 130 Figure 3: Bayesian phylogeny of (A) nucleotide alignment of 16S rRNA sequences and (B) the translation- guided alignment of nucleotide sequences of aroF Node numbers represent posterior probabilities of Bayesian analysis. 5.3.2 Localisation of the three strains of P. truncatus in the bacteriome In a previous study, the bacteriomes of R. dominica and P. truncatus were localised via FISH by targeting the 16S rRNA of the symbionts10. We repeated the FISH for both species, yielding very similar bacteriome structures and no information on the distribution of the three different strains in P. truncatus (Figure 5 f). Thus, we used the cloned 16S rRNA sequences of the three PTRU strains to design strain-specific FISH probes to distinguish the single strains in the bacteriome. We could show that the designed probes for PTRU strain A (SymType2), PTRU strain B 131 (SymType1) and PTRU strain C (SymType3) indeed highlighted three distinct types of localisations in the bacteriome (Figure 6 A). All cells labelled as strain B also showed a signal with the probe targeting strain C, but 50-70% of strain C-labelled cells were not stained for strain B. PTRU strain A-stained cells were dominant and mainly found alone, but often in contact with strain C. Figure 4: Titers of the three PTRU strains across life stages of Prostephanus truncatus, measured via qPCR with strain-specific primers targeting aroA (strain A), aroF (strain B) and aroC (strain C). In addition, we made use of the hybridisation chain reaction technique (HCR) to label strains on the gene level (Figure 6 B and C). We used genes of the shikimate pathway as GOIs to identify the individual strains in the bacteriome (Figure 6 B). The probes visualizing the shikimate gene aroA (strain A) and aroC (strain C) were often co-localised, while aroF signals (strain B) occurred alone. As aroA is only encoded by the genome of PTRU strain A and aroC is only encoded in the genome of PTRU strain C, this result is indicating a co-localisation of these two strains in the bacteriome. Further, we used the sequences of rProteins which are uniquely encoded in the individual PTRU strain genomes (Figure 6 C). Here, the signal of rProteins encoded by PTRU 132 strain A is co-localised with the signal of rProteins encoded by PTRU strain B, while the signal of the rProteins encoded by the genome of PTRU strain C showed no co-localisation with any of the two other strains. Figure 5: Images of Rhyzopertha dominica (a) and Prostephanus truncatus (e) (kindly provided by Thomas Hörren). Fluorescence in situ hybridisation micrographs of R. dominica (b-d) and P. truncatus (f-h) stained with a Shikimatogenerans bostrichidophilus specific probe (orange) and DAPI targeting DNA in general (white). Scale bars represent 20 µm. 133 Figure 6: (A) Fluorescence in situ hybridisation micrographs of a Prostephanus truncatus bacteriome stained with strain-specific 16S rRNA probes: strain A (SymType2, yellow), strain B (SymType1, cyan) and strain C (SymType3, magenta). (B) HCR-FISH on an isolated bacteriome of P. truncatus, to visualize strains of Shikimatogenerans bostrichidophilus PTRU using probes against unique sequences of shikimate pathway genes aroA (strain A, yellow), aroF (strain B, cyan) and aroC (strain C, magenta) and (C) unique ribosomal protein (rProtein) sequences of strain A (yellow) strain B (cyan) and strain C (magenta). DNA of host and symbiont stained with DAPI (white). Scale bar representing 10 µm. 134 5.1 Discussion In this study, we sequenced the metagenomes of the larger grain borer Prostephanus truncatus. Contrary to our expectations, we assembled not only one but three genomes of the Bacteroidota symbiont Shikimatogenerans bostrichidophilus PTRU. We compared the genomes of the three PTRU strains to the close relative S. bostrichidophilus RDOM, the endosymbiont of the lesser grain borer Rhyzopertha dominica (Chapter 4), revealing that all three strains of PTRU together contain the same metabolic capacity as RDOM in R. dominica. The genomes encode only genes of the shikimate pathway apart from gene families involved in the core processes of replication, transcription, and translation. It is noteworthy, that the genes of the shikimate pathway are spread over all three strains, however, two of the strains are sufficient to complete the pathway. Of the six genes of the shikimate pathway (aroA, aroB, aroC, aroF, aroK and aroQ), only aroF is encoded in the genome of every strain. Although aroB, aroC, aroK and aroQ are encoded in the genome of PTRU strain C, aroA is uniquely encoded in the genome of PTRU strain A. The lack of the shikimate pathway gene aroE was also described in other tyrosine-supplementing bacterial endosymbionts: S. silvanidophilus of O. surinamensis, Nardonella EPO of the sweetpotato weevil Euscepes postfasciatus (Curculionidae: Cryptorhynchinae) and Carsonella ruddii in the gall-forming psyllid Pachypsylla venusta (Aphalaridae: Pachypsyllinae). However, the shikimate pathway remained functional16,17 (Chapter 2), suggesting that the function of aroE can be taken over by other enzymes, either from the host or the endosymbiont—that have yet to be identified. Besides genes of the shikimate pathway, several other genes are encoded only by one or two of the PTRU strains. Among these, the ribosomal proteins (rProteins) are particularly noteworthy. From the whole set of rProteins, some are uniquely encoded in one of the three strains, resulting presumably in each strain being required to assemble functional ribosomes. This scenario is similar to what has been seen in Hodgkinia in some species of the cicada genus Tettigades, where rProteins tended to only be encoded on the most abundant genome18. The symbionts are localised in the cytoplasm of the bacteriocytes. As the genomes of S. bostrichidophilus in R. dominica and P. truncatus lack genes for the synthesis of peptidoglycan, the symbiont cells show amorphous morphology, known as the L-form11,19–21. Also, several studies on other bacteriome-localised endosymbionts found that the symbionts are often polyploid, stating that there are multiple copies of the genome per cell22–24. If we assume the same for 135 S. bostrichidophilus, then the genomes of the three PTRU strains can either be mixed in the same symbiont cell, or they can be segregated into different cells. In the latter case, the symbiont strains must exchange either mRNAs, proteins/enzymes and/or entire ribosomes and metabolites to be functional. Okude et al. (2017), while researching the endosymbionts of R. dominica, already showed via FISH microscopy (Okude et al 2017, Figure 1 g-j) and transmission electron microscopic (TEM) images (Okude et al 2017, Figure 5 b) of the bacteriome that the symbiont cells are connected in a grape-like pattern. In our FISH micrographs, we see the same pattern for both R. dominica and similar ones in P. truncatus, where different endosymbiont cells are connected. These connections between single cells could be a prerequisite for the evolution of separate strains and the starting point for S. bostrichidophilus to diverge into multiple strains. If they can at least still exchange metabolites via these connections, there would not be any drastic negative consequences if the genome of every cell no longer encoded all the individual enzymes and ribosomal proteins. In P. truncatus, likely, cells of the three PTRU strains are still connected in this manner, building a continuum where all necessary gene products