Dissecting the Molecular Basis of the Drosophila melanogaster Anti-nematode Immune Response
by Shruti Yadav
M.Sc.in Medical Molecular Biology, September 2008, The University of Westminster, UK
A Dissertation submitted to
The Faculty of The Columbian College of Arts and Sciences of The George Washington University in partial fulfillment of the requirements for the degree of Doctor of Philosophy
May 20, 2018
Dissertation directed by
Ioannis Eleftherianos Associate Professor of Biology
The Columbian College of Arts and Sciences of The George Washington University certifies that Shruti Yadav has passed the Final Examination for the degree of Doctor of Philosophy as of March 22, 2018. This is the final and approved form of the dissertation.
Dissecting the Molecular Basis of the Drosophila melanogaster Anti-nematode Immune Response
Shruti Yadav
Dissertation Research Committee:
Ioannis Eleftherianos, Associate Professor of Biological Sciences, Dissertation Director
L. Courtney Smith, Professor of Biological Sciences, Committee Member
Damien O’Halloran, Assistant Professor of Biological Sciences, Committee Member
ii
© Copyright 2018 by Shruti Yadav All rights reserved
iii
Dedication
For my parents, who are the wind beneath my wings.
iv Acknowledgements
I’m grateful to my research mentor, Dr. Ioannis Eleftherianos, for his continuous guidance throughout my time here. I appreciate him for allowing me to plan and conduct my research work. Without his support and continued guidance, this dissertation wouldn’t be what it is today.
I would like to thank members of my committee, Dr. L. Courtney Smith, Dr.
Damien O’Halloran and Dr. John Hawdon for all the guidance, suggestions and encouragement. I would also like to thank Dr. Mollie Manier and Dr. Douglas Nixon for providing vital recommendations towards my dissertation. I would like to extend my gratitude to all the faculty, students and members of the GWU Biology Department for creating a positive learning and enjoyable environment.
I am indebted to Dr. Tara Scully for all her support and encouragement. I would also like to thank Julio Castillo and Upasana Shokal for teaching me technical skills and guiding me in my early Ph.D years. I’m grateful to Sneh Harsh and Eric Kenney for their inputs and suggestions. I also want to acknowledge all the past and present lab members for creating a collegial research environment. I’m grateful for the pleasure of interacting with all these wonderful people.
Finally, I would like to thank my family and friends for their unconditional love and support.
v Abstract of Dissertation
Dissecting the Molecular Basis of the Drosophila melanogaster Anti-nematode Immune Response
Drosophila melanogaster is an excellent model to study the molecular basis of anti-pathogen mechanisms. Currently, microbial pathogenesis in Drosophila and the insect antimicrobial immune response are well understood. However, our knowledge of nematode infections and anti-nematode immune mechanisms is still in its infancy. In this work, I investigated the interaction between the Drosophila immune response with the entomopathogenic nematode parasite, Steinernema carpocapsae. I report a new protocol for generating Steinernema nematodes lacking their mutualistic Xenorhabdus nematophila bacteria. Infection with Steinernema symbiotic (carrying Xenorhabdus) or axenic (lacking Xenorhabdus) results in Drosophila larval death at the similar rates.
Interestingly, the Drosophila immune response to Steinernema infection can be modulated by the presence of the endosymbiotic bacteria, Wolbachia and Spiroplasma. I report that presence of Wolbachia alone or together with Spiroplasma promote the survival of Drosophila larvae against Steinernema symbiotic nematodes. The presence or absence of endosymbiotic bacteria differentially modulate the immune signaling and phenoloxidase response, and alter the metabolic status and fat body lipid droplet size in the nematode infected larvae. These data demonstrate an interaction between the endosymbiotic bacteria, Wolbachia and Spiroplasma, and the Drosophila immune and metabolic responses in the context of Steinernema nematode infection.
vi Then, I investigated the transcriptomic profile of Drosophila larvae during infection with Steinernema symbiotic or axenic nematodes. I have found that infection with symbiotic nematodes induce genes associated with immune functions, whereas axenic nematodes induce genes associated with chitin binding and metabolic functions. I also report that the Drosophila genes with potential anti-nematode activity are conserved in the lepidopteran insect host, Manduca sexta, as well as in humans. These findings have led to the identification of genes with potential roles in nematode recognition and anti- nematode activity in Drosophila.
I also investigated the participation of the chitinase-like imaginal disc growth factor encoding genes, Idgf2 and Idgf3, in the Drosophila response to Steinernema nematode infection. I report that Idgf2 and Idgf3 are upregulated in Drosophila larvae responding to Steinernema nematodes and inactivation of Idgf2 provides a survival advantage to the larvae against axenic nematodes. I show that Idgf2 or Idgf3 modulate distinct immune signaling pathways and inactivation of Idgf3 induces the recruitment of
Drosophila larval hemocytes in response to Steinernema nematodes. These data demonstrate that Idgf2 and Idgf3 are involved in different yet crucial anti-nematode immune functions in Drosophila.
Next, I report the participation of Jonah66Ci that encodes a serine protease, in the
Drosophila response to Steinernema infection. I show that Jonah66Ci is strongly upregulated in the gut of Drosophila larvae responding to Steinernema. Jonah66Ci differentially modulates Toll, Imd and Wnt/Wg signaling in the gut of Drosophila larvae upon Steinernema infection. Inactivation of Jonah66Ci increases the numbers of replicating cells in uninfected larvae and nematode infection reduces cell replication.
vii These results indicate that Jonah66Ci is essential in maintaining homeostasis in
Drosophila larvae during Steinernema nematode infection.
In conclusion, my work has demonstrated that Steinernema infection differentially regulates a large number of genes in Drosophila larvae, endosymbiotic bacteria participate in the interaction between the Drosophila immune system with the nematode parasites, and Idgf2, Idgf3 and Jonah66Ci genes encode factors with distinct yet important anti-nematode activity in Drosophila larvae against Steinernema challenge.
These findings generate novel insights towards deciphering the molecular and functional basis of the Drosophila anti-nematode immune response and unraveling previously unknown aspects of the Drosophila innate immune system.
viii Table of Contents
Dedication………………………………………………………………………………...iv
Acknowledgements……………………………………………………………………...... v
Abstract of Dissertation ………………………………………………………………….vi
List of Figures……………………………………………………………………………..x
List of Supplementary Materials………………………………………………………..xiii
List of Abbreviations……………………………………………………………………xiv
Dissertation Introduction………………………………………………………………….1
Chapter 1: An improved method for generating axenic entomopathogenic nematodes……………………………………………………………26
Chapter 2: Endosymbiont-based immunity in Drosophila against parasitic nematode infection……………………………………………………………………….42
Chapter 3: RNAseq analysis of the Drosophila response to the entomopathogenic nematode Steinernema………………………………………………74
Chapter 4: The Imaginal Disc Growth Factors 2 and 3 regulate the Drosophila anti-nematode immunity………………………………………………………………..117
Chapter 5: Participation of a serine protease gene Jonah66Ci in the Drosophila anti-nemtaode immune response……………………………………….……………….143
Thesis General Discussion……………………………………………………………...175
References………………………………………………………………………………181
ix List of Figures
Chapter 1
Figure 1. Flow diagram of the method for obtaining axenic nematodes………………..39
Figure 2. Validation of nematode axenicity status………………………………………40
Figure 3. Survival results for Drosophila larvae infected by Steinernema nematodes….41
Chapter 2
Figure 1. Survival of Drosophila melanogaster larvae carrying or lacking endosymbionts in response to nematode infection…..…………………………………..67
Figure 2. Numbers for endosymbiotic and pathogenic bacteria in Drosophila melanogaster larvae responding to nematode infection…………………………………68
Figure 3. Transcript levels of immune genes in Drosophila melanogaster larvae carrying or lacking endosymbionts upon nematode infection..………………………….69
Figure 4. Phenoloxidase activity and melanization response in uninfected and nematode-infected Drosophila melanogaster larvae carrying or lacking endosymbionts…………………………………………………………………...70
Figure 5. Metabolic activity in Drosophila melanogaster larvae carrying or lacking endosymbionts following nematode infection……...…………………………...71
Figure 6. Lipid droplet size in Drosophila melanogaster larvae carrying or lacking endosymbionts upon nematode infection……………...………………………...72
Table 1. Primer sequences and annealing temperatures used for quantitative
RT-PCR (qRT- PCR)………………..…………………………………………………...73
x Chapter 3
Figure 1. Infection of wild-type Drosophila melanogaster third instar larvae with Steinernema carpocapsae symbiotic or axenic nematodes induces a large number of transcripts………………………………………………..………………….103
Figure 2. Infection with S. carpocapsae symbiotic or axenic nematodes induces distinct and shared transcriptomic profiles in D. melanogaster larvae…………………104
Figure 3. Infection with S. carpocapsae symbiotic or axenic nematodes induces diverse physiological responses and biological activities in D. melanogaster larvae….105
Figure 4. Infection with S. carpocapsae symbiotic or axenic nematodes differentially regulates the transcription of a variety of immune and developmental genes in D. melanogaster larvae…………………………………….…108
Figure 5. Orthologs of the top 55 differentially transcribed Drosophila melanogaster (Dm) genes in Manduca sexta (Ms) and Homo sapiens (Hs).…………...109
Table 1. List of primers used for qRT-PCR……………………………………………110
Chapter 4
Figure 1. Upreglation of Idgf2 and Idgf3 in Drosophila larvae following
Steinernema nematode infection………………………………...……………………...137
Figure 2. Drosophila Idgf2 mutants exhibit delayed mortality following
Steinernema infection………………...………………………………………………...138
Figure 3. Drosophila Idgf2 and Idgf3 mutants induce Toll, Imd and Jak/Stat signaling following Steinernema infection……………………………………………..139
Figure 4. Upregulation of Drosophila Idgf2 in Imd and Jak/Stat mutants
xi following Steinernema infection……………………………………………………...... 140
Figure 5. Increased hemocyte numbers in Drosophila Idgf3 mutants following Steinernema infection…………………………………………………………………..141
Table 1. Primers used for quantitative RT-PCR analysis………………………………142
Chapter 5
Figure 1. Relative gene transcript levels of Jonah66Ci in Drosophila larvae upon infection with Steinernema nematodes…………………………...………………168
Figure 2. Survival response of Drosophila Jonah66Ci mutant larvae upon infection with Steinernema nematodes……………….………………………………...169
Figure 3. Transcript levels of immune pathway read-out genes in Drosophila
Jonah66Ci mutant larvae infected with Steinernema nematodes……….……………...170
Figure 4. Transcript levels of Toll and Imd pathway read-out genes in the gut of Drosophila Jonah66Ci mutant larvae infected with Steinernema nematodes………….171
Figure 5. Replication in the intestinal cells of Drosophila Jonah66Ci mutant larvae infected with Steinernema nematodes……………………………………..…….172
Figure 6. Nitric oxide levels and feeding rates of Drosophila Jonah66Ci mutant larvae infected with Steinernema nematodes……………………………………..…….173
Table 1. Primers used for quantitative RT-PCR…………………………………….….174
xii List of Supplementary Material
Chapter 3
Figure S1. Infection by Steinernema symbiotic and axenic nematodes induces the expression of diverse classes of genes in
D. melanogaster larvae………………………………………………………………....112
Figure S2. Quantitative real time RT-PCR validation of seven
D. melanogaster genes (CG31508, CG31698, CG16844, CG7248, CG1934,
CG7596 and CG3906) selected from the transcriptome dataset………………………..116
xiii
List of Abbreviations
EPN Entomopathogenic nematode
GNBP Gram-negative binding protein
IDGF Imaginal Disc Growth Factors
IJ Infective juvenile
Imd Immune deficiency
Jak/Stat Janus kinase/ Signal transducer and activator of transcription
Jnk c-Jun-N-terminal kinase
NF휅B Nuclear Factor-Kappa B
NO Nitric Oxide
PAMPs Pathogen associated molecular patterns
PO Phenoloxidase
PGRPs Peptidoglycan recognition proteins
PRRs Pattern recognition receptors
TLR Toll-like receptor
Tot-A Turandot-A
xiv
Dissertation Introduction
The immune system
Every organism is under threat from a diverse range of infectious agents, affecting the fitness of the host. To combat infection, the host is under constant selection to evolve mechanisms to defend against or avoid the encounter with pathogenic organisms. These selective mechanisms include detection and subsequent elimination of the pathogen. The role of the immune system is to survey the host body and to combat infections, all the while distinguishing self from non-self (Hoffmann & Reichhart, 2002).
Infectious agents come in different shapes and sizes, leading to deployment of a variety of host immune responses to combat the infection. In vertebrates, the immune responses are classified as innate and adaptive (acquired) (Murphy, 2012). Innate immune responses are the host’s first line of defense, and they are broadly specific and lack memory. Innate immunity is the ancient form of immune response and is found in all multicellular organisms (Janeway & Medzhitov, 2002). The adaptive immune response, on the other hand, is largely reliant on the varied antigen specific receptors present on lymphocytes (Medzhitov & Janeway, 1998). The innate immune system interacts with and induces the adaptive immune responses (Fearon & Locksley, 1996).
There are an estimated 10 million species that rely exclusively on innate immune responses (Hoffmann & Reichhart, 2002). Of these, the fruit fly, Drosophila
1 melanogaster, has garnered attention due to its relative simplicity and significant structural and functional homology to mammalian innate immune signaling pathways
(Hoffmann & Reichhart, 2002). The conservation of the signaling pathways in both fruit flies and mammals is suggestive of an ancient origin in evolution, making the fruit fly a remarkable model system for innate immunity studies. In addition, Drosophila flies and larvae feed on decaying matter and are exposed to large numbers and varieties of pathogenic microbes. The robustness of their innate immune response enables them to defend against pathogenic assaults (Lemaitre and Hoffmann 2007). Previous studies have led to the identification and characterization of two distinct immune signaling pathways in Drosophila, Toll [mammalian Interleukin (IL)-1/ Toll-like receptor (TLR) pathway] and Immune deficiency (Imd) [Tumor Necrosis Factor (TNF)-α signaling pathway), in response to bacterial and fungal pathogens (Kaneko & Silverman, 2005; Ligoxygakis et al., 2002).
Defense systems in Drosophila
The Drosophila innate immune system is multi-dimensional consisting of epithelial barriers that are responsible for the production of antimicrobial peptides, reactive oxygen species (ROS), and nitric oxide (NO), as well as efficient humoral and cellular immune reactions (Royet & Dziarski, 2007). The activation of ROS and NO activates AMPs against microbial infections (Foley & O'Farrell, 2003; S. C. Wu et al.,
2012). Different Drosophila organs such as the trachea, fat body (functionally equivalent to the mammalian liver) and Malpighian tubules (functionally equivalent to kidneys)
2 produce AMPs that inhibit microbial infection (Kounatidis & Ligoxygakis, 2012).
Furthermore, hemocytes equivalent of blood cells) regulate the cellular immune response.
Plasmatocytes are the hemocyte type that perform phagocytosis and coagulation responses, whereas lamellocytes encapsulate larger pathogens such as nematode parasites and parasitoid wasp eggs (Vlisidou & Wood, 2015).
Drosophila humoral immune responses
Synthesis and secretion of the AMPs form the hallmark of the Drosophila humoral immune response. Followed by immunological activation of the hemocytes, the fat body is the primary organ that produces and secretes AMPs into the hemolymph
(insect blood). There are seven categories of AMPs (Diptericin, Drosomycin, Attacin,
Cecropin, Defensin, Drosocin and Metchnikowin) and a total of 20 different AMPs
(isoforms), which are cationic in nature (Hetru et al., 2003; Imler & Bulet, 2005). The first step in every immune response is recognition of the pathogen and culminates in the production of effector molecules such as AMPs, stress factors and serpins (Fig. 1).
Pathogen recognition is processed through the germ-line encoded receptors, called pattern recognition receptors (PRRs), which detect the pathogen associated molecular patterns (PAMPs) that are present on the microbial surface or secreted by microbes
(Hetru et al., 2003; Imler & Bulet, 2005; Pal & Wu, 2009). The Drosophila Toll pathway is activated in response to infection by bacteria containing Lys-type peptidoglycan (PGN) and fungi whereas DAP-type PGN containing bacteria induce the Imd pathway (Kleino &
Silverman, 2014; Valanne et al., 2011). The intracellular signaling events culminate in the nuclear translocation of the NF-κB transcription factor homologues leading to the
3 transcriptional activation of AMP encoding genes, a similarity shared by both pathways
(Govind, 2008).
Pathogen recognition
Immune signaling commences with the recognition of pathogens. As mentioned above, the Drosophila PRRs are responsible for detecting PAMPs, which are found on evolutionarily distant organisms such as bacteria but absent on eukaryotic cells (Murphy,
2012). For example, the bacterial cell wall component peptidoglycan (PGN), lipopolysaccharide (LPS) from Gram-negative bacteria or fungal β 1, 3-glucans are some of the PAMPs associated with microbes (Royet & Dziarski, 2007; Stokes et al., 2015).
Two families of PRRs are identified in Drosophila immune signaling, the peptidoglycan recognition proteins (PGRPs) and the Gram-negative binding proteins (GNBPs) (Stokes et al., 2015). The specific host PRRs differentially detect these differences in the bacterial
PAMPs. For example, PGN from Gram-negative bacteria contains meso-diaminopimelic acid (DAP), which is replaced by lysine (Lys) in Gram-positive bacteria (Lemaitre &
Hoffmann, 2007; Leulier et al., 2003). Using loss-of-function mutants of one of the
PGRPs, PGRP-SA, demonstrated that it is involved in the activation of the Toll pathway in response to Gram-positive bacteria, but not to fungal pathogens (Michel et al., 2001).
Additionally, PGRP-LE functions in the activation of the Imd pathway. Additionally, experiments using PGRP-LE and PGRP-LC double mutants demonstrated that these mutants are more susceptible to Gram-negative bacteria compared to either of the single mutant, thereby implying that these two receptor proteins function together in the
4 detection of Gram-negative bacteria and the subsequent Imd pathway activation
(Takehana et al., 2004). Additionally, GNBP1 is involved in the detection of Gram- negative bacteria, but only those that have Lys-type PGN. Loss-of-function GNBP3 mutant flies exhibit significantly reduced Drosomycin expression and infection with
Gram-positive bacteria did not affect the survival ability of the flies, thus indicating that
GNBP3 functions as a receptor for fungal pathogens (Gottar et al., 2006). Finally, when activated, these PRRs induce intracellular signaling resulting in the activation of genes involved in host defense.
Toll pathway
Pathogen recognition in the Drosophila Toll pathway takes place upstream of the
Toll receptor. Drosophila Toll receptors are responsible for binding to endogenous ligands that are generated through a proteolytic cascade in response to infection (Janssens
& Beyaert, 2003; Tauszig et al., 2000). Thus indicating that they function as cytokine receptors and are not true PRRs. The activated Toll receptor binds to MyD88, an adaptor protein, through the intracellular TIR domains (Horng & Medzhitov, 2001). This in turn recruits Tube (an adaptor protein) and the kinase Pelle, forming a heterodimer through interactions mediated by the death domain. The two distinct death domain surfaces in
Tube are responsible for the separate binding of MyD88 and Pelle (Sun et al., 2002;
Valanne et al., 2011). The downstream events include the phosphorylation of Cactus, an
IκB homologue, which binds to the NF-κB transcription homologues, Dorsal and/or Dif.
The nuclear translocation of the transcription factors induces the production of AMPs
5 such as Drosomycin (Valanne et al., 2011; Wu & Anderson, 1998). PGN-recognition proteins (PGRP)-SA and –SD, and Gram-negative binding protein (GNBP)-1 and 3, carry out pathogen recognition (Michel et al., 2001). In order to activate the Toll pathway, these extracellular recognition proteins are responsible for initiating a protease cascade to activate the Toll receptor ligand Spatzle (Schneider et al., 1994). In non-signaling conditions, Spatzle is found as an inactive precursor and upon activation it dimerizes and binds to the two Toll receptors to initiate downstream signaling events (Arnot et al.,
2010). Upon pathogen recognition, the Spatzle-processing enzyme is responsible for
Spatzle cleavage, thus activating it (Jang et al., 2006). Depending on the type of activating microorganism, SPE is activated via one of three upstream protease cascades
(Valanne et al., 2011). Toll pathway also establishes the dorsal-ventral patterning in the
Drosophila embryo and also controls immune responses against Gram-positive and fungal pathogens (Brennan & Anderson, 2004; Valanne et al., 2011). Interestingly, Toll-7 is essential for viral recognition and activation of antiviral autophagy (Nakamoto et al.,
2012) and the Toll pathway read-out gene encoding the AMP Drosomycin is induced in response to nematode infection (Castillo et al., 2011).
Imd pathway
The Drosophila Imd pathway is activated when the Gram-negative bacterial PGN binds to the receptor (PGRPs) (Dziarski & Gupta, 2006). This binding leads to the recruitment of a signaling complex consisting of Imd (death domain protein homologous to mammalian RIP1 of the TNF-receptor), the adaptor dFADD and Dredd (a caspase-8
6 homolog). When Dredd is activated, it cleaves Imd and creates a binding site for lap2
(E3-ligase inhibitor of apoptosis 2) allowing for ubiquitination of Imd. The activation of
Imd then leads to the recruitment and activation of Tab2/Tak1 complex (Myllymaki et al., 2014). This complex is responsible for the phosphorylation and activation of the
Drosophila IκB complex. This complex then phosphorylates and activates Relish, a NF-
κB protein (Silverman et al., 2000). The nuclear translocation of the active N-terminal part activates the transcription of genes coding for AMPs such as Diptericin (De Gregorio et al., 2002). The Imd pathway effector gene Diptericin is induced in response to nematode infections (Castillo et al., 2011). Loss-of-function mutations in several Imd pathway genes were responsible for increased sensitivity of Drosophila adults to virus infection (Costa et al., 2009).
JAK/STAT pathway
In insects, the Janus kinase/signal transducers and activators of transcription pathway (JAK/STAT) (Fig. 1) was first studied in the mosquito, Anopheles gambiae
(Barillas-Mury et al., 1999). The core components of the JAK/STAT pathway are the three extracellular ligands called Unpaired (Upd), four transmembrane Janus kinase receptors and seven genes coding for STATs (Kisseleva et al., 2002). The signaling initiates with binding of Upd to the transmembrane receptor, Domeless, leading to the activation of receptor associated JAKs called Hopscotch. Upon activation, the JAK tyrosine kinases phosphorylate themselves and create docking sites for STAT binding.
This results in phosphorylation of STAT, which then form dimers and translocate into the
7 nucleus to activate transcription of target genes (Shaposhnikov et al., 2013; Zeidler et al.,
2000). The JAK/STAT pathway was first studied in Drosophila as an embryonic developmental pathway (Zeidler et al., 2000), and is also directed against infections by certain viruses (Kingsolver et al., 2013), and parasitoid wasps in Drosophila larvae
(Yang et al., 2015) and adult flies (Schlenke et al., 2007).
JNK pathway
The c-Jun-N-terminal Kinase (JNK) pathway is mainly stimulated in response to stress signals (Ramet et al., 2002; Stronach & Perrimon, 1999). (Fig.1). The sole JNK in flies, Basket, is phosphorylated by its upstream JNK kinase, Hemipterous, and a JNKK kinase upstream of Basket. This activation is a result of triggers such as cytokines, apoptosis or wounding in Drosophila. The phosphorylation of the transcription factors
Jun and Fos (the AP-1 complex) by Basket, forms a heterodimer of Jun/Fos. This heterodimer then translocates into the nucleus and induces the activation of target genes
(Paul et al., 1997; Stronach & Perrimon, 1999).
8
Figure 1. Overview of the Drosophila humoral and cellular immune responses. Detection of pathogens by PRRs activates the signaling pathways (humoral response). The downstream signaling events induce the production of defense modules (AMPs, serpins, stress factors or clotting factors) in specific immune responsive tissues. Pathogen detection also induces the production of specific hemocytes (cellular response) that respond to a particular pathogen (green). The synergistic activation of immune molecules is responsible for pathogen elimination and tissue repair. Adapted from (Hoffmann & Reichhart, 2002).
9 Drosophila cellular immune responses
Hematopoiesis in Drosophila is responsible for the production of hemocytes, which consist of two populations: circulating and sessile. The circulating hemocytes are free-floating and the sessile cells are found attached to the epithelial tissues. Both hemocyte populations include three types of hemocytes: plasmatocytes, crystal cells, and lamellocytes (Williams, 2007). Each hemocyte type originates from a common precursor expressing the GATA transcription factor, Serpent (Jung et al., 2005; Rehorn et al.,
1996). Plasmatocytes closely resemble the mammalian macrophage/monocyte lineage.
They make up about 90-95% of the total hemocyte population and function in the phagocytic removal of invading microbes and dead cells in adult flies (Fig. 1). In the larval stage, plasmatocytes are primarily involved in the recognition and elimination of microbes whereas in the pupal stage, they are involved in clearing the larval tissues and remodeling of adult structures. Crystal cells make up about 5% of the total population and are involved in the melanization response (Fig. 1). Lamellocytes are rarely observed in the Drosophila embryo or larval stages (Lackie, 1988; Lanot et al., 2001). They participate in the encapsulation of large pathogens such as wasp eggs or nematode parasites (Anderl & Hultmark, 2015; Arefin et al., 2015)(Fig. 1). In addition to the specific transcription factors that regulate hematopoiesis in the Drosophila lymph glands, certain signaling pathways such as the JAK/STAT, Toll, and Notch pathways also play important roles in hematopoiesis (Meister & Lagueux, 2003).
Phenoloxidase response
10 The Drosophila phenoloxidase (PO) response constitutes the production of melanin (a black pigment) at the site of injury or infection. It is an immediate reaction that occurs at the end of both encapsulation of pathogens and coagulation reactions
(wound healing). The melanization reaction initiates with the cleavage of an inactive enzyme called pro-phenoloxidase (proPO) into the active form PO by a serine protease called phenoloxidase-activating enzyme (PPAE) (Soderhall & Cerenius, 1998). This active PO catalyzes the oxidation of phenols leading to non-enzymatic polymerization to produce melanin. The cascade is activated by physical injury and recognition of microbial elicitors such as glucan, lipopolysaccharide (LPS) or PGN by the insect host recognition molecules (PRRs) such as PGRP-SA, GNBP1 and GNBP3 which are also
Toll pathway specific receptors (Cerenius & Soderhall, 2004; Eleftherianos & Revenis,
2011). These receptors are found in the hemolymph and activate PO through serine proteases distinct from those involved in Toll pathway activation. Loss-of-function MP1 or MP2 flies demonstrated reduced PO levels upon microbial challenge indicating that these serine proteases function upstream in the melanization cascade and are involved in the proteolytic cleavage and activation of certain PPOs (An et al., 2013; Binggeli et al.,
2014). Three proPO (PPO)-encoding genes are identified in Drosophila: PPO1 and
PPO2 are expressed in the crystal cells and PPO3 is expressed in the lamellocytes
(Dudzic et al., 2015).
In Drosophila larvae, several studies have demonstrated the effects of melanization on infestation of parasitoid wasp eggs. Mutations affecting the melanization reaction significantly compromised immunity against the eggs of the endoparasitic wasp
Leptopilina boulardi (Nappi et al., 1992; Rizki RM., 1990). A gene encoding a serpin,
11 LbSPɣ, highly expressed in the venom of Leptopilina, specifically targets and inhibits the
PO cascade in the hemolymph of Drosophila yakuba host larvae (Colinet et al., 2009).
Steinernema nematodes are capable of activating the PO response in Drosophila larvae and its Xenorhabdus bacteria are known to interfere with its activation (Pena et al.,
2015).
Nitric oxide
In insects, nitric oxide (NO) has emerged as an important molecule due to its participation in several physiological processes including immune responses to microbial pathogens and parasites (Nappi et al., 2000; Stasiv et al., 2001). NO is a short-lived and highly soluble second messenger that is capable of traveling freely within and to nearby cells. NO is produced by the oxidation of L-arginine to citrulline mediated by the enzyme nitric oxide synthase (NOS). In Drosophila, a single NOS-encoding gene is present in the genome, which can generate several protein isoforms by alternative splicing (Stasiv et al.,
2001). In vertebrates, however, there are three NOS genes. They are classified into two main types: constitutive and inducible. Constitutive NOS is mainly involved in facilitating transmission of cellular signals and is further classified as neuronal and endothelial, whereas inducible NOS is rarely observed in resting cells. In response to proinflammatory cytokines, inducible NOS catalyzes NO synthesis allowing it to grow to toxic concentrations, and this NO reacts with oxygen and oxygen-related reactive intermediates, resulting in toxic species that cause enzymatic and DNA damage (Rivero,
12 2006). Studies have shown that NO is toxic to a wide range of pathogens such as bacteria, fungi, viruses as well as metazoan parasites (Colasanti et al., 2002).
In insects, NO is involved in the cellular and humoral immune responses (Ishii et al., 2013; Sadekuzzaman et al., 2018). NO is essential in the survival of Drosophila flies following infection with gram-negative bacterial pathogens (Eleftherianos et al.,
2014). In addition, in the mosquito, Anopheles stephensi, NO is directly involved in limiting the development of the malaria parasite, Plasmodium berghei (Luckhart et al.,
1998). Although several studies have demonstrated the importance of NO in the insect immune response to pathogens, the mode of action or cellular signals that trigger NO production are not well understood (Rivero, 2006).
Endosymbionts and the Drosophila immune response
In insects, endosymbiotic bacteria exist in two forms, the primary and the secondary form (Dale & Moran, 2006; Kikuchi, 2009). Primary form endosymbionts are vertically transmitted and are essential in the growth, development and fecundity of the hosts in which they reside (Buchner, 1965). These endosymbionts provide essential amino acids and have a long evolutionary history with the insect host (Buchner, 1965;
Dale & Moran, 2006; Eleftherianos et al., 2013). Secondary form endosymbionts can be transmitted vertically or horizontally (Russell & Moran, 2005). These endosymbionts provide a variety of functions in the host such as conferring fitness benefits and increasing resistance to parasitic wasps (Eleftherianos et al., 2017; Jaenike, Unckless, et
13 al., 2010; Oliver et al., 2003). Finally, they have a short evolutionary history with their host (Buchner, 1965).
Wolbachia and Spiroplasma endosymbiontic bacteria are found in the mushroom fly species, Drosophila neotestacea and are commonly infected with the parasitic nematode, Howardula aoronymphium. Infection with Howardula nematodes decreases the mating ability of male flies, causes sterility in female flies and reduces the survival of adult flies (Jaenike & Brekke, 2011). However presence of Spiroplasma in these flies provides protection against infection with Howardula nematodes (Jaenike & Brekke,
2011). In Drosophila adults carrying Wolbachia and Spiroplasma, infection with Gram- negative bacteria does not alter the survival rates of flies, however the transcription of certain immune related genes is upregualted, which was attributed to the presence of these endosymbionts (Shokal et al., 2016).
Entomopathogenic nematodes
Entomopathogenic nematodes (EPNs) infect several different hosts ranging from humans to plants. Parasitic nematode infections are one of the leading causes of gastrointestinal infection in most of the developing countries. These infections are responsible for causing life-threatening conditions (Stepek et al., 2006). Despite their effects on humans, nematode infections are not as well studied compared to other pathogens such as viruses, bacteria, or fungi.
14 EPNs belonging to Steinernematidae and Heterorhabditidae families are increasingly used in biological control practices to manage noxious insect pests (Lacey &
Georgis, 2012). Steinernema carpocapsae lives in mutualism with the Gram-negative bacterium Xenorhabdus nematophila and both of these partners, together or separately, are potent pathogens of insect hosts (Martens et al., 2003; Pena et al., 2015). The EPNs infect insect hosts during their infective juvenile (IJ) stage. To locate the insects, the nematodes have evolved chemosensory features that allow them to detect CO2 released by the insects (Hallem et al., 2011). The IJ stage is developmentally arrested and is similar to the Caenorhabditis elegans dauer stage (Goodrich-Blair, 2007; Hallem et al.,
2011). During infection, the IJs enter the insect host through the cuticle or natural openings (Arefin et al., 2014; Pena et al., 2015). Upon entry into the host, the nematodes expel their mutualistic bacteria (Fig. 2), which then produce toxins and virulence factors thereby suppressing the insect defense systems, eventually resulting in rapid insect death.
The nematodes then develop into the feeding third stage juveniles, which use their proliferated mutualistic bacteria as food and undergo molting to become fourth stage juveniles thereby becoming the first generation adult males and females. Adult nematodes mate, and the females lay eggs that hatch into first stage juveniles and successively molt to the second, third and fourth stages eventually becoming second- generation adult males and females. The cycle of egg laying and juvenile stage continues until the food source (insect) is depleted. At this point, the adult nematodes lay eggs, which develop into IJs. The second stage juveniles reacquire the mutualistic bacteria and develop to the pre-infective juvenile stage and eventually into the IJ stage, in the absence
15 of food (Fig. 2). These IJs then exit the host in search of new susceptible insects and can survive without food for several months (Adams, 2002; Dillman et al., 2012).
Figure 2. An overview of the Steinernema life cycle. The EPN IJs carry their mutualistic Xenorhabdus bacteria and search for insect hosts. They enter the insects and gain access to the hemolymph where they expel the bacteria. The bacteria causes septicemia, allowing the nematodes to undergo development and complete its life cycle. Once the resources from the insect biomass is consumed, new IJs develop and they carry the bacteria in search of new insect hosts. Adapted from (Dillman et al., 2012).
Insect immune response to EPNs
16 Several insect species are identified as natural hosts of EPNs. These hosts allow the EPNs to infect and complete their life cycle. Studies have identified specific host immune responses directed against these EPNs. For example, the larvae of Popillia japonica, a Japanese beetle, exhibit a potent encapsulation and melanization response against the EPNs Heterorhabditis bacteriophora, Steinernema carpocapsae and
Steinernema scapterisci but not against Steinernema glaseri. Additionally, the adults of
Acheta domesticus, the house cricket, direct a strong immune response against
Heterorhabditis, S. carpocapsae and S. glaseri but not against S. scapterisci (Wang et al., 1994). S. glaseri nematodes are more virulent to P. japonica larvae compared to
Heterorhabditis nematodes. This virulence is attributed to the lower encapsulation and melanization reponse against S. glaseri nematodes (Wang et al., 1994). Interestingly, the coleopteran Rhynchophorus ferrugineus larvae fail to encapsulate S. carpocapsae nematodes. With the exception of a sharp decrease in the number of circulating hemocytes, no other type of immune response activation was observed (Manachini et al.,
2013). Heterorhabditis and Steinernema feltiae nematodes are encapsulated significantly earlier and at higher levels in the prepupae of the Colorado potato beetle Leptinotarsa decelineata than in the Galleria mellonella larvae. This led to the conclusion that
Leptinotarsa is a more immunologically responsive host compared to Galleria (Ebrahimi et al., 2011).
17 Drosophila immune response to EPNs
In recent years, the effects of insect immune activation and response against EPN infections have also been studied in Drosophila. Because Drosophila is a widely used model organism to study host-pathogen interactions (Hallem et al., 2007)and due to its significant homology to the mammalian innate system signaling pathways, the study of entomopathogenic nematodes in Drosophila is on the rise. The effect of various microbial pathogens is well understood in Drosophila and from these studies we have learned that each type of pathogen is capable of inducing distinct immune responses in the host (Govind, 2008). Thus, the study of EPN infections in Drosophila is warranted. A major advantage of studying the mutualistic-pathogenic partnership in EPNs such as
Steinernema nematodes is that these EPNs are viable and potent pathogens without their mutualistic Xenorhabdus bacteria (axenic nematodes) (Hallem et al., 2007; Yadav et al.,
2015). Thus, each partner in this mutualistic relationship can be separated and studied individually or in combination. This facilitates the exploration of the host immune responses directed against each pathogen separately or against both partners together
(Eleftherianos, ffrench-Constant, et al., 2010; Hallem et al., 2007; Yadav et al., 2017).
Recently, protocols have been developed for generating viable axenic nematodes
(McMullen & Stock, 2014; Yadav et al., 2015) and studies have demonstrated that these nematodes are potent pathogens of Drosophila adults and larvae (Castillo et al., 2015;
Hallem et al., 2007).
Transcriptomic studies of the insect immune response to EPN infections have identified several molecules involved in the interaction with nematodes (Arefin et al.,
18 2014; Yadav et al., 2017). Recently, a transcriptomic study reported the induction of several known immune genes such as AMPs and PRRs as well as gene families previously known only to function in Drosophila development and metabolism such as chitinases and lipid metabolic processes in Drosophila larvae infected with Steinernema symbiotic or axenic nematodes (Yadav et al., 2017). A genome-wide microarray analysis of the transcriptional response of Drosophila larvae to infection by Heterorhabditis symbiotic nematodes identified strong upregulation of several known immune genes and lower induction of genes participating in oocyte maturation, Wnt and ubiquitin-mediated pathways, and genes coding for cuticular molecules, Tep proteins, and clotting factors
(Arefin et al., 2014). RNA-sequencing analysis of Drosophila adult flies infected with symbiotic or axenic Heterorhabditis nematodes, or their Photorhabdus luminescens bacteria showed that axenic Heterorhabditis nematodes and Photorhabdus bacteria produce distinct transcriptomic profiles and infection with symbiotic nematodes produces a transcriptomic profile that is a combination of the transcriptional profiles of axenic nematodes and its symbiotic bacteria. Also, other differentially expressed genes in
Drosophila in response to Heterorhabditis nematodes included genes involved in stress response, nociception, lipid metabolism as well as potential nematode recognition genes
(Castillo et al., 2015).
In addition to transcriptomic analysis, the functional role of specific Drosophila genes in response to EPN infections has been investigated. For example, inactivation of transglutaminase, a conserved component of clotting factor that also functions in humans, increases the susceptibility of larvae (Wang et al., 2010). The participation of two clotting factors, gp150 and fondue (Hyrsl et al., 2011) as well as a homolog of Tep3, a
19 basement membrane protein (glutactin) and a pathogen recognition protein (GNBP3) contribute towards the control of Heterorhabditis nematodes in Drosophila larvae (Arefin et al., 2014). Infection of Drosophila larvae with Heterorhabditis symbiotic nematodes induces the transcription of four AMP genes (Hallem et al., 2007). Because
Heterorhabditis axenic nematodes fail to induce the transcription of these genes, it was concluded that this antimicrobial response is specific to the mutualistic bacteria,
Photorhabdus.
Steinernema nematodes are more virulent towards Drosophila larvae compared to
Heterorhabditis nematodes and they can upregulate the expression of certain AMP genes and activate the melanization pathway (Pena et al., 2015). Recently, a study reported that the imaginal disc growth factor-3 (Idgf3) functions as a regulatory molecule in the epithelial immune response to Heterorhabditis and is an essential component of the clot matrix. Inactivation of Idgf3 increases susceptibility of larvae and Idgf3 mutant larvae exhibit defects in hemolymph clotting and wound healing responses (Kucerova et al.,
2016). Results from transcriptomic and functional studies in Drosophila model provide valuable information about the role and regulation of insect genes that participate in the interaction with EPN.
