Toll-8/Tollo Negatively Regulates Antimicrobial Response in the Drosophila Respiratory Epithelium

Barrier epithelia that are persistently exposed to microbes have evolved potent immune tools to eliminate such pathogens. If mechanisms that control Drosophila systemic responses are well-characterized, the epithelial immune responses remain poorly understood. Here, we performed a genetic dissection of the cascades activated during the immune response of the Drosophila airway epithelium i.e. trachea. We present evidence that bacteria induced-antimicrobial peptide (AMP) production in the trachea is controlled by two signalling cascades. AMP gene transcription is activated by the inducible IMD pathway that acts non-cell autonomously in trachea. This IMD-dependent AMP activation is antagonized by a constitutively active signalling module involving the receptor Toll-8/Tollo, the ligand Spätzle2/DNT1 and Ect-4, the Drosophila ortholog of the human Sterile alpha and HEAT/ARMadillo motif (SARM). Our data show that, in addition to Toll-1 whose function is essential during the systemic immune response, Drosophila relies on another Toll family member to control the immune response in the respiratory epithelium.


Introduction
Although the innate immune system is a primitive host defense mechanism, it involves a sophisticated repertoire of humoral and cellular responses both acting systemically and locally [1]. In recent years, the Drosophila model organism has proven to be an invaluable system in dissecting in great details the genetics and cellular mechanisms regulating the innate immunity [2][3]. One fundamental mechanism common to humans and Drosophila immunity involves signaling by receptors of the Toll family. Upon microbial infection, human TLRs activate the synthesis of cytokines and other regulatory molecules that stimulate the adaptive immune system [4]. In Drosophila, Toll signalling leads to the activation of the systemic immune response, which is characterized by the synthesis of AMPs by the fat body cells [5]. Upon secretion into the circulating blood, these AMPs provide systemic protection against fungi and bacteria. In the mammalian innate immune response, bacteria are directly sensed by TLRs [6]. In contrast, these microorganisms are detected in Drosophila by another class of proteins, the Peptidoglycan Recognition Proteins (PGRPs), also present in the human proteome [7][8][9]. Recognition of Lys-type peptidoglycan (PGN) (mainly found in Gram-positive bacteria cell wall) by the circulating PGRP-SA protein triggers a protease cascade involving successively Mod-Sp, Grass, and the Spä tzle-Processing-Enzyme (SPE) [8,[10][11]. Upon activation, SPE becomes competent to transform the zymogen pro-Spä tzle into an active ligand for the Toll receptor, inducing its dimerization and intracellular signalling [12]. Production of AMPs after infection by Gram-negative bacteria, is however largely independent of the Toll pathway but rather relies on another NF-kB signalling cascade named IMD [13]. Sensing of Gram-negative bacteria upstream of the IMD pathway takes place at the plasma membrane, through PGN recognition by the transmembrane PGRP-LC receptor. Binding of DAP-type PGN (present in Gramnegative bacterial cell wall) to PGRP-LC induces its dimerization, which, in turn, triggers IMD-dependent intracellular events enabling the nuclear translocation of the NF-kB transcription factor Relish [14][15][16][17].
Although penetration of infectious microbes into the body cavity, and consequently, activation of a systemic immune response, is a rare event, interactions between microbes and epithelia take place constantly throughout the life of all metazoan. This implies that these barrier epithelia must be armed with efficient systems for microbial detection and elimination. However, these epithelia that act as interfaces with the external environment, share some characteristics that could be seen as detrimental for the needs of an effective immune system. Indeed, they usually have large surface areas and consist of thin structural layers, thus representing ideal entry points for pathogens. In this respect, the airway epithelium is unique among all epithelia, since it has a very delicate structure and is constantly exposed to a plethora of airborne pathogens. This could explain the occurrence of a great variety of inflammatory lung diseases, including asthma, chronic obstructive pulmonary disease or cystic fibrosis [18][19]. Elucidation of the primary steps that lead to chronic inflammation of the mammalian lung is obstructed by the complexity of inflammatory responses in this organ. Animals with a much simpler organization, such as the fruit fly, might help us clarify the basic architecture of this epithelial immune response, thereby helping to unravel the mechanisms that lead to chronic inflammation of the airways. Although of much simpler organi-zation, the fly's airway system shows striking similarities with the mammalian lung regarding both its architecture and its physiology [20][21][22][23][24][25].
In this report, we present a detailed description of the mechanisms that regulate AMP production in the Drosophila respiratory epithelium. We show that, in contrast to systemic fat body immune response, the IMD pathway can be activated noncell autonomously in the tracheal network. We present evidence that IMD pathway activation is tightly regulated in the cells of the respiratory epithelium. We demonstrate that the molecular mechanisms underlying IMD down-regulation following infection, are different from those previously reported in the gut and in the fat body, and rely on a dialog between two antagonist pathways. The production of AMPs in the trachea is positively regulated by the IMD pathway, which is counterbalanced by a negative regulation from a signalling cassette, whose upstream receptor is a member of the Toll family, Toll-8/Tollo. Our data suggest that the Spz2/DNT1 cytokine is a putative Tollo ligand in this process, and that Ect4, the Drosophila ortholog of the human Toll/ Interleukin-1 Receptor (TIR) domain-containing protein SARM mediates Tollo signalling during tracheal immune response.