would be present and accessible for every strain, even when specific genes are not encoded in its genome. Intriguingly, this includes genes encoding enzymes involved in genetic information processing, such as dnaE (the DNA polymerase III alpha subunit) and rpoA (the RNA polymerase subunit alpha), which are missing in PTRU strain C (Figure 4 C). Thus, the products of genes involved in essential bacterial cellular functions need to be transported among cells in this continuum. This raised the question of how the shikimate pathway as well as the ribosomes can still be functional when the necessary genes are distributed across different symbiont genomes. To approach this question, we visualized the different strains in the bacteriome. The FISH based on the 16S rRNA sequences of the three PTRU strains showed that the strains matching SymType2 and SymType3 are mainly co-localised, while the 16S rRNA matching SymType1 shows little to no co-localisation. The HCR FISH with probes targeting genes of the shikimate pathway showed that aroA and aroC are mostly co-localised, indicating a co-localisation of strain A and strain C. In terms of the shikimate pathway, this would make perfect sense, since the co-localisation of these two strains would complete the pathway. In the same image, the signal detecting aroF is present in some cells too, but mostly it is localised on its own. Regarding the qPCR, in which the titers of the three strains via individual genes of the shikimate pathway were measured, a relatively high signal for aroF can also be seen here. This could be caused by the method of HCR 136 itself. While primers were used in the qPCR which specifically amplifies the sequence of the aroF from PTRU strain B, this sequence was divided into eleven small fragments for the HCR to serve as a probe. Some of these fragments may also have non-specifically aligned to the aroF sequence of PTRU strains A and C, resulting in false positive signals. In contrast, the HCR with the sequences of the rProteins as a probe shows a completely different picture. Here, it appears that the rProtein sequences encoded by PTRU strain A reside in the same symbiont cells as the rProtein sequences encoded by PTRU strain B, while the rProtein sequences encoded in the genome of PTRU strain C stand alone. The reason for this contrasting picture again could be based on the HCR method itself. The respective sequences of the single ribosomal proteins alone are not long enough to design sufficient single-gene probes. Therefore, we selected several ribosomal proteins per strain, based on which the probes were generated. Since the three genomes only differ in individual genes, but they are largely identical, it can happen in the assembly that individual positions were assembled incorrectly. This could be improved by using long reads by third- generation sequencing technologies in addition. Nevertheless, our findings of three sequences of the shikimate pathway gene aroF in addition to the three 16S rRNA sequences strongly indicate the presence of three strains in the bacteriome of P. truncatus. In direct comparison, both the 16S rRNA FISH and the two HCRs performed show the same picture: two of the three strains are usually co-localised, while the third shows little to no co-localisation. The only question that remains open is how the individual sequences and genes are distributed across the three genomes. After metamorphosis, there is no more pressure on the endosymbiont as they uniquely contribute to precursor synthesis for cuticle formation. However, bacteriomes are retained for symbiont transmission in female beetles. During this period, the quality of the symbionts may then no longer be controlled at host-level and less fit symbiont genotypes could accumulate25. Something similar was observed in cicadas of the genus Tettigades (Hemiptera: Cicadidae) which are associated with the endosymbiont Hodgkinia cicadicola (α-proteobacteria). Łukasik et al. (2017) found that in Tettigades the ancestral Hodgkinia has split independently six times over the last 4 Mya, which results in a complex of two to six Hodgkinia lineages per host25,43. The authors argue that the long dormant period without host-level selection on symbiont functionality (relative to the period of larval growth where the symbionts are important) may allow for slightly deleterious processes to occur, such as lineage splitting18. Also, they tested the idea of this being adaptive by allowing for adjusting expression levels by gene copy numbers - instead of transcriptional 137 regulation - which is likely impaired in genome-eroded symbionts due to the loss of most or all transcription factors. In contrast, in P. truncatus the adult lifespan is very long with females living approximately 61 days and males living approximately 45 days26. Complete development from egg to adult took approximately 37 day27, whereas the pupation time is only five days until emerging of the adult beetle26. Following, in female adults, the endosymbionts spend a long time being functionally irrelevant to the host, which may allow for lineage splitting. In different species such non-adaptive and rather degenerative and finally probably “costly” processes can take place in symbioses. Campbell et al. (2018) showed in cicadas associated with the symbionts Sulcia muelleri and several Hodgkinia linages that the host has increased the number of transmitted Hodgkinia cells six-fold, whereas the number of S. muelleri cells is unaltered44. This example shows that there is a dose adaption in transmission of strains, i.e. those with many strains pass on significantly more symbionts via the eggs to ensure that really all strains, including the rare ones, are successfully transferred to the next generation. 138 5.2 Material & Methods 5.2.1 Insect cultures The initial Rhyzopertha dominica and Prostephanus truncatus cultures (strain JKI) were obtained from the Julius-Kühn-Institute/Federal Research Centre for Cultivated Plants (Berlin, Germany) in 2014 and kept in culture since then. Continuous cultures were maintained in 1.8-L plastic containers, filled with 50 g spelt (R. dominica) or corn (P. truncatus), at 28°C, 60% relative humidity, and a day and night cycle of 16 to 8 hours. 5.2.2 Symbiont genome sequencing, assembly, and annotation Total DNA was isolated using the Epicentre MasterPureTM Complete DNA and RNA Purification Kit (Illumina Inc., Madison, WI, USA) including RNase digestion. Library preparation and sequencing for R. dominica (SRR19201380 and SRR19201381) and P. truncatus (SRR19638530 and SRR19638531) was performed at the Max-Planck-Genome-Centre (Cologne, Germany) on a HiSeq3000 Sequencing System (Illumina Inc., Madison, WI, USA). Adaptor and quality trimming was performed with Trimmomatic28. Assembly of Illumina reads was performed using SPAdes (v3.15.0) with the default settings29. The resulting contigs were then binned using BusyBee Web30 and screened for GC content and taxonomic identity to Bacteroidota bacteria. The extracted contigs were de novo assembled in Geneious Prime 2019 (v2019.1.3, https://www.geneious.com). The resulting genomes were then automatically annotated with PROKKA15 using the app Annotate Assembly and Re-annotate Genomes (v1.14.5) on KBase31. The annotated genomes were plotted using CIRCOS (v0.69-6) for the visualisation of gene locations, GC content, and coverage32. 