Drosophila immune evasion by EPNs
To parasitize the host successfully, the EPNs must either suppress or evade the insect humoral and cellular immune responses. Nematode evasion strategies include: 1)
20 anatomical seclusion- migrating to tissues where they can avoid detection, 2) camouflage or mimicry- secreting molecules or using host immunocompetent cells and sequestering them on the cuticle to avoid detection by host, or 3) interference- producing molecules that actively suppress the host immune defenses (Brivio MF, 2005; Castillo et al.,
2011). Certain molecules from Steinernema and Heterorhabditis nematodes that promote host immune evasion have been identified. Proteases compose the majority of the immunomodulatory factors (Trap & Boireau, 2000). An extracellular protease from
Heterorhabditis and phase 1 variants of Photorhabdus bacteria are able to digest the
AMP cecropin from Galleria and other lepidopteran insects (Fig. 3) (Jarosz, 1998). The nematode cuticle has the ability to prevent host antibacterial defenses. This was attributed to the binding of S. feltiae cuticular lipids to the hemolymph proteins (particularly those involved in proPO cascade) of Galleria (Brivio et al., 2004) (Fig. 3). Steinernema glaseri nematodes produce a surface coat protein (SCP3a) that protects the nematodes from encapsulation in Popillia japonica larvae (Japanese beetle) and this protein could potentially be involved in protecting its bacterial symbiont from phagocytosis (Wang et al., 1999).
Steinernema nematodes have developed strategies to disarm the insect immune system, even before they expel their associated bacteria in the hemolymph. This is demonstrated in Galleria, the hemocytes of which can detect the presence of
Heterorhabditis, but not S. carpocapsae or S. glaseri nematodes (Ebrahimi et al., 2011).
This suggests that Steinernema nematodes employ immunomodulatory factors that protect them from the host cellular response (Fig. 3). To evade host immune recognition,
S. feltiae nematodes use their cuticular surface lipids to remove certain host hemolymph
21 proteins to coat themselves, and thus enabling the parasites to escape detection (Fig. 3).
This strategy allows the nematodes to persist in the insect while also suppressing host
hemocyte aggregation and encapsulation responses (Mastore & Brivio, 2008). In addition
to evasion, the nematodes also produce serine proteases that function as virulence factors
(Fig. 3). A chymotrypsin-like serine protease purified from S. carpocapsae nematodes
induces early apoptosis and disrupt the host gut (Toubarro et al., 2009). Another trypsin-
like from S. carpocapsae nematodes blocks hemocyte spreading and causes severe
morphological changes to G. mellonella hemocytes (Balasubramanian et al.,
2010). Similar functional characterization of host-virulence gene interactions will help to
better understand the insect anti-nematode immune functions.
Figure 3: Strategies employed by entomopathogenic nematodes to evade host immune responses.
Steinernema nematodes use their body surface lipids to sequester certain host hemolymph proteins and expel mutualistic bacteria to avoid encapsulation and inhibit phagocytosis by host hemocytes, respectively. Serpins from Steinernema protect the nematode from host proteolytic enzymes. The nematodes also secrete proteases and unknown factors that help them evade the insect immune defenses. From (Eleftherianos et al., 2017).
22 Thesis structure and composition
The general aim of this thesis was to investigate the modulation of the Drosophila melanogaster immune response against infection with the entomopathogenic nematode
Steinernema carpocapsae. The five chapters are divided as follows:
Chapter 1 ‘An improved method for generating axenic entomopathogenic nematodes’ is published in ‘BMC Research Notes’ 2015, 8: 461. In this chapter, I report a protocol for generating axenic Steinernema carpocapsae nematodes using a mutant
Xenorhabdus nematophila strain (ΔrpoS) and antibiotics. I also report that Steinernema symbiotic or axenic nematodes cause Drosophila larval death at the same rates.
Chapter 2 ‘Endosymbiont-based immunity in Drosophila against parasitic nematode infection’ is published in ‘PLoS One’ 2018, 13: e0192183. Here, I explore the role of the endosymbionts Wolbachia and Spiroplasma in the Drosophila response against
Steinernema nematodes. I find that the presence of Wolbachia alone or together with
Spiroplasma promote the survival of larvae against Steinernema symbiotic nematodes. I also report that presence or absence of endosymbiotic bacteria differentially regulates the immune signaling, phenoloxidase response, and alters metabolic status and fat body lipid droplet size in Drosophila larvae responding to Steinernema symbiotic or axenic nematodes. Finally, these results suggest an interaction between the endosymbiotic bacteria, Wolbachia or Spiroplasma, with the Drosophila immune and metabolic response against infection with the entomopathogenic nematode Steinernema.
23 Chapter 3 ‘RNAseq analysis of the Drosophila response to the entomopathogenic nematode Steinernema’ is published in ‘G3 (Bethesda)’ 2017, 7: 1955-1967. In this chapter, I report the number and nature of Drosophila genes induced in response to infection with Steinernema symbiotic or axenic nematodes. Infection with Steinernema nematodes up- and down-regulates genes with known immune roles as well as genes associated with development, stress response, metabolic functions, peritrophic membrane and chitin binding. I also report genes with potential anti-nematode activity in Drosophila larvae as well as genes conserved in Manduca and humans.
Chapter 4 ‘The imaginal disc growth factors 2 and 3 regulate the Drosophila anti- nematode immunity’ is under review at ‘Parasite Immunology’. Here I report that the chitinase-like Idgf2 and Idgf3 genes are involved in different immune functions in the
Drosophila anti-nematode immune response. I find that inactivation of Idgf2 induces certain humoral signaling pathways, whereas inactivation of Idgf3 not only induces immune signaling pathways, but also hemocyte numbers in response to Steinernema nematode infections.
Chapter 5 ‘Participation of a serine protease gene Jonah66Ci in the Drosophila anti- nematode immune response’ explores the physiological participation of Jonah66Ci in the Drosophila response to Steinernema symbiotic or axenic nematodes. I report that
Jonah66Ci is strongly upregulated in Drosophila larvae responding to Steinernema nematodes. I also report that Jonah66Ci interacts closely with the gut specific processes such as Toll and Imd signaling and Wg/Wnt signaling, the activation of which is differentially altered in response to Steinernema infections. Finally, I report that
24 inactivation of Jonah66Ci reduces nitric oxide and feeding rates of larvae, and infection with Steinernema nematodes does not affect its levels. The outcomes suggest that
Jonah66Ci is essential in maintaining homeostasis of specific physiological functions in
Drosophila larvae and Steinernema nematode infection affects some of these responses.
25 *Chapter 1: An improved method for generating axenic entomopathogenic nematodes
ABSTRACT
Background: Steinernema carpocapsae are parasitic nematodes that invade and kill insects. The nematodes are mutualistically associated with the bacteria Xenorhabdus nematophila and together form an excellent model to study pathogen infection processes and host anti-nematode/antibacterial immune responses. To determine the contribution of
S. carpocapsae and their associated X. nematophila to the successful infection of insects as well as to investigate the interaction of each mutualistic partner with the insect immune system, it is important to develop and establish robust methods for generating nematodes devoid of their bacteria.
Findings: To produce S. carpocapsae nematodes without their associated X. nematophila bacteria, we have modified a previous method, which involves the use of a X. nematophila rpoS mutant strain that fails to colonize the intestine of the worms. We confirmed the absence of bacteria in the nematodes using a molecular diagnostic and two rounds of an axenicity assay involving appropriate antibiotics and nematode surface sterilization. We used axenic and symbiotic S. carpocapsae to infect Drosophila melanogaster larvae and found that both types of nematodes were able to cause insect death at similar rates.
Conclusion: Generation of entomopathogenic nematodes lacking their mutualistic bacteria provides an excellent tool to dissect the molecular and genetic basis of nematode
* This chapter has been published in BMC Res Notes (2015) 8:461
26 parasitism and to identify the insect host immune factors that participate in the immune response against nematode infections.
INTRODUCTION
The entomopathogenic (or insect pathogenic) nematodes Steinernema carpocapsae form an obligate mutualistic association with the Gram-negative bacteria Xenorhabdus nematophila in the family Enterobacteriaceae (Forst et al., 1997). The S. carpocapsae-X. nematophila nematode-bacteria complex has emerged as a biological control agent of diverse insect pest species (Shapiro-Ilan, 2006). Nematodes of the infective juvenile (IJ) stage, which is the only stage that is able to survive outside of the host, enter insects through natural openings or by piercing the body wall (Herbert & Goodrich-Blair, 2007;
Waterfield et al., 2009). Once inside the insect body cavity, the IJ releases the bacteria into the hemolymph where they divide exponentially and produce a wide range of toxins and virulence factors that result in insect death (Nielsen-LeRoux et al., 2012). The nematodes feed on the bacterial biomass, and insect tissues, and nematode reproduction continues over 2-3 generations until the nutrient status of the cadaver deteriorates whereupon progeny IJs colonized with X. nematophila disperse in search of new hosts.
Transmission of mutualistic bacteria by IJ nematodes to the insect is essential for the nematodes to parasitize insects successfully and to reproduce (Clarke, 2014; Richards &
Goodrich-Blair, 2009). Instead the nematodes provide nutrients to their associated bacteria by permitting access to the insect host (Chaston et al., 2011).
27 A major advantage of this mutualistic-pathogenic complex is that S. carpocapsae nematodes, like other entomopathogenic nematodes, are viable in the absence of their mutualistic X. nematophila bacteria (axenic nematodes) (Han & Ehlers, 2000) .
Consequently, each partner in the mutualistic relationship can be separated and studied in isolation or in combination enabling host immune responses to be studied against each partner separately, and against both partners together (Castillo et al., 2012; Eleftherianos,
Joyce, et al., 2010; Hallem et al., 2007; Pena et al., 2015). Therefore, this extremely efficient relationship is an excellent model for simultaneously investigating the molecular and functional basis of anti-nematode and anti-bacterial immune responses in the insect host, as well as for analyzing factors that promote nematode parasitism and bacterial pathogenicity (Castillo et al., 2011; Eleftherianos et al., 2010).
Here we describe a modification of a previous protocol for generating S. carpocapsae entomopathogenic nematodes without the presence of their X. nematophila bacteria
(Mitani, 2004). A recent study has reported in vivo and in vitro laboratory procedures for maintaining entomopathogenic nematodes and a method that precludes the use of antibiotics for generating nematodes free of their mutualistic bacteria (McMullen &
Stock, 2014). To generate S. carpocapsae axenic nematodes, we use X. nematophila mutant bacteria that support the growth of their nematode hosts but are not naturally acquired by the parasites (Vivas & Goodrich-Blair, 2001). This method can be readily used in combination with a wide range of molecular/genetic and physiological techniques to study nematode parasitism and humoral/cellular anti-nematode immune reactions in model insects as well as in insects of agricultural or medical importance.
28 METHODS
Bacterial strains
The mutant bacteria X. nematophila ΔrpoS (Vivas & Goodrich-Blair, 2001) were used for generating S. carpocapsae axenic nematodes. For inoculation of liquid cultures, the bacteria were grown in 2 ml Luria-Bertani broth (BD Difco), overnight at 30°C in a shaker-incubator at 220 rpm. ΔrpoS bacterial cultures were supplemented with 50 μg/ml ampicillin (Fisher Scientific) and 30 μg/ml kanamycin (Corning) because ΔrpoS mutants contain a kanamycin cassette and an ampicillin resistant plasmid (Vivas & Goodrich-
Blair, 2001). An aliquot of 250 μl of the overnight culture was added to fresh 5 ml LB and the mix was incubated at 30°C with shaking for 22-24 hours.
Oily Agar Plates
For preparation of twenty oily agar plates, we mixed 100 ml of growth media containing 0.8% w/v of nutrient broth (BD Difco), 1.5% w/v of bacteriological agar
(Amresco), 0.5% w/v of yeast extract (Amresco) in 89 ml of distilled water. The mix was autoclaved and the following components were then added to the media: 1% v/v of 0.98
M MgCl2, 1.42% v/v sterile corn syrup and 0.4% v/v sterile corn oil. After autoclaving the solution, ampicillin and kanamycin were added to the media and the mix was stirred and then poured into one side of the bi-plates (culture plates with two compartments).
The X. nematophila ΔrpoS bacterial culture (100 μl) was pipetted onto the oily agar media and spread evenly with a sterile spreader. The plates were incubated at 30°C for 24 hours.
29 Nematode surface sterilization
S. carpocapsae worms resuspended in 1 ml of sterile water were pipetted into a 1.5 ml
Eppendorf tube and the solution was spun at 13,000 rpm (17,981 g; Eppendorf 5430R) for 10 seconds at room temperature to obtain a concentrated nematode pellet. The supernatant was discarded and 1 ml of freshly prepared 1% bleach solution was added to the nematode pellet. The suspension was mixed well and the nematode pellet was washed in 1 ml of sterile distilled water to remove the bleach residue. The washing step was repeated 5 times. The nematode pellet was resuspended in appropriate volume of distilled water and the number of nematodes was counted using a stereoscope.
Nematode collection
For nematode collection, 500-700 surface-sterilized S. carpocapsae nematodes were transferred to the bacterial plates that were kept in a cabinet lined with moist paper towels at room temperature. After approximately 10 days, the plates were observed under a stereoscope to monitor the age and condition of the nematodes. When the IJ stage was reached in approximately 2-3 weeks, water traps were prepared and the nematodes were collected in cell culture flasks (Round 1) (Vivas & Goodrich-Blair, 2001).
The nematodes from Round 1 were used to set up a new batch of nematodes, which was done by following the same steps as mentioned above. The resultant nematodes were labelled as Round 2.
Axenicity test
For testing the presence or absence of X. nematophila bacterial cells in S. carpocapsae
30 nematodes, 1 ml of sterile water containing highly concentrated nematodes
(approximately 50 worms/μl) was pipetted into a 1.5 ml Eppendorf tube. The solution was centrifuged at 13,000 rpm (17,981 g; Eppendorf 5430R) for 10 seconds at room temperature. The supernatant was discarded and the nematode pellet was homogenized using a small plastic pestle. The nematode homogenate was plated onto LB agar plates
(one plate per treatment), which were incubated at 30°C for 24 hours. Growth of bacterial colonies on the plates indicated that S. carpocapsae nematodes contained their mutualistic X. nematophila bacteria (symbiotic nematodes) whereas lack of bacterial colonies on the plates indicated the absence of bacteria in the nematodes (axenic nematodes). The experiment was repeated at least five times.
For diagnosing the axenicity status of S. carpocapsae IJ nematodes, 100 μl pellets containing worms from Round 1 and Round 2 of the axenicity assay, and symbiotic nematodes (as positive control) were used. The nematodes were crushed using a pestle and DNA was extracted using the DNeasy Blood and Tissue Kit (Qiagen) by following the manufacturer’s instructions. Set of primers (Forward:
GCCTGGAAAGAGTGGACGAA, Reverse: GTAAGACCAAGGGGCACTCC) specific for X. nematophila XptA2 gene were used for PCR amplification using the HotMasterMix
(5 Prime) (Sheets et al., 2011). The cycling program was as follows: 95°C for 2 minutes,
34 cycles of 95°C for 30 seconds, annealing temperature of 61°C for 1 minute and 73°C for 1 minute followed by 72°C for 10 minutes (Castillo et al., 2011). The samples were viewed on a 1.5% agarose gel to determine the presence or absence of XptA2 bands.
31 Infecting D. melanogaster with nematodes
For infection of Drosophila melanogaster larvae with S. carpocapsae IJ nematodes,
100 μl of 1.5% agarose gel were added to the wells of a 96-well microtitre plate. The agarose was allowed to cool for 3 hours prior to use. Third instar D. melanogaster larvae
(Oregon strain) were transferred onto a Whatman filter paper using a fine soft bristle paintbrush and then washed by pipetting a small drop of sterile water to remove any food debris from their surface. Prior to infection, the symbiotic IJ nematodes were washed with sterile distilled water and the axenic nematodes were surface sterilized using bleach and then washed with distilled water, as mentioned above. The washed nematodes were then suspended in fresh sterile distilled water. A drop of 10 μl of water containing 100 symbiotic or axenic S. carpocapsae IJ nematodes and a single D. melanogaster larva were added to each well of the microtitre plate. Treatment with sterile distilled water (10
μl) served as control. Each row of the 96-well plate was covered with a strip of PCR clear film (Eppendorf) and holes were poked to allow air circulation. Thirty larvae were used per treatment and fresh batches of nematodes for each experiment. The results represent at least three independent experiments conducted on three different days. Values were expressed as means ± the standard deviation. Comparisons between survival curves were performed using a long-rank (Mantel-Cox) test in GraphPad Prism 5.0 software.
RESULTS
The protocol described here reports a modified method for generating S. carpocapsae entomopathogenic nematodes lacking their X. nematophila mutualistic bacteria (Figure
1). Using a standard plating technique, we found that completion of the first round of the
32 process resulted in nematodes containing their X. nematophila bacteria (Round 1, Figure
2A). We also found that surface sterilized nematodes subjected to the first round of the axenicity assay still contained X. nematophila bacteria (Round 1 - SS, Figure 2A). To eliminate all X. nematophila cells from S. carpocapsae nematodes, we repeated the entire method using the nematodes that were generated from Round 1. Repeating the method cleared the nematodes from their associated bacteria, which was further confirmed by surface sterilization of the worms (Round 2 - SS, Figure 2A) leading to the generation of
S. carpocapsae axenic nematodes. Importantly, we found that addition of the nematode sterilization step was crucial for removing the X. nematophila cells from the surface of the worms (Round 2, Figure 2A).
Using a PCR diagnostic method, we amplified the X. nematophila XptA2 gene (213 bp) from DNA samples extracted from bacteria associated with S. carpocapsae nematodes, which had been generated through Round 1, Round 1 - SS, and Round 2 of the axenicity assay. However, there was no amplification of XptA2 sequences from
Round 2 - SS samples (Figure 2B). These results suggested that the axenicity assay was efficient in clearing X. nematophila bacteria from S. carpocapsae nematodes; therefore resulting in the generation of axenic worms.
We have used the symbiotic and axenic S. carpocapsae nematodes in infection assays to assess their potency against D. melanogaster larvae. We found that infection of D. melanogaster third instar larvae with the two types of nematodes resulted in insect death within 4.5 days post challenge with the parasites. Interestingly, we found no significant differences between the survival curves of fruit fly larvae following infection with axenic or symbiotic worms (Figure 3; P>0.1, Log-Rank Test).
33 DISCUSSION
Entomopathogenic nematodes are widely used in crop protection for effective control of soil-borne insect pests, and they are excellent models for dissecting the molecular and genetic basis of parasitism and host anti-nematode immune function (Eleftherianos,
Joyce, et al., 2010; Shapiro-Ilan, 2006). Because the nematode-bacteria complex dissociates once inside the insect (Goodrich-Blair, 2007), it is possible that the host activates distinct immune responses against each mutualistic partner. There is also potential that the nematodes and their associated bacteria employ different strategies to evade or suppress the host immune system. To investigate these possibilities it is important to use robust methods for generating nematode parasites lacking their mutualistic bacteria (axenic nematodes).
Here we report a modified and improved method for the production and experimental use of S. carpocapsae nematodes without their mutualistic X. nematophila bacteria. The current method is based on a previously published procedure (Vivas & Goodrich-Blair,
2001). The relationship between S. carpocapsae and X. nematophila is highly specific and nematodes will only maintain mutualistic associations with their cognate bacteria
(Forst et al., 1997). Therefore, to produce S. carpocapsae nematodes without X. nematophila bacteria we used a X. nematophila strain containing a mutation in the rpoS gene that codes for the transcription factor sigma(S), which regulates survival of the bacteria, resistance to stress and interactions with their nematode host (Vivas &
Goodrich-Blair, 2001). X. nematophila rpoS mutants have been shown previously to abolish the ability of the bacteria to colonize the intestine of S. carpocapsae IJ, which negates the mutualistic relationship between the two partners (Vivas & Goodrich-Blair,
34 2001). A recently described method involves the inoculation of agar plates with surface- sterilized eggs and does not require the addition of antibiotics (McMullen & Stock,
2014). The main differences between the current protocol and previous methods is the use of surface-sterilized IJ nematodes and the incorporation of antibiotics into the media to generate axenic worms. We consider the latter as an important step toward preventing the growth of other unwanted bacteria or fungal contamination in the nematode preparations.
We have used 1% bleach solution for surface sterilization of the nematodes. This method eliminates all X. nematophila bacteria from the surface of the worms. The IJ stage is the developmentally arrested stage of most entomopathogenic nematodes and is analogous to the Caenorhabditis elegans dauer stage and the developmentally arrested infective third stage larva (L3) of many important parasitic nematodes (Goodrich-Blair,
2007). During the IJ stage the nematode mouth is closed (Ciche, 2007; Ciche et al.,
2006), thus treatment with bleach eliminates only the bacterial cells that are present on the surface of the worms without affecting nematode infectivity.
We have found no differences in pathogenicity between axenic and symbiotic nematode infections of D. melanogaster larvae. Given that X. nematophila bacteria are potent pathogens of fruit flies and other insects (Ciche et al., 2006; Herbert & Goodrich-
Blair, 2007), we would have expected to find increased pathogenicity of symbiotic nematodes toward D. melanogaster larvae compared to infections with axenic worms.
The reason for this unexpected result is currently unknown and requires further investigation. Previous studies involving D. melanogaster and Manduca sexta larvae have reported that Heterorhabditis bacteriophora nematodes without their mutualistic
35 Photorhabdus luminescens bacteria are less pathogenic than symbiotic nematodes (Han
& Ehlers, 2000; Pena et al., 2015). However, another study has shown that S. carpocapsae IJ with or without their mutualistic X. nematophila bacteria are equally pathogenic to Spodoptera exigua larvae in laboratory and greenhouse experiments
(Mitani, 2004), and we have recently found that H. bacteriophora symbiotic nematodes are as pathogenic as axenic worms following infection of D. melanogaster adult flies
(Castillo et al., 2012). It is worth noting that all infection experiments in the current study used D. melanogaster Oregon strain larvae whereas infection assays in previous investigations used the Canton-S strain (Hallem et al., 2007; Pena et al., 2015). We have recently found that different D. melanogaster wild-type strains can exhibit strong variation in their immune response against microbial infections (Eleftherianos et al.,
2014).
Our current results suggest that the presence of X. nematophila mutualistic bacteria in
S. carpocapsae nematodes is probably not imperative for the ability of the worms to infect efficiently and to kill D. melanogaster wild-type larvae. Therefore it is possible that X. nematophila contribute to the reproductive fitness of S. carpocapsae nematodes without providing an additional advantage to the pathogenicity of the worms (Vivas &
Goodrich-Blair, 2001). Alternatively, the nematodes may produce certain molecules that could enhance pathogenicity or molecules that could potentially mask the activity of X. nematophila virulence factors that are secreted during infection of insects (Bode, 2009; ffrench-Constant & Bowen, 2000). It is also possible that migration and constant movement of S. carpocapsae nematodes, even in the absence of their mutualistic
36 bacteria, within D. melanogaster larvae could result in severe physical damage of vital insect tissues and organs, which could ultimately lead to insect death.
CONCLUSION
Here we describe a modified method for the generation of parasitic nematodes without their mutualistic bacteria. This method involves the completion of two rounds of an axenicity protocol, the use of appropriate antibiotics and nematode surface sterilization treatment to eliminate the presence of bacterial cells on the surface of the worms. This method is expected to promote studies on the molecular basis of nematode parasitism, host anti-nematode immunity and host-microbial mutualism, and it will assist in the identification of nematode genes that participate in these important biological processes.
FIGURE LEGENDS
Figure 1 Flow diagram of the method for obtaining axenic nematodes. Xenorhabdus nematophila ΔrpoS mutant bacteria are grown overnight and then subcultured before plating on oily agar plates containing antibiotics. Surface-sterilized Steinernema carpocapsae nematodes are transferred to the plates covered by the mutant bacteria and after 3-4 weeks infective juvenile progeny are collected in water-traps. These steps consist the first round (Round 1) of the method. The entire procedure is repeated (Round
2) and the newly emerged nematodes are tested for the presence or absence of mutualistic
X. nematophila bacteria.
37 Figure 2 Validation of nematode axenicity status. (A) To estimate the presence of
Xenorhabdus nematophila bacterial cells in Steinernema carpocapsae nematodes, a nematode pellet is homogenized and the homogenate is spread onto agar plates. The absence of X. nematophila colonies on the plates denotes that the nematodes are free of bacterial cells. Bacterial Colony forming Units (CFU, log scale) are shown in Round 1 and Round 2 of the axenicity assay. SS: Surface sterilized nematodes. Diagnostic PCR for detecting the presence or absence of X. nematophila bacteria in surface-sterilized or non-surface-sterilized S. carpocapsae nematodes that were subjected to a single round of the axenicity assay (Round 1 and Round 1 - SS) or two rounds of the procedure (Round
2 and Round 2 - SS). Symbiotic nematodes served as control. The size of the PCR amplified X. nematophila XptA2 gene is indicated.
Figure 3 Survival results for Drosophila larvae infected by Steinernema nematodes.
Drosophila melanogaster Oregon third instar larvae were infected by axenic (lacking
Xenorhabdus nematophila bacteria) or symbiotic (containing X. nematophila bacteria)
Steinernema carpocapsae infective juvenile nematodes. Treatment with sterile distilled water served as negative control. Survival was monitored every 12 hours. Results showed that axenic and symbiotic nematodes were equally pathogenic to D. melanogaster larvae
(P>0.1, Log-Rank Test; GraphPad Prism 5).
38
Figure 1.
39 A B
M
213b p XptA
2 Figure 2.
40 100
l 80
a
v
i
v r 60
u
s
t
n
e 40 Control
c
r
e Symbiotic
P 20 Axenic 0 0 12 24 36 48 60 72 84 96 108 Time (hours)
Figure 3.
41
*Chapter 2: Endosymbiont-based immunity in Drosophila against parasitic nematode infection
ABSTRACT
Associations between endosymbiotic bacteria and their hosts represent a complex ecosystem within organisms ranging from humans to protozoa. Drosophila species are known to naturally harbor Wolbachia and Spiroplasma endosymbionts, which play a protective role against certain microbial infections. Here, we investigated whether the presence or absence of endosymbionts affects the immune response of Drosophila melanogaster larvae to infection by Steinernema carpocapsae nematodes carrying or lacking their mutualistic Gram-negative bacteria Xenorhabdus nematophila (symbiotic or axenic nematodes, respectively). We find that the presence of Wolbachia alone or together with Spiroplasma promotes the survival of larvae in response to infection with S. carpocapsae symbiotic nematodes, but not against axenic nematodes. We also find that
Wolbachia numbers are reduced in Spiroplasma-free larvae infected with axenic compared to symbiotic nematodes, and they are also reduced in Spiroplasma-containing compared to Spiroplasma-free larvae infected with axenic nematodes. We further show that S. carpocapsae axenic nematode infection induces the Toll pathway in the absence of
Wolbachia, and that symbiotic nematode infection leads to increased phenoloxidase activity in D. melanogaster larvae devoid of endosymbionts. Finally, infection with either type of nematode alters the metabolic status and the fat body lipid droplet size in D. melanogaster larvae containing only Wolbachia or both endosymbionts. Our results
* This chapter has been published in PLoS One (2018) 13: e0192183
42 suggest an interaction between Wolbachia endosymbionts with the immune response of
D. melanogaster against infection with the entomopathogenic nematodes S. carpocapsae.
Results from this study indicate a complex interplay between insect hosts, endosymbiotic microbes and pathogenic organisms.
INTRODUCTION
The soil dwelling nematode parasite Steinernema carpocapsae together with the
Gram-negative bacteria Xenorhabdus nematophila form a mutualistic complex that is pathogenic to insects (Dillman & Sternberg, 2012). X. nematophila bacteria are localized in the gut of S. carpocapsae nematodes, which complete their life cycle in insect hosts
(Martens et al., 2003). The nematodes cause infections at the infective juvenile (IJ) stage, which is the developmentally arrested third larval stage analogous to the dauer stage of the non-pathogenic nematode, Caenorhabditis elegans (Goodrich-Blair, 2007). Upon entry into the insect host, the nematodes release their bacteria into the hemolymph (insect blood), where the latter divide and produce a wide range of toxins and virulence factors that kill the host (Akhurst, 1982; Goodrich-Blair & Clarke, 2007). Although little is known about the contribution of nematode virulence factors to this process, we and others have shown that entomopathogenic (or insect pathogenic) nematodes lacking their mutualistic bacteria are still pathogenic to insects (Castillo et al., 2015; Castillo et al.,
2013; Pena et al., 2015; Yadav et al., 2017; Yadav et al., 2015). Recent studies have demonstrated that the nematodes produce certain molecules that suppress or promote evasion of certain insect immune responses allowing them to survive and reproduce in the insect host (Toubarro, Avila, Hao, et al., 2013; Toubarro, Avila, Montiel, et al., 2013;
43 Toubarro et al., 2010).
Insects have developed a diverse range of immune defenses to combat infection by nematode parasites (Castillo et al., 2011). Most studies have mainly focused on the immune response of insect larvae against entomopathogenic nematodes and the immune response of mosquitoes and black flies against filarial nematodes (Arefin et al., 2014;
Castillo et al., 2011; Erickson et al., 2009; Klager et al., 2002; Pena et al., 2015; Yadav et al., 2017). Insects activate both humoral and cellular immune responses to nematode infections as well as phenoloxidase (PO) and coagulation cascades that lead to melanotic encapsulation of the parasites (Bidla et al., 2005; Ebrahimi et al., 2011; Krautz et al.,
2014; Pena et al., 2015). Certain entomopathogenic nematodes have developed strategies to evade or suppress the insect immune system by preventing or disrupting the activation of immune responses to promote their survival in the host (Castillo et al., 2011; Cooper
& Eleftherianos, 2016; Maizels et al., 2001; Schmid-Hempel, 2009). The fruit fly
Drosophila melanogaster is an outstanding model for innate immunity studies. Its major benefit is the availability of a wide range of genetic tools that permit dissection of the molecular basis of the innate immune response to a range of pathogens (Bier & Guichard,
2012; Dionne & Schneider, 2008; Ramet, 2012). Recent transcriptomic studies have demonstrated the power of using D. melanogaster for identifying the molecular components of the insect immune system that are directed against entomopathogenic nematode infections (Arefin et al., 2014; Yadav et al., 2017). It was recently shown that
Steinernema nematodes are able to upregulate the expression of certain antimicrobial peptide genes and induce the melanization pathway, the activation of which is suppressed by Xenorhabdus bacteria (Pena et al., 2015).
44 Wolbachia and Spiroplasma are the most common and widespread maternally- transmitted facultative endosymbiotic bacteria in insects, and they are naturally harbored by certain D. melanogaster strains (Jaenike, 2012; Jaenike, Stahlhut, et al., 2010;
Kikuchi, 2009; Montenegro et al., 2006). Recent studies have led to the proposition of endosymbiont-based strategies for the control of vector borne diseases (Blagrove et al.,
2012; Glaser & Meola, 2010; Hughes et al., 2011; Moreira et al., 2009; Zele et al., 2012).
D. melanogaster is an excellent system to investigate the effect of endosymbionts on host immune function. Previous studies have shown that the presence of certain Wolbachia strains in D. melanogaster confers resistance to infection by various RNA viruses, fungi and parasitoid wasps (Fytrou et al., 2006; Hedges et al., 2008; Kremer et al., 2012;
Osborne et al., 2009; Teixeira et al., 2008), but not by entomopathogenic bacteria
(Rottschaefer & Lazzaro, 2012; Shokal et al., 2016). The presence of Spiroplasma endosymbionts in D. melanogaster flies does not activate the fly immune system, but induction of Toll or immune deficiency (Imd) immune signaling increases Spiroplasma numbers in the fly hemolymph (Herren & Lemaitre, 2011). Furthermore, mushroom- feeding flies D. neotestacea carrying Spiroplasma have increased tolerance against their natural nematode parasite Howardula aoronymphium (Jaenike & Brekke, 2011), which is probably due to an unknown mechanism that reduces the growth and reproduction of the nematodes in the Spiroplasma-carrying flies. Alternatively, flies carrying Spiroplasma are more sensitive to some Gram-negative bacterial pathogens (Herren & Lemaitre, 2011;
Shokal et al., 2016).
The goal of this research is to investigate whether the presence of heritable endosymbiont Wolbachia alone or together with Spiroplasma can modulate the D.
45 melanogaster immune and metabolic response against S. carpocapsae nematodes that either carry (symbiotic) or lack (axenic) their associated X. nematophila bacteria. For this, we use D. melanogaster strains with or without their heritable endosymbionts for infections with S. carpocapsae symbiotic or axenic nematodes. We explore certain aspects of the immune response and estimate levels of triglyceride, glucose, trehalose, and glycogen in all D. melanogaster strains in the presence or absence of nematode infection. We also investigate the involvement of lipid droplets in the D. melanogaster anti-nematode immune response in the context of host endosymbionts. We find that the presence of Wolbachia in D. melanogaster larvae enhances the survival ability against S. carpocapsae symbiotic nematodes, Wolbachia numbers are reduced in larvae responding to symbiotic nematodes while Xenorhabdus numbers are unaffected, the absence of
Wolbachia induces Toll pathway activation in response to axenic nematodes, and that endosymbionts can affect the metabolic state, and in particular the lipid droplet size, of
D. melanogaster during parasitic nematode infection. Current findings reveal that
Wolbachia and Spiroplasma interact closely with the D. melanogaster immune system and are able to modulate certain aspects of the larval response to infection against a potent nematode parasite.
MATERIALS AND METHODS
Fly stocks
Drosophila melanogaster third instar larvae carrying both Wolbachia pipientis
(strain wMel) and Spiroplasma poulsonii (strain MSRO, designated as W+S+), no endosymbiotic bacteria (W-S-), or Wolbachia only (W+S-) were used in all experiments,
46 as previously described (Shokal et al., 2016). All fly strains were amplified for experimentation with approximately 2.5 g of Carolina Formula 4-24 Instant Drosophila media (Carolina Biological Supply, USA), 10 ml of deionized water, and a few granules
(approximately 0.003 g) of dry baker’s yeast. All fly strains were grown at 25°C and a
12:12-hour light:dark cycle.
Nematode stocks
The entomopathogenic nematode Steinernema carpocapsae harboring the Gram- negative bacteria Xenorhabdus nematophila (symbiotic nematodes) were amplified in the larvae of the wax moth Galleria mellonella. Nematodes lacking X. nematophila (axenic nematodes) were generated as described (Yadav et al., 2015). Prior to use, axenic nematodes were surface sterilized in 1% bleach and washed five times with sterile distilled water to remove any residual bacteria from their surface. Infective Juvenile (IJ) stage nematodes 2-4 weeks old were used in all experiments.
Larval survival
To each well of a 96-well plate (Corning), 100 µl of 1.25% agarose were added.
Sterile water (10 µl) suspensions containing 100 S. carpocapsae symbiotic or axenic nematodes were transferred to each well together with an individual D. melanogaster third instar larva. To remove any food particles from the cuticle, each larva was washed with sterile distilled water prior to infection. The wells were covered with a Masterclear
47 real-time PCR film (Eppendorf, USA) and two holes were pierced for ventilation. For control treatment, 10 µl of sterile water were applied to each larva and survival was monitored every 12 h for up to 108 h post-infection. Twenty larvae per strain per treatment were used and the experiment was repeated three times.
Endosymbiont numbers
Four larvae from each fly strain were infected with S. carpocapsae symbiotic or axenic nematodes and subsequently frozen at 12, 36 and 60 h post infection. DNA samples were extracted from the frozen larvae using DNeasy Blood and Tissue kit
(Qiagen) following the manufacturer’s protocol. For estimation of endosymbiont load, all
DNA samples were normalized to 300 ng. Quantitative PCR was performed in twin-tech. semi skirted- 96 well plates (Eppendorf) in a Mastercycler® ep realplex2 (Eppendorf). The experiments were repeated three times and samples were run as technical duplicates.
Wolbachia and Spiroplasma CFUs were determined using the standard curves generated using plasmid DNA and PCR conditions were followed as described (Shokal et al.,
2016). Relative numbers of Wolbachia and Spiroplasma cells were determined as a ratio of the endosymbiont number in larvae infected with S. carpocapsae symbiotic or axenic nematodes and in control larvae treated with water.
Xenorhabdus nematophila standard curve
DNA from X. nematophila bacteria was extracted using the Invitrogen™ Ambion™
TRIzol™ Reagent. PCR amplifications were performed using the X. nematophila 16S
48 rRNA primer sequences (Table 1). The cycling protocol used was described (Shokal et al., 2016). The samples were run as technical duplicates. Standard curve for X. nematophila 16S rRNA was used to estimate bacterial load in infected larvae using the same method as described before (Shokal et al., 2016).
Xenorhabdus nematophila quantification
Four larvae from each fly strain were infected with S. carpocapsae symbiotic IJs and frozen at 12, 36 and 60 h post infection. DNA samples were extracted from the frozen larvae using the Invitrogen™ Ambion™ TRIzol™ Reagent. DNA samples (300 ng) were used in a total reaction volume of 20 μl. The cycling protocol was the same as described (Shokal et al., 2016). X. nematophila CFUs were calculated using the standard curve. The experiment was repeated three times.
Immune gene signaling
Four larvae from each fly strain were infected with S. carpocapsae symbiotic or axenic IJs and frozen at 12, 36, and 60 h after infection. Controls were treated with water.
Total RNA was extracted using the PrepEase RNA spin kit (Affymetrix USB) following the manufacturer’s instructions and adjusted to 300 ng. Complementary DNA (cDNA) synthesis and qRT-PCR were performed as described (Shokal et al., 2016). Primers were purchased from Eurofin MWG Operon (Table 1). Relative gene transcript levels are calculated relative to the housekeeping ribosomal gene, RpL32, and expressed as a ratio
49 compared to mRNA values of uninfected control samples. The experiment was repeated three times and values represent mean and error bars show standard deviations.
Melanization and PO response
For assessing melanization, 10 larvae from each strain were heat treated to visualize blackening of the crystal cells, as described previously (Duvic et al., 2002). For estimating PO activity, larvae from each fly strain were infected with 10 S. carpocapsae symbiotic or axenic nematodes. Larvae were collected 24 h post infection, washed with
1X cold PBS and hemolymph was collected in 2.5X protease inhibitor (Sigma) by puncturing the larvae with a needle. The hemolymph was then loaded onto a spin column
(Pierce, ThermoFisher) and spun at 13,000 rpm at 4°C for 10 min. Protein concentrations were estimated using a BCA test (Pierce, ThermoFisher) and a mixture of 15 μg of protein (diluted in 2.5X protease inhibitor) with 5 mM CaCl2 was added to L-DOPA (15 mM in phosphate buffer, pH 6.6) for a final volume of 200 μl. The samples were measured at absorbance 492 nm after 34 min of incubation at 29°C in the dark and compared to a blank. Each experiment was run in technical duplicates and repeated three times.