AMP activation followed a stereotypical pattern in the Drosophila tracheal network
The tracheal system is a relatively simple model system that has provided an important insight into the biology of branching morphology [22]. It is a tubular structure covered by a lumenal cuticular lining that forms a physical barrier against dehydration and invading microorganisms [20]. This network consists of a monolayer epithelium made up of two dorsal trunks (DT) connected to several visceral branches (VB) bringing oxygen to internal tissues. Upon infection by the entomopathogenic bacterium Erwinia carotovora carotovora (Ecc), these epithelial cells produce a cocktail of AMPs including Drosomycin, Drosocin and Attacin [26][27]. In order to perform a detailed spatiotemporal analysis of this epithelial response, we used a Drosomycin-GFP reporter transgene (Drs-GFP) and monitored the response of this tissue after infection. When reared on conventional medium, a few larvae showed sporadic tracheal Drs-GFP expression mainly in VB, and rarely in the posterior part of the DT, namely the posterior spiracles (PS) ( Figure 1A and 1B). This basal Drs-GFP tracheal expression was qualitatively and quantitatively similar in larvae reared conventionally or under axenic conditions (data not shown). Upon infection with Ecc, Drs-GFP activation followed a somewhat stereotypical pattern. Responding larvae were categorized into three classes according to their Drs-GFP expression pattern ( Figure 1A and 1C), namely, larvae expressing GFP in PS and in posterior VB only (class I), in PS and all VB (class II) and in VB and DT (class III). Kinetic experiments showed that GFP signal was first detected in PS and then spread into VB ( Figure 1A, C). The reporter was only later activated in the DT and, unexpectedly, first in the anterior half and then in the entire trunk ( Figure 1A). Although we appreciate that the expression patterns of reporter transgenes can slightly deviate from those of the actual AMP, the Drs-GFP expression patterns observed cannot easily be attributed to a progressive diffusion of bacterial elicitors (such as PGN) from the spiracles into the tracheal network, but rather speak for more complicated mechanisms in tracheal AMP activation.