5.2.3 16S rRNA cloning and phylogenetic analyses Bacterial 16S rRNA amplicons were obtained from single bacteriomes of four P. truncatus individuals and cloned into Escherichia coli for separation and subsequent Sanger sequencing of vector inserts. DNA was extracted from individual bacteriomes with the Epicentre MasterPureTM Complete DNA and RNA Purification Kit (Illumina Inc., Madison, WI, USA) and amplified with the Phusion high-fidelity DNA polymerase (ThermoFisher Scientific) and general bacterial 16S rRNA primers fD1 and rP233. The PCR parameters were as follows: after an initial denaturation step for 30 seconds at 98°C, 35 cycles were run for 10 seconds at 98°C, 30 seconds at 58°C and 30 seconds at 72°C followed by a final extension of 5 minutes. Amplicons were 139 purified from 1.5% agarose gels with the InnuPrep gel extraction kit (Analytik Jena GmbH, Jena, Germany) and cloned with the pMiniT 2.0 vector into Escherichia coli K12 (NEB PCR Cloning Kit; New England Biolabs, MA, USA). Vector insertion sequences of successfully transformed colonies were amplified by another PCR using the vector-specific cloning analysis primers (NEB PCR Cloning Kit; New England Biolabs, MA, USA) with PCR conditions as above, and entire cells from clone colonies were added to the PCR reaction mix as a template. Bidirectional Sanger sequencing was performed in-house on an Applied Biosystems 3730xl DNA Analyzer (ThermoFisher Scientific, Germany) to obtain the full sequence of the amplified 16S fragments using the cloning analysis forward and reverse primers. 16S rRNA sequences were aligned to representative Bacteroidota 16S rRNA sequences obtained from the NCBI database, using the SILVA algorithm10,34,35. Sequences of the shikimate pathway gene aroF encoded in the genomes of R. dominica and P. truncatus were extracted and aligned to representative Bacteroidota aroF sequences obtained from the NCBI database, using MAFFT36. Phylogenetic reconstruction for all alignments was done by Bayesian inference applying a GTR+G+I model using MrBayes (v3.2.7)37–40. The analysis ran for 20,000,000 generations with a “Burnin” of 25%, a sampling frequency of 1,000 generations, and we confirmed that split frequencies converged to <0.01. The obtained trees were visualized using FigTree (v1.4.4, http://tree.bio.ed.ac.uk/software/figtree/). 5.2.4 Comparative genomics Endosymbiont genomes were annotated with PROKKA15 in KBase31 to compare the bacteria and to estimate the genome-wide nucleotide sequence divergence level. Therefore, we identified single- copy orthologs in each genome pair using OrthoMCL (v2.0)41 in KBase31. KEGG categories were then assessed via GhostKOALA (v2.2)42 of each gene’s amino acid sequence. CIRCOS (v0.69-6)32 was used to link orthologous genes. Heatmaps were visualized using ‘gplots’ package in Rstudio (V 1.1.463 with R V3.6.3). 5.2.5 Quantitative PCR DNA of eight to ten Prostephanus truncatus eggs, larvae, pupae, and adults was isolated individually using the Epicentre MasterPureTM Complete DNA and RNA Purification Kit following the manufacturer’s instruction (Illumina Inc., USA) to evaluate symbiont titers. For the adult beetles, the abdomen (without wings) was used. The DNA was dissolved in 30 µL low 140 TE buffer (1:10 dilution of 1x TE buffer: 10 mM Tris-HCl + 1 mM EDTA). qPCRs were carried out in 25 µL reactions using EvaGreen (Solis BioDyne, Estonia), including 0.5 µM of each primer (Table 1) and 1 µL template DNA. All reagents were mixed, vortexed and centrifugated in 0.1 mL reaction tubes (Biozym, 711200). qPCRs were carried out on a Rotor-Gene Q thermal cycler (Qiagen, Hilden, Germany). The initial temperature was 95°C for 12 minutes, followed by 60 cycles of 95°C for 40 seconds followed by 20 seconds at 60°C. A melting curve analysis was used to assess the specificity of the qPCR reaction by a gradual increase of temperature from 60 to 95°C, with 0.25 K per second. The qPCR results were analysed using the Rotor Gene Q Software (Qiagen, Hilden, Germany). Standard curves with defined copy numbers of the genes were created by amplifying the fragment first via PCR, followed by purification and determination of the DNA concentration via NanoDrop1000 (Peqlab, Germany). After the determination of the DNA concentration, a standard containing 1010 copies/µL was generated and 1:10 serial dilutions down to 101 copies/µL were prepared. 1 µL of each standard was included in a qPCR reaction to standardize all measurements. The plot was visualized using ‘ggplot2’ in RStudio (V 1.1.463 with R V3.6.3). Table 1: Used Primer for qPCR on PTRU strains. Name Target Sequence 5´-3 Orientation Ptru_A_Shiki6_f aroA / strain A TGTAGGAGCTATATCAGGAGTAGA fwd Ptru_A_Shiki6_r aroA / strain A TGATTGGAGTTCAGCATCTTATTTT rev Ptru_B_Shiki1_f aroF / strain B CCATTTTAAACCTTCTTCCCCATAC fwd Ptru_B_Shiki1_r aroF / strain B GGGCCATGTAGTGCAGAAAAT rev Ptru_C_Shiki7_f aroC / strain C ACAATTGGTACTGCTCTTGGT fwd Ptru_C_Shiki7_r aroC / strain C GGGGAATTCAAGGAGGTATATCA rev 5.2.6 Fluorescence in situ hybridisation To localise the bacteriomes in R. dominica and P. truncatus, fluorescence in situ hybridisation (FISH) was performed, targeting the 16S rRNA sequence. Adult beetles were fixated in tertiary butanol (80%; Roth, Karlsruhe, Germany), paraformaldehyde (37-40%; Roth, Karlsruhe, Germany) and glacial acetic acid (Sigma-Aldrich, Germany) in proportions 6:3:1 for 2 hours, followed by post- fixation in alcoholic formaldehyde (paraformaldehyde (37-40%) and tertiary butanol (80% in proportion 1:2). After dehydration, the specimens were embedded in Technovit 8100 (Kulzer, 141 Germany)45 and cut into 8 µm sagittal sections using a Leica HistoCore AUTOCUT R microtome (Leica, Wetzlar, Germany) equipped with glass knives. The obtained sections were mounted on silanised glass slides. Each slide was covered with 100 µL of hybridisation mix, consisting of hybridisation buffer (0.9 M NaCl, 0.02 M Tris/HCl pH 8.0, 0.01% SDS; Roth, Germany) and 0.5 µM of each specific probe (Table 2). DAPI (0.5 µg/mL) was included as a general counterstain for DNA. Slides were covered with glass cover slips and incubated in a humid chamber at 50°C overnight. After washing and incubating them for two hours at 50°C in wash buffer (0.1 M NaCl, 0.02 M Tris/HCl, 5 mM EDTA, 0.01% SDS), they were washed in deionized water for 20 minutes and mounted with Vectashield (Vector Laboratories, Burlingame, CA, USA). The sections were either observed under a Zeiss AxioImager Z2 with Apotome.2 (Zeiss, Jena, Germany) illuminated by a SOLA Light Engine (Lumencor, Beaverton, OR, USA), or a Leica THUNDER imager Cell Culture 3D (Leica, Wetzlar, Germany). Images obtained on the Leica microscope were processed with the instant and small volume computational clearing algorithm using standard settings in the Leica Application Suite X software (Leica, Wetzlar, Germany). Table 2: Used FISH probes targeting 16S rRNA. Name Sequence 5´-3 Marker Bostrichidae_Sym2 CTTCCTACACGCGAAATAG Cy5 SymType1 TATTACTAAGACTATCTTTT Cy3 SymType2 AATTAATAAGACTATTCTTC Cy5 SymType3 ATTTAATAAAACTATTCTTT RhoGreen 5.2.7 HCR in situ staining To localise the strains in the bacteriome of P. truncatus, hybridisation chain reaction (HCR) in situ staining was performed. The bacteriomes of P. truncatus were fixated in 4% PFA in 80% butanol for 24 hours immediately after collection. For pre-embedding, 1% aqueous agar was liquefied in the microwave and cooled down to approximately 40°C. With the solution, a layer of ager was placed on a slide into which the bacteriomes were transferred individually while the solution was still liquid. After approximately 5 minutes, the solidified agar was cut with a scalpel into little blocks, each containing one bacteriome, and transferred to 80% butanol. Dehydration of the samples was performed in a series of butanol at concentrations of 90%, 96% and 3 x 100%. As an intermediate, 3 steps of 100% isopropanol were used. The samples remained 142 in the solutions for 2 hours each at room temperature, excluding the steps in 100% butanol, which were incubated at 30°C. This was followed by infiltration with ROTI®Plast paraffin wax with DMSO (Carl Roth, Karlsruhe, Germany) in two batches for 1 x 6 hours and 1 x overnight at 62°C, then the blocks were produced. 