Metabolic activity
Ten-fifteen larvae from each fly strain were infected with 10 S. carpocapsae symbiotic or axenic nematodes, or treated with sterile distilled water and larvae were collected 24 h later. Samples were processed using a previously published protocol
50 (Tennessen et al., 2014). Protein quantification was performed using the PierceTM BCA protein assay kit (ThermoFisher Scientific) following manufacturer’s instructions. The plate containing the samples and BCA reagents was covered and placed in a 37°C incubator for 20 min. Absorbance was measured at 562 nm and protein concentrations of samples were calculated from the standard curve.
For estimating triglycerides in nematode-infected and uninfected larvae, samples were diluted 1:1 in PBS-Tween to which 200 μl of the InfinityTM Triglycerides Liquid
Stable Reagent (ThermoFisher Scientific) had been added. The 96-well plate containing the samples was covered and incubated at 37°C for 30 min, and absorbance was measured at 540 nm. Standard curve was generated using the Glycerol Standard Solution
(Sigma) and Triglyceride content in samples was calculated using the glycerol standard curve.
For estimating trehalose levels in the larvae, samples were diluted 1:8 in
Trehalase Buffer (5 mM Tris pH 6.6, 137 mM NaCl, 2.7 mM KCl). Free glucose was estimated from samples diluted in Trehalase Buffer (TB) whereas trehalose content was calculated from samples digested in Trehalase Stock (3 μl of porcine trehalase in 1 ml of
TB) by subtracting the amount of free glucose from the standard curve.
For estimating glucose and glycogen levels, samples were diluted 1:3 in PBS.
Samples were further divided into two sets; the first set was diluted 1:1 in amyloglucosidase stock (1.5 μl of amyloglucosidase in 1 ml of PBS) and the second set was diluted 1:1 in PBS. Samples (30 μl) from each set were added to individual wells of a
96-well plate and allowed to incubate at 37°C for 60 min. To each well, 100 μl of HK
(Glucose Assay Reagent, Sigma) were added and absorbance was measured at 340 nm
51 after 15 min at room temperature. The amount of glucose was calculated from the samples in PBS using the glucose standard curve. For glycogen, the absorbance of glucose in PBS was subtracted from the absorbance of the samples digested with amyloglucosidase (Sigma). Glycogen content was calculated from the glycogen standard curve.
For all metabolic assays, each experiment was run in technical duplicates and repeated four times. The amounts of triglyceride, trehalose, glucose and glycogen are expressed relative to the total protein content in each sample.
Lipid Droplet (LD) staining
Ten larvae from each fly strain were infected with 10 S. carpocapsae symbiotic or axenic nematodes and samples were collected 24 h post-infection. Dissections were performed in 1X PBS, and fat body tissues were separated from the rest of the larval carcass. They were then fixed in 4% paraformalydehyde prepared in PBS for 30 min at room temperature followed by rinsing with 1X PBS twice. They were then incubated in the dark for 30 min in 1:1000 dilution of 0.05% Nile Red in 1 mg/ml of methanol. These tissues were then mounted in ProLongTM Diamond AntiFade Mountant with DAPI (Life
Technologies). Images were obtained using a Confocal Olympus FluoViewTM FV1000 imaging system. Data were collected from fat body tissues of each of the 10 larvae. LD area was assessed using ImageJ software (National Institutes of Health). A minimum of five random regions were selected for LD size quantification from each fat body tissue.
52 Statistical analysis
All values were expressed as means ± standard deviation. Survival experiments were analyzed using a log-rank (Mantel-Cox) and Chi square tests. Bacterial load and endosymbiont numbers were analyzed using unpaired two-tailed t-test. Gene expression,
PO activity, metabolic activity and lipid droplet sizes were analyzed using one-way analysis of variance (ANOVA) with a Tukey post-hoc test for multiple comparisons. All figures were generated using GraphPad Prism7 software.
RESULTS
Presence of Wolbachia in Drosophila enhances the survival response to symbiotic nematode infection
We first estimated the survival ability of D. melanogaster W+S-, W+S+ and W-S- larvae in response to S. carpocapsae symbiotic or axenic nematodes. We found significant differences in the survival among each D. melanogaster strain infected by either symbiotic or axenic nematodes and the uninfected controls (Fig. 1). We also found that
W+S+ larvae infected with symbiotic nematodes survived significantly better than W-S- larvae (Log-rank test, P<0.0001; Fig. 1A), and this result was reversed upon infection with axenic nematodes (log-rank test, P<0.0001; Fig. 1A). We further observed that the
W+S- larvae survived the infection with symbiotic nematodes longer than the W-S- individuals (log-rank test, P<0.0001; Fig. 1B), but there were no statistically significant differences in survival between W-S- and W+S- larvae upon infection with axenic nematodes (log-rank test, P=0.6154, Fig. 1B). These results suggest that the presence of
53 Spiroplasma does not affect the survival of D. melanogaster against symbiotic S. carpocapsae when Wolbachia is also present; however, the presence of Spiroplasma together with Wolbachia is detrimental to the larvae upon axenic nematode infection.
Wolbachia numbers are reduced in Drosophila responding to symbiotic nematodes while Xenorhabdus numbers are unaffected
To estimate whether infection by S. carpocapsae nematodes affected the numbers of endosymbiotic bacteria in D. melanogaster, we infected W+S-, W+S+ and W-S- larvae with S. carpocapsae symbiotic or axenic nematodes and estimated the endosymbiont numbers at different time points post infection. Interestingly, Wolbachia numbers were significantly reduced in W+S- larvae infected with axenic nematodes compared to symbiotic nematodes at 36 h post-infection (P=0.0076; Fig. 2A), and they were also significantly lower than those in W+S+ larvae infected with axenic nematodes at 36 h post infection (P=0.0143; Fig. 2A). There were no significant changes in the numbers of
Spiroplasma in W+S+ larvae infected by either symbiotic or axenic nematodes at any time-point (P>0.05; Fig. 2B). These results imply that the presence of X. nematophila bacteria in S. carpocapsae and the presence of Spiroplasma in D. melanogaster larvae can affect the number of Wolbachia endosymbionts at late times after infection with the nematode parasites.
To estimate whether X. nematophila replication is affected in the presence or absence of the endosymbionts in D. melanogaster, we infected W-S-, W+S+ and W+S- larvae with S. carpocapsae symbiotic nematodes and estimated the X. nematophila numbers at three time-points post infection. We found no significant differences in the number of X.
54 nematophila Colony Forming Units (CFUs) in W+S+ larvae among the different time- points post infection (P>0.05; Fig. 2C). In W-S- larvae, the increase in X. nematophila
CFUs was not statistically different (P>0.05; Fig. 2C). In addition, larvae carrying only
Wolbachia (W+S-) contained fewer X. nematophila CFUs at 60 h post symbiotic nematode infection compared to 12 and 36 h, but again no statistically significant difference was observed (P>0.05; Fig. 2C). These results suggest that the presence of
Wolbachia and Spiroplasma endosymbionts in D. melanogaster does not have a significant impact on X. nematophila load during S. carpocapsae nematode infection.
Absence of Wolbachia in Drosophila can induce Toll pathway activation in response to axenic nematode infection
The transcriptional activation of immune signaling pathway read-out genes forms the hallmark of the D. melanogaster humoral immune response. Here, we followed this approach to examine immune signaling pathway activation upon infection with S. carpocapsae symbiotic or axenic nematodes in the presence or absence of endosymbionts
(Fig. 3). For this, we used real-time quantitative reverse transcription polymerase chain reaction (qRT-PCR) and gene-specific primers to determine the transcript levels of
Diptericin as a readout for the Imd pathway, Drosomycin for the Toll pathway, Turandot
A (TotA) for the Jak/Stat pathway and Puckered for the Jnk pathway (Brun et al., 2006;
Imler & Bulet, 2005; Kaneko & Silverman, 2005; McEwen & Peifer, 2005). We found that induction of Diptericin was higher in W-S- larvae infected with symbiotic nematodes at all three time points post infection, but this induction was not statistically significant compared to axenic nematode infections (Fig. 3A, B, C). We also found no changes in
55 Drosomycin transcript levels between the different strains at 12 and 60 h post nematode infection (P>0.05; Fig. 3D and F), but Drosomycin transcript levels were significantly higher in W-S- larvae than in W+S+ and W+S- individuals infected with axenic nematodes at 36 h (P=0.0165 and P=0.0141, respectively) and compared to control uninfected larvae (P=0.0212; Fig. 3E). There were no significant differences in TotA and
Puckered transcript levels among the three D. melanogaster strains infected by either symbiotic or axenic nematodes compared to uninfected controls (P>0.05; Fig. 3G-I and J-
L). These results suggest that the absence of Wolbachia endosymbionts in D. melanogaster larvae allowed for the Toll signaling activation in the context of axenic nematode infection.
Endosymbionts do not affect the PO response to nematode infection in Drosophila
We first examined the melanization response of each D. melanogaster strain carrying or lacking endosymbionts after heat treatment (Duvic et al., 2002), and observed that larvae developed dark spots indicating PO activation in crystal cells (Fig. 4A). Upon nematode infection, PO activity in W-S- larvae was significantly higher in response to symbiotic nematodes compared to axenic nematodes and uninfected controls (P=0.0392; Fig. 4B).
In W+S+ larvae, PO activity was also significantly higher upon symbiotic nematode infection compared to uninfected larvae (P=0.0111; Fig. 4B). However, in W+S- larvae,
PO activity was higher upon infection with axenic nematodes compared to symbiotic nematode infections, but this difference was not statistically significant (P=0.9972; Fig.
4B). These results suggest that S. carpocapsae symbiotic nematode infection in D.
56 melanogaster larvae can induce PO activity, which is not significantly affected by the presence or absence of endosymbiotic bacteria.
Endosymbionts can alter the metabolic state of Drosophila upon nematode infection
To estimate whether the presence or absence of endosymbionts affects the metabolic functions of D. melanogaster in response to nematode infection, we infected W-S-,
W+S+ and W+S- larvae with S. carpocapsae symbiotic or axenic nematodes and measured various metabolic processes 24 h post infection. We found that upon axenic or symbiotic nematode infection, changes in triglyceride concentrations were not statistically significant (P>0.05; Fig. 5A). Also, trehalose levels were not statistically different among the various treatments (P>0.05; Fig. 5B). Interestingly, the amount of glucose was significantly lower in W+S- larvae compared to W+S+ larvae upon infection with symbiotic or axenic nematodes (P=0.0251 and P=0.0469, respectively) as well as in uninfected controls (P=0.0422; Fig. 5C). We further found a significant decrease in glucose levels in W+S- larvae compared to W-S- larvae when responding to axenic nematodes (P=0.0032; Fig. 5C). In the context of symbiotic nematode infection, the amount of glycogen in W+S- larvae was significantly higher than in W+S+ larvae
(P=0.0324; Fig. 5D). These findings indicate that Wolbachia and Spiroplasma can affect glucose and glycogen levels in D. melanogaster larvae upon S. carpocapsae nematode infection, but have no effect on triglyceride or trehalose levels.
57 Presence of both endosymbionts can alter lipid droplet size in Drosophila larvae responding to parasitic nematodes
Recent studies have demonstrated an interaction between the host and pathogen metabolism. The supply of metabolites from the commensal bacteria to its host can be consumed by the pathogen, which leads to an increase in the lipid droplet size in the insect fat body (A. C. Wong et al., 2016). Here, we evaluated whether the presence or absence of endosymbionts influences the size of lipid droplets in D. melanogaster during infection with entomopathogenic nematodes. We found that in uninfected individuals, the size of lipid droplets was significantly larger in W+S+ larvae compared to W-S- individuals (P=0.0081; Fig. 6A, B). We also found that W+S+ larvae contained lipid droplets of larger size upon infection with symbiotic nematodes compared to W-S- and
W+S- larvae (P<0.0001) and to uninfected controls (P<0.0001; Fig. 6A, B). In contrast,
W+S+ larvae contained reduced size lipid droplets upon axenic nematode infection compared to controls (P=0.0066; Fig. 6A, B). The lipid droplet size in W+S- larvae was unaffected by nematode infection. These results suggest that the presence or absence of both endosymbionts might alter the lipid droplet size in response to S. carpocapsae axenic or symbiotic infections. On the contrary, the presence of Wolbachia alone has no effect on the size of lipid droplets upon nematode infection.
DISCUSSION
Previous studies in D. melanogaster adult flies have shown a protective role for
Wolbachia, but not Spiroplasma, in response to certain viral infections (Hedges et al.,
2008; Teixeira et al., 2008), but not against bacterial infections (Herren & Lemaitre,
58 2011; Rottschaefer & Lazzaro, 2012; Shokal et al., 2016; Wong et al., 2011). Here, we explore the modulation of the D. melanogaster immune and metabolic responses, in the presence of Wolbachia alone or together with Spiroplasma, against S. carpocapsae nematodes. We find that the presence of Wolbachia alone or together with Spiroplasma in D. melanogaster larvae increases their survival upon infection with symbiotic S. carpocapsae; whereas the presence of both endosymbionts reduces larval survival in response to axenic worms. Interestingly, Drosophila neotestacea flies carrying
Spiroplasma show delayed mortality when parasitized with Howardula aoronymphium nematodes; however, Wolbachia does not participate in the survival response to these nematodes (Hamilton et al., 2016; Jaenike, Unckless, et al., 2010). Similarly, the presence of Wolbachia in Aedes pseudoscutellaris has no effect on the mosquito survival to Brugia pahangi filarial nematodes (Dutton & Sinkins, 2005). Our current results indicate that the effect of Wolbachia alone or together with Spiroplasma on S. carpocapsae during infection of D. melanogaster larvae depends on the presence or absence of the mutualistic X. nematophila bacteria in the nematode parasites.
We then investigated whether infection of D. melanogaster with S. carpocapsae alters the number of endosymbiotic bacteria in the infected larvae. Although compared to larvae not exposed to nematodes, Spiroplasma numbers remain unaffected in W+S+ larvae upon infection with S. carpocapsae, infection with either type of nematode
(symbiotic or axenic) reduces Wolbachia numbers in W+S+ and W+S- D. melanogaster larvae. This could imply that Wolbachia, but not Spiroplasma, forms a target for S. carpocapsae pathogenesis. In agreement with the current findings, we have found previously that infection of D. melanogaster adult flies with Photorhabdus luminescens,
59 the mutualistic bacterium of the entomopathogenic nematodes Heterorhabditis bacteriophora, has no effect on Spiroplasma numbers (Shokal et al., 2016). Interestingly,
P. luminescens infection caused a reduction in Wolbachia numbers in flies carrying this endosymbiont only. This suggests that certain entomopathogenic nematodes and their mutualistic bacteria employ currently unknown strategies to interfere with the growth of endosymbionts in certain insect hosts. Microbial infection can also increase endosymbiont numbers in D. melanogaster, as demonstrated by the rise of Spiroplasma in flies infected with Micrococcus luteus or Erwinia cartovora (Herren & Lemaitre,
2011). Together these findings indicate species-specific interactions between exogenous microbes and endosymbiotic bacteria in D. melanogaster.
A previous study has also estimated the impact of endosymbionts on pathogen load in infected flies and found that the presence of Wolbachia does not influence the replication of Pseudomonas aeruginosa (Wong et al., 2011). Similarly, we have observed recently that the presence of endosymbionts in D. melanogaster adult flies do not affect
P. luminescens numbers (Shokal et al., 2016). Our current data are in agreement with these findings because we also find no changes in X. nematophila cell numbers in any of the strains used in the experiments. This suggests that the growth of this entomopathogenic bacterium is independent of the presence of Wolbachia alone, the simultaneous presence of Wolbachia and Spiroplasma, or the absence of both endosymbionts in D. melanogaster.
The transcriptional induction of genes encoding antimicrobial peptides or other effector molecules serves as an indicator of immune signaling activation in D. melanogaster (Ferrandon et al., 2007). We have investigated whether endosymbionts in
60 D. melanogaster larvae can affect the induction of immune-related genes in the context of nematode infection. Our results demonstrate that in the absence of both endosymbionts,
S. carpocapsae axenic nematodes upregulate the Toll pathway. It was previously shown that infection with H. bacteriophora axenic nematodes also upregulated Drosomycin transcript levels compared to symbiotic nematodes in D. melanogaster flies (Castillo et al., 2013). These results show that induction of Toll signaling is not specific to S. carpocapsae nematodes only. In addition, failure of S. carpocapsae symbiotic and axenic nematodes to provoke immune gene upregulation in larvae containing Wolbachia only or both Wolbachia and Spiroplasma suggests a potential interference with the activation of immune signaling in response to nematode attack. The nature of the molecular mechanism through which endosymbionts might interact with the D. melanogaster immune signaling during nematode infection requires further investigation.
PO is the primary enzyme that regulates melanization at the wound sites and around invading microbes in the hemolymph (Eleftherianos & Revenis, 2011); however,
X. nematophila bacteria released from S. carpocapsae nematodes can suppress the melanization response (Pena et al., 2015). Here we show that in the absence of both
Wolbachia and Spiroplasma, S. carpocapsae nematodes and their associated X. nematophila bacteria fail to suppress the PO activity in D. melanogaster larvae, but they are able to suppress the activity of the enzyme in the presence of Wolbachia alone or in the presence of both endosymbionts. This implies that the suppression of PO activity by this nematode-bacteria complex is strongly dependent on the presence of Wolbachia and probably Spiroplasma endosymbiotic bacteria. Of note, the introduction of certain
Wolbachia strains into D. melanogaster and Drosophila simulans flies as well as Aedes
61 aegypti mosquitoes triggers hemolymph melanization in the absence of infection with exogenous pathogenic microbes (Thomas et al., 2011), suggesting a complex interaction between endosymbiotic bacteria and the insect phenoloxidase response.
Dietary macronutrients are one of the essential factors that promote host- endosymbiont interactions (Ponton et al., 2015) and that the host metabolism may be altered in the presence of endosymbionts and in the context of nematode infection. D. melanogaster flies carrying Wolbachia have elevated insulin signaling (Ikeya et al.,
2009), and in Brugia malayi that contain Wolbachia, the endosymbiont relies on the nutrients (glucose) and energy stores (glycogen) of its host filarial nematode (Voronin et al., 2016). Here we show that D. melanogaster larvae carrying Wolbachia have increased levels of glycogen and trehalose, whereas levels of triglyceride and glucose are unchanged. Our results are consistent with the notion that these endosymbionts confer little to no beneficial fitness effect to their host. Spiroplasma also relies on host lipid availability for its own proliferation (Herren et al., 2014). Lipid metabolism and storage in D. melanogaster occurs in lipid droplets, which are mainly localized in the fat body tissue, although recent evidence indicates that lipid droplets perform additional functions through interactions with pathogenic microbes (Anand et al., 2012). We show that the presence of both Wolbachia and Spiroplasma, but not Wolbachia alone, results in an increase in the number and size of lipid droplets in the fat body, suggesting increased lipid accumulation in the fat body. We also show that larvae carrying or lacking both endosymbionts have increased lipid droplet size upon symbiotic nematode infection, which correlates with higher levels of triglycerides, whereas infection with axenic nematodes has the opposite effect. These results suggest that despite the presence or
62 absence of Wolbachia and Spiroplasma, X. nematophila mutualistic bacteria may affect fatty acid concentrations during infection; however, these changes in the host do not promote pathogen replication.
In spite of recent advances in the insect innate immunity field, our understanding of the role of endosymbiotic bacteria in the host immune response to entomopathogenic nematode infections remains largely unexplored. Results from the research presented here will improve our understanding of the complex symbiotic interactions between eukaryotic hosts and microbial organisms in the context of parasitic infections. From the practical point of view, a better understanding of insect-endosymbiont relationships could potentially lead to the development of alternative strategies for the efficient management of agricultural insect pests and vectors of human diseases.
FIGURE LEGENDS
Figure 1. Survival of Drosophila melanogaster larvae carrying or lacking endosymbionts in response to nematode infection. Survival of D. melanogaster third- instar larvae upon infection with Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes. Sterile distilled water served as control (C) treatment. (A) Survival response of D. melanogaster strains lacking both Wolbachia and Spiroplasma (W-S-) and strains carrying both endosymbionts (W+S+), (B) Survival response of D. melanogaster strains lacking both endosymbionts (W-S-) and strains carrying Wolbachia only (W+S-).
Survival was tracked every 12 h for 96 h and is represented as percent survival on the graph. Data were analyzed using the Log-Rank test (GraphPad Prism7 software). The
63 experiment was repeated three times and bars represent standard errors (****P<0.001,
****P<0.0001).
Figure 2. Numbers for endosymbiotic and pathogenic bacteria in Drosophila melanogaster larvae responding to nematode infection. D. melanogaster third instar larvae carrying no endosymbionts (W-S-), both endosymbionts (W+S+) or only
Wolbachia (W+S-) were infected with Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes. Relative number of cells for (A) Wolbachia at 12 and 36 h, and
(B) Spiroplasma at 12, 36 and 60 h were determined using quantitative PCR. (C)
Numbers of colony forming units of Xenorhabdus nematophila were estimated at 12, 36 and 60 h post infection using quantitative PCR. Data were analyzed using an unpaired two-tailed t-test. Means from three independent experiments are shown and standard deviations are represented by error bars (*P<0.05, **P<0.01).
Figure 3. Transcript levels of immune genes in Drosophila melanogaster larvae carrying or lacking endosymbionts upon nematode infection. Gene transcript levels for (A, B and C) Diptericin, (D, E and F) Drosomycin, (G, H and I) Turandot-A (Tot-A), and (J, K and L) Puckered in D. melanogaster larvae containing no endosymbionts (W-S-
), both Wolbachia and Spiroplasma (W+S+), or Wolbachia only (W+S-) at 12, 36 and 60 h after infection with Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes. Sterile distilled water served as control (C) treatment. The experiment was repeated three times and error bars show standard deviations. Data were analyzed using one-way analysis of variance with a Tukey post hoc test (*P<0.05).
64 Figure 4. Phenoloxidase activity and melanization response in uninfected and nematode-infected Drosophila melanogaster larvae carrying or lacking endosymbionts. (A) Melanization response in D. melanogaster larvae containing no endosymbionts (W-S-), both Wolbachia or Spiroplasma (W+S+), or Wolbachia only
(W+S-) following heat treatment. (B) Relative phenoloxidase (PO) activity was measured in the larval hemolymph of the three D. melanogaster strains at 24 h post-infection with
Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes. Sterile distilled water served as control (C) treatment. The experiment was repeated three times and error bars show standard deviations. Data analysis was performed using one-way analysis of variance with a Tukey post hoc test (*P<0.05).
Figure 5. Metabolic activity in Drosophila melanogaster larvae carrying or lacking endosymbionts following nematode infection. D. melanogaster third instar larvae lacking both endosymbionts (W-S-), carrying both Wolbachia and Spiroplasma (W+S+), or containing Wolbachia only (W+S-) were infected with Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes. Application of sterile distilled water served as control (C) treatment. The relative amount of (A) Triglycerides, (B) Trehalose, (C)
Glucose and (D) Glycogen was estimated 24 h post-infection. The experiment was repeated three times and error bars show standard deviations. Data were analyzed using one-way analysis of variance with a Tukey post hoc test (*P<0.05, **P<0.01).
Figure 6. Lipid droplet size in Drosophila melanogaster larvae carrying or lacking endosymbionts upon nematode infection. (A) Representative images of lipid droplets
65 (LD) labeled with Nile Red and DAPI (blue) in fat body tissues of D. melanogaster third instar larvae lacking both endosymbionts (W-S-), containing both Wolbachia and
Spiroplasma (W+S+), or carrying Wolbachia only (W+S-) followed infection with
Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes. Sterile distilled water served as control (C) treatment. Magnification: 40X. (B) Quantification of LD area in the fat body tissues obtained from 10 D. melanogaster larvae per treatment using
ImageJ. Values show the means from three independent experiments and error bars show standard deviations. Data were analyzed using one-way analysis of variance with a Tukey post hoc test (*P<0.05, **P<0.01, ****P<0.0001).
66 A 100
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Figure 1.
67 Wolbachia A B Spiroplasma
10
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12 h 36 h 12 h 36 h 60 h
Xenorhabdus C 2×1015 2×1015
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0 - + - - + - - + - S S S S S S S S S - + + - + + - + + W W W W W W W W W
12 h 36 h 60 h
Figure 2.
68 12 h 36 h A B C 60 h 800 1500 8000
e
c 6000
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n 2000 u 500 b 400 Diptericin a 300
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a
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D E F 80 350 400
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Figure 3.
69
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71 C Sy Ax
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Figure 6.
72 Table 1: Primer sequences and annealing temperatures used for quantitative RT-
PCR (qRT-PCR).
Gene Accession No Primer Sequence (5’-3’) Tm (°C)
Diptericin CG10794 Forward TGCGCAATCGCTTCTAC 56
Reverse GTGGAGTGGGCTTCATG
Drosomycin CG10810 Forward TGAGAACCTTTTCCAATATGATG 56
Reverse CCAGGACCACCAGCAT
Turandot-A CG31509 Forward AGATCGTGAGGCTGACAAC 61
Reverse CCTGGGCGTTTTTGATAA
Puckered CG7850 Forward GGCCTACAAGCTGGTGAAAG 61
Reverse AGTTCAGATTGGGCGAGATG
RpL32 CG7939 Forward GATGACCATCCGCCCAGCA 61
Reverse CGGACCGACAGCTGCTTGGC
X. nematophila 16s rRNA Forward GCTTGCTGTTTTGCTGACGA 61
Reverse CCGAAGGTCCCCCACTTTAC
73 *Chapter 3: RNAseq analysis of the Drosophila response to the entomopathogenic nematode Steinernema
ABSTRACT
Drosophila melanogaster is an outstanding model to study the molecular and functional basis of host-pathogen interactions. Currently our knowledge of microbial infections in
D. melanogaster is well understood, however the response of flies to nematode infections is still in its infancy. Here we have used the potent parasitic nematode Steinernema carpocapsae that lives in mutualism with its endosymbiotic bacteria Xenorhabdus nematophila to examine the transcriptomic basis of the interaction between D. melanogaster and entomopathogenic nematodes. We have employed next generation
RNA-sequencing to investigate the transcriptomic profile of D. melanogaster larvae in response to infection by S. carpocapsae symbiotic (carrying X. nematophila) or axenic
(lacking X. nematophila) nematodes. With the help of bioinformatic analyses we have found that genes with different biological functions are induced in D. melanogaster larvae responding to either symbiotic or axenic nematodes. We have also identified the strong induction of genes that are associated with the peritrophic membrane, function in stress responses as well as several genes that participate in developmental processes in response to nematode infections. We further show that while symbiotic nematode infection induced certain known immune-related genes, axenic nematode infection
* This chapter has been published in G3 (2017) 7: 1955-1967
74 induced several genes associated with chitin binding, lipid metabolic functions and neuroactive ligand-receptors. In addition, we have identified genes with potential role in nematode recognition and genes with potential anti-nematode activity. Findings from this study will undoubtedly set the stage for identifying key regulators of anti-nematode immune mechanisms in D. melanogaster as well as in other insects of socioeconomic importance.
INTRODUCTION
Parasitic nematodes infect both vertebrate and invertebrate animals and cause serious diseases of socioeconomic importance (Krecek & Waller, 2006; Stock, 2005). The lack of good animal models has hindered the study of parasitic nematode infections in humans and agricultural pests (Hawdon, 2014; Hotez, 2009). Insects have emerged as convenient models to study host response to nematode parasitism because they share considerable homology to certain mammalian molecular factors (Loker, 1994). The common fruit fly,
Drosophila melanogaster, has been established as a supreme model organism to investigate the molecular basis of the interactions between hosts and microbes and to identify the genetic pathways that participate in the host response to pathogenic microorganisms due to the substantial similarities it shares with the physiological processes of vertebrate animals, including humans (Bier & Guichard, 2012; Dionne &
Schneider, 2008; Ramet, 2012).
Entomopathogenic nematodes are facultative parasites of insects, and members of the Steinernematidae family are potent pathogens of a wide range of insect species
75 (Dillman et al., 2012; Gaugler, 1995; Poinar, 1972). Steinernema carpocapsae nematodes associate mutualistically with the Gram-negative bacteria Xenorhabdus nematophila to invade and kill insects (Pena et al., 2015). These nematodes cause infections at the infective juvenile (IJ) stage, which is the developmentally arrested third larval stage analogous to the dauer stage of the non-pathogenic nematode, Caenorhabditis elegans
(Goodrich-Blair, 2007). IJs gain access into the host either by entering through natural openings or by penetrating through the insect cuticle (Arefin et al., 2014; Pena et al.,
2015). Once inside the host, the IJs release their mutualistic Xenorhabdus bacteria, which secrete a wide range of toxins, some of which interfere with the host immune response
(Goodrich-Blair & Clarke, 2007). The nematodes also produce molecules that suppress or evade certain insect immune functions in order to survive and complete their life cycle in their insect host (Castillo et al., 2011). The nematodes reproduce using the insect cadaver as food source and once the resources are depleted, they reacquire the bacteria and exit as
IJs in search of a new prey (Goodrich-Blair, 2007).
Recent studies have shown that S. carpocapsae is more pathogenic to D. melanogaster larvae compared to Heterorhabditis bacteriophora nematodes, which could lead to changes in the transcriptome profile of the host. Therefore S. carpocapsae can be used to explore the interplay between certain aspects of the insect immune response and nematode parasitism strategies (Castillo et al., 2011; Pena et al., 2015). In a previous microarray study, the transcriptome of D. melanogaster larvae infected with H. bacteriophora nematodes was analyzed. The authors identified the participation of tep and Imaginal Disc Growth Factor (Idgf) genes, Peptidoglycan Recognition Proteins
(PGRP-LC) and some unknown genes with putative immune function against H.
76 bacteriophora nematodes (Arefin et al., 2014). A later study used whole genome mRNA sequencing (RNAseq) to analyze the transcriptome of D. melanogaster adult flies responding to H. bacteriophora symbiotic or axenic nematodes or their mutualistic bacteria, Photorhabdus luminescens. This study revealed the participation of several different types of genes encoding lipases and heat shock proteins as well as genes that are involved in stress response, metabolism, and neuronal functions against these pathogens
(Castillo et al., 2015).
Previous and recent work has demonstrated the power of using Drosophila for studying the molecular/genetic basis of insect immune responses against infections by entomopathogenic nematodes. Infection of D. melanogaster larvae with H. bacteriophora symbiotic nematodes results in the transcriptional activation of four antimicrobial peptide-coding genes (Hallem et al., 2007). The antimicrobial peptide response is specific to P. luminescens bacteria because axenic nematodes fail to induce AMP gene transcription. We recently found that H. bacteriophora symbiotic and axenic nematodes induce transcription of several immune-related genes in adult flies, but injection of P. luminescens bacteria alone results in lower levels of gene transcription in the fly (Castillo et al., 2012). Inactivation of D. melanogaster transglutaminase, a conserved component of clotting cascades in insects and humans, results in decreased aggregation of zymosan beads and increased sensitivity of larvae to infection by H. bacteriophora symbiotic nematodes (Wang et al., 2010). Two clotting factors (gp150 and fondue), a homolog of thioester-containing complement protein 3, a basement membrane component (glutactin), a recognition protein (Gram-Negative Binding Protein-like 3; GNBP3) and several small peptides contribute to the immune response of D. melanogaster larvae against H.
77 bacteriophora symbiotic nematodes (Arefin et al., 2014; Hyrsl et al., 2011). It has further been shown that S. carpocapsae symbiotic nematodes upregulate the expression of certain AMP genes and induce the melanization pathway in D. melanogaster larvae (Pena et al., 2015). More recently, we have found that infection with Heterorhabditis nematodes regulates the TGF-beta pathway in D. melanogaster adults and inactivation of certain TGF-beta ligands modulates the survival of flies to nematode infection and the persistence of the parasites in the mutant flies (Eleftherianos et al., 2016).
Here we used RNAseq analysis to investigate the transcriptomic profiles of D. melanogaster larvae responding to infection by S. carpocapsae symbiotic or axenic nematodes. Our goal was to identify the number and nature of D. melanogaster genes that are differentially regulated upon S. carpocapsae nematode infection. We have found that the S. carpocapsae nematode infection induces distinct types of D. melanogaster genes compared to infection by microbial pathogens or other nematode parasites. We have determined several genes with putative roles in the interaction between D. melanogaster and S. carpocapsae parasitic nematodes. These results set the scene for identifying the molecular determinants of the insect immune response to entomopathogenic nematodes.
MATERIALS AND METHODS
Fly stocks
Oregon R third instar larvae were used for the transcriptomic analysis. Flies were reared on instant Drosophila diet (Formula 4-24 Drosophila medium) supplemented with yeast
78 (Carolina Biological Supply, Burlington, NC), and maintained at 25°C and a 12:12 h light:dark photoperiodic cycle.
Nematodes
Steinernema carpocapsae entomopathogenic nematodes carrying their mutualistic bacteria Xenorhabdus nematophila were amplified in the larvae of the wax moth Galleria mellonella using the water trap technique (White, 1927). Axenic nematodes were cultured using the Oily-Agar plates protocol (Yadav et al., 2015). To confirm the absence of X. nematophila bacteria in these nematodes, a fresh pellet of IJs was collected, washed once with 1% bleach and five times with sterile distilled water, homogenized, and the lysate was spread on LB-Agar plates. Absence of bacterial growth after 24-48 h confirmed the axenicity status of S. carpocapsae nematodes. Nematodes with or without
X. nematophila bacteria were used 1-3 weeks after collection and nematode density was estimated in 10 µl of suspension.
Infection assay
Microtiter 96-well plates were used for carrying out nematode infections. The plates were prepared by adding 100 µl of 1.25% agarose to each well. Sterile distilled water (10 µl) containing 100 nematodes was pipetted into the wells and an individual D. melanogaster larva was transferred to each well. The plate was covered with a Masterclear real-time
PCR film (Eppendorf, USA) and holes were pierced for ventilation. Treatment with sterile water served as control.
79
RNA isolation
Four larvae per treatment were collected at 6 h and 24 h post-infection. Total RNA was extracted using the PrepEase RNA spin kit (Affymetrix, Santa Clara, CA) following the manufacturer’s instructions. Total RNA was eluted in 40 µl of nuclease-free water and
RNA concentration was measured using a NanoDrop Microvolume Spectrophotometer
(Thermo Scientific, Waltham, MA). RNA integrity and quality were estimated using a
Bioanalyzer (Agilent Technologies, Santa Clara, CA).
Library Preparation and RNA sequencing
Separate libraries for the three experimental conditions (larvae infected with S. carpocapsae symbiotic or axenic as well as uninfected water controls) belonging to three independent experiments were prepared with the TruSeq RNA Sample Prep kit (Illumina,
San Diego, CA) per manufacturer’s protocol. Adapters containing seven nucleotide indexes were ligated to the double-stranded cDNA. The DNA was purified between enzymatic reactions and the size selection of the library was performed with AMPure XT beads (Beckman Coulter Genomics, Danvers, MA).
Libraries were assessed for concentration and fragment size using the DNA High
Sensitivity Assay on the LabChip GX (Perkin Elmer, Waltham, MA). The library concentrations were also assessed by qPCR using the KAPA Library Quantification Kit
(Complete, Universal) (Kapa Biosystems, Woburn, MA). The libraries were pooled and sequenced on a 100PE Illumina HiSeq 2500 run (Illumina, San Diego, CA).
80
Alignment reads and coverage analysis
The reads obtained from the sequencing platforms were fed into the TopHat read alignment tool to be aligned to the Drosophila melanogaster genomic reference sequence for each of the sequencing datasets. The reference genomic sequences were downloaded from the Ensembl project website (useast.ensembl.org). The TopHat alignment tool developed at the University of Maryland Center for Bioinformatics and Computational
Biology was used to align the raw sequencing reads. TopHat v1.4 is a fast splice junction mapper for RNA-Seq reads (Trapnell et al., 2009). It aligns RNA-Seq reads to the reference genome using the ultra high-throughput short read aligner Bowtie, and then analyzes the mapping results to identify splice junctions between exons. The output from
TopHat was obtained as BAM format files that consist of information on where the individual reads align within the reference genome and the splicing information of that read. In the alignment phase, we allowed up to two mismatches per 25 bp segment and removed reads that aligned to more than 20 genomic locations.
Differential gene expression analysis
The Tophat alignments were then used to generate read counts for each gene in the reference genome annotation using HTSeq (Anders, 2015). The counts generated by
HTSeq were subsequently used to generate the differential expression results using the R package DESeq.
81
Transcript analysis using CUFFLINKS
Transcript abundances and splice variant identification for each sample was done using
Cufflinks version 1.3 using the BAM alignment files obtained from TopHat (Ghosh &
Chan, 2016).
Differential Transcript analysis using Cuffdiff
The BAM files from TopHat and the gtf files generated by cufflinks were used to identify differentially expressed transcripts using CuffDiff (Trapnell et al., 2012). The results were filtered by FDR of less than .05, a FPKM value of greater than 10 and a fold change of +/- 2.
Gene Ontology (GO)
The Database for Annotation, Visualization and Integrated Discovery (Derer et al.)
(Huang da et al., 2009a, 2009b) web service was used for GO analysis using the list of differentially expressed genes. The p-value cut-off to determine enriched pathways was
0.1
Quantitative real-time RT-PCR validation of genes
82 To validate differentially expressed genes, we selected seven candidate genes based on significant fold differences and analyzed their mRNA levels using qRT-PCR. Four larvae from each treatment were collected at 6 and 24 h after infection and total RNA was extracted using the PrepEase RNA spin kit (Affymetrix) following the manufacturer’s instructions. RNA concentration was measured using a Nanodrop (Thermo Scientific) and samples were normalized to 350 µg. Complementary DNA (cDNA) was synthesized using the High Capacity cDNA reverse transcription kit (Applied Biosystems) on a
C1000 Thermal Cycler (Bio-Rad). cDNA samples were diluted 1:10 in nuclease-free water and 1 µl was used as a template for qRT-PCR experiments using the
SsoAdvanced™ Universal SYBR® Green Supermix (Bio-Rad). All experiments were carried out on a CFX96TM Real-Time System (Bio-Rad). Primers (Table 1) for individual genes were designed using primer blast (NCBI) and annealing temperatures for each primer pair were estimated using a gradient PCR. All primers produced a single amplicon, and this was confirmed by both melting curve analysis and by visualizing the
PCR product on the gel. Samples were run as technical duplicates and a total of three biological replicates were used for each treatment and time-point. The cycling conditions included 95°C for 2 mins, 40 cycles of 95°C for 15 s and an annealing step for 30 s. The melting curve analysis consisted of an initial denaturation step at 95°C for 15 s, followed by an incremental temperature gradient from 65°C to 95°C for 15 s at each temperature, with a ramp of 20 min from the lowest to the highest temperature. For each sample, the amount of mRNA detected was normalized to mRNA values of the housekeeping gene
RpL32 (ribosomal protein encoding gene). The relative level of a given gene is represented as a ratio of 2^CT(RpL32)/ 2^CT(Gene).