IMD pathway activation is not strictly cell autonomous in the trachea
Next, we compared the mode of activation of AMP following forced immune pathway activation in the trachea and in another immune tissue, the fat body. Although activation of Drosomycin transcription is mainly controlled by the Toll pathway in the fat body (but can be activated by ectopic IMD pathway triggering, see later), it is strictly IMD-dependent in the trachea [5,26,28]. Indeed, over-activation of the IMD (UAS-PGRP-LCa, UAS-IMD), but not of the Toll pathway (UAS-spz act, Tl 3 ), is sufficient to induce tracheal expression of Drs-GFP in non-infected larvae ( Figure S1A and S1B). Concomitantly, loss-of-function mutations in IMD pathway components (Relish, PGRP-LC, IMD) prevent Drs-GFP tracheal activation in infected larvae, whereas Toll signalling mutants such as spz or Dif do show a wild-type tracheal response upon infection. To analyze whether all tracheal cells were competent to trigger AMP production upon IMD pathway activation, we induced UAS-IMD expressing clones in tracheal cells, using fat body clones as controls. Overexpression of IMD led to a strictly cell-autonomous and fully penetrant activation of both Drs-GFP and Dipt-Cherry in the fat body ( Figure 2B and 2C). In the trachea, although most IMD-expressing cells showed Drs-GFP expression, a fraction did not ( Figure 2A). In addition, Drs-GFP activation was not always associated with the expression of the UAS-IMD transgene (Figure 2A), suggesting that IMD pathway activation in trachea is not strictly cell autonomous. These results were confirmed by using a UAS-PGRP-LCa transgene that activated Drs-GFP both autonomously and non-autonomously in trachea cells ( Figure 2E) but strictly cell-autonomously in the fat body ( Figure 2F). We next addressed whether PGRP-LC function was required cell-autonomously for IMD pathway activation in the trachea upon infection. Analysis of MARCM loss-of-function clones for PGRP-LC indicates that tracheal cells mutant for PGRP-LC were totally impaired in their ability to trigger Drs-GFP expression, following Ecc infection ( Figure 2D and H). These results indicate that, although PGRP-LC is essential in tracheal cells for IMD pathway triggering, IMD pathway activation in one tracheal cell can spread to neighboring cells. This contrasts with the strictly cell-autonomous IMD-dependent immune response observed in fat body cells.

Tollo is expressed apically in the Drosophila trachea cells
In order to get a further insight into the mechanisms that control AMP induction in trachea, we looked for putative immune

Author Summary
Invertebrates solely rely on innate immune responses for defense against microbial infections. Taking advantage of its powerful genetics, the fly Drosophila melanogaster has been extensively used as a model system to dissect the molecular mechanisms that control innate immunity. This work led to the discovery of the essential role of the Toll-1 receptor in triggering the systemic immune response in flies, and paved the way for the discovery of the function of members of the Toll-like receptor (TLR) family in mammalian immunity. Whereas all TLRs are implicated in the mammalian immune response, Toll-1 was, so far, the only Drosophila Toll family member to be involved in the regulation of the immune response. In the present study, we show that another Toll family member, Toll-8 (Tollo), plays an important role in controlling the respiratory epithelium immune response. Our data indicate that, by antagonizing the IMD pathway, Tollo is preventing overactivation of the antibacterial response in the airway epithelium.
genes expressed in this tissue. A recent report identified the repertoire of all immune genes expressed in the trachea [29]. One of the striking data of this study, [confirmed by FlyAtlas (http:// flyatlas.org/)] was that, in addition to Toll itself, two other Toll family members, 18-Wheeler (Toll-2) and Tollo (Toll-8), are strongly expressed in trachea. 18-wheeler being implicated in developmental processes with indirect impacts the immune response [30], we focused our study on the putative function of the Tollo transmembrane protein in the tracheal immune response. Using Lac-Z reporter lines (data not shown) and q- RT-PCR ( Figure 3A), we confirmed that Tollo mRNA is highly enriched in the tracheal epithelium, and expressed at lower levels in other tissues. To investigate the subcellular localization of the Tollo protein, we genetically associated a UAS-Tollo::Myc construct with the trachea-specific Breathless-Gal4 driver (Btl-Gal4). Anti-Myc antibody staining suggested that Tollo was localized apically at the cell membrane facing the airway lumen ( Figure 3B). Double staining experiments showed that Tollo::Myc partially co-localized with the apical marker Cadherin::GFP, but was mutually exclusive with Viking::GFP, a basal membrane-associated protein ( Figure 3B). These results indicate that Tollo is a protein enriched in the tracheal epithelium with an apical subcellular localization.