5 µm thick sections were made using a rotary microtome HistoCore AUTOCUT R (Leica, Nussloch, Germany) with disposable metal blades Feather R35 (PFM medical, Köln, Germany). Sections were briefly stretched in 40°C water, mounted on silanised slides, and dried overnight at 40°C. The dried sections were deparaffinized and rehydrated in a series of 2x xylene, 2x ethanol abs, ethanol 96%, 70% and water for 5 minutes each. Finally, the sections were post-fixated in 4% neutral buffered aqueous formaldehyde solution for 20 minutes, followed by washing in water for 5 minutes and drying at 40°C for 1 hour. Hybridisation chain reaction (HCR) in situ staining (Molecular Instruments, USA) was performed according to the manufacturer’s instructions following the HCR v3.0 FFPE human tissue sections protocol with the following modifications (provided by Andrew Gillis, Olivia Tidswell and Tobias Engl) on isolated bacteriome of P. truncatus: 5% dextran sulphate (instead of 10%) in both the 30% probe hybridisation buffer and amplification buffer. Before pre-hybridisation, the section was washed in 0.2 M HCl for 10 minutes at room temperature, afterwards washed in 1x PBS and incubated with Proteinase K (0.1 mg/mL in TE buffer) for 5 minutes at room temperature and then washed twice in probe hybridisation buffer. Also, after incubation overnight, the slides were heated at 75°C for 25 minutes to open the DNA strands and further maximize probe attachment and then cooled down to 37°C. HCR probe sets for each target gene with specific amplifier sequences were designed and synthesized by Molecular Instruments (Los Angeles, CA, USA; Table 3). To reduce autofluorescence of the tissue, the Vector® TrueVIEW® Autofluorescence Quenching Kit (Vector Laboratories, Burlingame, CA, USA) was used following the manufacturer’s instructions. Sections were mounted with Invitrogen™ ProLong™ Gold with DAPI (Fisher Scientific, Schwerte, Germany) and observed under a Zeiss AxioImager Z2 with Apotome.2 (Zeiss, Jena, Germany) illuminated by a SOLA Light Engine (Lumencor, Beaverton, OR, USA). 143 Table 3: Used HCR probes for HCR in situ staining. Complementary probe pairs (25 bp + linker) from each sequence were designed by Molecular Instrument. Target Number of probes Amplifier Marker aroA 17 B3 Atto647 aroC 19 B1 Atto488 aroFb 11 B2 Atto546 rProteins strain A 15 B1 Atto488 rProteins strain B 14 B3 Atto647 rProteins strain C 20 B2 Atto546 144 5.3 Data Accessibility Statement Raw sequence libraries of Prostephanus truncatus were uploaded to the NCBI Sequence Read Archive (SRR19638530 and SRR19638531; BioProject PRJNA848820). The annotated genomes are available on GenBank (CP104094 - CP104096). 5.4 Acknowledgments We thank Benjamin Weiss for technical assistance in histology, Eugen Bauer for genomic support, and Cornel Adler for the original provisioning of the beetle cultures. We thank Bruno Hüttel and the Max Planck-Genome-Centre Cologne (http://mpgc.mpipz.mpg.de/home/) for performing library preparation and sequencing, Olivia Tidswell for her introduction to the HCR, the Johannes Gutenberg-University Mainz for computation time granted on the supercomputer ‘MOGON’, and Christian Meesters for administrative assistance on ‘MOGON’. Additionally, we would like to thank Thomas Hörren for kindly sharing pictures for Figure 5. We gratefully acknowledge financial support from the Max Planck Society (to T.E. and M.K.) and the European Research Council through a Consolidator Grant to M.K. (ERC CoG 819585 “SYMBeetle”). 5.5 Contributions J.S.T.K. and T.E. designed the project, and J.S.T.K., E.B., and T.E. sequenced and assembled the symbiont genome. J.S.T.K. and E.B. annotated the genomes and performed symbiont genomic analysis. J.S.T.K. and T.E. wrote the paper, with input from M.K. 145 5.6 References 1. McCutcheon, J. P. & Moran, N. A. 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Weiss, B. & Kaltenpoth, M. Bacteriome-localised intracellular symbionts in pollen-feeding beetles of the genus Dasytes (Coleoptera, Dasytidae). Front. Microbiol. 7, 1486 (2016). 147 Chapter 6 General Discussion In this thesis, I demonstrate that beetles of the two families Silvanidae and Bostrichidae are associated with multiple bacterial symbionts. The sawtoothed grain beetle Oryzaephilus surinamensis (Silvanidae) harbours the Bacteroidota endosymbiont Shikimatogenerans silvanidophilus OSUR (Chapter 2) and the α-proteobacteria Wolbachia strain wSur (Chapter 3). In the Bostrichidae family, the Bacteroidota endosymbiont Shikimatogenerans bostrichidophilus is present in almost all examined species. In addition, species of the genera Dinoderus and Lyctus harbour the co-obligate symbiont Bostrichicola ureolyticus of the Bacteroidota phylum (Chapter 4). Furthermore, in the large grain borer Prostephanus truncatus the endosymbiont diverged into three, metabolically partially complementary strains (Chapter 5). The Shikimatogenerans endosymbionts supplement the host’s cuticle biosynthesis by the shikimate pathway, while Bostrichicola is capable of recycling nitrogen and supplementing their host’s diet with essential amino acids. 6.1 Ecology of symbioses in Oryzaephilus surinamensis The genome of the Bacteroidota endosymbiont of O. surinamensis, S. silvanidophilus, is highly eroded, encoding only genes of the shikimate pathway and glycolysis apart from gene families involved in the core processes of replication, transcription, and translation. The genome also lacks any genes for peptidoglycan biosynthesis, indicating that the symbiont can no longer synthesize its own cell wall. This raises the question of how the host controls its symbiont. In Chapter 2, the symbiotic relationship between host and symbiont was experimentally manipulated by inhibition of the shikimate pathway of S. silvanidophilus via glyphosate, and by adding tyrosine to the food source of the host. The outcome of these experiments demonstrates that Shikimatogenerans provisions tyrosine which also impacts the cuticle development of the host. In addition, it indicates that it is the production of the tyrosine precursors chorismate/prephenate via the shikimate pathway by which the host measures the symbionts’ contribution. Further, this 149 indicates that the symbiont's contribution is at least partially regulated on the host level via symbiont titer and not by gene expression, as most symbionts can presumably no longer control the expression level of their genes1,2. This control mechanism on the host level can be observed when the host has direct access to tyrosine via a food source, which led to a drastic reduction of the symbiont population (Chapter 2). From this finding, it can be concluded that it is the availability of tyrosine by which the host senses and controls the symbiont population. This