83
Statistical Analysis
Results from qRT-PCR tests are represented as means and standard deviations of relative values from three biological replicates. Data were analyzed using a one-way analysis of variance with a Tukey post-hoc test for multiple comparisons (GraphPad Prism 7).
RESULTS
Steinernema carpocapsae nematodes induce a large number of genes in D. melanogaster larvae
We infected D. melanogaster wild-type Oregon larvae with 100 S. carpocapsae symbiotic or axenic IJs and generated the transcriptomic profile of larvae infected at an early (6 h) and a late (24 h) time-point. Gene induction in infected larvae was relative to gene expression levels in uninfected larvae. The number of sequence reads mapped to an average of 74% of the D. melanogaster genome (Fig. 1A). We found that at 6 h, symbiotic nematodes induced the expression of 170 genes and axenic nematodes induced
109 genes. We also found that at 24 h, the number of genes induced by symbiotic nematodes increased slightly to 183 and those induced by axenic nematodes decreased to
103 (Fig. 1B). We also found that a large number of isoforms was differentially regulated upon infection with symbiotic nematodes compared to axenic nematodes. Symbiotic
84 nematode infections up regulated 23 isoforms at 6 h and 45 isoforms at 24 h, while only zero and five isoforms were down regulated at 6 and 24 h, respectively. In contrast, the number of isoforms induced by axenic nematodes was substantially lower. We found that infection with axenic nematodes up regulated only two isoforms at 6 h and seven isoforms at 24 h, and down regulated two and 10 isoforms at 24 h, respectively (Fig. 1C).
These results depict a summary of the numbers of genes induced in D. melanogaster larvae responding to S. carpocapsae nematode infection.
Steinernema carpocapsae nematodes modulate the induction of similar or different genes in D. melanogaster
We first investigated the number of common and distinct genes that are differentially regulated in D. melanogaster larvae upon infection with S. carpocapsae nematodes carrying or lacking X. nematophila bacteria. Upon infection with symbiotic nematodes, we found 121 and 127 up regulated genes at 6 and 24 h, respectively. We also found 8 and 23 down regulated genes at 6 h and 24 h post infection, respectively. Interestingly, between the two time points, there were 36 shared genes, 31 of which were up regulated at 6 and 24 h, one gene (CG2229) was down regulated at 6 h and up regulated at 24 h, two genes (CR32658 and CG31091) were down regulated at both time points, and two genes (CG42500 and CG3763) were up regulated at 6 h but down regulated at 24 h (Fig.
2A). We also found that at 6 h post infection with axenic nematodes, 23 genes were up regulated and 54 were down regulated at 6 h compared to 67 up regulated and 11 down regulated genes at 24 h. Also, among the commonly regulated genes by axenic nematodes, 21 genes were up regulated and only two genes (CG9070 and CG44956)
85 were down regulated at 6 h and up regulated at 24 h, whereas four genes (CG2559,
CG6806, CG3292 and CR32658) were down regulated at each time point (Fig. 2B).
These results demonstrate the number of genes differentially expressed in Drosophila larvae upon Steinernema symbiotic or axenic nematode infection.
To identify changes in the number and types of genes that were differentially regulated by symbiotic and axenic nematodes, we compared the D. melanogaster genes that were induced early and late upon infection by the two types of nematodes.
Interestingly, at 6 h we found 142 up regulated and nine down regulated genes in symbiotic nematode infections versus 40 up regulated and 50 down regulated genes in axenic nematode infections. Also, of the common genes at 6 h, four genes (CG18444,
CG16772, CG16844 and CG33337) were up regulated and two genes (CG32658 and
CG32071) were down regulated by both symbiotic and axenic nematodes, and eight genes (CG11650, CG3440, CG8502, CG7342, CG11089, CG7592, CG10078 and
CG42500) were up regulated by symbiotic nematodes and down regulated by axenic nematodes (Fig. 2C). We then compared the number of induced genes between symbiotic and axenic nematode infections at 24 h and found that although the number of up regulated genes was lower than those at the 6 h time point, the number of commonly regulated genes between the two types of nematode infections was higher at 24 h.
Symbiotic nematode infections up regulated 105 genes and down regulated 23 genes. In contrast, 37 genes were up regulated and 10 genes (such as CG3292, CG2736 and
CG8745) were down regulated upon axenic nematode infections. Among the commonly regulated genes at 24 h, both types of nematodes up regulated 53 genes, four genes
(CG2559, CG4181, CG10513 and CR32658) were down regulated by either axenic or
86 symbiotic nematodes, and only one gene (CG6271) was up regulated by symbiotic nematodes and down regulated by axenic nematodes (Fig. 2D). These results indicate that
S. carpocapsae axenic and symbiotic nematodes regulate a large variety of genes at early and late times post infection of D. melanogaster larvae.
Steinernema carpocapsae infection regulates several molecular pathways and biological activities in D. melanogaster
We conducted the Gene Ontology (GO) analysis using the DAVID database to identify the molecular pathways and biological activities that are involved in the D. melanogaster larval response to infection by S. carpocapsae nematodes (Fig. 3). We found that infection of D. melanogaster larvae with symbiotic or axenic nematodes elicited the enrichment of several specific and overlapping categories of genes at each time-point post infection. For example, at 6 h we found that infection with symbiotic nematodes induced the enrichment of genes involved in the humoral immune response and serine- type endopeptidase activity (Fig. 3A and Supplementary Fig.1A), whereas infection with axenic nematodes up regulated genes with transmembrane transporter activity and down regulated genes in the chitin based cuticle pathway (Fig. 3B and Supplementary Fig.1B).
We also found that infection with symbiotic nematodes at 24 h up regulated genes associated with immune system, polysaccharide binding, aminoglycan metabolic process, and cell wall macromolecule catabolic process. Down regulated genes were associated with carbohydrate metabolic process and apoptosis signaling (Fig. 3C and Supplementary
Fig.1C). Infection with axenic nematodes at 24 h up regulated genes that function in similar pathways as well as genes related to neuroactive ligand-receptor interaction. The downregulated genes were mainly associated with larval serum protein complex, lipid
87 particle and nutrient reservoir activity (Fig. 3D and Supplementary Fig.1D). We next compared the gene set enrichment between symbiotic and axenic nematode infections at
6 and 24 h time points. At 6 h post infection, we observed that a large number of up regulated genes function in immune defense responses and the downregulated genes encoded metalloproteases and hydrolases (Fig. 3E and Supplementary Fig.1E).
Conversely, at 24 h, we found that all gene categories were upregulated and with the exception of lipid metabolic processes, the rest of the upregulated genes belonged to a variety of immune response categories, such as immune system process and antimicrobial humoral response (Fig. 3F and Supplementary Fig.1F). Thus, the pathway analysis revealed that infection with symbiotic nematodes induced genes related to immune functions in D. melanogaster, whereas infection with axenic nematodes induced genes belonging to a variety of categories ranging from polysaccharide binding to chitin metabolic process. These results provide novel insights into the molecular processes that take place in D. melanogaster larvae upon infection by entomopathogenic nematodes carrying or lacking their associated bacteria and contribute toward a better understanding of the molecular events that take place in the host during nematode infection.
Steinernema carpocapsae nematodes affect key immune and developmental processes in D. melanogaster larvae
To identify which immune and developmental genes were regulated upon infection by S. carpocapsae nematodes, we generated a heat-map to illustrate the differential gene transcription levels for both types of nematode infections for both time points (Fig. 4A,
B). For the heat map with the immunity-related genes, we included genes from the four
88 known immune signaling pathways (Immune Deficiency, IMD; Janus Kinase and Signal
Transducer and Activator of Transcription, JAK/STAT; cJun- N-terminal Kinase, JNK; and Toll), genes involved in cellular immune responses and hematopoiesis, immune induced molecules (IIM), genes with immune receptor activity, and genes with general immune functions, which also included genes with putative immune roles (Fig. 4A).
In the Toll pathway (Valanne et al., 2011), the GNBP3 gene was down regulated in larvae infected by axenic nematodes at 6 h compared to the uninfected control larvae, and it was up regulated at 24 h post infection in larvae infected by either symbiotic or axenic nematodes. We found that at 6 h, the Toll pathway protein, Serpin-27A, was up regulated by symbiotic nematodes and down regulated by axenic nematodes. Conversely, this serpin gene was up regulated at 24 h by either symbiotic or axenic nematodes. The
Toll immune regulated gene Fondue was down regulated by both types of nematodes at 6 h, but its expression increased at 24 h post infection. The AMP-coding gene Drosomycin
(Zhang & Zhu, 2009) was up regulated at both 6 and 24 h in larvae infected by symbiotic nematodes, but it was down regulated at 6 h and up regulated at 24 h by axenic nematodes.
In the IMD pathway (Kaneko et al., 2005), certain genes encoding recognition proteins were induced at different levels by either symbiotic or axenic S. carpocapsae.
We found that PGRP-SC1a/b (Garver et al., 2006) was slightly down regulated at 6 h in larvae responding to symbiotic or axenic nematodes, it was up regulated at 24 h in response to axenic nematodes, and showed no changes in expression with symbiotic nematodes. In contrast, PGRP-SC2 (Bischoff et al., 2006) and PGRP-LB (Zaidman-
Remy et al., 2006) were strongly induced by both symbiotic and axenic nematodes at 24
89 h post infection, but showed little to no change at the 6 h time point. We also found that kenny (Silverman et al., 2000) and the IMD pathway transcription factor, Relish (Stoven et al., 2000), were highly induced in larvae infected for 24 h by either type of nematode.
The IMD controlled AMP encoding genes Attacin-A, Attacin-B and Attacin-C were strongly up regulated by symbiotic nematodes at both 6 and 24 h and only slightly up regulated at 24 h by axenic nematodes.
We also found that Jak/Stat and JNK pathway genes were not induced as strongly as genes in the Toll and IMD signaling pathways. We found that Turandot-A (Tot-A)
(Ekengren & Hultmark, 2001) expression increased by symbiotic nematodes at 6 h and decreased at 24 h compared to the uninfected control. However, axenic nematode infections caused the down regulation of Tot-A at 6 h and its up regulation at 24 h. The
JNK pathway gene, puckered (McEwen & Peifer, 2005), was significantly up regulated at
24 h in axenic nematode infected larvae, but it was slightly up regulated by symbiotic nematodes. Interestingly, the expression of genes with receptor activity, such as peste
(Nakanishi, 2007), scavenger receptor class V, type 1 and Toll 5 (Tauszig et al., 2000) was increased highly by axenic nematodes at 24 h with lower levels of expression in response to symbiotic nematodes. In contrast, the expression of genes such as Lapsyn,
Gp150, diuretic hormone 31 and CG5096 was significantly increased only by axenic nematode infections at 24 h. We also found that genes involved in cellular immune responses were up regulated by both symbiotic and axenic nematodes. The zinc finger protein jing and singed were strongly induced by symbiotic nematodes at 6 h, whereas serine protease 7 was strongly induced at 24 h. We further noticed that other genes such as nitric oxide synthase and ATP dependent RNA helicase p62 were up regulated by
90 axenic nematodes at 24 h, whereas hemese (Kurucz et al., 2003) was up regulated at both
6 and 24 h. Interestingly, Tep1 and Tep2 (Bou Aoun et al., 2011) were strongly induced by symbiotic nematodes at both 6 and 24 h. These results identify some of the known immune genes induced in the D. melanogaster larvae in response to infection with S. carpocapsae symbiotic or axenic nematodes.
To identify the D. melanogaster differential regulated developmental genes due to
S. carpocapsae infection, we generated a heat map to illustrate the differential gene expression levels in larvae infected by symbiotic or axenic nematodes at each time point post infection (Fig. 4B). We included genes belonging to the imaginal disc growth factors
(Idgf) encoding gene family, and pathways functioning in multicellular organism development, organ development and Notch and Wnt signaling. In the Idgf category, axenic nematodes strongly induced the expression of tenectin at 6 h, and lamina ancestor and bursicon genes at 24 h. The gene E(spl) region transcript m2 of Notch signaling was up regulated by symbiotic nematodes at 6 h, and lethal malignant blood neoplasm was up regulated by both symbiotic and axenic nematodes at 24 h. We also found differential expression of genes regulating organ development functions. Matrix metalloproteinase 1 was induced by symbiotic nematodes at 6 h, and the gene lonely heart and LDLa domain containing chitin binding protein-1 was induced by axenic nematodes at 24 h.
Interestingly, pericardin and scab were strongly induced by both axenic and symbiotic nematodes at 24 h; however, induction levels by axenic nematodes were higher compared to those by symbiotic nematodes, with the exception of CG17278 (Wnt signaling), which was highly induced by symbiotic nematodes at 24 h. We also estimated the expression of other developmental genes and found that the gene punch was up regulated by symbiotic
91 nematodes at 6 h and down regulated by axenic nematodes at 24 h. Yellow-F and glial cells missing was induced only by symbiotic nematodes at 6 h, and the tissue inhibitor of metalloproteases was induced by axenic nematodes at 24 h. Several other genes were induced by both symbiotic and axenic nematodes at 24 h. For example, viking, ejaculatory bulb III, collagen type IV and SPARC were all induced at higher levels by axenic nematodes, while symbiotic nematodes were responsible for the stronger induction of CG7714. Thus these results identify the D. melanogaster genes with known functions in the immune or developmental processes induced in response to S. carpocapsae nematode infections.
Steinernema carpocapsae infection affects D. melanogaster genes conserved in M. sexta and humans
To investigate whether the D. melanogaster genes induced by S. carpocapsae infection had known functions in other organisms, we selected the top 55 most differentially expressed genes in nematode-infected larvae. We included results based on the UniProt database for a natural host of entomopathogenic nematodes, the tobacco hornworm
Manduca sexta, and human (Homo sapiens). We used Venn Diagrams to depict the conservation of genes based on shared or distinct protein domains in these three organisms. We observed that of the 55 D. melanogaster protein-coding genes, 20 were exclusive to D. melanogaster, 1 to M. sexta and 9 to H. sapiens. We also found that 17 of these 55 protein-coding genes in D. melanogaster were also present in M. sexta and they belong to proteins with scorpion toxin-like activity, chitin binding properties, hemocyanin, attacin, and pheromone binding domains. However, the five domains shared
92 between D. melanogaster and H. sapiens belong to lipase, pyridoxal phosphate- dependent decarboxylase, EGF-like, tetraspanin and heat shock proteins. Interestingly, the 13 common elements among all three organisms possess certain domains, such as amidase, kazal, serpin or kunitz. The results from this analysis showed that certain D. melanogaster genes induced upon infection with S. carpocapsae nematodes have orthologs in two other organisms, M. sexta (insect host) and H. sapiens (humans) indicating conservation in their potential roles against parasitic nematodes.
DISCUSSION
Here we present the transcriptional profile of D. melanogaster larvae infected by the potent nematode parasite S. carpocapsae containing or lacking its mutualistic X. nematophila bacteria. We report the identification of several types of D. melanogaster genes that are differentially regulated in the larval stage during interaction with either type of nematode. Initial characterization of the D. melanogaster transcriptome reveals that the number and nature of genes induced upon infection by axenic or symbiotic nematodes is substantially different. This suggests that S. carpocapsae nematodes in the absence or presence of their associated bacteria elicit different types of immune reactions because they employ distinct strategies to infect insects and interfere with their immune system. We have also identified specific genes that are significantly up or down regulated during nematode infection. We have found that these genes have conserved functions in the natural host of the entomopathogenic nematodes S. carpocapsae, larvae of the lepidopteran M. sexta, as well as in humans and therefore they might possess conserved anti-nematode properties.
93 A recent transcriptome analysis on D. melanogaster adult flies infected by H. bacteriophora symbiotic or axenic nematodes, or their associated P. luminescens bacteria alone, has identified a wide variety of genes that are differentially regulated in response to the pathogens. These genes are mainly related to stress response, lipid homeostasis, metabolic processes, and neuronal functions (Castillo et al., 2015). Interestingly, some of these genes were reported to form factors with potential roles in host anti-nematode and anti-bacterial immune responses. Also, a previous transcriptome study on D. melanogaster larvae infected by symbiotic H. bacteriophora nematodes only, show that genes encoding complement factors as well as recognition and extracellular matrix proteins are expressed at high levels (Arefin et al., 2014). Here we include infections of
D. melanogaster larvae with axenic S. carpocapsae nematodes to identify the D. melanogaster genes that are differentially regulated in response to the nematodes without the input of their mutualistic X. nematophila bacteria. We show that most D. melanogaster genes and isoforms are differentially regulated in response to symbiotic nematodes compared to axenic worms, suggesting the additional contribution of mutualistic X. nematophila in the interaction with the insect immune system during infection with the nematode-bacteria complexes.
Our analysis shows that a subset of D. melanogaster induced genes is common between the two types of nematode infections compared to a larger number of genes that are distinct either to axenic or symbiotic S. carpocapsae infection. In addition, early in the infection process, axenic nematodes down regulate a larger number of genes compared to those down regulated by symbiotic nematodes, but as the infection progresses the number of down regulated genes increases in larvae infected by symbiotic
94 worms. This suggests that the insect immune system can be compromised by entomopathogenic nematodes devoid of their associated bacteria, especially during the initial stages of infection.
Upon infection with axenic or symbiotic S. carpocapsae, we found strong induction of several Heat Shock Protein (hsp) coding genes, which can be attributed to the insect response to stress conditions during nematode penetration, invasion and migration in the insect, which is accompanied by severe tissue damage (Feder &
Hofmann, 1999; Sorensen et al., 2005). Heterorhabditis bacteriophora nematodes use the specialized buccal protruding tooth to penetrate through the D. melanogaster larval cuticle and the gut epithelium, thus causing extensive wounding to those tissues (Arefin et al., 2014; Ciche et al., 2008). Based on our findings, certain hsp genes, such as Hsp23 and Hsp27, are strongly up regulated by S. carpocapsae symbiotic nematodes and show little to no change in response to axenic nematodes at 6 h post-infection. In contrast, at 24 h post infection, these genes are up regulated in response to axenic nematodes only.
These results indicate that both the nematode-bacteria complexes as well as the nematodes alone are capable of causing physical damage to the larvae thereby leading to the strong induction of hsp genes. We further observed a strong induction of TotC, an immune and stress response gene of the Turandot family, upon infection by either symbiotic or axenic nematodes (Ekengren & Hultmark, 2001). Similarly, Hsp and TotC are also detected in D. melanogaster adult flies upon infection by either symbiotic or axenic H. bacteriophora nematodes as well as P. luminescens bacteria only (Castillo et al., 2015). These results confirm that entomopathogenic nematode infection in D.
95 melanogaster adult flies as well as larvae leads to the potent induction of several stress factors.
Classifying the genes that are induced by either type of S. carpocapsae nematode reveals that symbiotic nematodes primarily induce genes with immune related functions whereas axenic nematodes induce genes encoding peptidases, chitin binding and structural components of the larval cuticle. A particular class of genes that is induced by axenic nematodes can be grouped into the category ‘structural constituent of the insect peritrophic membrane’. This membrane consists of chitin and peritrophin-like proteins that line the insect gut and modulate gut immune responses in the host against bacterial infections (Buchon, Broderick, et al., 2009; Hegedus et al., 2009; Kuraishi et al., 2011;
Lehane, 1997). The strong induction of genes that are mainly expressed in the peritrophic membrane (peritrophin-15a, -15b) upon axenic nematode infection suggests that they may be involved in the insect response against nematodes free from X. nematophila bacteria. This induction could be a result of damage of the gut epithelium caused by the movement of nematode in the insect host.
In the D. melanogaster gut, the Imd pathway is responsible for the induction of anti-microbial peptides in response to microbial infections (Myllymaki et al., 2014). The
AMP induction is the result of the interaction between peptidoglycan (PGN)-recognition proteins (PGRPs) and the pathogen specific PGN, thereby initiating the intracellular molecular cascade (Casanova-Torres & Goodrich-Blair, 2013). Both PGRP-LB and
PGRP-SC2 are among the five PGRPs that can process DAP-type PGN (Dziarski &
Gupta, 2006), and their induction suggests that S. carpocapsae axenic nematodes or certain molecules that they produce are recognized by these IMD pathway receptors, but
96 apparently fails to induce certain AMP effectors or induces Attacin-A, -B or –C at low levels. Conversely, these AMPs are strongly induced in response to symbiotic nematodes, suggesting that the detection of nematode-bacteria complexes was likely due to the identification of X. nematophila by the insect PGRPs.
Contrary to the number and induction level of genes related to humoral immune responses, we found very few genes with known function in cellular immune processes that are differentially regulated by S. carpocapsae nematode infection. This suggests that the D. melanogaster cellular immune response may not be crucial against infection by S. carpocapsae or that the molecules secreted by these nematodes may be effective in suppressing or preventing hemocyte action that regulate insect cellular immune processes
(Brivio MF, 2005). Interestingly, G. mellonella hemocytes respond to infection by H. bacteriophora, but not to S. carpocapsae (Ebrahimi et al., 2011), suggesting that these nematodes are able to evade the insect cellular immune response. The encapsulation response in insects is facilitated by the ability of hemocytes to spread and adhere to the nematode surface (Stanley et al., 2012). Steinernema carpocapsae nematodes produce certain proteases and other factors that impair clot formation thereby evading the insect melanization response and eicosanoid biosynthesis (Stanley et al., 2012; Toubarro et al.,
2013). Eicosanoids and their related lipids participate in the immune response of D. melanogaster larvae in response to infection by H. bacteriophora nematodes (Hyrsl et al., 2011). Here we observe the induction of certain genes belonging to the melanization response in larvae infected by S. carpocapsae symbiotic or axenic nematodes. The induction of ppo1, ppo2 and pro-PO A1 in response to both symbiotic and axenic nematodes is in contrast to the up regulation of genes such as black cells encoding
97 prophenoloxidase (Gajewski et al., 2007), and phenoloxidase subunit A3 in response to axenic nematodes only. Taken together, it can be argued that the wound healing and the clotting response in the D. melanogaster larvae upon infection by S. carpocapsae nematodes may not be entirely dependent on the action of the ppo genes and might involve the contributions of other genes that have not yet been identified or fully characterized.
Categorizing the strongly induced genes into those with immune or developmental related function reveals the nature of genes from each category. Certain genes previously reported to function in developmental processes are highly induced in
D. melanogaster larvae in response to infection by S. carpocapsae axenic or symbiotic nematodes. One of those genes is pericardin (prc), a mammalian collagen IV homolog
(Chartier et al., 2002). Interestingly, at 24 h post-infection, there is a strong up regulation of prc in response to symbiotic nematode infections and an even stronger induction in response to axenic nematodes. The function of prc in the D. melanogaster organ development is to regulate heart morphogenesis, and to maintain cardiac integrity
(Chartier et al., 2002; Zaffran et al., 1995). We also find strong induction of the gene lonely heart (loh), which is responsible for the recruitment of PRC to the extracellular matrices of different tissues to regulate the assembly of the matrices. The normal functioning of both prc and loh is crucial for cellular behavior and proper functioning of the organs in D. melanogaster (Drechsler et al., 2013). We find that loh exhibits stronger induction levels in response to S. carpocapsae axenic nematodes compared to symbiotic nematodes at the late stage of infection. We also find strong induction of the collagen homologs Collagen type IV alpha 1 and Viking at a later time point post infection by both
98 symbiotic and axenic nematodes. Viking as well as the basement membrane protein glutactin function together in would healing in the D. melanogaster larvae infected by H. bacteriophora nematodes (Arefin et al., 2014). Therefore it can be argued that prc and loh, may also be involved in wound healing or clotting responses in D. melanogaster larvae against infection by S. carpocapsae nematodes.
Although S. carpocapsae can infect a wide range of insect species naturally
(Lacey et al., 2015), D. melanogaster does not act as host to this nematode species.
Examination of the transcriptional regulation of the immune response of certain
Drosophila species to natural parasites and microbial pathogens suggests that Drosophila adult flies and larvae can trigger different immune genes and pathways against natural viral pathogens (Carpenter et al., 2009; Habayeb et al., 2006; Kemp et al., 2013), bacterial pathogens (Vodovar et al., 2005), and endoparasitoid wasps (Wertheim et al.,
2005). The Drosophila immune response can vary from activating PGRPs and AMPs through NF-κB signaling pathways to reactions that are restricted to specific types of natural pathogens (Keebaugh & Schlenke, 2014). Here we show that S. carpocapsae nematodes interfere with the expression of genes with known immune roles in D. melanogaster, but also with genes with unexplored function in the fly immune system.
These findings imply that D. melanogaster has particular mechanisms to respond to S. carpocapsae nematodes, likely due to the lack of host-parasite coadaptation and coevolution.
In conclusion, we show here that S. carpocapsae nematodes are able to trigger the
D. melanogaster larval immune system even in the absence of their X. nematophila mutualistic bacteria. We show that D. melanogaster larvae activate several different types
99 of genes in response to S. carpocapsae nematode infection. These include genes with known immune function, genes involved in developmental processes as well as genes with unknown mechanistic roles, especially in the interaction of the insect immune system with entomopathogenic nematodes. Our transcriptome study expanded our understanding of the nature of insect genes that are induced in response to potent nematode parasites. Infection of D. melanogaster larvae by axenic or symbiotic S. carpocapsae nematodes reveals the induction of unique genes, which are not shared with other infection models. Similar transcriptome studies will lay the foundation for testing the candidate genes through functional studies that will promote our understanding of the molecules that modulate the interaction between insects and parasitic nematodes.
FIGURE LEGENDS
Figure 1 Infection of wild-type Drosophila melanogaster third instar larvae with
Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes induces a large number of transcripts. (A) Transcriptome summary (total number of reads, total number of mapped reads and percentage reads mapped to the D. melanogaster genome) from larvae infected with S. carpocapsae symbiotic (blue) or axenic nematodes at 6 and 24 h post infection. (B) Number of differentially expressed transcripts from larvae infected by
S. carpocapsae symbiotic or axenic nematodes at 6 and 24 h post infection. (C)
CUFFLINKs analysis of the upregulated (blue) or downregulated (Herren et al.) isoforms in D. melanogaster larvae infected with S. carpocapsae symbiotic or axenic nematodes at
6 and 24 h post infection.
100
Figure 2 Infection with S. carpocapsae symbiotic or axenic nematodes induces distinct and shared transcriptomic profiles in D. melanogaster larvae. Venn diagrams showing the number of D. melanogaster differentially expressed genes upon infection with S. carpocapsae (A) symbiotic nematodes at 6 and 24 h, (B) axenic nematodes at 6 and 24 h,
(C) symbiotic and axenic nematodes at 6 h and (D) symbiotic and axenic nematodes at 24 h. Expression patterns are indicated (up/up: gene up regulation at both 6 and 24 h, up/down: gene up regulation at 6 h and down regulation at 24 h, down up: gene down regulation at 6 h and up regulation at 24 h, down/down: gene down regulation at both 6 and 24 h).
Figure 3 Infection with S. carpocapsae symbiotic or axenic nematodes induces diverse physiological responses and biological activities in D. melanogaster larvae. This is characterized by the enrichment of pathway specific genes based on their molecular and biological functions using the DAVID classification database. Representative categorization of genes in larvae infected with S. carpocapsae (A) symbiotic nematodes at 6 h, (B) symbiotic nematodes at 24 h, (C) axenic nematodes at 6 h, (D) axenic nematodes at 24 h, and comparison of axenic and symbiotic infections at (E) 6 h and (F)
24 h post infection.
Figure 4 Infection with S. carpocapsae symbiotic or axenic nematodes differentially regulates the transcription of a variety of immune and developmental genes in D. melanogaster larvae. Genes selected from the Gene Ontology (GO) analysis have a positive expression level as an indication of their up regulation upon infection with the nematode parasites. The selected genes are categorized into (A) Immune Genes and they
101 are further grouped into genes with cellular immune functions, genes encoding immune induced molecules, genes with receptor activity or genes that are regulated by IMD, Toll,
JAK /STAT, JNK pathways, and (B) Developmental Genes and they are further grouped into genes that have functions in multicellular organism reproduction and organ development, genes that are regulated by the Notch or Wnt signaling pathways, genes that belong to the Imaginal Disc Growth Factors or genes with other functions in development.
Figure 5 (A) Orthologs of the top 55 differentially transcribed Drosophila melanogaster genes (Dm) in Manduca sexta (Ms) and Homo sapiens (Hs). D. melanogaster genes were selected from the Gene Ontology (GO) analysis that were either up or down regulated upon infection by S. carpocapsae axenic or symbiotic nematodes and the protein domains were selected based on the UniProt I.Ds. (B) Table showing the protein domains shared between Dm and Ms, and those that are shared between Dm and Hs.
102
A
200 B 180 160 140 120 100 80 60
40 Number ofgenes Number 20 0 Sym Ax Sym Ax
50 C 40
30
20
10
0
Number of isoforms of Number -10 Sym -20 Ax Sym Ax
103 Figure 1.
Figure 2.
104
Fold Enrichment
10
20
30 0 A humoral immune response response to other organism defense response to bacterium antimicrobial humoral response antibacterial humoral response response to biotic stimulus extracellular region response to bacterium immune response defense response
Fold Enrichm
-2 ent
20
40
60
0 0 B extracellular region defense response defense response to bacterium response to bacterium antimicrobial humoral response antibacterial humoral response immune response humoral immune response cytoplasm cytoplasmic part
Figure 3.
105 °
Fold En -1 -1 richment 10 -5 C 5 0 0 5 active transmembrane transporter activity transmembrane transporter activity 2° active transmembrane transporter activity transporter activity extracellular region structural constituent of chitin-based cuticle structural constituent of cuticle structural constituent of chitin-based larval cuticle structural molecule activity extracellular region
Fold Enrich
-1 ment
10
20
30
0 0
D extracellular region chitin binding chitin metabolic process pattern binding polysaccharide binding aminoglycan metabolic process polysaccharide metabolic process larval serum protein complex lipid particle nutrient reservoir activity
Figure 3.
106
Fold Enrichme -2
-1 nt
10
20
30
0
0 0 E response to other organism defense response to bacterium response to biotic stimulus antibacterial humoral response 2° active transmembrane transporter activity active transmembrane transporter activity metallopeptidase activity transmembrane transporter activity peptidase activity, acting on L-amino acid peptides
Fold Enrichment
100
150
50 0 F defense response antibacterial humoral response immune response defense response to bacterium response to bacterium immune system process innate immune response antimicrobial humoral response humoral immune response response to other organism
Figure 3.
107
Figure 4.
108
Figure 5.
109 Table 1: List of primers used for qRT-PCR
Gene Accession Comments Sequences Name no. TotC CG31508 Stress induced Forward 5’-ACGTTGTCCCCTGAACAAAGG-3’ humoral factor Reverse 5’-TCCGACGTACTTGGTCTTTCG-3’ Unknown CG31698 Unknown Forward 5’-CCAAACTTCCACCTCGGGAT-3’ function Reverse 5’-GATTCACGGGTTTGCTGTCG-3’
IM3 CG16844 Immune induced Forward 5’-TTGGGTCTGCTGGCTCTG-3’ molecule Reverse 5’-TTCAACTGGCATCCTTCATTC-3’
Unknown CG7248 Unknown Forward 5’-CAACACCTTCACCCACAGAAT-3’ function- Reverse 5’-TTCACGCACAAGTAGAACTCATT- contains a chitin 3’ binding domain
ImpE2 CG1934 Unknown Forward 5’-AAGCCCGTTGCCTTGATCC-3’ function Reverse 5’-CTACTGGTGGCTCCTTATCCT-3’
Sgs5 CG7596 Unknown Forward 5’-TCAGAGCCTGAAATTGAATCCG- function 3’ Reverse 5’-AAGAGCCCATTGGTAGTTCCT-3’ Unknown CG3906 Unknown Forward 5’-AGCCACATTACATTGAGGTGTC-3’ function- insect Reverse 5’-CGTGATCGGTTCTATTCGGATTG- allergen related 3’
RpL32 CG7939 Ribosomal Forward 5’-GATGACCATCCGCCCAGCA-3’ Protein L32 Reverse 5’-CGGACCGACAGCTGCTTGGC-3’
110 SI FIGURE LEGENDS
Figure S1 Infection by S. carpocapsae symbiotic and axenic nematodes induces the expression of diverse classes of genes in D. melanogaster larvae. Results are shown for
A. Symbiotic nematode infections at 6 h, B. Symbiotic nematode infections at 24 h, C.
Axenic nematode infections at 6 h, D. Axenic nematode infections at 24 h, and comparison between symbiotic and axenic nematode infections at E. 6 h, and F. 24 h.
Figure S2 Quantitative real time RT-PCR validation of seven D. melanogaster genes
(CG31508, CG31698, CG16844, CG7248, CG1934, CG7596 and CG3906) selected from the transcriptome dataset. Results are shown for seven of the D. melanogaster genes from larvae infected with A. S. carpocapsae symbiotic nematodes at 24 h and B. S. carpocapsae axenic nematodes at 24 h. The solid bars represent the transcript levels obtained from the RNAseq study and qRT-PCR results are shown as hollow bars. Larvae used for both RNAseq and qRT-PCR were subject to the same experimental conditions.
Error bars represent the mean of three experimental samples.
111
10 20 30 0 A humoral immune response antimicrobial humoral response extracellular region defense response innate immune response multicellular organism adhesion molting cycle, chitin-based cuticle molting cycle, protein-based cuticle imaginal disc eversion response to fungus instar larval/pupal development post-embryonic development death De novo purine biosynthesis serine-type endopeptidase activity
biological adhesion
-2
20
40
60
0 0 B extracellular region antimicrobial humoral response immune system process multi-organism process extracellular region part cytolysis molting cycle polysaccharide binding acyl groups transferase activity cell wall macromolecule catabolic process aminoglycan metabolic process phospholipase activity carbohydrate binding glycosyl bonds hydrolase activity carbohydrate metabolic process Apoptosis signaling pathway
Supplementary Figure 1.
112
Fold -1
-1 Enrichment
10
-5
5
0 0 C 5
active transmembrane transporter transmembrane transporter 2° active transmembrane transporter transporter activity extracellular region substrate transmembrane transporter substrate-specific transporter ion transmembrane transporter chitin-based cuticle structural constituent of cuticle chitin-based larval cuticle structural molecule activity extracellular region extracellular space
extracellular region part
-1
10
20
30
0 0 D extracellular region chitin binding chitin metabolic process pattern binding polysaccharide binding aminoglycan metabolic process polysaccharide metabolic process carbohydrate binding carbohydrate metabolic process amine metabolic process peritrophic membrane Neuroactive ligand-receptor extracellular region part acyltransferase activity transferase,not amino-acyl groups extracellular space acyl groups transferase activity antibacterial humoral response larval serum protein complex lipid particle nutrient reservoir activity
Supplementary Figure 1.
113
-4
-2
20
40
0 0 E 0 extracellular region response to bacterium structural constituent of cuticle multicellular organism adhesion molting cycle instar larval/pupal morphogenesis imaginal disc eversion cell death neuron fate commitment pigment metabolic process salivary gland histolysis cellular ketone metabolic process endopeptidase activity peptidase, L-amino acid peptides cation transmembrane transporter hydrolase activity extracellular region part
ion binding
100
200
300 0 F defense response antibacterial humoral response immune response defense response to bacterium response to bacterium immune system process innate immune response antimicrobial humoral response humoral immune response response to other organism response to stress response to biotic stimulus extracellular region multi-organism process defense response to Gram-neg bacterium extracellular space phospholipase A1 activity triacylglycerol lipase activity extracellular region part response to stimulus lipase activity carboxylesterase activity defense response to Gram-pos bacterium phospholipase activity hydrolase activity defense response to fungus response to fungus hydrolase activity, ester bonds catalytic activity lipid metabolic process
Supplementary Figure 1.
114
Supplementary Figure 2
115 *Chapter 4: The Imaginal Disc Growth Factors 2 and 3 regulate the Drosophila anti-
nematode immunity
SUMMARY
The Drosophila imaginal disc growth factors (IDGFs) induce the proliferation of imaginal disc cells and terminate cell proliferation at the end of larval development.
However, the participation of Idgf encoding genes in other physiological processes of
Drosophila including the immune response to infection is not fully understood. Here we show the contribution of Idgf2 and Idgf3 in the Drosophila response to infection with
Steinernema carpocapsae nematodes carrying or lacking their mutualistic Xenorhabdus nematophila bacteria (symbiotic or axenic nematodes, respectively). We find that Idgf2 and Idgf3 are upregulated in Drosophila larvae infected with symbiotic or axenic
Steinernema and inactivation of Idgf2 confers a survival advantage to Drosophila larvae against axenic nematodes. Inactivation of Idgf2 induces the Imd and Jak/Stat pathways, whereas inactivation of Idgf3 induces the Imd and Toll pathways. We also show that inactivation of the Imd pathway receptor PGRP-LE upregulates Idgf2 against
Steinernema nematode infection. Finally, we demonstrate that inactivation of Idgf3 induces the recruitment of larval hemocytes in response to Steinernema nematodes. Our results indicate that Idgf2 and Idgf3 might be involved in different yet crucial immune functions in the Drosophila anti-nematode immune response. Similar findings will promote the development of new targets for species-specific pest control strategies.
* This chapter in under review at Parasite Immunology
116 INTRODUCTION
Understanding the molecular mechanisms of insect response to nematode infection is a crucial step towards interpreting the anti-parasitic properties of the host innate immune system (Li et al., 2012). Entomopathogenic (or insect pathogenic) nematodes have emerged as excellent models for studying the molecular basis of nematode parasitism and elucidating the function of molecules that promote nematode persistence in the host
(Balasubramanian et al., 2012; Hao et al., 2010; Toubarro, Avila, Hao, et al., 2013;
Vadnal et al., 2017). A major advantage of entomopathogenic nematodes is that they are viable in the absence of their mutualistic bacteria (axenic nematodes) (Hallem et al.,
2007; Yadav et al., 2015); consequently, each partner in the mutualistic relationship can be separated and studied in isolation or in combination enabling host immune responses to be studied against each pathogen separately, and against the nematode-bacteria complex together (Castillo et al., 2015; Castillo et al., 2013; Hallem et al., 2007).