hypothesis is supported by our own experience in the laboratory, where the symbiont titer increases when the host is maintained on ground oat that has been “diluted” by 50% with starch (data not published). Thus, the host can plastically adjust the symbiotic contribution to its needs via the regulation of the symbiont titer3. Similar results on diet-dependent symbiont regulation were also reported in the symbiosis between Wigglesworthia and its host the tsetse fly Glossina morsitans. Wigglesworthia provides B vitamins to the host, one of which is thiamine (vitamin B )41 . When the tsetse flies have access to thiamine from the food source, this resulted in reduced Wigglesworthia density in both male and female flies5. Both results - amino acids influencing S. silvanidophilus titer in O. surinamensis and thiamine influencing density of Wigglesworthia in G. morsitans - are examples of how supplementation of the host’s diet with nutrients derived from the symbiont affects the symbiont population negatively. Furthermore, the bacteriomes with the endosymbionts are slowly degraded in male adults of O. surinamensis after metamorphosis6. A similar yet drastically faster process was reported in the rice weevil Sitophilus oryzae, which is recycling its bacteriomes harbouring their endosymbiont Sodalis pierantonius in adults of both genders and only retains a separate, ovary associated symbiont pool for transmission7. In both systems, the benefit provisioned by the symbionts is over and by recycling the endosymbionts and their structures, the host can recoup some of the nutrients invested. Only in female adults, the symbiont is preserved at a constant level to ensure their transmission to next generations6,7. In addition to S. silvanidophilus, the silvanid O. surinamensis is host to Wolbachia, which is known as a reproductive manipulator in many insects8,9. In contrast to the Bacteroidota endosymbiont, the occurrence of the Wolbachia strain wSur is not restricted to the bacteriome but is localised throughout the entire body of the host. In Chapter 3, I demonstrate that wSur is indeed manipulating the reproduction of its host by inducing cytoplasmic incompatibility (CI), which leads to infertile embryos in wSur uninfected females that mate with wSur infected males10. In 150 consequence, the Wolbachia strain increases in frequency in the populations, which is reflected by its high prevalence in lab conditions. From our observations and experiments, there is no direct evidence that wSur also has a mutualistic effect on its host. Interestingly, the genome of wSur encodes for similar pathways to synthetic amino acids as Bostrichicola in the bostrichids, e.g. lysine, serine, and glutamine. It is possible that wSur complements the host's diets with those amino acids in addition to its reproductive manipulation, as it is reported for vitamin supplementation in other systems11,12. To clarify whether Wolbachia also has a mutualistic effect in O. surinamensis, experiments would have to be carried out with beetles that lack S. silvanidophilus but are infected with wSur. As I demonstrate in Chapter 2, it is possible to eliminate the Bacteroidota endosymbiont with glyphosate to get an O. surinamensis population with is associated with wSur alone, which could be used for experiments to elucidate a potential mutualistic effect. Also, the same experiments should be carried out with beetles that lack wSur but are infected with Shikimatogenerans, since the Bacteroidota endosymbiont is always present in the beetles in nature and only differ in whether they are infected with Wolbachia in addition. 6.2 Symbionts within Bostrichidae Beetles of the Bostrichidae family are associated with up to two Bacteroidota endosymbionts: Shikimatogenerans bostrichidophilus and Bostrichicola ureolyticus. Both endosymbionts are characterized by heavily eroded and A+T-biased genomes with extremely limited biosynthetic capabilities. S. bostrichidophilus and B. ureolyticus exhibit extremely reduced genomes. Apart from genes involved in replication, transcription, and translation, the genome of S. bostrichidophilus encodes exclusively genes for the shikimate pathway. The symbionts are localised in the cytoplasm of the bacteriocytes. As the genomes of S. bostrichidophilus lack genes for the synthesis of peptidoglycan, the symbiont cells show amorphous morphology, known as the L-form. Also, studies on other bacteriome-localised endosymbionts found that the symbionts are often polyploid, i.e. there are multiple copies of the genome per cell. In terms of genome size, the genome of Shikimatogenerans in the Dinoderinae Rhyzopertha dominica (173 kbp) and Prostephanus truncatus (avg. 156 kbp) are the most reduced. This, combined with the previously mentioned regulation of the symbiont population on the host level, could be the presupposition necessary to favour the split of the symbiont into different 151 linages in P. truncatus (Chapter 5). In R. dominica it can already be observed that there are no longer individual cells of the symbiont in the bacteriome, but rather several symbiont cells that are fused or no longer divided. Okude et al. (2017)13 appropriately described this as a grape-like pattern. From this point, it seems quite possible that in such a continuum of cells with multiple genomes some of them link together. The genomes of the three PTRU linages can either be mixed in the same symbiont cell, or they can be segregated into different cells. In the latter case, the symbiont strains must exchange either mRNAs, proteins/enzymes, and/or entire ribosomes and metabolites in order to be functional. After metamorphosis, only female beetles keep the bacteriomes to pass the symbionts on to the next generation6,7,79. At this point, there is probably no host-level selection on functionality single symbiont genomes in the adult stage. Although, it is also possible that females need the tyrosine supply as adults, while males do not. Quality check on the host level only takes place again in the next generation, when the beetle receives tyrosine precursors from the symbiont. As long as the symbiont or the symbionts in their continuum remain entirety functional and have a complete shikimate pathway to provide tyrosine precursors to the host, there is no immanent reason for host selection or even recognition of ongoing changes. A similar scenario - but even more complex - was observed in cicadas of the genus Tettigades (Hemiptera: Cicadidae) and their α-proteobacterial endosymbiont Hodgkinia cicadicola14–16. Also in this system, the question of how the linages evolved could not (yet) be finally clarified. It was shown, that to still ensure that each lineage is transferred to the next generation, these cicadas have increased the number of Hodgkinia cells six-fold17, whereas the number of Sulcia muelleri cells - the second endosymbiont - is unaltered. We have no data on the number of symbiont cells transmitted via egg to the next generation in P. truncatus, but that should be feasible with R. dominica and its endosymbiont S. bostrichidophilus RDOM as a comparison. Unfortunately, it is not possible to eliminate only a single PTRU lineage without also eliminating the other two as they are potentially treated and seen as one by the host. Only beetles of the genera Lyctus and Dinoderus harbour the second co-obligate endosymbiont Bostrichicola ureolyticus. The genome of Bostrichicola encodes genes to recycle nitrogen, as well as a glutamate dehydrogenase that allows integration of the resulting ammonium into the amino acid metabolism via glutamate. In addition, the genome encodes for an aspartate aminotransferase to 152 transfer the amino group from glutamate to oxaloacetate. As aminotransferase genes are known to be promiscuous, this specific encoded gene could also take over another aminotransferase activity18. The genome also encodes for an almost complete diaminopimelate pathway to synthesize the essential amino acid lysine from aspartate. Lysine is an important component of cuticular proteins as its ε-amino group represents an anchor point for cross-linking19. Grain diets are specifically limited in lysine20, which could be relevant for the stored product pest beetles of the genus Dinoderus, but also other species of the Dinoderinae subfamily. It also retained a methionine synthase to convert L-homoserine to L-methionine and is able to synthesise menaquinone. In addition, the genome encodes for a complete fatty acid and peptidoglycan biosynthesis, albeit other cell wall components apparently cannot be synthesized. S. bostrichidophilus was already integrated into the bostrichid beetles of the genera Lyctus and Dinoderus when the ancestor of B. ureolyticus was taken up at a later time point. The metabolic potential of B. ureolyticus and Blattabacterium derives from the same ancestor, which likely contained everything contained in the genome of all descendants since horizontal gene transfer is rare in intracellular symbionts. As B. ureolyticus lacks the shikimate pathway – and S. bostrichidophilus already fulfilled these needs, it is likely that it lost the genes for this pathway because of redundancy according to the black queen hypothesis21,22. As larvae accumulate a lot of uric acids, which are often deposited in the epidermis and cause the white colour of the larvae, B. ureolyticus likely plays a key role in nitrogen recycling during this period in the beetle’s life cycle23. Unfortunately, there is no data yet if B. ureolyticus faces the same destiny in adult males or if only the bacteriome harbouring S. bostrichidophilus is degraded and consumed by the host while the bacteriome harbouring B. ureolyticus persists as the host is still in need urea recycling or lysine from the endosymbiont. This would explain why beetles of the genera Lyctus and Dinoderus are associated with two closely related symbionts and not with just one with the metabolic potential of both combined. It can be assumed that B. ureolyticus takes over the urea recycling that was previously encoded in the early genome of Shikimatogenerans. However, by also encoding new genes for lysine synthesis and other individual enzymatic steps, B. ureolyticus could offer additional benefits. Currently, no other insect is known to be associated with two symbionts, of which one symbiont only supports cuticle synthesis, while the second symbiont recycles nitrogen and provides amino acids. In addition, the relationship between both Shikimatogenerans and 153 Bostrichicola remains unsolved. From the genomic point of view, both B. ureolyticus and S. bostrichidophilus complement each other in the symbiosis with their host (Chapter 4). Several insects are associated with two or more endosymbionts with complementary biosynthetic pathways to synthesize amino acids and/or vitamins4,24–30 or pathways for the essential amino acids are split up between the two symbionts27,31,32. Symbionts can also be replaced when they are no longer needed or capable of sufficiently supporting their host’s metabolism33–35. As the genome of an endosymbiont gets streamlined to fulfil the specific needs of the host, mutations and pseudogenes can accumulate, which can lead to reduced efficiency of the genome. The consequences are the rapid evolution of protein sequences and gene loss. This process is called Muller’s ratchet33,36. It’s solved by replacing the symbiont with the eroded genome with a new symbiont that brings in novel functions that the original symbiont did not provide and can therefore be selectively favoured, or the replacing symbiont fulfils the role of the original symbiont better or more efficiently. This could be the case e.g. if genome erosion has resulted in suboptimal codon composition and therefore stability of proteins (due to Muller’s ratchet). Evidence for this is the usually very high expression of chaperones in intracellular symbionts, which assists in refolding malfunctional proteins and is usually interpreted as a compensatory adaptation37. Replacement of endosymbionts was shown in the Auchenorrhyncha, which are associated with the Bacteroidota endosymbiont Sulcia muelleri and a β-proteobacterium. This β-proteobacterium has been proposed to be the ancient co-obligate partner of S. muelleri and has been replaced multiple times by – or evolved as a common ancestor into Zinderia insecticola in the spittlebugs (Hemiptera: Auchenorrhyncha: Cercopoidea), Nasuia deltocephalinicola of deltocephaline leafhoppers, and Vidania fulgoroideae of fulgorid planthoppers30,38. In addition to the before-mentioned Auchenorrhyncha, this is also known for the di-symbiotic Buchnera aphidicola and Serratia symbiotica in aphids (Hemiptera: Aphididae)28,32. Interestingly, the bostrichids represent the first case, where two closely related bacteria of the same family are complementing each other. One reason could be that bostrichid beetles of the Dinoderinae and Lyctinae subfamily were already adapted to one Bacteroidota, S. bostrichidophilus. Further research on the Dinoderinae Dinoderus porcellus should elucidate the relationship of this tripartite symbiosis. Given the results from the experiments in Chapter 2, it should be possible to remove S. bostrichidophilus from the beetle with a tyrosine-rich or glyphosate-containing diet and 154 thus eventually obtain a host population of D. porcellus that only harbours B. ureolyticus – as long as the two endosymbionts are not heavily dependent on each other. In addition, the functionality of the urease encoded in the genome of B. ureolyticus should be assessed. This would be possible with Isotope-labelled urea, which would make it possible to track whether and where it is metabolized39,40. It is worth mentioning that of the species-rich family of the Bostrichidae, only a couple of species are known as grain pests (e.g. Rhyzopertha dominica and Dinoderus porcellus) while the majority is still feeding on dead wood (e.g. Lyctus brunneus and Dinoderus minutus). Wood in general is a very poor diet for insects as due to the recalcitrant polymers in the (woody) cell walls (i.e. lignocellulose) it is difficult for the insect to utilize the carbon as energy sources and gain access to the more nutritious cell content41. To compensate for the nutritional deficiency, xylophagous (wood-feeding) beetles harbour gut bacteria that provide them with the missing essential amino acids, as well as recycle nitrogen42,43. The xylophagous bostrichids, e.g. the common powderpost beetle Lyctus brunneus harbour two endosymbionts that can synthesize a subset of amino acids as well as recycle nitrogen. While collecting samples for this work, it was observed that specimens of Trogoxylon impressum and Lyctus cavicollis were found on the same type of wood branches at the same time point. While both species belong to the subfamily of the Lyctinae, I could show that only Lyctus lives in a tripartite symbiosis with Shikimatogenerans and Bostrichicola, whereas T. impressum only