The entomopathogenic nematode Steinernema carpocapsae forms a mutualistic relationship with the Gram-negative bacteria Xenorhabdus nematophila (Martens et al.,
2003). Infective juveniles (IJs) gain entry into the insect through natural openings or by penetrating the cuticle (Arefin et al., 2014; Pena et al., 2015). Once inside the insect, the
IJ nematodes resume development and expel Xenorhabdus into the hemolymph (insect blood) where the bacteria begin to divide (Goodrich-Blair & Clarke, 2007). After 2-3 days of bacterial growth the insect succumbs to the infection with the concomitant conversion of the internal tissues into bacterial biomass, facilitated by a wide range of toxins, virulence factors and hydrolytic enzymes produced by the bacteria (Kumari et al.,
2014; Massaoud et al., 2010; Rodou et al., 2010). The developing nematodes feed on the
117 bacteria and nematode reproduction continues over 2-3 generations until the insect carcass is consumed, whereupon adult development is suppressed and the IJ stage accumulates (Goodrich-Blair, 2007). These non-feeding IJ containing their mutualistic
Xenorhabdus bacteria emerge into the soil to seek new hosts. Interestingly, Steinernema nematodes lacking their Xenorhabdus bacteria are still pathogenic to insects
(Eleftherianos, Joyce, et al., 2010; Pena et al., 2015; Yadav et al., 2015).
Recent work has demonstrated the power of using Drosophila for dissecting the molecular/genetic basis of insect immune response against entomopathogenic nematodes
(Arefin et al., 2014; Castillo et al., 2015; Castillo et al., 2013; Eleftherianos et al., 2016;
Kucerova et al., 2016; Pena et al., 2015; Yadav et al., 2017). Infection of Drosophila larvae with Heterorhabditis bacteriophora symbiotic nematodes results in the transcriptional activation of four antimicrobial peptide genes (Castillo et al. 2013). This response is specific to mutualistic Photorhabdus bacteria because axenic Heterorhabditis fails to upregulate antimicrobial peptide genes (Hallem et al., 2007). Heterorhabditis infection also upregulates several immune-related genes in adult flies, but injection of
Photorhabdus bacteria alone results in lower immune gene transcript levels in
Drosophila (Castillo et al,.2013). Inactivation of Drosophila transglutaminase, a conserved component of the clotting cascade in insects and humans, results in decreased aggregation of zymosan beads and increased sensitivity of larvae to Heterorhabditis infection (Wang et al., 2010). Two clotting factors (gp150 and fondue) were also shown to participate in the Drosophila anti-nematode immune response (Hyrsl et al., 2011), and more recently a homolog of thioester-containing complement protein 3, a basement membrane component (glutactin), a recognition protein (Gram Negative Binding Protein-
118 like 3) and several small peptides contribute to the control of Heterorhabditis infection in
Drosophila larvae (Arefin et al., 2014). In addition, Steinernema symbiotic nematodes upregulate the expression of certain antimicrobial peptide genes and induce the melanization pathway, the activation of which is suppressed by their mutualistic
Xenorhabdus bacteria (Pena et al., 2015). The Drosophila chitinase-like protein imaginal disc growth factor 3 (IDGF3) plays an immune-protective role against infections of larvae with Heterorhabditis nematodes (Kucerova et al., 2016).
Imaginal disc growth factors (IDGFs) are secreted glycoproteins mainly expressed in embryonic yolk and fat body insect cells. They are also abundantly secreted by various
Drosophila cell lines exhibiting macrophage properties (Kawamura et al., 1999;
Kirkpatrick et al., 1995), and they promote proliferation, polarization, and motility of
Drosophila cells in culture in cooperation with insulin-like growth factor (Kirkpatrick et al., 1995). In addition, certain IDGFs possess immune properties (Kucerova et al., 2016).
Previous proteomic and microarray analyses identified IDGF1-3 in Drosophila immune processes in both larvae and adult flies (De Gregorio et al., 2001; Irving et al., 2001;
Irving et al., 2005; Karlsson et al., 2004; Vierstraete et al., 2003). In particular, Idgf1 and
3 are induced upon septic injury (De Gregorio et al., 2001), and Idgf2 is upregulated early after infection with Gram-positive bacteria and yeast (Broz et al., 2017). Interestingly,
Idgf3 expression is regulated by Toll and Imd signalling (De Gregorio et al., 2001), and all IDGFs are part of Drosophila hemolymph clots (Irving et al., 2005; Karlsson et al.
2004; Vierstraete et al., 2003).
Here we have investigated the transcriptional regulation of Idgf2 and Idgf3 genes in
Drosophila larvae infected with Steinernema nematodes as well as the survival ability of
119 Idgf mutant larvae against the nematode parasite, Steinernema. We have found that Idgf2 and Idgf3 are upregulated in response to symbiotic and axenic Steinernema, and that
Drosophila Idgf2 loss-of-function mutants survive better the infection by axenic nematodes than the control larvae. Also, we have found that inactivation of Idgf2 alters the Imd and Jak/Stat signaling and inactivation of Idgf3 alters the Imd and Toll signaling.
These results suggest that Idgf2 and Idgf3 are essential components of the Drosophila anti-nematode immune defense. Molecular and functional characterization of the insect immune response to entomopathogenic nematodes will potentially reveal novel factors with anti-parasitic properties that could serve as targets for the development of alternative tactics to control insect pests of agricultural or medical importance.
MATERIALS AND METHODS
Fly Strains
Drosophila melanogaster strains used included the background strains Oregon, w1118, yw, and mutants for Idgf2 (Bloomington Drosophila Stock Centre; 18031; FBst0018031),
Idgf3 (Bloomington Drosophila Stock Centre; 26437; FBst0026437), PGRP-LE
(Bloomington Drosophila Stock Centre; 33055; FBst0033055), MyD88 (Tauszig-
Delamasure et al., 2002), Wengen (Vienna Drosophila Resource Centre; v330339;
FBst0490891) and UAS-DomeDN (Yang et al., 2015), which was crossed with the Actin
Gal4 driver (y[1] w[*]; P{w[+mC]=Act5C-GAL4}25FO1/CyO, y[+]). All strains were grown on approximately 2.5 g of Carolina Formula 4-24 Instant Drosophila media
(Carolina Biological Supply), 10 ml of deionized water, and approximately 10 granules
120 of dry baker’s yeast. All fly stocks were maintained at 25ºC with a 12:12-hour light:dark cycle. Late second to early third instar larvae were used for all experiments.
Nematodes and Bacteria
The insect pathogenic nematodes Steinernema carpocapsae harboring the Gram-negative bacteria Xenorhabdus nematophila (symbiotic nematodes) were amplified in the larvae of the waxworm Galleria mellonella, as previously described (McMullen & Stock, 2014).
Nematodes lacking their Xenorhabdus bacteria (axenic nematodes) were generated using a recently published protocol (Yadav et al., 2015). Prior to use, axenic nematodes were surface sterilized in 1% bleach and washed five times with sterile distilled water to remove any traces of bacteria on their surface. Infective juvenile (IJ) stage nematodes 2-4 weeks old were used for all experiments.
Transcriptomic Analysis
Reads Per Kilobase of transcript, per Million mapped reads (RPKM) values for Idgf2 and
Idgf3 genes were obtained from a recently published RNA-sequencing project involving transcriptomic analysis at 6 and 24 h post infection of Drosophila Oregon larvae with 100 symbiotic or axenic Steinernema IJ nematodes (Yadav et al., 2017). The uninfected control RPKM values were set to 1 and the RPKM values for nematode infected larvae were calculated relative to uninfected controls.
121 Survival Assays
Microtiter 96-well plates were prepared by adding 100 µl of 1.25% agarose to each well.
Ten symbiotic or axenic Steinernema IJ nematodes were placed into 10 µl of sterile distilled water. The nematode suspension (10 µl) was added to an individual larva in each well, the plate was covered with a Masterclear real-time PCR Film (Eppendorf, USA) and holes were pierced to allow ventilation. Addition of sterile water served as control.
Each larva was washed with sterile distilled water prior to infection. Each experiment consisted of 10 larvae per Idgf mutant or background control strain, per treatment.
Survival was estimated at 6-hour intervals and up to 66 h post infection.
Gene Transcript Levels
To validate the RNA-seq results, four larvae of the Oregon strain were infected with 10 or 100 Steinernema symbiotic or axenic IJ nematodes, and infected larvae were collected at 6 and 24 h post-infection. Control treatment involved application of sterile water to the larvae. Total RNA was extracted using the Invitrogen™ Ambion™ TRIzol™ Reagent.
Complementary DNA (cDNA) synthesis and qRT-PCR was performed as described previously (Shokal et al., 2016). All primer sequences used for qRT-PCR are shown in
Table 1. The amount of mRNA detected for each sample was normalized to the mRNA of the housekeeping gene RpL32 and represented relative to uninfected controls. Results demonstrate the mean and standard deviations of relative values from three biological replicates.
122 Hemocyte numbers
Hemolymph was collected from Drosophila larvae (n=2) at 6 and 24 h post infection with 10 Steinernema symbiotic or axenic IJ nematodes. Larvae treated with sterile distilled water served as controls. The procedure for hemolymph extraction followed a previously published protocol (Zettervall et al., 2004). Larvae were washed with 1X PBS and hemolymph was extracted by dissecting the larval cuticle with forceps. Hemolymph was diluted in 10 µl of 1X PBS. The hemolymph-PBS solution was then loaded onto a
Neubauer hemocytometer for hemocyte counting. Total numbers of cells were estimated at 40X magnification on a compound microscope (Olympus CX21). Two technical replicates were used for each strain and treatment per time-point and the experiment was repeated three times.
Statistical Analysis
Statistics were performed using the GraphPad Prism7 software. Statistical analysis of data from survival experiments was conducted using a log-rank (Mantel-Cox) and Chi- square test. P-values below 0.05 were considered statistically significant. Statistical analysis for gene transcription results and total hemocyte counts were performed using one-way analysis of variance (ANOVA) and a Tukey post-hoc test for multiple comparisons.
RESULTS
Steinernema nematode infection upregulates Idgf2 and Idgf3 in Drosophila larvae
123 To investigate the transcriptional activation of Idgf2 and Idgf3 in Drosophila larvae responding to Steinernema nematodes, we infected Drosophila larvae with 100 symbiotic or axenic Steinernema nematodes and estimated the relative expression (RPKM) of Idgf2 and Idgf3 at 6 and 24 h post infection using RNA-seq. We found that the number of reads for Idgf2 were higher than for Idgf3 in larvae infected with symbiotic or axenic IJ (Fig.
1A-D). To validate the Idgf gene expression results obtained through RNA-seq, we infected wild-type (Oregon) Drosophila larvae with Steinernema IJ and used qRT-PCR to estimate Idgf2 and Idgf3 gene transcript levels at 6 and 24 h post infection. We found that infection with 100 or 10 symbiotic nematodes resulted in similar transcript levels of
Idgf2 at 6 and 24 h (Fig. 1A). Also, at 6 h and 24 h post-infection with symbiotic nematodes, Idgf3 transcript levels obtained from RNA-seq or qRT-PCR were upregulated at similar levels (Fig. 1B). We found that at 24 h post-infection with 100 symbiotic nematodes, Idgf3 transcript levels were significantly higher compared to larvae infected with 10 symbiotic nematodes (Fig. 1B; p=0.0439). In addition, there were no significant differences in the upregulation of Idgf2 in larvae infected with 100 or 10 axenic nematodes at 6 h and 24 h. RNA-seq and qRT-PCR results showed that Idgf2 and Idgf3 are upregulated at similar levels at 6 and 24 h post infection with axenic nematodes (Fig.
1C, D). These findings suggest that Idgf2 and Idgf3 are upregulated in Drosophila larvae responding to symbiotic or axenic Steinernema nematode infections.
Drosophila Idgf2 mutants display increased survival upon Steinernema infection
To estimate the effect of inactivating Idgf2 or Idgf3 on the Drosophila survival response to nematode infection, we infected Idgf2 or Idgf3 loss-of-function mutant larvae and their
124 respective background controls with axenic or symbiotic Steinernema and estimated survival rates over time (Fig. 2). We found that Idgf2 mutants survived the infection with axenic nematodes at higher rates than their controls (Fig. 2A; p=0.0042). We found no differences in survival rates between Idgf2 mutant larvae and their controls upon infection with symbiotic nematodes (Fig. 2A). There were also no differences in survival between Idgf3 mutant larvae and control individuals following infection with either symbiotic or axenic nematodes (Fig. 2B). These results indicate that mutating Idgf2 in
Drosophila provides a survival advantage to larvae when responding to Steinernema nematodes lacking their mutualistic Xenorhabdus bacteria.
Drosophila Idgf2 and Idgf3 mutants induce Imd, Toll and Jak/Stat signaling upon
Steinernema infection
To estimate whether immune signaling pathway activation is affected in Drosophila Idgf mutant larvae in the context of nematode infection, we infected Idgf2 and Idgf3 mutants and their background strains with Steinernema symbiotic or axenic nematodes and quantified the transcript levels of read-out genes at 6 and 24 h post-infection. We used gene-specific primers to determine the transcript levels of Drosomycin as a read-out for the Toll pathway, Attacin for the Imd pathway, Puckered for the Jnk pathway and
Turandot-A (Tot-A) for the Jak-Stat pathway (Brun et al., 2006; McEwen & Peifer, 2005;
Myllymaki et al., 2014; Valanne et al., 2011). We found that in uninfected control larvae,
Drosomycin transcript levels were higher in Idgf2 mutants compared to the w1118 larvae, although this increase was not significant (Fig.3A). Also, Drosomycin transcript levels were significantly higher in Idgf3 mutant larvae compared to w1118 larvae (p=0.0137; Fig.
125 3A). We further found that Drosomycin mRNA levels were increased in Idgf2 mutants at
24 h post infection with symbiotic nematodes compared to axenic nematodes as well as in w1118 larvae infected with symbiotic nematodes, although this increase was not statistically significant (Fig. 3B). Similarly, Drosomycin levels were higher in Idgf3 mutant larvae at 24 h post infection with either symbiotic or axenic nematodes compared to the w1118 controls (Fig. 3B). In uninfected control larvae, transcript levels of Attacin were increased in Idgf2 mutants at 6 h compared to Idgf3 mutants and the w1118 controls
(Fig. 3C). Also at 6 h post-infection with axenic nematodes, we detected significantly higher levels of Attacin transcript levels in Idgf3 mutant larvae compared to Idgf2 mutants (p=0.0070) and w1118 (p=0.0016) larvae infected with axenic nematodes (Fig.
3C). At 24 h post-infection with symbiotic nematodes, Attacin transcript levels were significantly increased in Idgf2 mutants compared to w1118 larvae (p<0.001), Idgf3
(p<0.001) and Idgf2 larvae (p<0.0001) infected with axenic nematodes, as well as to uninfected controls (p<0.0001; Fig. 3D). In w1118 control larvae, Puckered transcript levels were significantly higher compared to Idgf2 (p=0.004) and Idgf3 mutant larvae
(p=0.004; Fig.3E). Also in the w1118 larvae, infection with symbiotic or axenic nematodes decreased Puckered transcript levels compared to the uninfected controls (p=0.0157 and p=0.0053 respectively; Fig.3E); however, at 24 h post-infection with symbiotic nematodes Puckered was upregulated in w1118 larvae compared to the uninfected controls
(p=0.0150) and larvae infected with axenic nematodes (p=0.0150). The increase in
Puckered transcript levels in w1118 larvae was also significantly higher compared to Idgf2
(p=0.0107) and Idgf3 (p=0.0054) mutant larvae upon infection with symbiotic nematodes
(Fig. 3F). Finally, we found that TotA was upregulated only in uninfected Idgf2 mutant
126 larvae at 6 and 24 h compared to uninfected w1118 (p=0.0071) and Idgf3 mutant larvae
(p=0.0155; Fig. 3G,H). These results suggest that inactivation of Idgf2 in Drosophila results in the Imd and Jak/Stat pathways activation whereas defective Idgf3 results in the
Toll and Imd pathways activation, in response to nematode infection.
Idgf2 is upregulated in Drosophila Imd and Jak/Stat mutants responding to
Steinernema infection
To investigate whether interfering with immune signaling in Drosophila regulates Idgf gene expression upon nematode infection, we infected Drosophila immune mutant larvae with Steinernema symbiotic or axenic nematodes and estimated Idgf2 and Idgf3 transcript levels at 6 and 24 h post-infection (Fig. 4). We used Drosophila strains mutant for
MyD88 (Toll), PGRP-LE (Imd), Wengen (Jnk) or Domeless (Jak/Stat). We found no differences in Idgf2 or Idgf3 induction compared to the background controls in uninfected or nematode infected MyD88 mutant larvae (Fig. 4A,B). We found increased Idgf2 transcript levels in uninfected PGRP-LE mutants and those infected for 6 h with axenic nematodes compared to the background controls (p=0.0013 and p=0.0431 respectively;
Fig. 4C). Idgf3 transcript levels were also increased in PGRP-LE mutants at 24 h post- infection with symbiotic or axenic nematodes compared to uninfected controls, but this increase was not statistically significant (Fig. 4D). In addition, Idgf2 and Idgf3 transcript levels were unaltered between uninfected or nematode-infected Wengen mutants and their background controls (Fig. 4E,F). In Domeless mutants, Idgf2 transcript levels decreased from 6 to 24 h in uninfected control larvae (p=0.0015), and in those infected with symbiotic (p=0.0234) or axenic nematodes (p=0.0612; Fig. 4G). The decrease in Idgf3
127 transcript levels in Domeless mutants infected with symbiotic nematodes was not statistically significant compared to the background controls (Fig. 4H). These results indicate that interfering with Imd or Jak/Stat signaling in Drosophila alters Idgf2 expression in response to Steinernema nematode infection.
Hemocyte numbers increase in Drosophila Idgf3 mutants responding to Steinernema infection
To investigate the cellular immune response in Drosophila larvae infected with
Steinernema nematodes, we looked for changes in the total number of hemocytes in
Drosophila larvae upon parasitic nematode infection. For this, we infected Drosophila
Idgf2 or Idgf3 mutant larvae and their background controls with Steinernema symbiotic or axenic nematodes and assessed the total number of hemocytes at two time-points post- infection (Fig.5). We found that the hemocyte numbers in Idgf2 mutants increased at 6 h post infection with symbiotic or axenic nematodes compared to uninfected control larvae, but this increase was not statistically significant (Fig. 5A). There were no changes in hemocyte numbers in Idgf2 mutant larvae infected with either type of nematode compared to uninfected larvae, at 24 h (Fig. 5A). However, hemocyte numbers in Idgf3 mutants were higher than in background controls at 24 h after infection with symbiotic
(p=0.0445) or axenic (p=0.0011) nematodes (Fig. 5B). Also, Idgf3 mutants infected with axenic nematodes contained significantly higher numbers of hemocytes at 24 h post- infection compared to uninfected controls (p=0.0009; Fig. 5B). These results imply that interfering with Idgf3 expression in Drosophila larvae increases hemocyte numbers in response to Steinernema nematode infection.
128 DISCUSSION
The Drosophila Imaginal Disc Growth Factors belong to family 18 of glycosyl hydrolases. Members of this protein family consist of true enzymes and chitinase-like proteins (lacking the enzymatic activity). The Drosophila IDGFs are structurally similar to chitinases, but lack their typical enzymatic activity (Arakane & Muthukrishnan, 2010;
Pesch et al., 2016). Various proteins have been identified in vertebrates with sequence homology to chitinase enzymes lacking the catalytic domain, which are typically secreted locally or in circulation during inflammation responses (Bussink et al., 2007). For example, the murine Yml protein, a member of the family 18 of glycosyl hydrolases, is expressed during allergic inflammation and nematode infection (Homer et al., 2006; Nair et al., 2005). This suggests that chitinase-like proteins may not be involved in chitin degradation in vertebrates and invertebrates. The Drosophila IDGFs are part of the larval hemolymph clot (De Gregorio et al., 2001; Karlsson et al., 2004; Vierstraete et al., 2003), and certain Idgf genes can function as pattern recognition molecules to detect carbohydrate (chitin) present on the surface of nematode parasites (De Gregorio et al.,
2001).
Transcriptional studies have implied the potential involvement of Idgf genes in the
Drosophila anti-nematode immune response. Infection with Heterorhabditis symbiotic nematodes as well as Steinernema axenic or symbiotic nematodes induces the expression of Idgf genes in Drosophila larvae (Arefin et al., 2014; Kucerova et al., 2016). In the current study, we have estimated the transcriptional induction of Idgf2 and Idgf3 in response to two types of Steinernema nematode infections. We show the transcriptional upregulation of these genes in response to two nematode parasites containing or lacking
129 their closely associated mutualistic bacteria Xenorhabdus. We find that infection with
100 or 10 Steinernema nematodes highly increases transcript levels of both Idgf2 and
Idgf3, suggesting the potential involvement of those genes in the interaction of
Drosophila with these parasitic nematodes.
Recent studies have shown that Idgf3 participates in wound response and protects
Drosophila larvae against infection with Heterorhabditis bacteriophora symbiotic nematodes (Kucerova et al., 2016), and Idgf2 is induced in response to septic (microbial) and aseptic injury (Broz et al., 2017). Here, we have investigated the participation of
Idgf2 and Idgf3 genes in the Drosophila response to Steinernema nematodes. We find that inactivation of Idgf2, but not Idgf3, delays Drosophila larval death upon infection with Steinernema axenic nematodes. This is in contrast to previous findings showing high sensitivity of Idgf3 mutants to Heterorhabditis symbiotic nematode infection (Kucerova et al., 2016). The contradiction between the previous and current findings suggests that certain Idgf genes can play distinct roles in the immune system of Drosophila responding to specific parasitic nematode species. Recently, an in vitro investigation using
Drosophila wing disc cell lines identified several potential downstream target genes of
Idgf2 mainly in the Imd pathway, including Attacin-A, -B, -D, Cecropin-A1, PGRP-LB and Relish (Broz et al., 2017). Interestingly, here we find that inactivation of Idgf2 strongly upregulates the Imd pathway read-out gene Attacin-A, in response to
Steinernema symbiotic nematodes.
We have also shown that interfering with Imd signaling by inactivating the Imd receptor gene PGRP-LE upregulates Idgf2 in response to Steinernema infection. These findings have prompted us to speculate that Idgf2 interacts with the Imd pathway, likely
130 functioning as a recognition molecule taking part in nematode detection or the detection of molecules secreted by parasitic nematodes during infection. Indeed, introduction of extracted cuticles from entomopathogenic nematodes into G. mellonella larvae can decrease hemocyte numbers and suppress aggregation as well as phagocytosis and encapsulation responses (Yi et al., 2016). It is therefore likely that Idgf2 is in some way involved in identifying the presence of cuticular chitin in the invading nematodes. Also, entomopathogenic nematodes secrete serine proteases, sc-sp-1 and sc-sp-3, that function as virulence factors that disarm the insect immune response by destroying the gut lumen and inducing apoptosis, respectively (Toubarro et al., 2009; Toubarro et al., 2010).
Nematode secreted proteins could also form identification targets for Idgf molecules in
Drosophila and potentially in other insects.
Idgf3 expression and Toll pathway signaling activity are essential in the wound response of Drosophila larvae and embryos respectively (Carvalho et al., 2014; Kucerova et al., 2016). We have hypothesized that Idgf3 and Toll pathway might work synergistically in response to nematode infections. We have found that inactivation of
Idgf3 induces the activation of the Toll pathway read-out drosomycin in response to
Steinernema; however, inactivation of the Toll pathway adaptor encoding gene MyD88 has no effect on Idgf3 induction in response to these nematodes. Thus, implying that the interaction between Idgf3 and Toll signaling probably takes place upstream of the Toll receptor and Idgf3 might function as an antagonist of the Toll pathway in response to entomopathogenic nematode infections.
We also find that infection with Steinernema symbiotic or axenic nematodes increases hemocyte numbers in Idgf3, but not Idgf2 mutant larvae. During infection,
131 hemocytes control insect cellular immune responses, which include hemocyte proliferation to phagocytose or encapsulate the invading pathogen (Cooper &
Eleftherianos, 2016). Although Idgf3 is essential in wound healing (Kucerova et al.,
2016), there is currently no indication that Idgf3 or members of this gene family is involved in hemocyte recruitment in Drosophila. Also, because Idgf3 mutants contain elevated numbers of hemocytes but they are as sensitive to Steinernema nematodes as the background controls, whereas Idgf2 mutants have increased survival against axenic nematodes with no changes in hemocytes implies that hemocyte density may not form an essential feature of the Drosophila response to Steinernema nematode infection. Further studies to deduce the exact role of Idgf3 in hemocyte recruitment and function are warranted.
In conclusion, we find that Idgf2 and Idgf3 regulate distinct immune functions in
Drosophila larvae in response to Steinernema nematode infection. We show that interfering with Idgf2 expression confers a protective effect against Steinernema axenic nematodes. We also show that Idgf2 and Idgf3 act in a distinct manner to regulate the
Drosophila immune signaling against Steinernema symbiotic or axenic nematodes. Thus, our findings indicate novel regulatory roles for Drosophila Idgf genes in contributing to the modulation of the anti-nematode immune response. Such findings contribute towards elucidating the molecular components involved in the interaction of parasitic nematodes with the insect innate immune system. Future investigations will focus on dissecting the mechanistic basis of Drosophila factors that oppose entomopathogenic nematode infection to understand their properties and exact mode of action, and their contribution to the overall insect anti-nematode immunity.
132
FIGURE LEGENDS
FIGURE 1 Upregulation of Idgf2 and Idgf3 in Drosophila larvae following
Steinernema nematode infection. The relative expression of (A,C) Idgf2 and (B,D)
Idgf3 at 6 and 24 h post infection in late second or early third instar larvae of the
Drosophila melanogaster (Oregon strain) with (A,B) symbiotic (Sy) or (C,D) axenic
(Ax) Steinernema carpocapsae nematodes was estimated by RNAseq (RPKM values) or quantitative RT-PCR analysis. Idgf gene transcript levels from qRT-PCR are shown as relative abundance of transcripts normalized to the housekeeping gene RpL32 and expressed as a ratio compared to uninfected control larvae. Values represent the means from three biological replicates and error bars represent standard deviations. Data were analyzed using one-way analysis of variance with a Tukey post-hoc test for multiple comparisons on GraphPad Prism 7. *P<0.05, non-significant differences (P>0.05) are not shown.
FIGURE 2 Drosophila Idgf2 mutants exhibit delayed mortality following
Steinernema infection. Survival of late second or early third instar Drosophila melanogaster larvae mutant for (A) Idgf2 and (B) Idgf3 following infection with
Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes. Survival was monitored every 6 h and up to 66 h post infection. Data were analyzed using Log-Rank test on GraphPad Prism7, and the values are the percent survival of the infected larvae.
The means from three independent experiments are shown and bars represent standard errors. **P<0.01, non-significant differences (P>0.05) are not shown.
133
FIGURE 3 Drosophila Idgf2 and Idgf3 mutants induce Toll, Imd and Jak/Stat signaling following Steinernema infection. Transcript levels of (A,B) Drosomycin,
(C,D) Attacin, (E,F) Puckered and (G,H) TotA were estimated in Drosophila melanogaster Idgf2 and Idgf3 mutant larvae at 6 and 24 h post-infection with
Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes. Transcript levels are shown as relative abundance normalized to the housekeeping gene RpL32 and expressed as a ratio that is compared to uninfected control larvae for each strain. Values represent means and error bars represent standard deviations. Data were analyzed using one-way analysis of variance with a Tukey post-hoc test for multiple comparisons on GraphPad
Prism 7. *P<0.05, **P<0.01, ***P<0.001, ****P<0.0001, non-significant differences
(P>0.05) are not shown.
FIGURE 4 Upregulation of Drosophila Idgf2 in Imd and Jak/Stat mutants following
Steinernema infection. Relative transcript levels of Idgf2 and Idgf3 were estimated in
(A,B) MyD88 (Toll), (C,D) PGRP-LE (Imd), (E,F) Wengen (Jnk) and (G,H) Domeless
(Jak/Stat) Drosophila melanogaster mutant larvae at 6 and 24 h post-infection with
Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes. Transcript levels are shown as relative abundance and normalized to the housekeeping gene RpL32, and expressed as a ratio compared to uninfected control larvae for each strain. Values represent means and error bars represent standard deviations. Data were analyzed using one-way analysis of variance with a Tukey post-hoc test for multiple comparisons on
134 GraphPad Prism 7. *P<0.05, **P<0.01, non-significant differences (P>0.05) are not shown.
FIGURE 5 Increased hemocyte numbers in Drosophila Idgf3 mutants following
Steinernema infection. Total numbers of hemocytes were assessed in Drosophila melanogaster (A) Idgf2 and (B) Idgf3 mutant larvae and their background controls at 6 and 24 h post-infection with Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes. Values represent means from three biological replicates and error bars represent standard deviations. Data were analyzed using one-way analysis of variance with a Tukey post-hoc test for multiple comparisons on GraphPad Prism 7. *P<0.05,
**P<0.01, ***P<0.001, non-significant differences (P>0.05) are not shown.
135 Idgf Idgf 2 RPKM 3 RPKM A 2.0 B 6 100 Sy 100 Sy *
e
e
c
c
n 1.5 10 Sy n 10 Sy
a
a
d d 4
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b 1.0 b
a
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i
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t 2
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l 0.5 l
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R
0.0 0 6 h 24 h 6 h 24 h
RPKM RPKM C 2.0 D 4 100 Ax 100 Ax
e
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n 1.5 10 Ax n 3 10 Ax
a
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0.0 0 6 h 24 h 6 h 24 h
Figure 1.
136 A 100 w1118 C
l 80 w1118 10Sy
a
v
i
v w1118 10Ax r 60
u
s
Idgf2 C
t
n
e 40 Idgf2 10Sy **
c
r
e
P 20 Idgf2 10Ax
0 0 6 12 18 24 30 36 42 48 54 60 66 B Time (hours) 100 w1118 C
l 80 w1118 10Sy
a
v i
v w1118 10Ax
r 60
u s
Idgf3 C
t n
e 40 Idgf3 10Sy
c
r e
P 20 Idgf3 10Ax
0 0 6 12 18 24 30 36 42 48 54 60 66 Time (hours)
Figure 2.
137 6 24 A 2.0×108 B 2.0×108
h*
e e h
c 8 c 8
n 1.5×10 n 1.5×10
a
a
d
d
n
n
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b 8 b 8
a 1.0×10
a 1.0×10
e e Drosomycin
v
v
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l 5.0×107 l 7 e 5.0×10
e
R R (Toll)
0.0 0.0 C y x C y x C y x C C C S A S A S A Sy Ax Sy Ax Sy Ax w1118 Idgf2 Idgf3 w1118 Idgf2 Idgf3
C 300 ** D **** 800 ****
e 200 **** ****
c 700
e n **
c a 600
n
d 100
a
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d u 500
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400 10 b
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v Attacin
i 2.0
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t 1.5 e 5
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R e 1.0 (Imd)
R 0 0.5 C y x C y x C y x 0.0 S A S A S A C C C Sy Ax Sy Ax Sy Ax w1118 Idgf2 Idgf3 w1118 Idgf2 Idgf3 *** E *** F ** ** * 6000 * * 8000 *
e 4000 6000
e
c
c
n
n 4000
a
2000 a
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d n 2000
n
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b
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50 a
Puckered
e 150
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i 40
v
t
i
t
a
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30 a 100
l
e
e R 20 (Jnk) 10 R 50 0 0 C y x C y x C y x S A S A S A C C C Sy Ax Sy Ax Sy Ax w1118 Idgf2 Idgf3 w1118 Idgf2 Idgf3
* ** G 4 ** H 4 *
e
e
c
c
n n 3 3
a
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R R (Jak/Stat) 0 0 C y x C y x C y x C C C S A S A S A Sy Ax Sy Ax Sy Ax w1118 Idgf2 Idgf3 w1118 Idgf2 Idgf3
Figure 3.
138 Idgf Idgf3 A B 2.5 2 2.5
e
e
c 2.0 c 2.0
n
n
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1.5 n
u 1.5
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1.0 e v 1.0
i
v
i t t Toll
a
l
a
l
e 0.5 e
R 0.5
R
0.0 0.0 C y x C y x C y x C y x y x y x y x y x S A S A S A S A C S A C S A C S A C S A w1118 MyD88 w1118 MyD88 w1118 MyD88 w1118 MyD88 6 h 24 h 6 h 24 h
* 4 C 2.0 ** D
e
c
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c n 3
n 1.5 a
a
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b 2
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1.0
a
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a 1 Imd
l 0.5 e
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R
R 0 0.0 C Sy Ax C Sy Ax C Sy Ax C Sy Ax C Sy Ax C Sy Ax C Sy Ax C Sy Ax w1118 PGRP-LE w1118 PGRP-LE w1118 PGRP-LE w1118 PGRP-LE 6 h 24 h 6 h 24 h
E 3 F 4
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i t 1
t
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e 1 e
R Jnk
R
0 0 C C C C Sy Ax Sy Ax Sy Ax Sy Ax C Sy Ax C Sy Ax C Sy Ax C Sy Ax w1118 Wengen w1118 Wengen w1118 Wengen w1118 Wengen 6 h 24 h 6 h 24 h
G 2.0 * H 2.0
**
e
e
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* c n 1.5 n 1.5
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R Jak/Sta
0.0 0.0 C C C C Sy Ax Sy Ax Sy Ax Sy Ax C Sy Ax C Sy Ax C Sy Ax C Sy Ax t w1118 Domeless w1118 Domeless w1118 Domeless w1118 Domeless 6 h 24 h 6 h 24 h
Figure 4.
139 Idgf Idgf 8×106 ** a 2 6
v 5×10 3 a
r B *** v
a r
l
/
a l
s 6 / 6
s 4×10 e 6×10
t
e t
y y
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4×10 m
e e
h h
6
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o o
r 6 r
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2×10 e
b 6 b 1×10
m m
u
u N N 0 0 y x y x y x y x C y x C y x C y x C y x C S A C S A C S A C S A S A S A S A S A
w1118 Idgf2 w1118 Idgf2 w1118 Idgf3 w1118 Idgf3 6 h 24 h 6 h 24 h
Figure 5.
140 TABLE 1 Primers used for quantitative RT-PCR analysis.
Gene Accession Primer Sequence (5´ – 3´) Tm (ºC)
Number
Idgf2 CG4475 Forward CATGAAGGCGTGGATCTGGT 54
Reverse CACCTCTCAGCCCAGCATAG
Idgf3 CG4559 Forward AGACATCTCCAGCACGCAAA 54
Reverse GGACAGACACAGAAGGAGGC
Drosomycin CG10810 Forward TGAGAACCTTTTCCAATATGATG 56
Reverse CCAGGACCACCAGCAT
Attacin-A CG10146 Forward CAATGGCAGACACAATCTGG 57
Reverse ATTCCTGGGAAGTTGCTGTG
Puckered CG7850 Forward GGCCTACAAGCTGGTGAAAG 61
Reverse AGTTCAGATTGGGCGAGATG
Turandot-A CG31509 Forward AGATCGTGAGGCTGACAAC 61
Reverse CCTGGGCGTTTTTGATAA
RpL32 CG7939 Forward GATGACCATCCGCCCAGCA 61
Reverse CGGACCGACAGCTGCTTGGC
141 Chapter 5: Participation of a serine protease gene Jonah66Ci in the Drosophila anti-
nematode immune response
ABSTRACT
Serine proteases and serine protease homologs form the second largest gene family in the Drosophila melanogaster genome. Several serine proteases are known to participate in immune functions in Drosophila. Jonah genes, which form a gene family of serine proteases, are implicated in the Drosophila anti-nematode and anti-viral immune response. However, the participation of these genes in the physiological processes of the
Drosophila immune response is still unknown. Here we report the involvement of
Jonah66Ci in the Drosophila immune defense against Steinernema carpocapsae nematode infection. We find that Drosophila Jonah66Ci is upregulated in response to symbiotic (carrying the mutualistic bacteria Xenorhabdus nematophila) or axenic
(lacking Xenorhabdus) Steinernema nematodes and is expressed exclusively in the gut of
Drosophila larvae. Inactivation of Jonah66Ci provides a survival advantage to
Drosophila larvae against axenic nematodes and results in differential expression of the
Toll and Imd pathway effector genes, specifically in the gut. We also find that inactivation of Jonah66Ci increases the rate of replication in the gut and infection with
Steinernema nematodes reduces replication rate. We show that absence of Jonah66Ci reduces nitric oxide levels and infection with Steinernema does not affect nitric oxide levels in Jonah66Ci mutant larvae. Finally, we show that absence of Jonah66Ci does not alter the feeding rates of Drosophila larvae, however infection with Steinenernema
142 axenic nematodes lowers larval feeding rates. In conclusion, we find that Jonah66Ci is essential in maintaining homeostasis of certain physiological processes in Drosophila larvae in the context of Steinernema nematode infection. Similar findings will take us a step further towards understanding the molecular mechanisms that take place during parasitic nematode infection in insects.
INTRODUCTION
Entomopathogenic nematodes of the genera Steinernema and Heterorhabditis are emerging as excellent models for studying insect-nematode interactions (Castillo et al.,
2011; Stock, 2005). They are natural obligate parasites of a wide range of insects that they infect to complete their life cycle. These nematode parasites infect susceptible insects as infective juveniles, a developmentally arrested stage, analogous to the
Caenorhabditis elegans dauer stage (Goodrich-Blair, 2007). A distinct feature of entomopathogenic nematodes is the presence of mutualistic bacteria that are localized to their intestines (Ciche et al., 2008; Martens & Goodrich-Blair, 2005). Steinernema carpocapsae forms a mutualistic relationship with the Gram-negative bacteria
Xenorhabdus nematophila (symbiotic nematodes) and together they form potent pathogenic complexes that infect insects (Goodrich-Blair, 2007; Pena et al., 2015; Yadav et al., 2017). The nematodes enter the insect body cavity through the cuticle or natural openings and subsequently expel their bacteria into the insect open circulatory system
(Pena et al., 2015). The bacteria secrete toxins, virulence factors and degradative enzymes that target several insect tissues and interfere with the insect immune response, which eventually leads to rapid insect death (Castillo et al., 2011; Waterfield et al.,
143 2009). The bacteria also provide nutrients to the nematodes, which promote the completion of their reproductive cycle (Richards & Goodrich-Blair, 2009). Once the food source is depleted, the nematodes reacquire the bacteria, and exit the insect cadaver in search of new insect hosts (Goodrich-Blair, 2007).
Drosophila melanogaster is an established model for dissecting the molecular and cellular basis of host-pathogen interactions (Kounatidis & Ligoxygakis, 2012). Extensive studies have led to the identification and understanding of evolutionarily conserved signaling pathways that are activated in response to different types of microbial infections
(Hetru & Hoffmann, 2009; Hoffmann & Reichhart, 2002; Tanji et al., 2007). Drosophila has been employed recently to dissect the molecular mechanisms that occur in insects responding to parasitic nematode infections (Arefin et al., 2014; Hallem et al., 2007;
Pena et al., 2015; Yadav et al., 2017). The use of Drosophila and Steinernema to unravel the insect immune response has certain advantages. The Drosophila immune system shares significant homology to the mammalian innate immune system, which facilitates modeling parasitic processes and anti-nematode immune reactions in humans (Castillo et al., 2011; Silverman & Maniatis, 2001; Stock, 2005). Steinernema symbiotic or axenic nematodes are pathogenic to Drosophila and interestingly they are capable of killing larvae at similar rates (Yadav et al., 2015). In addition, Steinernema nematode infection activates the expression of antimicrobial peptide genes and the melanization pathway, and mutualistic Xenorhabdus bacteria suppress the latter response (Pena et al., 2015).