harbours Shikimatogenerans (Chapter 4). While both species share the same habitat, harbouring a second symbiont with additional metabolic capacities also brings the opportunity to avoid intraspecific competition by switching from one food source to another. It is likely that T. impressum - lacking Bostrichicola and therefore a source for amino acids – is feeding on nutrient-rich phloem whereas L. cavicollis favours the nutrient-poor xylem and hard-wood. Experiments with Lyctus africanus – harbouring both Shikimatogenerans and Bostrichicola - showed, the beetle favoured starch and sugar as vital nutrients for oviposition44. The same can be hypothesized about the Dinoderinae subfamily, where Rhyzopertha dominica harbours Shikimatogenerans but species of the Dinoderus genus harbour Bostrichicola in addition. Still, as they are known as grain pests, they seem to feed on the same food source, whereas it is likely that R. dominica aims at the grain embryo, which is rich in nutrients, whereas Dinoderus sp. is satisfied with the surrounding nutrient-poor starch, which the R. dominica beetles dig out of the grain on the way for oviposition. Beetles of the genus Dinoderus are in general less specialised as some infest 155 cereals as well as carbohydrate-rich products, e.g. yam roots (D. porcellus and D. bifoveolatus), while others are bamboo pests (D. minutus). As R. dominica and P. truncatus are really specialised on grain where they feed on the nutrient-rich embryo, Bostrichicola could have been lost as a result. Interestingly, Li et al. (2015) detected Wolbachia in both R. dominica and P. truncatus45. However, I have no data for Wolbachia in our cultures. Given the results from Chapter 3, it would make sense to also examine the existing metagenomes for Wolbachia. Since wSur encodes for genes to synthesise essential amino acids in O. surinamensis, it seems possible that in cases where B. ureolyticus is not present, Wolbachia takes over this mutualistic part instead. At the moment, however, this is pure speculation. 6.3 Evolution of beetle-associated Flavobacteria The insect-associated Flavobacteria constitute a single clade that is about 400 Mya old46. Since then, the bacteria have diverged, and the clade contains symbionts with manifold metabolic abilities. They developed from the probably most ancestral lifestyle of male-killing Flavobacteria, which form the sister group to this clade and still are reproductive manipulators in ladybird beetles (Coleoptera: Coccinellidae)47 - of which the genome is yet not available. Presumably, originally reproductive manipulators, the genomes of Blattabacterium spp., Walczuchella monophlebidarum, Uzinura diaspidicola, probably Brownia, to Sulcia muelleri were able to spread within insects and co-evolve in as little as 100 Mya into nutritional mutualists46, leading to a whole clade of Flavobacteria associated with several insect families. While Bostrichicola ureolyticus still encodes genes for pathways to produce essential amino acids as well as urease for the recycling of nitrogen – similar to Blattabacterium, Walczuchella, and Uzinura48-50 - Shikimatogenerans bostrichidophilus became an exclusive chorismate supplier. Evolving from a free-living bacterium to an ‘enslaved’ symbiont, the genome of Shikimatogenerans underwent a drastic reduction in a host-beneficial manner. The genome of Shikimatogenerans exclusively encodes genes for the shikimate pathway. Although the metabolic potential of S. silvanidophilus is already limited as the genome still encodes genes for glycolysis and potentially nitrogen recycling, the genome of S. bostrichidophilus is even more reduced exclusively encoding the shikimate pathway besides general cell processes - levelling Shikimatogenerans to a status comparable with the mitochondria in eukaryotic cells51. Mentioning mitochondria, it is noteworthy that in comparison to Bostrichicola, the genome of S. bostrichidophilus does not encode 156 any genes for glycolysis or any ATP synthetase genes. The same is true for the genomes of Uzinura and Hodgkinia52 - to name some that have already been investigated previously. S. silvanidophilus, on the other hand, has genes for glycolysis and ATP synthesis. Further experiments will have to show whether glycolysis in S. silvanidophilus is functional and whether the symbiont itself determines it, or whether it is already being taken over by the host – which is more likely, as the glycolysis also needs products as input that already must be delivered by the host (Chapter 2). The complete absence of genes for glycolysis and ATP synthase in S. bostrichidophilus indicates a total dependency on energy production since the host's environment must supply some necessary metabolites directly to the symbionts. Phylogenetic analyses showed that the three Bacteroidota endosymbionts examined in both beetle families are closely related. According to Engl et al. (2018), the common ancestors of the three Bacteroidota endosymbionts - S. silvanidophilus, S. bostrichidophilus, and B. ureolyticus -, Blattabacterium, and S. muelleri lived 409 Mya46. Subsequently, Bostrichicola and Shikimatogenerans diverged around 15 Mya later and another 63 Mya later S. silvanidophilus and S. bostrichidophilus diverged from each other. S. silvanidophilus combines the metabolic capacity of Bostrichicola to recycle nitrogen, and S. bostrichidophilus to supplement the tyrosine precursor prephenate - between which it also clusters phylogenetically. Following, it can be assumed that the ancestor of the three endosymbionts combined these abilities, and afterwards individual pathways were lost in each of the lineages. 6.4 Grain pest beetles as non-model model organisms The example of O. surinamensis demonstrates that the investigation of insects associated with multiple symbionts are crucial for understanding the symbiont-influenced eco-evolutionary dynamics of their host. Sharing the same habitat, the Bacteroidota endosymbionts of the silvanid and bostrichid beetle grain pests got shaped accordingly to the specific needs of the host. The ancestor of these Bacteroidota endosymbionts had richer repertoire of pathways from which the host and its habitat could “select”. According to the symbiont metabolic capabilities, both beetle families are in demand for tyrosine. Likely, the ancestors of both the silvanid and the bostrichid beetles compared to their symbiont-free conspecifics had an immediate benefit because of the enhanced tyrosine supply after they acquired the ancestor of Shikimatogenerans. However, the 157 symbiotic beetles might have had to eat more because they now had to feed their costly symbionts as well6,7. In return, they had more tyrosine available, which allowed them to build up a cuticle that is harder and thicker - which means they were better protected from desiccation and predators46,53. And in the case of the bostrichids, this may even have led to mandibles that made it easier to penetrate harder materials such as hardwood. Under the current climate crises, in which droughts are becoming more frequent due to global warming, a symbiosis with Shikimatogenerans or a similar symbiont would be beneficial for insects in nature, since Shikimatogenerans contribute via the shikimate pathway to cuticle biosynthesis, which leads to a thicker cuticle to protects the beetle from desiccation46 and predators53. However, it was also shown in several studies that the Bacteroidota endosymbionts of the silvanid and bostrichid