The imaginal disc growth factor-3 (Idgf3) and two clotting factors (gp150 and fondue) are found to participate specifically in the Drosophila anti-nematode immune response.
144 Knockdown of Idgf3, gp150 or fondue increases susceptibility of larvae responding to
Heterorhabditis bacteriophora nematodes (Hyrsl et al., 2011; Kucerova et al., 2016).
The Jonah gene family consists of approximately 20 genes organized in small clusters on different chromosomal sites and exhibit complex expression patterns (Carlson
& Hogness, 1985a; Hafen et al., 1983). In situ hybridization identified the expression of
Jonah25Bi, Jonah65Ai and Jonah99Cα in the Drosophila midgut. These Jonah genes are expressed during the larval and adult stage of Drosophila, but not in the pupal stage
(Carlson & Hogness, 1985a). Low level Jonah expression is also detected in the presumptive midgut from 18 h embryos (Hafen et al., 1983). Because Jonah genes are exclusively expressed in the Drosophila gut, Jonah proteases are implicated in the breakdown of dietary proteins due to their homology to mammalian serine proteases, trypsin and chymotrypsin (Fernandez-Ayala et al., 2010). More recently, transcriptomic studies have reported the induction of several Jonah genes in Drosophila responding to viral or nematode infections (Carpenter et al., 2009; Chtarbanova et al., 2014; Yadav et al., 2017). Jonah genes are conserved in the genomes of other Drosophila species, such as Drosophila simulans, but not in other insects (Carlson & Hogness, 1985a).
In this study, we have investigated the transcriptional regulation of Jonah66Ci in
Drosophila larvae infected with Steinernema symbiotic or axenic nematodes. In uninfected and nematode infected Drosophila larvae, Jonah66Ci is solely expressed in the gut. We report that infection with Steinernema axenic nematodes provides a survival advantage to Drosophila Jonah66Ci mutant larvae. Interestingly, interfering with
Jonah66Ci expression results in the downregulation of Imd signaling in whole larvae in response to Steinernema nematode infection. We also find that the read-out genes of the
145 Toll pathway, Drosomycin and Defensin, and the Imd pathway, Diptericin and Cecropin, are differentially expressed in the gut of Jonah66Ci mutants, in response to Steinernema nematode infection. In addition, inactivation of Jonah66Ci increases cell replication in the gut of Drosophila larvae responding to Steinernema infection. We show that in the absence of nematode infection, inactivation of Jonah66Ci decreases nitric oxide levels in the gut of larvae. Finally, we show that infection with Steinernema axenic nematodes reduces the feeding rates of Jonah66Ci mutant larvae. These results indicate that absence of Jonah66Ci in Drosophila is responsible for modulating the interaction with entomopathogenic nematodes. Identification and characterization of gene function with anti-nematode immune properties in Drosophila sets the stage for uncovering conserved genes encoding anti-nematode factors in vertebrates.
MATERIALS AND METHODS
Fly strains
Drosophila melanogaster yellow white (Lacey et al., 2015) and mutant Jonah66Ci
(v103008; Vienna Drosophila Resource Centre; FBst0474871) strains were used. The
Jonah66Ci RNAi line was crossed with the Actin Gal4 driver (y[1] w[*];
P{w[+mC]=Act5C-GAL4}25FO1/CyO, y[+]) (Ekengren et al., 2001) or the Escargot
Gal4 driver (w*; P{enG}esgG66/CyO, P{GAL4-Kr.C}DC3, P{UAS-GFP.S65T}DC7) (Le
Bras & Van Doren, 2006). All strains were reared on Drosophila media (Meidi
Laboratories) and sprinkled with approximately 10 g of baker’s yeast. Stocks were
146 maintained at 12:12 light: dark cycle at 25°C. Late second to early third instar larvae were used for all experiments.
Nematodes
Infective juveniles of Steinernema carpocapsae nematodes were used for all experiments.
Symbiotic nematodes carrying Xenorhabdus nematophila bacteria were reared in larvae of the wax-moth Galleria mellonella as described previously (White, 1927). Axenic nematodes lacking Xenorhabdus were generated following a previously established protocol (Yadav et al., 2015). Axenic nematodes were washed in 1% bleach solution to remove bacteria from the nematode surface and rinsed five times with water to remove the bleach residue. Infective juveniles of 2-5 weeks old were used for all experiments.
Gene transcript analysis with RNA-sequencing
Reads per Kilobase of transcript, per Million mapped reads (RPKM) values for
Jonah66Ci (CG7118) were obtained from a recent RNA-sequencing study (Yadav et al.,
2017). The reads were obtained at 6 and 24 h post-infection of Drosophila melanogaster
Oregon larvae with 100 Steinernema carpocapsae symbiotic or axenic infective juveniles. The RPKM values for nematode infected larvae are shown relative to the
RPKM values of uninfected control larvae, at each time-point.
147 Gene transcript analysis with quantitative RT-PCR
Four larvae each infected with 100 Steinernema symbiotic or axenic nematodes were collected at 6 and 24 h post-infection for measuring gene transcript levels using qRT-
PCR. For estimating Jonah66Ci transcript levels in the gut, 10 larvae infected with 10 symbiotic or axenic nematodes were dissected at 6 and 24 h post-infection to separate the gut from the rest of the larvae. In all cases, larvae treated with sterile distilled water served as the uninfected control treatment. Total RNA extraction was performed using the Invitrogen™ Ambion™ TRIzol™ Reagent, following the manufacturer’s instructions. RNA extraction, cDNA synthesis and qRT-PCR protocols were performed as described (Shokal et al., 2016). All primer sets used for qRT-PCR analyses and their respective annealing temperatures are listed in Table 1. Data are expressed as (ΔCt) of
2Ct(RpL32)/2Ct(gene) and presented as a ratio of infected larvae relative to uninfected controls.
Results represent mean and standard deviations from three biological replicates.
Survival experiments
A 96-well microtiter plate (Corning) was prepared by adding 100 µl of 1.5% agarose gel
(in 1X TAE buffer; Tris-Acetate-EDTA) to each well. A suspension (10 µl) containing
10 Steinernema symbiotic or axenic nematodes was added to each well. Application of sterile distilled water (10 µl) to larvae was used as uninfected control treatment. To each well, an individual Drosophila larva was added. The survival assay was performed as described previously (Yadav et al., 2017), and the experiment was repeated three times.
148 BrdU labeling
Ten Drosophila larvae from each strain were infected with 10 Steinernema symbiotic or axenic nematodes, or treated with sterile distilled water, and were subsequently collected at 24 h post-infection. The guts were dissected in Drosophila S2 medium and the intact gut tissues were then incubated for one hour at room temperature in 2 mg/ml 5-bromo-2- deoxyuridine (BrdU; Sigma). The fixation and antibody protocol for BrdU labeling have been described previously (Choubey & Roy, 2017). Fluorescent images were obtained using a LSCM-510 Meta confocal microscope (Carl Zeiss) at 40X magnification. Images were assembled using Adobe Photoshop (2018) and percentage of BrdU labeled nuclei were estimated.
Nitric oxide (NO) estimation
Gut samples from 10 Drosophila larvae infected with 10 Steinernema symbiotic or axenic nematodes were dissected 24 h post-infection. Gut samples from larvae treated with sterile distilled water served as uninfected controls. Gut samples were homogenized in PBS (1X phosphate buffered saline) by grinding with a sterile plastic pestle and then centrifuged at 10,000 g for 10 min at 4°C. The resultant supernatant was mixed 1:1 with
Greiss reagent (Sigma) and absorbance was measured at 595 nm using a plate-reader
(BioTek). NO levels were calculated from a silver nitrite standard curve.
149 Feeding rate
Ten Drosophila larvae from each strain were infected with 10 Steinernema symbiotic or axenic nematodes, or treated with sterile distilled water, and then collected 24 h post- infection. All larvae were fed on yeast paste containing 0.16% Erioglaucine disodium salt
(FD&C Blue No.1; Sigma) for 15 min. Larvae starved for 24 h served as background controls. The protocol for spectrophotometric detection of the food-dye has been described previously (Kaun et al., 2007). Whole larvae were used for all experiments.
The resultant supernatant (200 µl per sample) was loaded into a 96-well plate (Corning) and measured at 633 OD using a plate-reader (BioTek). The experiment was repeated three times.
Statistical analysis
For gene transcript level analysis, BrdU labeling, nitric oxide estimation and feeding rate, data analysis was performed using one-way analysis of variance (ANOVA) with a Tukey post-hoc test for multiple comparisons and unpaired two-tailed t-test. For survival experiments, a log-rank (Mantel-Cox) and Chi-square test was performed. P values lower than 0.05 were considered statistically significant. All figures were generated using
GraphPad Prism7 software.
150 RESULTS
Steinernema nematodes upregulate Jonah66Ci during the early stages of infection in
Drosophila
To investigate the transcriptional induction of Jonah66Ci in Drosophila larvae infected with Steinernema nematodes, we infected Drosophila larvae with 100 symbiotic or axenic nematodes and estimated the relative transcript levels of Jonah66Ci at 6 and 24 h post-infection. We compared the transcript levels (rpkm) of Jonah66Ci from a recent transcriptomic study (Yadav et al., 2017), and those from qRT-PCR analysis (ΔCt) (Fig.
1A). We found that at 6 h post-infection, Jonah66Ci transcript levels were higher in larvae infected with axenic nematodes compared to those infected with symbiotic nematodes, although values were not statistically significant.
A previous study identified members of the Jonah gene family, Jonah25Bi,
Jonah65Ai and Jonah99Cα in the Drosophila gut (Carlson & Hogness, 1985a). To determine whether Jonah66Ci is also expressed in the gut of Drosophila larvae, we estimated the transcript levels of Jonah66Ci in the Drosophila gut and the rest of the larval body at 6 and 24 h post-infection with Steinernema symbiotic or axenic nematodes.
We detected no mRNA levels of Jonah66Ci in the larval body without the gut in the absence or presence of nematode infection (Fig. 1B). At 6 h, we found that Jonah66Ci transcript levels were significantly higher in the gut of larvae infected with symbiotic nematodes compared to those in uninfected controls (p=0.0190; Fig. 1B). At 24 h, there were no differences in the transcript levels of Jonah66Ci in the gut of infected or uninfected larvae. We also found that Jonah66Ci transcript levels were significantly reduced from 6 h to 24 h in the gut of larvae infected with symbiotic (p=0.0073) or
151 axenic (p=0.0452) nematodes (Fig. 1B). These results indicate that Jonah66Ci is exclusively expressed in the gut of uninfected Drosophila larvae and infection with
Steinernema nematodes can increase its transcript levels.
Drosophila Jonah66Ci mutants display enhanced survival ability in response to infection with Steinernema axenic nematodes
To investigate whether inactivation of Jonah66Ci had an effect on the survival ability of
Drosophila larvae responding to Steinernema nematodes, we infected Jonah66Ci mutant and yw control Drosophila larvae with Steinernema symbiotic or axenic nematodes and measured larval survival every 8 h for three days (Fig. 2). We found that upon infection with axenic nematodes, yw larvae succumbed faster to infection compared to Jonah66Ci mutant larvae (p=0.0028; Fig.2). There were no differences in survival rates between
Jonah66Ci mutants and yw larvae infected with symbiotic nematodes. These results indicate that interfering with the expression of Jonah66Ci promotes the survival ability of
Drosophila larvae in response to Steinernema axenic nematode infections.
Imd pathway activation decreases in Drosophila Jonah66Ci mutants responding to
Steinernema axenic nematodes
To determine whether inactivation of Jonah66Ci in Drosophila has an effect on signaling pathway activation in response to Steinernema infection, we infected Jonah66Ci mutant and background control larvae with symbiotic or axenic nematodes and estimated transcript levels of Attacin (Imd pathway), Drosomycin (Toll pathway), Puckered (Jnk
152 pathway) and TotA (Turandot-A, Jak/Stat pathway) at two time-points post-infection
(Fig. 3). At 6 h, we found very low transcript levels of Attacin in both yw controls and
Jonah66Ci mutants infected with symbiotic or axenic nematodes. At 24 h, infection with symbiotic nematodes significantly upregulated Attacin in yw larvae compared to uninfected individuals (p=0.0022; Figure 3A). Attacin mRNA levels increased in yw larvae infected with axenic nematodes compared to uninfected controls; however, this increase was not statistically significant. Attacin transcript levels decreased in yw larvae infected with axenic nematodes compared to those infected with symbiotic nematodes
(p=0.0059; Fig. 3A). We also found that in yw larvae, symbiotic nematodes increased
Attacin transcript levels from 6 to 24 h post infection (p=0.0022; Fig. 3A). Conversely, infection of Jonah66Ci mutant larvae with symbiotic nematodes increased Attacin transcript levels, although this increase was not statistically significant. There were very low transcript levels of Attacin in uninfected Jonah66Ci mutant larvae or those infected with axenic nematodes. Estimation of Drosomycin in yw controls and Jonah66Ci mutant larvae revealed no significant changes in transcript levels upon infection with symbiotic or axenic nematodes at 6 and 24 h (Fig. 3B). There were no significant differences in transcript levels of puckered or Tot-A in yw or Jonah66Ci mutant larvae infected with symbiotic or axenic nematodes at any of the time-points (Fig. 3C and Fig. 3D, respectively). These results indicate that absence of Jonah66Ci in Drosophila larvae, axenic Steinernema nematodes fail to induce Imd signaling.
Toll and Imd pathways are differentially activated in Drosophila Jonah66Ci mutants responding to Steinernema symbiotic nematodes
153 Toll and Imd pathways regulate antimicrobial peptide production in the anterior midgut of Drosophila (Buchon et al., 2014). Jonah66Ci expression exclusively in the Drosophila gut (Fig. 1B), prompted us to investigate whether its inactivation affects Toll or Imd signaling in the context of Steinernema nematode infection. For this, we infected
Jonah66Ci mutants and their background controls with Steinernema symbiotic or axenic nematodes and 24 h later we estimated transcript levels of antimicrobial peptide encoding genes in the gut and the rest of the larva (Fig. 4). We found significant increase in
Drosomycin levels in the gut of yw larvae infected with symbiotic nematodes compared to axenic nematodes (p=0.005) and uninfected controls (p=0.0006; Fig. 4A). In the gutless larvae, Drosomycin transcript levels increased in response to symbiotic nematodes compared to uninfected controls, although this increase was not statistically significant. In yw larvae infected with Steinernema symbiotic nematodes, Drosomycin transcript levels were significantly higher in the gut compared to gutless larvae
(p=0.0391; Fig. 4A). Also, Drosomycin transcript levels increased in the gut of yw larvae compared to the gut of Jonah66Ci mutant larvae infected with symbiotic nematodes
(p=0.0004; Fig. 4A). There were few to no transcripts of Drosomycin in Jonah66Ci mutants, with or without nematode infection. In contrast, we found significant upregulation of Defensin transcripts in gutless larvae of Jonah66Ci mutants responding to symbiotic nematodes compared to uninfected larvae (p=0.0152; Fig. 4B). Upregulation of
Defensin in gutless Jonah66Ci larvae was significantly higher than in the gut of the mutants (p=0.171) and the yw gutless larvae (p=0.0188), in response to symbiotic nematode infection (Fig. 4B).
154 Infection with symbiotic nematodes consistently upregulated Diptericin in the gut and gutless larvae of yw and Jonah66Ci mutants (Fig. 4C). Diptericin was significantly upregulated in response to symbiotic nematode infection in yw gutless larvae compared to uninfected gutless larvae (p=0.0203), gutless larvae infected with axenic (p=0.0198) or gutless Jonah66Ci mutant larvae infected with symbiotic nematodes (p=0.0441; Fig. 4C).
Interestingly, Cecropin was significantly upregulated in the gut of Jonah66Ci mutants infected with symbiotic nematodes compared to axenic nematodes (p=0.0001) and uninfected control larval gut (p=0.0002; Fig. 4D). This increase was also statistically significant compared to Cecropin levels in gutless Jonah66Ci mutants infected with symbiotic nematodes (p=0.0005) as well as in the gut of yw larvae infected with symbiotic nematodes (p=0.0002; Fig. 4D). Cecropin was upregulated in yw gutless larvae previously infected with symbiotic nematodes, but this increase was not statistically significant (Fig. 4D). These results demonstrate that absence of Jonah66Ci in Drosophila larvae leads to differential expression of the Toll pathway genes Drosomycin and
Defensin and the Imd pathway genes Diptericin and Cecropin in the gut and the rest of the larval body in response to Steinernema nematodes.
Replication is reduced in Drosophila Jonah66Ci mutants in response to Steinernema symbiotic nematodes
Because Jonah66Ci is expressed in the gut of Drosophila larvae, we explored whether absence of Jonah66Ci influenced the activation of the gut specific Wnt/Wg signaling pathway (Swarup & Verheyen, 2012) (Fig. 5). For this, we infected yw and Jonah66Ci mutant larvae with Steinernema symbiotic or axenic nematodes and estimated transcript
155 levels of wingless, encoding a ligand of the Wnt/Wg signaling pathway, in the gut 24 h post-infection. We found that wingless was significantly upregulated in the gut of uninfected Jonah66Ci mutants compared to the background controls (Fig. 5A). We found that upon symbiotic nematode infection, wingless was upregulated in the gut of yw larvae compared to Jonah66Ci mutants (Fig. 5A). Infection with symbiotic or axenic nematodes had no effect on wingless transcript levels in the gut of yw larvae. Thus, these results suggest that inactivation of Jonah66Ci upregulates the Drosophila Wnt/Wg signaling.
Wnt/Wg signaling promotes tissue regeneration in the Drosophila gut after injury
(Liu et al., 2017). To investigate whether inactivation of Jonah66Ci in the gut affects tissue regeneration in response to nematode infection, we infected yw background controls and Jonah66i mutant larvae with Steinernema symbiotic or axenic nematodes and measured the number of cells undergoing replication (BrdU labeled) in the gut of infected and uninfected individuals. In uninfected guts, the numbers of BrdU labeled cells significantly increased in Jonah66Ci mutant larvae compared to yw controls (P=0.007;
Fig. 5C). Also, inactivation of Jonah66Ci significantly reduced the numbers of BrdU labeled cells in the gut of larvae infected with symbiotic nematodes compared to uninfected controls (P=0.0446; Fig. 5C). Thus, absence of Jonah66Ci in Drosophila larvae increases the numbers of gut cells undergoing replication and this effect is reduced in response to Steinernema symbiotic nematode infection.
Drosophila Jonah66Ci mutants exhibit reduced nitric oxide levels but not feeding rates
156 To determine whether certain physiological processes are affected by the absence of
Jonah66Ci in the context of nematode infection, we measured nitric oxide levels and feeding rates in Drosophila larvae 24 h post infection with Steinernema symbiotic or axenic nematodes (Fig. 6). We found that nitric oxide levels increased in yw larvae infected with symbiotic nematodes compared to those infected with axenic nematodes
(P=0.0320), and to Jonah66Ci mutant larvae infected with symbiotic nematodes
(P=0.0123; Fig. 6A). There were no changes in nitric oxide levels in Jonah66Ci mutant larvae in the presence or absence of nematode infection. We also measured the feeding rates in yw and Jonah66Ci mutant larvae at 24 h post nematode infection. We found that feeding rates increased in background control larvae infected with axenic nematodes, and this increase was significantly higher compared to yw larvae infected with symbiotic nematodes (P=0.0204), and to Jonah66Ci mutant larvae infected with axenic nematodes
(P=0.0249). These results indicate that absence of Jonah66Ci decreases NO levels in
Drosophila larvae, infection with symbiotic or axenic Steinernema nematodes does not alter this response, and presence or absence of Jonah66Ci does not affect the feeding rates of Drosophila larvae, with or without nematode infection.
DISCUSSION
Serine proteases and serine protease homologs constitute the second largest gene family in Drosophila (Ross et al., 2003). Within the serine protease family, those containing the trypsin-fold form one of the largest enzyme families that are distributed across several species (Di Cera, 2009). Over the years, studies have reported the participation of Drosophila serine proteases in development, aging, oxidative stress and
157 immune response (Veillard et al., 2016; Zou et al., 2000). Despite advances in the substrate specificity annotation of Drosophila serine proteases, our understanding of the exact function of these enzymes is still lacking (Ross et al., 2003). Jonah is part of a multi-member gene family (Carlson & Hogness, 1985b). Here we report the participation of Jonah66Ci in the interaction of the Drosophila immune response with a potent parasitic nematode.
Recent studies have shown the participation of serine proteases in the Drosophila immune responses to bacteria, fungi and parasitic nematode infections (Buchon,
Poidevin, et al., 2009; Castillo et al., 2015; Ligoxygakis et al., 2002; Patrnogic &
Leclerc, 2017; Yadav et al., 2017). Here, we find that inactivation of Jonah66Ci promotes the survival of Drosophila larvae upon infection with Steinernema axenic nematodes. This is in contrast to previous studies, showing that inactivation of the
Drosophila serine protease-encoding genes modSP and persephone confers susceptibility to flies responding to Gram-negative bacterial and fungal infections, respectively
(Buchon, Poidevin, et al., 2009; Ligoxygakis et al., 2002). This suggests that the function of certain serine protease genes in the Drosophila immune system depends on the developmental stage of Drosophila and on the type of pathogen encountered. The serine proteases Grass, spirit, spheroid, sphinx1/2 and persephone function upstream of the Toll receptor and are activated in response to Gram-positive bacteria or fungal pathogens, leading to the subsequent activation of Toll signaling (El Chamy et al., 2008; Kambris et al., 2006). In addition, a modular serine protease, ModSP, integrates signals from recognition molecules such as Gram-negative binding protein (GNBP3) and peptidoglycan recognition protein (PGRP)-SA and directs the activation of a protease
158 cascade involving Spatzle (Buchon, Poidevin, et al., 2009; Valanne et al., 2011) whereas the serine protease persephone acts independently of GNBP3 and identifies factors from live entomopathogenic fungi such as Beauveria bassinia (Gottar et al., 2006). This demonstrates that these particular serine proteases have crucial roles in recognition and immune activation in the Drosophila Toll pathway. We find that inactivation of the serine protease-encoding gene Jonah66Ci, generates mutants that fail to induce the transcriptional activation of the Drosophila Imd pathway read-out gene Attacin. This prompted us to speculate that Jonah66Ci interacts with the Imd pathway and could function as a regulator, possibly in a similar fashion to the upstream SPs functioning in the activation of the Drosophila Toll pathway (Valanne et al., 2011).
We found that Jonah66Ci is expressed in the gut of uninfected Drosophila larvae and infection with Steinernema nematodes alters its expression levels. The Drosophila lab-reared strains have varying levels of microbiota-associated guts (Wong et al., 2013) and a previous report has found that Jonah66Ci expression levels are comparatively lower in germ-free (no microbiotia) gut tissues (Erkosar et al., 2014). Hence it is likely that in response to Steinernema nematode infection, we would observe variation in the expression levels of Jonah66Ci in microbiota-associated and germ-free gut tissues of
Drosophila larvae. In addition, considering that Jonah66Ci is expressed only in the gut of uninfected and nematode infected Drosophila larvae, we hypothesized that Jonah66Ci controls processes and signaling pathways specific to this tissue. We find that inactivation of Jonah66Ci results in mutant larvae with differential expression of the Toll pathway read-out genes Drosomycin and Defensin, and the Imd pathway genes Diptericin and Cecropin in the gut of Drosophila larvae in response to Steinernema symbiotic or
159 axenic nematodes. A previous study has reported an increase in expression of the Toll pathway read-out Drosomycin and the Imd pathway read-out Diptericin in Drosophila flies injected with Xenorhabdus, the mutualistic bacterium of Steinernema nematodes
(Aymeric et al., 2010). Our data are in agreement with this study, because we find upregulation of Drosomycin and Diptericin in the gut of yw Drosophila larvae responding to Steinernema symbiotic nematodes, which carry mutualistic Xenorhabdus. In contrast, we find that in the gut of Jonah66Ci mutant larvae responding to Steinernema symbiotic nematodes, Drosomycin and Diptericin transcript levels are reduced. Thus, implying that
Jonah66Ci modulates the Drosophila Toll and Imd signaling activity in the context of
Steinernema symbiotic nematodes, and this is probably due to the presence of
Xenorhabdus bacteria. We also find that the Toll pathway read-out gene Defensin is upregulated in response to Steinernema axenic nematodes and at even higher levels in response to Steinernema symbiotic nematodes in the Jonah66Ci gutless mutant larvae, whereas Steinernema symbiotic or axenic nematodes failed to upregulate Defensin in wild-type larvae (Pena et al., 2015). This suggests that Jonah66Ci interacts with Toll signaling in larvae responding to Steinernema nematode infections. In Drosophila adult flies, Defensin is upregulated in the thioester-containing protein-4 mutants responding to
Photorhabdus luminescens or Photorhabdus asymbiotica infection, and this correlates with resistance of the mutant flies to infection (Shokal & Eleftherianos, 2017). Our findings are in agreement with this study, because we find upregulation of Defensin and higher survival for Jonah66Ci mutant larvae responding to Steinernema axenic nematodes. The differential induction of Toll and Imd pathway effector genes suggest that inactivation of Jonah66Ci not only modulates the immune signaling in the gut of
160 Drosophila larvae but in other tissues, probably the fat body or hemolymph, in response to Steinernema nematode infection. Another explanation could be the immune system compensating for the absence of Jonah66Ci in Drosophila larvae, in the context of nematode infection.
In case of gut epithelial damage or stress such as bacterial infection, the
Drosophila intestinal stem cells produce enteroblasts and enterocytes to regenerate the gut (Micchelli & Perrimon, 2006; Ohlstein & Spradling, 2006). Post-mitosis, enteroblasts endoreplicate their DNA, eventually differentiating into enterocytes (Takashima &
Hartenstein, 2012). A previous study using Drosophila adult flies has found that inactivation of Adenomatous polyposis coli (Apc), a tumor suppressor gene found in the intestinal epithelium, results in significant increase in the numbers of cells undergoing endoreplication (Lee et al., 2009). This finding is in agreement with our data because we also find that, in the absence of Steinernema nematode infection, the numbers of replicating cells are significantly increased in the gut of Jonah66Ci mutant larvae. Thus suggesting that Jonah66Ci, similar to Apc in Drosophila adults, is essential in maintaining homeostasis in the gut of Drosophila larvae responding to Steinernema nematodes. The upregulation of wingless in uninfected Jonah66Ci mutant larvae also suggests that Jonah66Ci functions synergistically with the Wnt/Wg pathway in cell proliferation and regulating asymmetric cell division (Bejsovec, 2013) and infection with
Steinernema nematodes affects this balance. A previous study reported that Pseudomonas entomophila secretes hemolysin that targets and lyses the enterocytes in the gut epithelium of Drosophila adults and larvae (Liehl et al., 2006; Xiang et al., 2017).
Similarly, Steinernema nematodes secrete a serine protease, sc-sp-1, that functions as a
161 virulence factor that disarms the immune system by destroying the gut lumen (Toubarro et al., 2010). Hence we speculate that the downregulation of the Wnt/Wg pathway gene wingless and reduction in the numbers of replicating cells in the gut of Jonah66Ci mutant larvae responding to Steinernema nematodes could be attributed to the virulence factors produced by Steinernema nematodes.
A previous study has reported the crucial role of NO in eliminating the eggs of the endoparasitic wasp Leptopilinia heterotoma in the Drosophila paramelancia larvae
(Carton et al., 2009). NO is essential for the survival of Drosophila flies responding to
Gram-negative bacterial infection (Eleftherianos et al., 2014). We find increased NO levels in yw larvae responding to Steinernema symbiotic, but not axenic, nematodes, which suggests that Drosophila larvae are capable of inducing NO response against the mutualistic Xenorhabdus bacteria. We also find a reduction in NO levels in the gut of uninfected and nematode infected Jonah66Ci mutant larvae, suggesting a role for
Jonah66Ci in regulating the NO anti-nematode response in Drosophila larvae.
We further find reduced feeding rates in Jonah66Ci mutant larvae responding to
Steinernema axenic nematodes. Infection of adult Drosophila with Drosophila C Virus
(DCV) increases the feeding rate of flies (Chtarbanova et al., 2014). Our data are in agreement with this finding, because we also find that yw larvae responding to
Steinernema axenic nematodes ingest significantly higher amounts of food. Because this effect is reduced in Jonah66Ci mutant larvae upon axenic nematode infection, we postulate that Jonah66Ci is associated with regulating the food uptake of larvae during infection with parasitic nematodes.
162 In conclusion, we find that inactivation of Jonah66Ci results in the modulation of immune signaling, replication and nitric oxide response in Drosophila larvae responding to Steinernema infection. We show that absence of Jonah66Ci confers partial protection to larvae against axenic nematodes. We also show that inactivation of Jonah66Ci results in the differential induction of the effector genes of the Drosophila Toll and Imd signaling in the gut of larvae responding to Steinernema symbiotic or axenic nematodes.
Finally, we show that inactivation of Jonah66Ci results in the modulation of gut specific processes including immune signaling, gut cell replication and nitric oxide levels in response to nematode attack. Our findings demonstrate a novel function for Drosophila serine protease Jonah66Ci in regulating the insect immune response to potent nematode parasites. We also find that presence of Jonah66Ci is essential in maintaining homeostasis of certain physiological processes in the gut of Drosophila larvae, particularly in the context of Steinernema nematode infection. Similar findings will pave the way towards a better understanding of the tissue-specific molecular players that modulate the insect immune response against parasitic nematodes.
FIGURE LEGENDS
Figure 1 Relative gene transcript levels of Jonah66Ci in Drosophila larvae upon infection with Steinernema nematodes. A. Relative transcript levels of Jonah66Ci using
RNA-sequencing and qRT-PCR analysis were estimated in Drosophila melanogaster late second or early third instar larvae (Oregon strain) at 6 and 24 h post-infection with 10
Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) infective juveniles. B. Relative
163 transcript levels for Jonah66Ci were estimated in the gut only and the rest of the larva in
Drosophila infected with Steinernema symbiotic or axenic nematodes. Application of water served as negative control treatment (C). Relative gene transcript levels for
Jonah66Ci were measured as a ratio compared to the uninfected control samples. Values represent the means from three separate experiments and error bars represent standard deviations. Data analysis was performed using one-way analysis of variance (ANOVA) with a Tukey’s post-hoc test on GraphPad Prism 7 software. *P<0.05, **P<0.01; non- significant differences are not shown.
Figure 2 Survival response of Drosophila Jonah66Ci mutant larvae upon infection with Steinernema nematodes. Survival rates of Drosophila melanogaster late second or early third instar yw control and Jonah66Ci mutant larvae following infection with 10
Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes. Application of water served as control (C) treatment. Survival results were monitored every 8 h and up to 72 h post-infection. Values are shown as percent survival of infected larvae and data analysis was performed using the Log-Rank (Mantel-Cox) test (GraphPad Prism7 software). The means from three independent experiments are shown and bars represent standard error.
**P<0.01, non-significant differences are not shown.
Figure 3 Transcript levels of immune pathway read-out genes in Drosophila
Jonah66Ci mutant larvae infected with Steinernema nematodes. Transcript levels of
A, B. Attacin, C, D. Drosomycin, E, F. Puckered and G, H. TotA in Drosophila
164 melanogaster yw control and Jonah66Ci mutant larvae at 24 h post-infection with 10
Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes, or treated with water
(C), at 6 and 24 h post-infection. Values are calculated relative to the housekeeping gene,
RpL32, and expressed as a ratio compared to the uninfected controls. Samples were run as technical duplicates and three biological replicates were performed. Bars represent standard deviations. Data analysis was performed using one-way analysis of variance
(ANOVA) with a Tukey’s post-hoc test on GraphPad Prism 7 software. Non-significant differences are not shown.
Figure 4 Transcript levels of Toll and Imd pathway read-out genes in the gut of
Drosophila Jonah66Ci mutant larvae infected with Steinernema nematodes.
Transcript levels of A. Drosomycin B. Defensin C. Diptericin D. Cecropin in Drosophila yw control and Jonah66Ci mutant larvae at 24 h post-infection with 10 Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes, or treated with water (C), at 6 and
24 h post-infection. Gene transcript levels are shown in gut tissue only and the rest of the larvae. Transcript level values are calculated relative to the housekeeping gene, RpL32, and are expressed as ratio compared to the uninfected control samples. Three independent experiments were performed and bars represent standard deviations. Data analysis was performed using one-way analysis of variance (ANOVA) with a Tukey’s post-hoc test on
GraphPad Prism 7 software. *P<0.05, **P<0.01, ***P<0.001; non-significant differences are not shown.
165 Figure 5 Replication in the intestinal cells of Drosophila Jonah66Ci mutant larvae infected with Steinernema nematodes. A. Relative wingless transcript levels, B.
Representative images of gut cells labeled with BrdU (red) and DAPI (blue) at 40X magnification, and C. Number of BrdU incorporated cells in the gut of Drosophila melanogaster yw control and Jonah66Ci mutant larvae at 24 h post infection with 10
Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes. Water treated larvae served as control (C). Transcript levels were estimated relative to the housekeeping gene,
RpL32, and as a ratio compared to uninfected control larvae. All experiments were repeated three times and data analysis was performed using one-way analysis of variance
(ANOVA) with a Tukey’s post-hoc test on GraphPad Prism 7. *P<0.05, **P<0.01; non- significant differences are not shown.
Figure 6 Nitric oxide levels and feeding rates of Drosophila Jonah66Ci mutant larvae infected with Steinernema nematodes. A. Relative nitric oxide (NO) levels in the gut and B. Spectrophotometric analysis of food intake in Drosophila melanogaster yw control and Jonah66Ci mutant larvae at 24 h post-infection with Steinernema carpocapsae symbiotic (Sy) or axenic (Ax) nematodes. Water treated larvae served as control (C). NO levels were measured relative to total protein. Experiments were repeated three times and analyzed using one-way analysis of variance (ANOVA) with a Tukey’s post-hoc test on GraphPad Prism 7. *P<0.05; non-significant differences are not shown.
166 A
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172 Table 1. Primers used for quantitative RT-PCR.
Accession Gene Primer Sequence (5´ – 3´) Tm (ºC) Number
Forward TTCATCACCCACGGATCTGC Jonah66Ci CG7118 57 Reverse GCACTCGGAGTTGTGGATGA
Forward CAATGGCAGACACAATCTGG Attacin-A CG10146 60 Reverse ATTCCTGGGAAGTTGCTGTG
CG10810 Forward TGAGAACCTTTTCCAATATGATG Drosomycin 60 Reverse CCAGGACCACCAGCAT
CG7850 Forward GGCCTACAAGCTGGTGAAAG Puckered 60 Reverse AGTTCAGATTGGGCGAGATG
CG31509 Forward AGATCGTGAGGCTGACAAC Turandot-A 60 Reverse CCTGGGCGTTTTTGATAA
Forward CGCATAGAAGCGAGCCACATG Defensin CG1385 60 Reverse GCAGTAGCCGCCTTTGAACC
Forward ACCGCAGTACCCACTCAATC Diptericin CG12763 60 Reverse CCCAAGTGCTGTCCATATCC
Forward TCTTCGTTTTCGTCGCTCTC Cecropin-A1 CG1365 60 Reverse CTTGTTGAGCGATTCCCAGT
Forward GATTATTCCGCAGTCTGGTC Wingless CG4889 60 Reverse CTATTATGCTTGCGTCCCTG
Forward GATGACCATCCGCCCAGCA RpL32 CG7939 60 Reverse CGGACCGACAGCTGCTTGGC
173 Dissertation General Discussion
Drosophila has emerged as a model insect host for investigating the molecular mechanisms that occur during parasitic nematode infection (Castillo et al., 2013; Dobes et al., 2012; Hallem et al., 2007), and therefore has been used in this collection of studies to investigate the immune response against infection with the entomopathogenic nematode, Steinernema carpocapsae. Steinernema nematodes are found in association with Xenorhabdus bacteria, and the ability to culture Steinernema axenic nematodes in the lab has permitted studies of the Drosophila immune response directed against the nemato-bacterial complex or the nematodes alone. The virulence of Steinernema axenic nematodes suggests that the presence of Xenorhabdus is not imperative for these nematodes to infect and kill Drosophila larvae efficiently. It is likely that Xenorhabdus contributes to the reproductive fitness of Steinernema without promoting pathogenicity of the complex (Vivas & Goodrich-Blair, 2001). Infection with Steinernema symbiotic or axenic nematodes kills Drosophila larvae at similar rates. This could be due to tissue damage caused by the migration of the nematodes in the insect body cavity or due to the production of virulence factors produced by the parasites (Bode, 2009; Castillo et al.,
2011; ffrench-Constant & Bowen, 2000).
Using the Drosophila-Steinernema infection model, I explored the participation of
Wolbachia and Spiroplasma endosymbiotic bacteria in the Drosophila immune response to entomopathogenic nematodes. Endosymbiotic bacteria are able to not only modulate physiological functions in their insect host, but also to manipulate the host immune response during infection. For instance, the presence of endosymbiotic bacteria confers
174 fitness benefits to their insect hosts D. neotestacea and Aedes polynesiensis in response to
Howardula and Brugia pahangi filarial nematodes, respectively (Andrews et al., 2012;
Jaenike & Brekke, 2011; Jaenike, Unckless, et al., 2010). Here I find that the presence of
Wolbachia and Spiroplasma promotes survival of Drosophila larvae in response to
Steinernema nematodes, however this effect was not observed when Drosophila adult flies were infected with the entomopathogenic bacterium Photorhabdus (Shokal et al.,
2016). These findings demonstrate that the endosymbiotic bacteria respond differently to the type of pathogen they encounter and this response might also be dependent on the developmental stage of Drosophila. In addition, I find that Wolbachia numbers are reduced in larvae infected with Steinernema symbiotic nematodes, but Spiroplasma numbers are unchanged. Interestingly, Spiroplasma numbers increase in Drosophila flies infected with Micrococcus luteus or Erwinia carotovora (Herren & Lemaitre, 2011), but they are not affected in Drosophila flies infected with Photorhabdus luminescens or
Escherichia coli bacteria (Shokal et al., 2016). These findings suggest a species-specific interaction between pathogens and endosymbiotic bacteria. Upon investigation of the specific host responses to nematode infections in which the endosymbiotic bacteria might participate, I observed modulation of the metabolic status, and down regulation of Toll signaling and increase in phenoloxidase levels in Drosophila larvae by Steinernema nematodes. This suggests a complex mechanism of interaction between insects, their endosymbiotic bacteria and parasitic nematodes. However, our understanding of the specific Drosophila host molecules that participate in response to nematode parasites remains unclear.
175 To gain further insights into the molecular basis of the Drosophila innate immune response to nematode infection, I analyzed the transcriptomic profile of Drosophila larvae infected with Steinernema symbiotic or axenic nematodes. I found induction of genes encoding proteins that function in the Toll and Imd signaling pathways as well as genes involved in cellular immune responses, suggesting that the Drosophila immune system is able to respond to parasitic nematodes. Transcriptomic analysis identified genes that are expressed in the peritrophic membrane and genes involved in chitin binding process are strongly induced in Drosophila larvae upon axenic nematode infection, suggesting that these genes might be involved in the Drosophila response to nematodes alone. Immune genes such as GNBP3, fondue, nimrod and croquemort are down regulated in response to Steinernema axenic nematode infection, suggesting that axenic nematodes are capable of disarming the insect immune system. In addition, induction of genes encoding proteins in the Notch and Wnt signaling pathways in response to
Steinernema nematodes suggest that these Drosophila signaling pathways, with known functions in development (Baonza & Garcia-Bellido, 2000; Bejsovec, 2013), could have a role in responding to Steinernema nematodes. I found orthologs of 55 strongly induced genes in the model lepidopteran insect host, Manduca sexta, and humans, which suggests their potential role in anti-nematode immune processes. Finally, using the Drosophila-
Steinernema infection model, I identified the imaginal disc growth factor (Idgf)-encoding genes, Idgf2 and Idgf3, as well as a gene that encodes a serine protease, Jonah66Ci, based on their upregulation in response to nematode infections, to explore their potential participation in the Drosophila immune response to nematode pathogens.
176 Drosophila IDGFs are structurally similar to chitinases but lack enzymatic activity. A murine protein, Yml, has sequence homology to Drosophila chitinases and is implicated in the immune response against nematode infections (Nair et al., 2005).
Previous studies have shown that Drosophila Idgf2 is induced in response to septic
(microbial) and aseptic injury, and IDGF3 serves as an important component in the formation of hemolymph clots in Drosophila larvae (Broz et al., 2017; Kucerova et al.,
2016). Here I find upregulation of Idgf2 and Idgf3 in Drosophila larvae responding to
Steinernema symbiotic or axenic nematodes, and loss-of-function Idgf2 mutants exhibit increased survival against axenic nematodes. Inactivation of Idgf2 strongly upregulates the Imd pathway effector gene Attacin, whereas inactivation of PGRP-LE upregulates
Idgf2 expression in response to Steinernema infection. These results indicate that Idgf2 expression and Imd signaling likely have an antagonistic relationship, in the context of nematode infection. It is also, however, likely that Imd pathway activation upon
Steinernema infection is an effect of Idgf2 inactivation, suggesting a correlation between the Imd signaling and Idgf2 expression. Furthermore, infection with Steinernema nematodes is responsible for increasing the number of hemocytes as well as upregulating the Toll pathway read-out gene Drosomycin in Idgf3 mutant larvae. Currently it remains unknown how Idgf3 modulates the Drosophila humoral and cellular immune responses against nematode infection. However, considering that IDGF3 is involved in wound repair, it is likely that it participates in hemocyte recruitment in response to nematode infections. Collectively, these findings suggest that both Idgf2 and Idgf3 have distinct roles in modulating the Drosophila anti-nematode immune response.
177 From the transcriptomic analysis, I also identified a serine protease-encoding gene, Jonah66Ci, which is strongly upregulated in Drosophila larvae infected by
Steinernema nematodes. I found that Jonah66Ci is specifically expressed in the gut of
Drosophila larvae, which is in agreement with the finding that other members of the
Jonah gene family, such as Jonah25Bi, Jonah65Ai and Jonah99Cα, are also expressed in the Drosophila gut (Carlson & Hogness, 1985a). Upon Steinernema nematode infection, inactivation of Jonah66Ci modulates specific responses in the gut of Drosophila larvae, including the differential expression of Imd and Toll signaling effector genes, reduction in the numbers of replicating gut cells, downregulation of the Wnt/Wg pathway gene wingless, and reduction of nitric oxide levels. In addition, inactivation of Jonah66Ci, similar to Idgf2, promotes the survival of larvae responding to Steinernema axenic nematodes, suggesting a potential similarity in their anti-nematode immune functions.
The potential similarity in their survival response to nematode infection could be attributed to the immune system compensating for the absence of these proteins. Notably, inactivation of Jonah66Ci strongly up regulates the expression of Attacin in response to
Steinernema symbiotic nematodes, an effect observed in Idgf2 mutants as well. These findings imply that Jonah66Ci and Idgf2 can modulate the Drosophila Imd signaling in response to Steinernema nematode infection.
In conclusion, my work dissects the molecular and functional basis of the
Drosophila immune response to the entomopathogenic nematode, Steinernema. These findings provide novel insights into the complex molecular mechanisms taking place in
Drosophila larvae infected by a multicellular eukaryotic pathogen and contribute to our understanding of how endosymbiotic bacteria influence the host immune response.
178 Further studies focusing on understanding the mechanistic basis of the interactions among Idgf2, Idgf3, Jonah66Ci, and specific molecules of the Toll and Imd signaling pathway as well as their potential contribution in the cellular immune response such as encapsulation of nematode parasites, will reveal their mode of action and contribution towards the Drosophila anti-nematode immune processes. Finally, it would be interesting to identify specific nematode molecular components to study “gene-for-gene” interactions between the insect host and nematode parasites to characterize the host anti- nematode defense. Understanding the molecular basis of the Drosophila response to nematode parasites has many advantages and applications such as development of strategies for biological control of insect pests, and even vectors of human diseases.
179
References
Adams, BJ, ; Nguyen, KB. (2002). Taxonomy and systematics. In G. R (Ed.), Entomopathogenic nematology. New York: CABI Akhurst, R. J. 1982. Antibiotic activity of Xenorhabdus spp., bacteria symbiotically associated with insect pathogenic nematodes of the families Heterorhabditidae and Steinernematidae. J Gen Microbiol, 128(12), 3061-3065. doi: 10.1099/00221287-128-12- 3061 An, C., Zhang, M., Chu, Y., & Zhao, Z. 2013. Serine protease MP2 activates prophenoloxidase in the melanization immune response of Drosophila melanogaster. PLoS One, 8(11), e79533. doi: 10.1371/journal.pone.0079533 Anand, P., Cermelli, S., Li, Z., Kassan, A., Bosch, M., Sigua, R., . . . Gross, S. P. 2012. A novel role for lipid droplets in the organismal antibacterial response. Elife, 1, e00003. doi: 10.7554/eLife.00003 Anderl, I., & Hultmark, D. 2015. New ways to make a blood cell. Elife, 4. doi: 10.7554/eLife.06877 Anders, S.; Pyl, PT. & Huber, W. 2015. HTSeq- a Python framework to work with high- throughput sequencing data. Bioinformatics, 31, 166-169. Andrews, E. S., Crain, P. R., Fu, Y., Howe, D. K., & Dobson, S. L. 2012. Reactive oxygen species production and Brugia pahangi survivorship in Aedes polynesiensis with artificial Wolbachia infection types. PLoS Pathog, 8(12), e1003075. doi: 10.1371/journal.ppat.1003075 Arakane, Y., & Muthukrishnan, S. 2010. Insect chitinase and chitinase-like proteins. Cell Mol Life Sci, 67(2), 201-216. doi: 10.1007/s00018-009-0161-9 Arefin, B., Kucerova, L., Dobes, P., Markus, R., Strnad, H., Wang, Z., . . . Theopold, U. 2014. Genome-wide transcriptional analysis of Drosophila larvae infected by entomopathogenic nematodes shows involvement of complement, recognition and extracellular matrix proteins. J Innate Immun, 6(2), 192-204. doi: 10.1159/000353734 Arefin, B., Kucerova, L., Krautz, R., Kranenburg, H., Parvin, F., & Theopold, U. 2015. Apoptosis in Hemocytes Induces a Shift in Effector Mechanisms in the Drosophila Immune System and Leads to a Pro-Inflammatory State. PLoS One, 10(8), e0136593. doi: 10.1371/journal.pone.0136593 Arnot, C. J., Gay, N. J., & Gangloff, M. 2010. Molecular mechanism that induces activation of Spatzle, the ligand for the Drosophila Toll receptor. J Biol Chem, 285(25), 19502-19509. doi: 10.1074/jbc.M109.098186 Aymeric, J. L., Givaudan, A., & Duvic, B. 2010. Imd pathway is involved in the interaction of Drosophila melanogaster with the entomopathogenic bacteria, Xenorhabdus nematophila and Photorhabdus luminescens. Mol Immunol, 47(14), 2342-2348. doi: 10.1016/j.molimm.2010.05.012 Balasubramanian, N., Toubarro, D., Nascimento, G., Ferreira, R., & Simoes, N. 2012. Purification, molecular characterization and gene expression analysis of an aspartic protease (Sc-ASP113) from the nematode Steinernema carpocapsae during the parasitic stage. Mol Biochem Parasitol, 182(1-2), 37-44. doi: 10.1016/j.molbiopara.2011.12.001 Balasubramanian, N., Toubarro, D., & Simoes, N. 2010. Biochemical study and in vitro insect immune suppression by a trypsin-like secreted protease from the nematode Steinernema carpocapsae. Parasite Immunol, 32(3), 165-175. doi: 10.1111/j.1365-3024.2009.01172.x
180 Baonza, A., & Garcia-Bellido, A. 2000. Notch signaling directly controls cell proliferation in the Drosophila wing disc. Proc Natl Acad Sci U S A, 97(6), 2609-2614. doi: 10.1073/pnas.040576497 Barillas-Mury, C., Han, Y. S., Seeley, D., & Kafatos, F. C. 1999. Anopheles gambiae Ag-STAT, a new insect member of the STAT family, is activated in response to bacterial infection. EMBO J, 18(4), 959-967. doi: 10.1093/emboj/18.4.959 Bejsovec, A. 2013. Wingless/Wnt signaling in Drosophila: the pattern and the pathway. Mol Reprod Dev, 80(11), 882-894. doi: 10.1002/mrd.22228 Bidla, G., Lindgren, M., Theopold, U., & Dushay, M. S. 2005. Hemolymph coagulation and phenoloxidase in Drosophila larvae. Dev Comp Immunol, 29(8), 669-679. doi: 10.1016/j.dci.2004.11.007 Bier, E., & Guichard, A. 2012. Deconstructing host-pathogen interactions in Drosophila. Dis Model Mech, 5(1), 48-61. doi: 10.1242/dmm.000406 Binggeli, O., Neyen, C., Poidevin, M., & Lemaitre, B. 2014. Prophenoloxidase activation is required for survival to microbial infections in Drosophila. PLoS Pathog, 10(5), e1004067. doi: 10.1371/journal.ppat.1004067 Bischoff, V., Vignal, C., Duvic, B., Boneca, I. G., Hoffmann, J. A., & Royet, J. 2006. Downregulation of the Drosophila immune response by peptidoglycan-recognition proteins SC1 and SC2. PLoS Pathog, 2(2), e14. doi: 10.1371/journal.ppat.0020014 Blagrove, M. S., Arias-Goeta, C., Failloux, A. B., & Sinkins, S. P. 2012. Wolbachia strain wMel induces cytoplasmic incompatibility and blocks dengue transmission in Aedes albopictus. Proc Natl Acad Sci U S A, 109(1), 255-260. doi: 10.1073/pnas.1112021108 Bode, H. B. 2009. Entomopathogenic bacteria as a source of secondary metabolites. Curr Opin Chem Biol, 13(2), 224-230. doi: 10.1016/j.cbpa.2009.02.037 Bou Aoun, R., Hetru, C., Troxler, L., Doucet, D., Ferrandon, D., & Matt, N. 2011. Analysis of thioester-containing proteins during the innate immune response of Drosophila melanogaster. J Innate Immun, 3(1), 52-64. doi: 10.1159/000321554 Brennan, C. A., & Anderson, K. V. 2004. Drosophila: the genetics of innate immune recognition and response. Annu Rev Immunol, 22, 457-483. doi: 10.1146/annurev.immunol.22.012703.104626 Brivio, M. F., Mastore, M., & Moro, M. 2004. The role of Steinernema feltiae body-surface lipids in host-parasite immunological interactions. Mol Biochem Parasitol, 135(1), 111-121. Brivio MF, Mastore M, Pagani M. 2005. Parasite-host relationship:a lesson from a professional killer. Invertebr Surv J, 2, 41-53. Broz, V., Kucerova, L., Rouhova, L., Fleischmannova, J., Strnad, H., Bryant, P. J., & Zurovec, M. 2017. Drosophila imaginal disc growth factor 2 is a trophic factor involved in energy balance, detoxification, and innate immunity. Sci Rep, 7, 43273. doi: 10.1038/srep43273 Brun, S., Vidal, S., Spellman, P., Takahashi, K., Tricoire, H., & Lemaitre, B. 2006. The MAPKKK Mekk1 regulates the expression of Turandot stress genes in response to septic injury in Drosophila. Genes Cells, 11(4), 397-407. doi: 10.1111/j.1365- 2443.2006.00953.x Buchner, Paul. (1965). Endosymbiosis of animals with plant microorganisms. New York, London, Sydney: Interscience Publishers. A Division of John Wiley. Buchon, N., Broderick, N. A., Poidevin, M., Pradervand, S., & Lemaitre, B. 2009. Drosophila intestinal response to bacterial infection: activation of host defense and stem cell proliferation. Cell Host Microbe, 5(2), 200-211. doi: 10.1016/j.chom.2009.01.003 Buchon, N., Poidevin, M., Kwon, H. M., Guillou, A., Sottas, V., Lee, B. L., & Lemaitre, B. 2009. A single modular serine protease integrates signals from pattern-recognition receptors upstream of the Drosophila Toll pathway. Proc Natl Acad Sci U S A, 106(30), 12442- 12447. doi: 10.1073/pnas.0901924106
181 Buchon, N., Silverman, N., & Cherry, S. 2014. Immunity in Drosophila melanogaster--from microbial recognition to whole-organism physiology. Nat Rev Immunol, 14(12), 796-810. doi: 10.1038/nri3763 Bussink, A. P., Speijer, D., Aerts, J. M., & Boot, R. G. 2007. Evolution of mammalian chitinase(- like) members of family 18 glycosyl hydrolases. Genetics, 177(2), 959-970. doi: 10.1534/genetics.107.075846 Carlson, J. R., & Hogness, D. S. 1985a. Developmental and functional analysis of Jonah gene expression. Dev Biol, 108(2), 355-368. Carlson, J. R., & Hogness, D. S. 1985b. The Jonah genes: a new multigene family in Drosophila melanogaster. Dev Biol, 108(2), 341-354. Carpenter, J., Hutter, S., Baines, J. F., Roller, J., Saminadin-Peter, S. S., Parsch, J., & Jiggins, F. M. 2009. The transcriptional response of Drosophila melanogaster to infection with the sigma virus (Rhabdoviridae). PLoS One, 4(8), e6838. doi: 10.1371/journal.pone.0006838 Carton, Y., Frey, F., & Nappi, A. J. 2009. Parasite-induced changes in nitric oxide levels in Drosophila paramelanica. J Parasitol, 95(5), 1134-1141. doi: 10.1645/GE-2091.1 Carvalho, L., Jacinto, A., & Matova, N. 2014. The Toll/NF-kappaB signaling pathway is required for epidermal wound repair in Drosophila. Proc Natl Acad Sci U S A, 111(50), E5373- 5382. doi: 10.1073/pnas.1408224111 Casanova-Torres, A. M., & Goodrich-Blair, H. 2013. Immune Signaling and Antimicrobial Peptide Expression in Lepidoptera. Insects, 4(3), 320-338. doi: 10.3390/insects4030320 Castillo, J. C., Creasy, T., Kumari, P., Shetty, A., Shokal, U., Tallon, L. J., & Eleftherianos, I. 2015. Drosophila anti-nematode and antibacterial immune regulators revealed by RNA- Seq. BMC Genomics, 16, 519. doi: 10.1186/s12864-015-1690-2 Castillo, J. C., Reynolds, S. E., & Eleftherianos, I. 2011. Insect immune responses to nematode parasites. Trends Parasitol, 27(12), 537-547. doi: 10.1016/j.pt.2011.09.001 Castillo, J. C., Shokal, U., & Eleftherianos, I. 2012. A novel method for infecting Drosophila adult flies with insect pathogenic nematodes. Virulence, 3(3), 339-347. doi: 10.4161/viru.20244 Castillo, J. C., Shokal, U., & Eleftherianos, I. 2013. Immune gene transcription in Drosophila adult flies infected by entomopathogenic nematodes and their mutualistic bacteria. J Insect Physiol, 59(2), 179-185. doi: 10.1016/j.jinsphys.2012.08.003 Cerenius, L., & Soderhall, K. 2004. The prophenoloxidase-activating system in invertebrates. Immunol Rev, 198, 116-126. Chartier, A., Zaffran, S., Astier, M., Semeriva, M., & Gratecos, D. 2002. Pericardin, a Drosophila type IV collagen-like protein is involved in the morphogenesis and maintenance of the heart epithelium during dorsal ectoderm closure. Development, 129(13), 3241-3253. Chaston, J. M., Suen, G., Tucker, S. L., Andersen, A. W., Bhasin, A., Bode, E., . . . Goodrich- Blair, H. 2011. The entomopathogenic bacterial endosymbionts Xenorhabdus and Photorhabdus: convergent lifestyles from divergent genomes. PLoS One, 6(11), e27909. doi: 10.1371/journal.pone.0027909 Choubey, P. K., & Roy, J. K. 2017. Rab11, a vesicular trafficking protein, affects endoreplication through Ras-mediated pathway in Drosophila melanogaster. Cell Tissue Res, 367(2), 269- 282. doi: 10.1007/s00441-016-2500-0 Chtarbanova, S., Lamiable, O., Lee, K. Z., Galiana, D., Troxler, L., Meignin, C., . . . Imler, J. L. 2014. Drosophila C virus systemic infection leads to intestinal obstruction. J Virol, 88(24), 14057-14069. doi: 10.1128/JVI.02320-14 Ciche, T. 2007. The biology and genome of Heterorhabditis bacteriophora. WormBook, 1-9. doi: 10.1895/wormbook.1.135.1 Ciche, T. A., Kim, K. S., Kaufmann-Daszczuk, B., Nguyen, K. C., & Hall, D. H. 2008. Cell Invasion and Matricide during Photorhabdus luminescens Transmission by
182 Heterorhabditis bacteriophora Nematodes. Appl Environ Microbiol, 74(8), 2275-2287. doi: 10.1128/AEM.02646-07 Ciche, T., Darby, C., Ehlers, R. U., Forst, S., & Goodrich-Blair, H. 2006. Dangerous liasions: The symbiosis of entomopathogenic nematodes and bacteria. Biological Control, 38, 22-46. Clarke, D. J. 2014. The genetic basis of the symbiosis between Photorhabdus and its invertebrate hosts. Adv Appl Microbiol, 88, 1-29. doi: 10.1016/B978-0-12-800260-5.00001-2 Colasanti, M., Gradoni, L., Mattu, M., Persichini, T., Salvati, L., Venturini, G., & Ascenzi, P. 2002. Molecular bases for the anti-parasitic effect of NO (Review). Int J Mol Med, 9(2), 131-134. Colinet, D., Dubuffet, A., Cazes, D., Moreau, S., Drezen, J. M., & Poirie, M. 2009. A serpin from the parasitoid wasp Leptopilina boulardi targets the Drosophila phenoloxidase cascade. Dev Comp Immunol, 33(5), 681-689. doi: 10.1016/j.dci.2008.11.013 Cooper, D., & Eleftherianos, I. 2016. Parasitic Nematode Immunomodulatory Strategies: Recent Advances and Perspectives. Pathogens, 5(3). doi: 10.3390/pathogens5030058 Costa, A., Jan, E., Sarnow, P., & Schneider, D. 2009. The Imd pathway is involved in antiviral immune responses in Drosophila. PLoS One, 4(10), e7436. doi: 10.1371/journal.pone.0007436 Dale, C., & Moran, N. A. 2006. Molecular interactions between bacterial symbionts and their hosts. Cell, 126(3), 453-465. doi: 10.1016/j.cell.2006.07.014 De Gregorio, E., Spellman, P. T., Rubin, G. M., & Lemaitre, B. 2001. Genome-wide analysis of the Drosophila immune response by using oligonucleotide microarrays. Proc Natl Acad Sci U S A, 98(22), 12590-12595. doi: 10.1073/pnas.221458698 De Gregorio, E., Spellman, P. T., Tzou, P., Rubin, G. M., & Lemaitre, B. 2002. The Toll and Imd pathways are the major regulators of the immune response in Drosophila. EMBO J, 21(11), 2568-2579. doi: 10.1093/emboj/21.11.2568 Derer, A., Bohm, C., Grotsch, B., Grun, J. R., Grutzkau, A., Stock, M., . . . David, J. P. 2016. Rsk2 controls synovial fibroblast hyperplasia and the course of arthritis. Ann Rheum Dis, 75(2), 413-421. doi: 10.1136/annrheumdis-2014-205618 Di Cera, E. 2009. Serine proteases. IUBMB Life, 61(5), 510-515. doi: 10.1002/iub.186 Dillman, A. R., Chaston, J. M., Adams, B. J., Ciche, T. A., Goodrich-Blair, H., Stock, S. P., & Sternberg, P. W. 2012. An entomopathogenic nematode by any other name. PLoS Pathog, 8(3), e1002527. doi: 10.1371/journal.ppat.1002527 Dillman, A. R., & Sternberg, P. W. 2012. Entomopathogenic nematodes. Curr Biol, 22(11), R430-431. doi: 10.1016/j.cub.2012.03.047 Dionne, M. S., & Schneider, D. S. 2008. Models of infectious diseases in the fruit fly Drosophila melanogaster. Dis Model Mech, 1(1), 43-49. doi: 10.1242/dmm.000307 Dobes, P., Wang, Z., Markus, R., Theopold, U., & Hyrsl, P. 2012. An improved method for nematode infection assays in Drosophila larvae. Fly (Austin), 6(2), 75-79. doi: 10.4161/fly.19553 Drechsler, M., Schmidt, A. C., Meyer, H., & Paululat, A. 2013. The conserved ADAMTS-like protein lonely heart mediates matrix formation and cardiac tissue integrity. PLoS Genet, 9(7), e1003616. doi: 10.1371/journal.pgen.1003616 Dudzic, J. P., Kondo, S., Ueda, R., Bergman, C. M., & Lemaitre, B. 2015. Drosophila innate immunity: regional and functional specialization of prophenoloxidases. BMC Biol, 13, 81. doi: 10.1186/s12915-015-0193-6 Dutton, T. J., & Sinkins, S. P. 2005. Filarial susceptibility and effects of Wolbachia in Aedes pseudoscutellaris mosquitoes. Med Vet Entomol, 19(1), 60-65. doi: 10.1111/j.0269- 283X.2005.00557.x Duvic, B., Hoffmann, J. A., Meister, M., & Royet, J. 2002. Notch signaling controls lineage specification during Drosophila larval hematopoiesis. Curr Biol, 12(22), 1923-1927.
183 Dziarski, R., & Gupta, D. 2006. The peptidoglycan recognition proteins (PGRPs). Genome Biol, 7(8), 232. doi: 10.1186/gb-2006-7-8-232 Ebrahimi, L., Niknam, G., & Dunphy, G. B. 2011. Hemocyte responses of the Colorado potato beetle, Leptinotarsa decemlineata, and the greater wax moth, Galleria mellonella, to the entomopathogenic nematodes, Steinernema feltiae and Heterorhabditis bacteriophora. J Insect Sci, 11, 75. doi: 10.1673/031.011.7501 Ekengren, S., & Hultmark, D. 2001. A family of Turandot-related genes in the humoral stress response of Drosophila. Biochem Biophys Res Commun, 284(4), 998-1003. doi: 10.1006/bbrc.2001.5067 Ekengren, S., Tryselius, Y., Dushay, M. S., Liu, G., Steiner, H., & Hultmark, D. 2001. A humoral stress response in Drosophila. Curr Biol, 11(18), 1479. El Chamy, L., Leclerc, V., Caldelari, I., & Reichhart, J. M. 2008. Sensing of 'danger signals' and pathogen-associated molecular patterns defines binary signaling pathways 'upstream' of Toll. Nat Immunol, 9(10), 1165-1170. doi: 10.1038/ni.1643 Eleftherianos, I., Atri, J., Accetta, J., & Castillo, J. C. 2013. Endosymbiotic bacteria in insects: guardians of the immune system? Front Physiol, 4, 46. doi: 10.3389/fphys.2013.00046 Eleftherianos, I., Castillo, J. C., & Patrnogic, J. 2016. TGF-beta signaling regulates resistance to parasitic nematode infection in Drosophila melanogaster. Immunobiology, 221(12), 1362- 1368. doi: 10.1016/j.imbio.2016.07.011 Eleftherianos, I., ffrench-Constant, R. H., Clarke, D. J., Dowling, A. J., & Reynolds, S. E. 2010. Dissecting the immune response to the entomopathogen Photorhabdus. Trends Microbiol, 18(12), 552-560. doi: 10.1016/j.tim.2010.09.006 Eleftherianos, I., Joyce, S., Ffrench-Constant, R. H., Clarke, D. J., & Reynolds, S. E. 2010. Probing the tri-trophic interaction between insects, nematodes and Photorhabdus. Parasitology, 137(11), 1695-1706. doi: 10.1017/S0031182010000508 Eleftherianos, I., More, K., Spivack, S., Paulin, E., Khojandi, A., & Shukla, S. 2014. Nitric oxide levels regulate the immune response of Drosophila melanogaster reference laboratory strains to bacterial infections. Infect Immun, 82(10), 4169-4181. doi: 10.1128/IAI.02318- 14 Eleftherianos, I., & Revenis, C. 2011. Role and importance of phenoloxidase in insect hemostasis. J Innate Immun, 3(1), 28-33. doi: 10.1159/000321931 Eleftherianos, I., Yadav, S., Kenney, E., Cooper, D., Ozakman, Y., & Patrnogic, J. 2017. Role of Endosymbionts in Insect-Parasitic Nematode Interactions. Trends Parasitol. doi: 10.1016/j.pt.2017.10.004 Erickson, S. M., Xi, Z., Mayhew, G. F., Ramirez, J. L., Aliota, M. T., Christensen, B. M., & Dimopoulos, G. 2009. Mosquito infection responses to developing filarial worms. PLoS Negl Trop Dis, 3(10), e529. doi: 10.1371/journal.pntd.0000529 Erkosar, B., Defaye, A., Bozonnet, N., Puthier, D., Royet, J., & Leulier, F. 2014. Drosophila microbiota modulates host metabolic gene expression via IMD/NF-kappaB signaling. PLoS One, 9(4), e94729. doi: 10.1371/journal.pone.0094729 Fearon, D. T., & Locksley, R. M. 1996. The instructive role of innate immunity in the acquired immune response. Science, 272(5258), 50-53. Feder, M. E., & Hofmann, G. E. 1999. Heat-shock proteins, molecular chaperones, and the stress response: evolutionary and ecological physiology. Annu Rev Physiol, 61, 243-282. doi: 10.1146/annurev.physiol.61.1.243 Fernandez-Ayala, D. J., Chen, S., Kemppainen, E., O'Dell, K. M., & Jacobs, H. T. 2010. Gene expression in a Drosophila model of mitochondrial disease. PLoS One, 5(1), e8549. doi: 10.1371/journal.pone.0008549 Ferrandon, D., Imler, J. L., Hetru, C., & Hoffmann, J. A. 2007. The Drosophila systemic immune response: sensing and signalling during bacterial and fungal infections. Nat Rev Immunol, 7(11), 862-874. doi: 10.1038/nri2194
184 Feyzkhanova, G., Voloshin, S., Smoldovskaya, O., Arefieva, A., Filippova, M., Barsky, V., . . . Rubina, A. 2017. Development of a microarray-based method for allergen-specific IgE and IgG4 detection. Clin Proteomics, 14, 1. doi: 10.1186/s12014-016-9136-7 ffrench-Constant, R. H., & Bowen, D. J. 2000. Novel insecticidal toxins from nematode- symbiotic bacteria. Cell Mol Life Sci, 57(5), 828-833. doi: 10.1007/s000180050044 Foley, E., & O'Farrell, P. H. 2003. Nitric oxide contributes to induction of innate immune responses to gram-negative bacteria in Drosophila. Genes Dev, 17(1), 115-125. doi: 10.1101/gad.1018503 Forst, S., Dowds, B., Boemare, N., & Stackebrandt, E. 1997. Xenorhabdus and Photorhabdus spp.: bugs that kill bugs. Annu Rev Microbiol, 51, 47-72. doi: 10.1146/annurev.micro.51.1.47 Fytrou, A., Schofield, P. G., Kraaijeveld, A. R., & Hubbard, S. F. 2006. Wolbachia infection suppresses both host defence and parasitoid counter-defence. Proc Biol Sci, 273(1588), 791-796. doi: 10.1098/rspb.2005.3383 Gajewski, K. M., Sorrentino, R. P., Lee, J. H., Zhang, Q., Russell, M., & Schulz, R. A. 2007. Identification of a crystal cell-specific enhancer of the black cells prophenoloxidase gene in Drosophila. Genesis, 45(4), 200-207. doi: 10.1002/dvg.20285 Garver, L. S., Wu, J., & Wu, L. P. 2006. The peptidoglycan recognition protein PGRP-SC1a is essential for Toll signaling and phagocytosis of Staphylococcus aureus in Drosophila. Proc Natl Acad Sci U S A, 103(3), 660-665. doi: 10.1073/pnas.0506182103 Gaugler, R. and Kaya, H. . 1995. Entomopathogenic nematodes in biological control of insects. J Nematol, 27, 529-534. Ghosh, S., & Chan, C. K. 2016. Analysis of RNA-Seq Data Using TopHat and Cufflinks. Methods Mol Biol, 1374, 339-361. doi: 10.1007/978-1-4939-3167-5_18 Glaser, R. L., & Meola, M. A. 2010. The native Wolbachia endosymbionts of Drosophila melanogaster and Culex quinquefasciatus increase host resistance to West Nile virus infection. PLoS One, 5(8), e11977. doi: 10.1371/journal.pone.0011977 Goodrich-Blair, H. 2007. They've got a ticket to ride: Xenorhabdus nematophila-Steinernema carpocapsae symbiosis. Curr Opin Microbiol, 10(3), 225-230. doi: 10.1016/j.mib.2007.05.006 Goodrich-Blair, H., & Clarke, D. J. 2007. Mutualism and pathogenesis in Xenorhabdus and Photorhabdus: two roads to the same destination. Mol Microbiol, 64(2), 260-268. doi: 10.1111/j.1365-2958.2007.05671.x Gottar, M., Gobert, V., Matskevich, A. A., Reichhart, J. M., Wang, C., Butt, T. M., . . . Ferrandon, D. 2006. Dual detection of fungal infections in Drosophila via recognition of glucans and sensing of virulence factors. Cell, 127(7), 1425-1437. doi: 10.1016/j.cell.2006.10.046 Govind, S. 2008. Innate immunity in Drosophila: Pathogens and pathways. Insect Sci, 15(1), 29- 43. doi: 10.1111/j.1744-7917.2008.00185.x Habayeb, M. S., Ekengren, S. K., & Hultmark, D. 2006. Nora virus, a persistent virus in Drosophila, defines a new picorna-like virus family. J Gen Virol, 87(Pt 10), 3045-3051. doi: 10.1099/vir.0.81997-0 Hafen, E., Levine, M., Garber, R. L., & Gehring, W. J. 1983. An improved in situ hybridization method for the detection of cellular RNAs in Drosophila tissue sections and its application for localizing transcripts of the homeotic Antennapedia gene complex. EMBO J, 2(4), 617-623. Hallem, E. A., Dillman, A. R., Hong, A. V., Zhang, Y., Yano, J. M., DeMarco, S. F., & Sternberg, P. W. 2011. A sensory code for host seeking in parasitic nematodes. Curr Biol, 21(5), 377-383. doi: 10.1016/j.cub.2011.01.048
185 Hallem, E. A., Rengarajan, M., Ciche, T. A., & Sternberg, P. W. 2007. Nematodes, bacteria, and flies: a tripartite model for nematode parasitism. Curr Biol, 17(10), 898-904. doi: 10.1016/j.cub.2007.04.027 Hamilton, P. T., Peng, F., Boulanger, M. J., & Perlman, S. J. 2016. A ribosome-inactivating protein in a Drosophila defensive symbiont. Proc Natl Acad Sci U S A, 113(2), 350-355. doi: 10.1073/pnas.1518648113 Han, R., & Ehlers, R. U. 2000. Pathogenicity, development, and reproduction of Heterorhabditis bacteriophora and Steinernema carpocapsae under axenic in vivo conditions. J Invertebr Pathol, 75(1), 55-58. doi: 10.1006/jipa.1999.4900 Hao, Y. J., Montiel, R., Abubucker, S., Mitreva, M., & Simoes, N. 2010. Transcripts analysis of the entomopathogenic nematode Steinernema carpocapsae induced in vitro with insect haemolymph. Mol Biochem Parasitol, 169(2), 79-86. doi: 10.1016/j.molbiopara.2009.10.002 Hawdon, J. M. 2014. Controlling soil-transmitted helminths: time to think inside the box? J Parasitol, 100(2), 166-188. doi: 10.1645/13-412.1 Hedges, L. M., Brownlie, J. C., O'Neill, S. L., & Johnson, K. N. 2008. Wolbachia and virus protection in insects. Science, 322(5902), 702. doi: 10.1126/science.1162418 Hegedus, D., Erlandson, M., Gillott, C., & Toprak, U. 2009. New insights into peritrophic matrix synthesis, architecture, and function. Annu Rev Entomol, 54, 285-302. doi: 10.1146/annurev.ento.54.110807.090559 Herbert, E. E., & Goodrich-Blair, H. 2007. Friend and foe: the two faces of Xenorhabdus nematophila. Nat Rev Microbiol, 5(8), 634-646. doi: 10.1038/nrmicro1706 Herren, J. K., & Lemaitre, B. 2011. Spiroplasma and host immunity: activation of humoral immune responses increases endosymbiont load and susceptibility to certain Gram- negative bacterial pathogens in Drosophila melanogaster. Cell Microbiol, 13(9), 1385- 1396. doi: 10.1111/j.1462-5822.2011.01627.x Herren, J. K., Paredes, J. C., Schupfer, F., Arafah, K., Bulet, P., & Lemaitre, B. 2014. Insect endosymbiont proliferation is limited by lipid availability. Elife, 3, e02964. doi: 10.7554/eLife.02964 Hetru, C., & Hoffmann, J. A. 2009. NF-kappaB in the immune response of Drosophila. Cold Spring Harb Perspect Biol, 1(6), a000232. doi: 10.1101/cshperspect.a000232 Hetru, C., Troxler, L., & Hoffmann, J. A. 2003. Drosophila melanogaster antimicrobial defense. J Infect Dis, 187 Suppl 2, S327-334. doi: 10.1086/374758 Hoffmann, J. A., & Reichhart, J. M. 2002. Drosophila innate immunity: an evolutionary perspective. Nat Immunol, 3(2), 121-126. doi: 10.1038/ni0202-121 Homer, R. J., Zhu, Z., Cohn, L., Lee, C. G., White, W. I., Chen, S., & Elias, J. A. 2006. Differential expression of chitinases identify subsets of murine airway epithelial cells in allergic inflammation. Am J Physiol Lung Cell Mol Physiol, 291(3), L502-511. doi: 10.1152/ajplung.00364.2005 Horng, T., & Medzhitov, R. 2001. Drosophila MyD88 is an adapter in the Toll signaling pathway. Proc Natl Acad Sci U S A, 98(22), 12654-12658. doi: 10.1073/pnas.231471798 Hotez, PJ.; Fenwick, A.; Savioli, L. & Molyneux, DH. 2009. Rescuing the bottom billion through control of neglected tropical diseases. Lancet, 373, 1570-1575. Huang da, W., Sherman, B. T., & Lempicki, R. A. 2009a. Bioinformatics enrichment tools: paths toward the comprehensive functional analysis of large gene lists. Nucleic Acids Res, 37(1), 1-13. doi: 10.1093/nar/gkn923 Huang da, W., Sherman, B. T., & Lempicki, R. A. 2009b. Systematic and integrative analysis of large gene lists using DAVID bioinformatics resources. Nat Protoc, 4(1), 44-57. doi: 10.1038/nprot.2008.211