beetles are sensitive to heat46,54-57,79. To make matters worse, the same endosymbiont is sensitive to exposure to glyphosate and, as a result, the host suffers equally from exposure to glyphosate (Chapter 2). This was not only shown in beetles but also other insects including tsetse flies, bees, and ants4,58–61,80. Experiments with glyphosate show that this herbicide, as desired, indeed targets the shikimate pathway - which means that eukaryotic cells should not be the target, since they do not have that metabolic pathway59,60,62. However, it was ignored that many animals - including humans - are associated with bacteria63,64. Research in recent years has clearly shown that there is a connection between a person's microbiome and their health65. The same applies to the beetle and its endosymbiont - if the symbiosis is disturbed, both partners are affected. Also, when factors directly affect only one partner, it always affects the other in return. Especially when such a dependency has developed between the two that one is no longer able to live without the other. In the case of the silvanid and the bostrichid beetles and their Bacteroidota endosymbionts, this means that both partners would suffer under glyphosate treatments (Shikimatogenerans) as well as rising temperatures. Glyphosate itself acts on a specific enzyme in the shikimate pathway: AroA66,67. There are two classes of this enzyme, namely class 1, which is sensitive to glyphosate, and class 2, which is non- sensitive to glyphosate. The phylogenetic study of the aroA sequences of different bacterial symbionts shows that they all encode aroA genes that belong to the glyphosate-sensitive class 1. This means that upon contact, glyphosate permanently binds to aroA and blocks its enzymatic ability, interrupting the entire metabolic pathway. In this work, I showed that beetles of the 158 family Silvanidae and Bostrichidae are associated with Shikimatogenerans, which uniquely contribute products of the shikimate pathway. Further, I also demonstrate that the aroA for this pathway encoded by the genomes of these symbionts belongs to the glyphosate-sensitive class I. With O. surinamensis I could verify this sensitivity experimentally - the Shikimatogenerans titer decreased drastically after glyphosate treatment. Although it now sounds like glyphosate would be suitable to treat grain storage infested with grain pests such as O. surinamensis or R. dominica, it should be noted that we only work with pest species because they are easy to keep in the laboratory and lend themselves so well to simply having access to many specimens for experimentation and analysis. Moreover, it should be pointed out that in the species-rich Bostrichidae family, only a very small proportion are known as pests of crops (e.g. Rhyzopertha dominica) and wood (e.g. Lyctus brunneus), whereas this family members otherwise act beneficially with the ability to decompose hard-wood into fine drill dust which is then decomposed by microorganisms and fungi in the soil74 or used for camouflage by insects such as the dust bug Reduvius personatus76. In addition, bostrichids also serve as a food source, above all for the checkered beetles (Coleoptera: Cleroidea) like Tilloidea unifasciata, Tilloidea notata or Clerus mutillarius75,78, but also chalcid wasps (Hymenoptera: Chalcinidae) like Cerocephala aquila and braconid wasps (Hymenoptera: Braconidae) like Platyspathius dinoderi, as well as spiders such as the woodlouse hunting spider Harpactea hombergi and the jumping spider Salticus zebraneus75. But also, birds such as the red-backed shrike Lanius collurio and woodpeckers (Piciformes: Picidae) as well as mammals like the dormouse Glis glis sometimes feed on bostrichids77. In addition, the holes dug in the wood are used by other species for oviposition, e.g., the carrot wasp Gasteruption erythrostomum, which uses the boreholes of bostrichids to lay eggs76. This shows that the Bostrichidae are closely networked in their ecosystem and that a negative impact on the symbiosis via the symbiont by glyphosate not only has negative consequences for the host beetle but could also subsequently affect all other species. However, besides these two highly specialized symbionts of the Silvanidae and Bostrichidae beetle families, many other insects are associated with Bacteroidota but also γ-proteobacteria symbionts, many of which encode a class I aroA enzyme. Thus, even if shikimate derivatives do not represent the major contribution of these symbionts, their inhibition could negatively impact symbiont growth and thereby host fitness. Whereas O. surinamensis could survive without its endosymbionts 159 when the environment supports its tyrosine-demanding lifestyle, it is not clear if the same can be said about other insects. In fact, it is known that many insects cannot survive without their obligate symbionts, for example, the black hard weevil Pachyrhynchus infernalis and its symbiont Nardonella68, the thistle tortoise beetle Cassida rubiginosa and its symbiont Stammera69 or the pea aphid Acyrothosiphon pisum and its obligate symbiont Buchnera aphidicola70. In addition to the current climate crisis, a decline in insect species abundance and diversity can also be observed71,72. Further investigations into insect symbioses could clarify the question of whether there is a connection between glyphosate and/or other agrochemicals and a decrease in the number and diversity of insects due to the disruption of their interactions with beneficial symbionts. This necessity is exacerbated by the fact that many of these active ingredients have also been detected in nature reserves – places that have been placed to protect animals and plants73. 6.5 Conclusion In my thesis, I demonstrate the importance of the Bacteroidota endosymbiont Shikimatogenerans associated with the Bostrichidae and Silvanidae beetle families for the cuticle biosynthesis of the host. Considering that the order Coleoptera constitutes the largest insect order, the influence of symbioses in beetles is still understudied. My work shows the influence that symbionts can have on the ecology and evolution of their hosts. In addition, I show how widespread this clade of Bacteroidota symbionts is in insects, where both convergent symbioses with different partners and divergent functions of closely related symbionts in different hosts can be found. 160 6.6 References 1. Hansen, A. K. & Moran, N. A. 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Ecol Entomol 47, 273–283 (2022). 163 Danksagung 165 166 Erklärung gemäß § 13 Abs. 4 der Promotionsordnung des Fachbereiches Biologie der Johannes Gutenberg-Universität Mainz Hiermit erkläre ich, dass ich die vorliegende Dissertation selbstständig angefertigt und keine anderen als die angegebenen Quellen oder Hilfsmittel verwendet habe. Personen, die mich bei der Auswahl und Auswertung des Materials sowie bei der Fertigstellung der Manuskripte unterstützt haben, sind am Beginn eines jeden Kapitels genannt. Es wurde weder die Hilfe eines Promotionsberaters in Anspruch genommen, noch haben Dritte für Arbeiten, welche im Zusammenhang mit dem Inhalt der vorliegenden Dissertation stehen, geldwerte Leistungen erhalten. Die vorgelegte Dissertation wurde außerdem weder als Prüfungsarbeit für eine staatliche oder andere wissenschaftliche Prüfung noch als Dissertation an einer anderen Hochschule eingereicht. Bremen, den 26. September 2022 167 Curriculum Vitae Julian Simon Thilo Kiefer 169 170