186 Hughes, G. L., Koga, R., Xue, P., Fukatsu, T., & Rasgon, J. L. 2011. Wolbachia infections are virulent and inhibit the human malaria parasite Plasmodium falciparum in Anopheles gambiae. PLoS Pathog, 7(5), e1002043. doi: 10.1371/journal.ppat.1002043 Hyrsl, P., Dobes, P., Wang, Z., Hauling, T., Wilhelmsson, C., & Theopold, U. 2011. Clotting factors and eicosanoids protect against nematode infections. J Innate Immun, 3(1), 65-70. doi: 10.1159/000320634 Ikeya, T., Broughton, S., Alic, N., Grandison, R., & Partridge, L. 2009. The endosymbiont Wolbachia increases insulin/IGF-like signalling in Drosophila. Proc Biol Sci, 276(1674), 3799-3807. doi: 10.1098/rspb.2009.0778 Imler, J. L., & Bulet, P. 2005. Antimicrobial peptides in Drosophila: structures, activities and gene regulation. Chem Immunol Allergy, 86, 1-21. doi: 10.1159/000086648 Irving, P., Troxler, L., Heuer, T. S., Belvin, M., Kopczynski, C., Reichhart, J. M., . . . Hetru, C. 2001. A genome-wide analysis of immune responses in Drosophila. Proc Natl Acad Sci U S A, 98(26), 15119-15124. doi: 10.1073/pnas.261573998 Irving, P., Ubeda, J. M., Doucet, D., Troxler, L., Lagueux, M., Zachary, D., . . . Meister, M. 2005. New insights into Drosophila larval haemocyte functions through genome-wide analysis. Cell Microbiol, 7(3), 335-350. doi: 10.1111/j.1462-5822.2004.00462.x Ishii, K., Adachi, T., Hamamoto, H., Oonishi, T., Kamimura, M., Imamura, K., & Sekimizu, K. 2013. Insect cytokine paralytic peptide activates innate immunity via nitric oxide production in the silkworm Bombyx mori. Dev Comp Immunol, 39(3), 147-153. doi: 10.1016/j.dci.2012.10.014 Jaenike, J. 2012. Population genetics of beneficial heritable symbionts. Trends Ecol Evol, 27(4), 226-232. doi: 10.1016/j.tree.2011.10.005 Jaenike, J., & Brekke, T. D. 2011. Defensive endosymbionts: a cryptic trophic level in community ecology. Ecol Lett, 14(2), 150-155. doi: 10.1111/j.1461-0248.2010.01564.x Jaenike, J., Stahlhut, J. K., Boelio, L. M., & Unckless, R. L. 2010. Association between Wolbachia and Spiroplasma within Drosophila neotestacea: an emerging symbiotic mutualism? Mol Ecol, 19(2), 414-425. doi: 10.1111/j.1365-294X.2009.04448.x Jaenike, J., Unckless, R., Cockburn, S. N., Boelio, L. M., & Perlman, S. J. 2010. Adaptation via symbiosis: recent spread of a Drosophila defensive symbiont. Science, 329(5988), 212- 215. doi: 10.1126/science.1188235 Janeway, C. A., Jr., & Medzhitov, R. 2002. Innate immune recognition. Annu Rev Immunol, 20, 197-216. doi: 10.1146/annurev.immunol.20.083001.084359 Jang, I. H., Chosa, N., Kim, S. H., Nam, H. J., Lemaitre, B., Ochiai, M., . . . Lee, W. J. 2006. A Spatzle-processing enzyme required for toll signaling activation in Drosophila innate immunity. Dev Cell, 10(1), 45-55. doi: 10.1016/j.devcel.2005.11.013 Janssens, S., & Beyaert, R. 2003. Role of Toll-like receptors in pathogen recognition. Clin Microbiol Rev, 16(4), 637-646. Jarosz, J. 1998. Active resistance of entomophagous rhabditid Heterorhabditis bacteriophora to insect immunity. Parasitology, 117 ( Pt 3), 201-208. Jung, S. H., Evans, C. J., Uemura, C., & Banerjee, U. 2005. The Drosophila lymph gland as a developmental model of hematopoiesis. Development, 132(11), 2521-2533. doi: 10.1242/dev.01837 Kambris, Z., Brun, S., Jang, I. H., Nam, H. J., Romeo, Y., Takahashi, K., . . . Lemaitre, B. 2006. Drosophila immunity: a large-scale in vivo RNAi screen identifies five serine proteases required for Toll activation. Curr Biol, 16(8), 808-813. doi: 10.1016/j.cub.2006.03.020 Kaneko, T., Golenbock, D., & Silverman, N. 2005. Peptidoglycan recognition by the Drosophila Imd pathway. J Endotoxin Res, 11(6), 383-389. doi: 10.1179/096805105X76823 Kaneko, T., & Silverman, N. 2005. Bacterial recognition and signalling by the Drosophila IMD pathway. Cell Microbiol, 7(4), 461-469. doi: 10.1111/j.1462-5822.2005.00504.x
187 Karlsson, C., Korayem, A. M., Scherfer, C., Loseva, O., Dushay, M. S., & Theopold, U. 2004. Proteomic analysis of the Drosophila larval hemolymph clot. J Biol Chem, 279(50), 52033-52041. doi: 10.1074/jbc.M408220200 Kaun, K. R., Hendel, T., Gerber, B., & Sokolowski, M. B. 2007. Natural variation in Drosophila larval reward learning and memory due to a cGMP-dependent protein kinase. Learn Mem, 14(5), 342-349. doi: 10.1101/lm.505807 Kawamura, K., Shibata, T., Saget, O., Peel, D., & Bryant, P. J. 1999. A new family of growth factors produced by the fat body and active on Drosophila imaginal disc cells. Development, 126(2), 211-219. Keebaugh, E. S.& Schlenke, T. A. 2014. Insights from natural host-parasite interactions: the Drosophila model. Dev. Comp. Immunol. 42: 111-123. Kemp, C., Mueller, S., Goto, A., Barbier, V., Paro, S., Bonnay, F., . . . Imler, J. L. 2013. Broad RNA interference-mediated antiviral immunity and virus-specific inducible responses in Drosophila. J Immunol, 190(2), 650-658. doi: 10.4049/jimmunol.1102486 Kikuchi, Y. 2009. Endosymbiotic bacteria in insects: their diversity and culturability. Microbes Environ, 24(3), 195-204. Kingsolver, M. B., Huang, Z., & Hardy, R. W. 2013. Insect antiviral innate immunity: pathways, effectors, and connections. J Mol Biol, 425(24), 4921-4936. doi: 10.1016/j.jmb.2013.10.006 Kirkpatrick, R. B., Matico, R. E., McNulty, D. E., Strickler, J. E., & Rosenberg, M. 1995. An abundantly secreted glycoprotein from Drosophila melanogaster is related to mammalian secretory proteins produced in rheumatoid tissues and by activated macrophages. Gene, 153(2), 147-154. Kisseleva, T., Bhattacharya, S., Braunstein, J., & Schindler, C. W. 2002. Signaling through the JAK/STAT pathway, recent advances and future challenges. Gene, 285(1-2), 1-24. Klager, S. L., Watson, A., Achukwi, D., Hultmark, D., & Hagen, H. E. 2002. Humoral immune response of Simulium damnosum s.l. following filarial and bacterial infections. Parasitology, 125(Pt 4), 359-366. Kleino, A., & Silverman, N. 2014. The Drosophila IMD pathway in the activation of the humoral immune response. Dev Comp Immunol, 42(1), 25-35. doi: 10.1016/j.dci.2013.05.014 Kounatidis, I., & Ligoxygakis, P. 2012. Drosophila as a model system to unravel the layers of innate immunity to infection. Open Biol, 2(5), 120075. doi: 10.1098/rsob.120075 Krautz, R., Arefin, B., & Theopold, U. 2014. Damage signals in the insect immune response. Front Plant Sci, 5, 342. doi: 10.3389/fpls.2014.00342 Krecek, R. C., & Waller, P. J. 2006. Towards the implementation of the "basket of options" approach to helminth parasite control of livestock: emphasis on the tropics/subtropics. Vet Parasitol, 139(4), 270-282. doi: 10.1016/j.vetpar.2006.04.018 Kremer, N., Charif, D., Henri, H., Gavory, F., Wincker, P., Mavingui, P., & Vavre, F. 2012. Influence of Wolbachia on host gene expression in an obligatory symbiosis. BMC Microbiol, 12 Suppl 1, S7. doi: 10.1186/1471-2180-12-S1-S7 Kucerova, L., Broz, V., Arefin, B., Maaroufi, H. O., Hurychova, J., Strnad, H., . . . Theopold, U. 2016. The Drosophila Chitinase-Like Protein IDGF3 Is Involved in Protection against Nematodes and in Wound Healing. J Innate Immun, 8(2), 199-210. doi: 10.1159/000442351 Kumari, P., Kant, S., Zaman, S., Mahapatro, G. K., Banerjee, N., & Sarin, N. B. 2014. A novel insecticidal GroEL protein from Xenorhabdus nematophila confers insect resistance in tobacco. Transgenic Res, 23(1), 99-107. doi: 10.1007/s11248-013-9734-3 Kuraishi, T., Binggeli, O., Opota, O., Buchon, N., & Lemaitre, B. 2011. Genetic evidence for a protective role of the peritrophic matrix against intestinal bacterial infection in Drosophila melanogaster. Proc Natl Acad Sci U S A, 108(38), 15966-15971. doi: 10.1073/pnas.1105994108
188 Kurucz, E., Zettervall, C. J., Sinka, R., Vilmos, P., Pivarcsi, A., Ekengren, S., . . . Hultmark, D. 2003. Hemese, a hemocyte-specific transmembrane protein, affects the cellular immune response in Drosophila. Proc Natl Acad Sci U S A, 100(5), 2622-2627. doi: 10.1073/pnas.0436940100 Lacey, L. A., & Georgis, R. 2012. Entomopathogenic nematodes for control of insect pests above and below ground with comments on commercial production. J Nematol, 44(2), 218-225. Lacey, L. A., Grzywacz, D., Shapiro-Ilan, D. I., Frutos, R., Brownbridge, M., & Goettel, M. S. 2015. Insect pathogens as biological control agents: Back to the future. J Invertebr Pathol, 132, 1-41. doi: 10.1016/j.jip.2015.07.009 Lackie, AM. 1988. Haemocyte Behaviour. Advances in Insect Physiology, 21, 85-178. Lanot, R., Zachary, D., Holder, F., & Meister, M. 2001. Postembryonic hematopoiesis in Drosophila. Dev Biol, 230(2), 243-257. doi: 10.1006/dbio.2000.0123 Le Bras, S., & Van Doren, M. 2006. Development of the male germline stem cell niche in Drosophila. Dev Biol, 294(1), 92-103. doi: 10.1016/j.ydbio.2006.02.030 Lee, W. C., Beebe, K., Sudmeier, L., & Micchelli, C. A. 2009. Adenomatous polyposis coli regulates Drosophila intestinal stem cell proliferation. Development, 136(13), 2255-2264. doi: 10.1242/dev.035196 Lehane, M. J. 1997. Peritrophic matrix structure and function. Annu Rev Entomol, 42, 525-550. doi: 10.1146/annurev.ento.42.1.525 Lemaitre, B., & Hoffmann, J. 2007. The host defense of Drosophila melanogaster. Annu Rev Immunol, 25, 697-743. doi: 10.1146/annurev.immunol.25.022106.141615 Leulier, F., Parquet, C., Pili-Floury, S., Ryu, J. H., Caroff, M., Lee, W. J., . . . Lemaitre, B. 2003. The Drosophila immune system detects bacteria through specific peptidoglycan recognition. Nat Immunol, 4(5), 478-484. doi: 10.1038/ni922 Li, R. W., Choudhary, R. K., Capuco, A. V., & Urban, J. F., Jr. 2012. Exploring the host transcriptome for mechanisms underlying protective immunity and resistance to nematode infections in ruminants. Vet Parasitol, 190(1-2), 1-11. doi: 10.1016/j.vetpar.2012.06.021 Liehl, P., Blight, M., Vodovar, N., Boccard, F., & Lemaitre, B. 2006. Prevalence of local immune response against oral infection in a Drosophila/Pseudomonas infection model. PLoS Pathog, 2(6), e56. doi: 10.1371/journal.ppat.0020056 Ligoxygakis, P., Pelte, N., Hoffmann, J. A., & Reichhart, J. M. 2002. Activation of Drosophila Toll during fungal infection by a blood serine protease. Science, 297(5578), 114-116. doi: 10.1126/science.1072391 Liu, X., Hodgson, J. J., & Buchon, N. 2017. Drosophila as a model for homeostatic, antibacterial, and antiviral mechanisms in the gut. PLoS Pathog, 13(5), e1006277. doi: 10.1371/journal.ppat.1006277 Loker, WM. & Grandin, B. . 1994. Culture and Agriculture. Anthropology News, 35, 23-24. Luckhart, S., Vodovotz, Y., Cui, L., & Rosenberg, R. 1998. The mosquito Anopheles stephensi limits malaria parasite development with inducible synthesis of nitric oxide. Proc Natl Acad Sci U S A, 95(10), 5700-5705. Maizels, R. M., Gomez-Escobar, N., Gregory, W. F., Murray, J., & Zang, X. 2001. Immune evasion genes from filarial nematodes. Int J Parasitol, 31(9), 889-898. Manachini, B., Schillaci, D., & Arizza, V. 2013. Biological responses of Rhynchophorus ferrugineus (Coleoptera: Curculionidae) to Steinernema carpocapsae (Nematoda: Steinernematidae). J Econ Entomol, 106(4), 1582-1589. Martens, E. C., & Goodrich-Blair, H. 2005. The Steinernema carpocapsae intestinal vesicle contains a subcellular structure with which Xenorhabdus nematophila associates during colonization initiation. Cell Microbiol, 7(12), 1723-1735. doi: 10.1111/j.1462- 5822.2005.00585.x
189 Martens, E. C., Heungens, K., & Goodrich-Blair, H. 2003. Early colonization events in the mutualistic association between Steinernema carpocapsae nematodes and Xenorhabdus nematophila bacteria. J Bacteriol, 185(10), 3147-3154. Massaoud, M. K., Marokhazi, J., Fodor, A., & Venekei, I. 2010. Proteolytic enzyme production by strains of the insect pathogen xenorhabdus and characterization of an early-log-phase- secreted protease as a potential virulence factor. Appl Environ Microbiol, 76(20), 6901- 6909. doi: 10.1128/AEM.01567-10 Mastore, M., & Brivio, M. F. 2008. Cuticular surface lipids are responsible for disguise properties of an entomoparasite against host cellular responses. Dev Comp Immunol, 32(9), 1050- 1062. doi: 10.1016/j.dci.2008.02.003 McEwen, D. G., & Peifer, M. 2005. Puckered, a Drosophila MAPK phosphatase, ensures cell viability by antagonizing JNK-induced apoptosis. Development, 132(17), 3935-3946. doi: 10.1242/dev.01949 McMullen, J. G., 2nd, & Stock, S. P. 2014. In vivo and in vitro rearing of entomopathogenic nematodes (Steinernematidae and Heterorhabditidae). J Vis Exp(91), 52096. doi: 10.3791/52096 Medzhitov, R., & Janeway, C. A., Jr. 1998. Innate immune recognition and control of adaptive immune responses. Semin Immunol, 10(5), 351-353. Meister, M., & Lagueux, M. 2003. Drosophila blood cells. Cell Microbiol, 5(9), 573-580. Micchelli, C. A., & Perrimon, N. 2006. Evidence that stem cells reside in the adult Drosophila midgut epithelium. Nature, 439(7075), 475-479. doi: 10.1038/nature04371 Michel, T., Reichhart, J. M., Hoffmann, J. A., & Royet, J. 2001. Drosophila Toll is activated by Gram-positive bacteria through a circulating peptidoglycan recognition protein. Nature, 414(6865), 756-759. doi: 10.1038/414756a Mitani, D. K.; Kaya, H. K. & Goodrich-Blair, H. . 2004. Comparative study of the entomopathogenic nematode, Steinernema carpocapsae, reared on mutant and wild-type Xenorhabdus nematophila. Biological Control, 29, 382-391. Montenegro, H., Petherwick, A. S., Hurst, G. D., & Klaczko, L. B. 2006. Fitness effects of Wolbachia and Spiroplasma in Drosophila melanogaster. Genetica, 127(1-3), 207-215. doi: 10.1007/s10709-005-3766-4 Moreira, L. A., Iturbe-Ormaetxe, I., Jeffery, J. A., Lu, G., Pyke, A. T., Hedges, L. M., . . . O'Neill, S. L. 2009. A Wolbachia symbiont in Aedes aegypti limits infection with dengue, Chikungunya, and Plasmodium. Cell, 139(7), 1268-1278. doi: 10.1016/j.cell.2009.11.042 Murphy, K.; Travers, P.; Walport, M.; Janeway, C. . (2012). Janeway's Immunobiology (8th ed.). New York: Garland Science. Myllymaki, H., Valanne, S., & Ramet, M. 2014. The Drosophila imd signaling pathway. J Immunol, 192(8), 3455-3462. doi: 10.4049/jimmunol.1303309 Nair, M. G., Gallagher, I. J., Taylor, M. D., Loke, P., Coulson, P. S., Wilson, R. A., . . . Allen, J. E. 2005. Chitinase and Fizz family members are a generalized feature of nematode infection with selective upregulation of Ym1 and Fizz1 by antigen-presenting cells. Infect Immun, 73(1), 385-394. doi: 10.1128/IAI.73.1.385-394.2005 Nakamoto, M., Moy, R. H., Xu, J., Bambina, S., Yasunaga, A., Shelly, S. S., . . . Cherry, S. 2012. Virus recognition by Toll-7 activates antiviral autophagy in Drosophila. Immunity, 36(4), 658-667. doi: 10.1016/j.immuni.2012.03.003 Nakanishi, Y. & Siratsuchi, A. 2007. Mechanisms and roles of phagocytosis in Drosophila and Caenorhabditis elegans. Invertebr Surv J, 3, 89-96. Nappi, A. J., Carton, Y., & Vass, E. 1992. Reduced cellular immune competence of a temperature-sensitive dopa decarboxylase mutant strain of Drosophila melanogaster against the parasite Leptopilina boulardi. Comp Biochem Physiol B, 101(3), 453-460. Nappi, A. J., Vass, E., Frey, F., & Carton, Y. 2000. Nitric oxide involvement in Drosophila immunity. Nitric Oxide, 4(4), 423-430. doi: 10.1006/niox.2000.0294
190 Nielsen-LeRoux, C., Gaudriault, S., Ramarao, N., Lereclus, D., & Givaudan, A. 2012. How the insect pathogen bacteria Bacillus thuringiensis and Xenorhabdus/Photorhabdus occupy their hosts. Curr Opin Microbiol, 15(3), 220-231. doi: 10.1016/j.mib.2012.04.006 Ohlstein, B., & Spradling, A. 2006. The adult Drosophila posterior midgut is maintained by pluripotent stem cells. Nature, 439(7075), 470-474. doi: 10.1038/nature04333 Oliver, K. M., Russell, J. A., Moran, N. A., & Hunter, M. S. 2003. Facultative bacterial symbionts in aphids confer resistance to parasitic wasps. Proc Natl Acad Sci U S A, 100(4), 1803-1807. doi: 10.1073/pnas.0335320100 Osborne, S. E., Leong, Y. S., O'Neill, S. L., & Johnson, K. N. 2009. Variation in antiviral protection mediated by different Wolbachia strains in Drosophila simulans. PLoS Pathog, 5(11), e1000656. doi: 10.1371/journal.ppat.1000656 Pal, S., & Wu, L. P. 2009. Pattern recognition receptors in the fly: lessons we can learn from the Drosophila melanogaster immune system. Fly (Austin), 3(2), 121-129. Patrnogic, J., & Leclerc, V. 2017. The serine protease homolog spheroide is involved in sensing of pathogenic Gram-positive bacteria. PLoS One, 12(12), e0188339. doi: 10.1371/journal.pone.0188339 Paul, A., Wilson, S., Belham, C. M., Robinson, C. J., Scott, P. H., Gould, G. W., & Plevin, R. 1997. Stress-activated protein kinases: activation, regulation and function. Cell Signal, 9(6), 403-410. Pena, J. M., Carrillo, M. A., & Hallem, E. A. 2015. Variation in the susceptibility of Drosophila to different entomopathogenic nematodes. Infect Immun, 83(3), 1130-1138. doi: 10.1128/IAI.02740-14 Pesch, Y. Y., Riedel, D., Patil, K. R., Loch, G., & Behr, M. 2016. Chitinases and Imaginal disc growth factors organize the extracellular matrix formation at barrier tissues in insects. Sci Rep, 6, 18340. doi: 10.1038/srep18340 Poinar, G. 1972. Nematodes as facultative parasites of insects. Annu Rev Entomol, 17, 103-122. Ponton, F., Wilson, K., Holmes, A., Raubenheimer, D., Robinson, K. L., & Simpson, S. J. 2015. Macronutrients mediate the functional relationship between Drosophila and Wolbachia. Proc Biol Sci, 282(1800), 20142029. doi: 10.1098/rspb.2014.2029 Ramet, M. 2012. The fruit fly Drosophila melanogaster unfolds the secrets of innate immunity. Acta Paediatr, 101(9), 900-905. doi: 10.1111/j.1651-2227.2012.02740.x Ramet, M., Lanot, R., Zachary, D., & Manfruelli, P. 2002. JNK signaling pathway is required for efficient wound healing in Drosophila. Dev Biol, 241(1), 145-156. doi: 10.1006/dbio.2001.0502 Rehorn, K. P., Thelen, H., Michelson, A. M., & Reuter, R. 1996. A molecular aspect of hematopoiesis and endoderm development common to vertebrates and Drosophila. Development, 122(12), 4023-4031. Richards, G. R., & Goodrich-Blair, H. 2009. Masters of conquest and pillage: Xenorhabdus nematophila global regulators control transitions from virulence to nutrient acquisition. Cell Microbiol, 11(7), 1025-1033. doi: 10.1111/j.1462-5822.2009.01322.x Rivero, A. 2006. Nitric oxide: an antiparasitic molecule of invertebrates. Trends Parasitol, 22(5), 219-225. doi: 10.1016/j.pt.2006.02.014 Rizki RM., Rizki TM. 1990. Encapsulation of parasitoid eggs in phenoloxidase-deficient mutants of Drosophila melanogaster. J Insect Physiol, 38(7), 523-529. Rodou, A., Ankrah, D. O., & Stathopoulos, C. 2010. Toxins and secretion systems of Photorhabdus luminescens. Toxins (Basel), 2(6), 1250-1264. doi: 10.3390/toxins2061250 Ross, J., Jiang, H., Kanost, M. R., & Wang, Y. 2003. Serine proteases and their homologs in the Drosophila melanogaster genome: an initial analysis of sequence conservation and phylogenetic relationships. Gene, 304, 117-131.
191 Rottschaefer, S. M., & Lazzaro, B. P. 2012. No effect of Wolbachia on resistance to intracellular infection by pathogenic bacteria in Drosophila melanogaster. PLoS One, 7(7), e40500. doi: 10.1371/journal.pone.0040500 Royet, J., & Dziarski, R. 2007. Peptidoglycan recognition proteins: pleiotropic sensors and effectors of antimicrobial defences. Nat Rev Microbiol, 5(4), 264-277. doi: 10.1038/nrmicro1620 Russell, J. A., & Moran, N. A. 2005. Horizontal transfer of bacterial symbionts: heritability and fitness effects in a novel aphid host. Appl Environ Microbiol, 71(12), 7987-7994. doi: 10.1128/AEM.71.12.7987-7994.2005 Sadekuzzaman, M., Stanley, D., & Kim, Y. 2018. Nitric oxide mediates insect cellular immunity via phospholipase A2 activation. J Innate Immun, 10, 70-81. Schlenke, T. A., Morales, J., Govind, S., & Clark, A. G. 2007. Contrasting infection strategies in generalist and specialist wasp parasitoids of Drosophila melanogaster. PLoS Pathog, 3(10), 1486-1501. doi: 10.1371/journal.ppat.0030158 Schmid-Hempel, P. 2009. Immune defence, parasite evasion strategies and their relevance for 'macroscopic phenomena' such as virulence. Philos Trans R Soc Lond B Biol Sci, 364(1513), 85-98. doi: 10.1098/rstb.2008.0157 Schneider, D. S., Jin, Y., Morisato, D., & Anderson, K. V. 1994. A processed form of the Spatzle protein defines dorsal-ventral polarity in the Drosophila embryo. Development, 120(5), 1243-1250. Shapiro-Ilan, D. I.; Gouge, D. H.; Piggott, S. J.; Fife, J. P. 2006. Application technology and environmental considerations for use of entomopathogenic nematodes in biological control. Biological Control, 38(1), 124-133. Shaposhnikov, A. V., Kryndushkin, A. S., Nikolenko Iu, V., Panov, V. V., Nabirochkina, E. N., Lebedeva, L. A., & Shidlovskii Iu, V. 2013. [Activation of JAK/STAT signaling pathway in S2 Drosophila melanogaster cell culture]. Mol Biol (Mosk), 47(3), 486-491. Sheets, J. J., Hey, T. D., Fencil, K. J., Burton, S. L., Ni, W., Lang, A. E., . . . Aktories, K. 2011. Insecticidal toxin complex proteins from Xenorhabdus nematophilus: structure and pore formation. J Biol Chem, 286(26), 22742-22749. doi: 10.1074/jbc.M111.227009 Shokal, U., & Eleftherianos, I. 2017. Thioester-Containing Protein-4 Regulates the Drosophila Immune Signaling and Function against the Pathogen Photorhabdus. J Innate Immun, 9(1), 83-93. doi: 10.1159/000450610 Shokal, U., Yadav, S., Atri, J., Accetta, J., Kenney, E., Banks, K., . . . Eleftherianos, I. 2016. Effects of co-occurring Wolbachia and Spiroplasma endosymbionts on the Drosophila immune response against insect pathogenic and non-pathogenic bacteria. BMC Microbiol, 16, 16. doi: 10.1186/s12866-016-0634-6 Silverman, N., & Maniatis, T. 2001. NF-kappaB signaling pathways in mammalian and insect innate immunity. Genes Dev, 15(18), 2321-2342. doi: 10.1101/gad.909001 Silverman, N., Zhou, R., Stoven, S., Pandey, N., Hultmark, D., & Maniatis, T. 2000. A Drosophila IkappaB kinase complex required for Relish cleavage and antibacterial immunity. Genes Dev, 14(19), 2461-2471. Soderhall, K., & Cerenius, L. 1998. Role of the prophenoloxidase-activating system in invertebrate immunity. Curr Opin Immunol, 10(1), 23-28. Sorensen, J. G., Nielsen, M. M., Kruhoffer, M., Justesen, J., & Loeschcke, V. 2005. Full genome gene expression analysis of the heat stress response in Drosophila melanogaster. Cell Stress Chaperones, 10(4), 312-328. Stanley, D., Haas, E., & Miller, J. 2012. Eicosanoids: Exploiting Insect Immunity to Improve Biological Control Programs. Insects, 3(2), 492-510. doi: 10.3390/insects3020492 Stasiv, Y., Regulski, M., Kuzin, B., Tully, T., & Enikolopov, G. 2001. The Drosophila nitric- oxide synthase gene (dNOS) encodes a family of proteins that can modulate NOS activity
192 by acting as dominant negative regulators. J Biol Chem, 276(45), 42241-42251. doi: 10.1074/jbc.M105066200 Stepek, G., Buttle, D. J., Duce, I. R., & Behnke, J. M. 2006. Human gastrointestinal nematode infections: are new control methods required? Int J Exp Pathol, 87(5), 325-341. doi: 10.1111/j.1365-2613.2006.00495.x Stock, S. P. 2005. Insect-parasitic nematodes: from lab curiosities to model organisms. J Invertebr Pathol, 89(1), 57-66. doi: 10.1016/j.jip.2005.02.011 Stokes, B. A., Yadav, S., Shokal, U., Smith, L. C., & Eleftherianos, I. 2015. Bacterial and fungal pattern recognition receptors in homologous innate signaling pathways of insects and mammals. Front Microbiol, 6, 19. doi: 10.3389/fmicb.2015.00019 Stoven, S., Ando, I., Kadalayil, L., Engstrom, Y., & Hultmark, D. 2000. Activation of the Drosophila NF-kappaB factor Relish by rapid endoproteolytic cleavage. EMBO Rep, 1(4), 347-352. doi: 10.1093/embo-reports/kvd072 Stronach, B. E., & Perrimon, N. 1999. Stress signaling in Drosophila. Oncogene, 18(45), 6172- 6182. doi: 10.1038/sj.onc.1203125 Sun, H., Bristow, B. N., Qu, G., & Wasserman, S. A. 2002. A heterotrimeric death domain complex in Toll signaling. Proc Natl Acad Sci U S A, 99(20), 12871-12876. doi: 10.1073/pnas.202396399 Swarup, S., & Verheyen, E. M. 2012. Wnt/Wingless signaling in Drosophila. Cold Spring Harb Perspect Biol, 4(6). doi: 10.1101/cshperspect.a007930 Takashima, S., & Hartenstein, V. 2012. Genetic control of intestinal stem cell specification and development: a comparative view. Stem Cell Rev, 8(2), 597-608. doi: 10.1007/s12015- 012-9351-1 Takehana, A., Yano, T., Mita, S., Kotani, A., Oshima, Y., & Kurata, S. 2004. Peptidoglycan recognition protein (PGRP)-LE and PGRP-LC act synergistically in Drosophila immunity. EMBO J, 23(23), 4690-4700. doi: 10.1038/sj.emboj.7600466 Tanji, T., Hu, X., Weber, A. N., & Ip, Y. T. 2007. Toll and IMD pathways synergistically activate an innate immune response in Drosophila melanogaster. Mol Cell Biol, 27(12), 4578- 4588. doi: 10.1128/MCB.01814-06 Tauszig, S., Jouanguy, E., Hoffmann, J. A., & Imler, J. L. 2000. Toll-related receptors and the control of antimicrobial peptide expression in Drosophila. Proc Natl Acad Sci U S A, 97(19), 10520-10525. doi: 10.1073/pnas.180130797 Tauszig-Delamasure, S., Bilak, H., Capovilla, M., Hoffmann, J. A., & Imler, J. L. 2002. Drosophila MyD88 is required for the response to fungal and Gram-positive bacterial infections. Nat Immunol, 3(1), 91-97. doi: 10.1038/ni747 Teixeira, L., Ferreira, A., & Ashburner, M. 2008. The bacterial symbiont Wolbachia induces resistance to RNA viral infections in Drosophila melanogaster. PLoS Biol, 6(12), e2. doi: 10.1371/journal.pbio.1000002 Tennessen, J. M., Barry, W. E., Cox, J., & Thummel, C. S. 2014. Methods for studying metabolism in Drosophila. Methods, 68(1), 105-115. doi: 10.1016/j.ymeth.2014.02.034 Thomas, P., Kenny, N., Eyles, D., Moreira, L. A., O'Neill, S. L., & Asgari, S. 2011. Infection with the wMel and wMelPop strains of Wolbachia leads to higher levels of melanization in the hemolymph of Drosophila melanogaster, Drosophila simulans and Aedes aegypti. Dev Comp Immunol, 35(3), 360-365. doi: 10.1016/j.dci.2010.11.007 Toubarro, D., Avila, M. M., Hao, Y., Balasubramanian, N., Jing, Y., Montiel, R., . . . Simoes, N. 2013. A serpin released by an entomopathogen impairs clot formation in insect defense system. PLoS One, 8(7), e69161. doi: 10.1371/journal.pone.0069161 Toubarro, D., Avila, M. M., Montiel, R., & Simoes, N. 2013. A pathogenic nematode targets recognition proteins to avoid insect defenses. PLoS One, 8(9), e75691. doi: 10.1371/journal.pone.0075691
193 Toubarro, D., Lucena-Robles, M., Nascimento, G., Costa, G., Montiel, R., Coelho, A. V., & Simoes, N. 2009. An apoptosis-inducing serine protease secreted by the entomopathogenic nematode Steinernema carpocapsae. Int J Parasitol, 39(12), 1319- 1330. doi: 10.1016/j.ijpara.2009.04.013 Toubarro, D., Lucena-Robles, M., Nascimento, G., Santos, R., Montiel, R., Verissimo, P., . . . Simoes, N. 2010. Serine protease-mediated host invasion by the parasitic nematode Steinernema carpocapsae. J Biol Chem, 285(40), 30666-30675. doi: 10.1074/jbc.M110.129346 Trap, C., & Boireau, P. 2000. [Proteases in helminthic parasites]. Vet Res, 31(5), 461-471. doi: 10.1051/vetres:2000132 Trapnell, C., Pachter, L., & Salzberg, S. L. 2009. TopHat: discovering splice junctions with RNA-Seq. Bioinformatics, 25(9), 1105-1111. doi: 10.1093/bioinformatics/btp120 Trapnell, C., Roberts, A., Goff, L., Pertea, G., Kim, D., Kelley, D. R., . . . Pachter, L. 2012. Differential gene and transcript expression analysis of RNA-seq experiments with TopHat and Cufflinks. Nat Protoc, 7(3), 562-578. doi: 10.1038/nprot.2012.016 Vadnal, J., Ratnappan, R., Keaney, M., Kenney, E., Eleftherianos, I., O'Halloran, D., & Hawdon, J. M. 2017. Identification of candidate infection genes from the model entomopathogenic nematode Heterorhabditis bacteriophora. BMC Genomics, 18(1), 8. doi: 10.1186/s12864- 016-3468-6 Valanne, S., Wang, J. H., & Ramet, M. 2011. The Drosophila Toll signaling pathway. J Immunol, 186(2), 649-656. doi: 10.4049/jimmunol.1002302 Veillard, F., Troxler, L., & Reichhart, J. M. 2016. Drosophila melanogaster clip-domain serine proteases: Structure, function and regulation. Biochimie, 122, 255-269. doi: 10.1016/j.biochi.2015.10.007 Vierstraete, E., Cerstiaens, A., Baggerman, G., Van den Bergh, G., De Loof, A., & Schoofs, L. 2003. Proteomics in Drosophila melanogaster: first 2D database of larval hemolymph proteins. Biochem Biophys Res Commun, 304(4), 831-838. Vivas, E. I., & Goodrich-Blair, H. 2001. Xenorhabdus nematophilus as a model for host- bacterium interactions: rpoS is necessary for mutualism with nematodes. J Bacteriol, 183(16), 4687-4693. doi: 10.1128/JB.183.16.4687-4693.2001 Vlisidou, I., & Wood, W. 2015. Drosophila blood cells and their role in immune responses. FEBS J, 282(8), 1368-1382. doi: 10.1111/febs.13235 Vodovar, N., Vinals, M., Liehl, P., Basset, A., Degrouard, J., Spellman, P., . . . Lemaitre, B. 2005. Drosophila host defense after oral infection by an entomopathogenic Pseudomonas species. Proc Natl Acad Sci U S A, 102(32), 11414-11419. doi: 10.1073/pnas.0502240102 Voronin, D., Bachu, S., Shlossman, M., Unnasch, T. R., Ghedin, E., & Lustigman, S. 2016. Glucose and Glycogen Metabolism in Brugia malayi Is Associated with Wolbachia Symbiont Fitness. PLoS One, 11(4), e0153812. doi: 10.1371/journal.pone.0153812 Wang, Y., Gaugler, R., & Cui, L. 1994. Variations in Immune Response of Popillia japonica and Acheta domesticus to Heterorhabditis bacteriophora and Steinernema Species. J Nematol, 26(1), 11-18. Wang, Y.; Gaugler, R. 1999. Steinernema glaseri surface coat protein suppresses immune response of Popillia japonica (Coleoptera: Scarabaeidae) larvae. Biological Control, 14, 111-121. Wang, Z., Wilhelmsson, C., Hyrsl, P., Loof, T. G., Dobes, P., Klupp, M., . . . Theopold, U. 2010. Pathogen entrapment by transglutaminase--a conserved early innate immune mechanism. PLoS Pathog, 6(2), e1000763. doi: 10.1371/journal.ppat.1000763 Waterfield, N. R., Ciche, T., & Clarke, D. 2009. Photorhabdus and a host of hosts. Annu Rev Microbiol, 63, 557-574. doi: 10.1146/annurev.micro.091208.073507
194 Wertheim, B., Kraaijeveld, A. R., Schuster, E., Blanc, E., Hopkins, M., Pletcher, S. D., . . . Godfray, H. C. 2005. Genome-wide gene expression in response to parasitoid attack in Drosophila. Genome Biol, 6(11), R94. doi: 10.1186/gb-2005-6-11-r94 White, G. F. 1927. A Method for Obtaining Infective Nematode Larvae from Cultures. Science, 66(1709), 302-303. doi: 10.1126/science.66.1709.302-a Williams, M. J. 2007. Drosophila hemopoiesis and cellular immunity. J Immunol, 178(8), 4711- 4716. Wong, A. C., Chaston, J. M., & Douglas, A. E. 2013. The inconstant gut microbiota of Drosophila species revealed by 16S rRNA gene analysis. ISME J, 7(10), 1922-1932. doi: 10.1038/ismej.2013.86 Wong, A. C., Vanhove, A. S., & Watnick, P. I. 2016. The interplay between intestinal bacteria and host metabolism in health and disease: lessons from Drosophila melanogaster. Dis Model Mech, 9(3), 271-281. doi: 10.1242/dmm.023408 Wong, Z. S., Hedges, L. M., Brownlie, J. C., & Johnson, K. N. 2011. Wolbachia-mediated antibacterial protection and immune gene regulation in Drosophila. PLoS One, 6(9), e25430. doi: 10.1371/journal.pone.0025430 Wu, L. P., & Anderson, K. V. 1998. Regulated nuclear import of Rel proteins in the Drosophila immune response. Nature, 392(6671), 93-97. doi: 10.1038/32195 Wu, S. C., Liao, C. W., Pan, R. L., & Juang, J. L. 2012. Infection-induced intestinal oxidative stress triggers organ-to-organ immunological communication in Drosophila. Cell Host Microbe, 11(4), 410-417. doi: 10.1016/j.chom.2012.03.004 Xiang, J., Bandura, J., Zhang, P., Jin, Y., Reuter, H., & Edgar, B. A. 2017. EGFR-dependent TOR-independent endocycles support Drosophila gut epithelial regeneration. Nat Commun, 8, 15125. doi: 10.1038/ncomms15125 Yadav, S., Daugherty, S., Shetty, A. C., & Eleftherianos, I. 2017. RNAseq Analysis of the Drosophila Response to the Entomopathogenic Nematode Steinernema. G3 (Bethesda), 7(6), 1955-1967. doi: 10.1534/g3.117.041004 Yadav, S., Shokal, U., Forst, S., & Eleftherianos, I. 2015. An improved method for generating axenic entomopathogenic nematodes. BMC Res Notes, 8, 461. doi: 10.1186/s13104-015- 1443-y Yang, H., Kronhamn, J., Ekstrom, J. O., Korkut, G. G., & Hultmark, D. 2015. JAK/STAT signaling in Drosophila muscles controls the cellular immune response against parasitoid infection. EMBO Rep, 16(12), 1664-1672. doi: 10.15252/embr.201540277 Yi, Y., Wu, G., Lv, J., & Li, M. 2016. Eicosanoids mediate Galleria mellonella immune response to hemocoel injection of entomopathogenic nematode cuticles. Parasitol Res, 115(2), 597-608. doi: 10.1007/s00436-015-4776-5 Zaffran, S., Astier, M., Gratecos, D., Guillen, A., & Semeriva, M. 1995. Cellular interactions during heart morphogenesis in the Drosophila embryo. Biol Cell, 84(1-2), 13-24. Zaidman-Remy, A., Herve, M., Poidevin, M., Pili-Floury, S., Kim, M. S., Blanot, D., . . . Lemaitre, B. 2006. The Drosophila amidase PGRP-LB modulates the immune response to bacterial infection. Immunity, 24(4), 463-473. doi: 10.1016/j.immuni.2006.02.012 Zeidler, M. P., Bach, E. A., & Perrimon, N. 2000. The roles of the Drosophila JAK/STAT pathway. Oncogene, 19(21), 2598-2606. doi: 10.1038/sj.onc.1203482 Zele, F., Nicot, A., Duron, O., & Rivero, A. 2012. Infection with Wolbachia protects mosquitoes against Plasmodium-induced mortality in a natural system. J Evol Biol, 25(7), 1243-1252. doi: 10.1111/j.1420-9101.2012.02519.x Zettervall, C. J., Anderl, I., Williams, M. J., Palmer, R., Kurucz, E., Ando, I., & Hultmark, D. 2004. A directed screen for genes involved in Drosophila blood cell activation. Proc Natl Acad Sci U S A, 101(39), 14192-14197. doi: 10.1073/pnas.0403789101 Zhang, Z. T., & Zhu, S. Y. 2009. Drosomycin, an essential component of antifungal defence in Drosophila. Insect Mol Biol, 18(5), 549-556. doi: 10.1111/j.1365-2583.2009.00907.x
195 Zou, S., Meadows, S., Sharp, L., Jan, L. Y., & Jan, Y. N. 2000. Genome-wide study of aging and oxidative stress response in Drosophila melanogaster. Proc Natl Acad Sci U S A, 97(25), 13726-13731. doi: 10.1073/pnas.260496697
196