AMPs are specifically over-produced in infected Tollo mutant trachea
The rather restricted expression pattern of Tollo mRNA in the trachea and the apical subcellular distribution of Tollo protein prompted us to investigate its putative function in the immune response. For that purpose, we used two previously characterized hypomorphic alleles (Tollo 145 and Tollo R5A ) together with a complete loss-of-function allele (Tollo C5 ) that we generated by Pelement mediated homologous recombination [31][32] ( Figure  S2A). All Tollo mutants were viable with no obvious developmental defects and gave rise to phenotypically normal pharate adults indicating that Tollo has no essential role in Drosophila development. We tested the ability of these Tollo mutant larvae to mount an immune response. In the absence of infection, approximately 5% of wild-type larvae showed Drs-GFP expression in VB and/or PS ( Figure S3). Similar Figures were obtained with Tollo mutants suggesting that Tollo is not required to set up the basal level of AMP production in the absence of infection ( Figure S3). After bacterial infection, however, the immune response was much stronger in Tollo mutants than in wild-type sibling larvae ( Figure 4A and 4B and Figure S3). While we could identify the three previously described classes of Drs-GFP positive larvae in both control and Tollo mutants, the relative proportion of these was significantly different between genotypes ( Figure S3). The percentage of larvae showing no GFP expression was reduced to 5-10% in Tollo mutant (compared to 40% in controls), whereas Class III larvae, which represented 15% of controls, reached up to 50% in the Tollo mutant larvae ( Figure S3). Similar results were obtained with three independent Tollo alleles and in transheterozygous allelic combination demonstrating that this phenotype was, indeed, due to Tollo inactivation and not to other mutations on the chromosome (data not shown). The effects were not only qualitative but also quantitative. In most infected Tollo mutant larvae, Drs-GFP expression was intense, whereas it was rarely the case in controls ( Figure 4A). q-RT-PCR experiments indicate that Drosomycin, Drosocin and Attacin mRNA levels were respectively increased by 6, 7 and 2.6 fold after infection in Tollo mutants compared to wild-type trachea ( Figure 4B). To ensure that this effect was indeed a consequence of Tollo inactivation in the tracheal network itself, we combined the Btl-Gal4 driver with a UAS-Tollo IR RNA interference construct. As shown in Figure 4A and Figure S3, larvae, in which Tollo was eliminated specifically in trachea, also showed increased Drs-GFP expression in this tissue after infection.
In addition to regulating Drosomycin expression in the trachea, the IMD pathway also controls Diptericin transcription locally in the gut and systemically in the fat body [13,26]. To test whether Tollo acts as a general negative regulator of IMD-dependent mechanisms in other immune tissues, we analyzed the effects of inactivating Tollo on IMD pathway activation in the gut and the fat body. Using the Dipt-Cherry reporter construct for the larval stage ( Figure 4D) and q-RT-PCR for both larval and adult stages ( Figure 4E and 4F), we showed that Tollo was not implicated in IMD negative regulation in either tissue. This was the case for both immune responses induced by septic injury or by oral ingestion. In addition, we showed that Tollo mutants were unaffected in their ability to activate the Toll pathway during a Gram-positive bacteria-mediated systemic immune response ( Figure 4F). Altogether these results demonstrate that the Tollo receptor is specifically required to dampen IMD pathwaydependent responses in the tracheal network after infection.
AMP over-production in Tollo mutants is not secondary to trachea defects Since AMPs are induced upon cellular stress, we tested whether Drosomycin expression in Tollo mutants was a secondary consequence of a possible implication of Tollo in tracheal formation. The following reasons led us to believe that it was not the case. 1) Tollo mutant embryos gave rise to viable adults, suggesting that Tollo mutant trachea are fully functional in larvae and adults. 2) Tracheal cell morphology of Tollo and control larvae appeared similar when observed under transmission electron microscopy ( Figure 5A). 3) No constitutive AMP transcription was detected in non-infected Tollo mutant larvae ( Figure 4B and Figure S3). We then wondered whether Drosomycin over-activation could be linked to the presence of higher levels of potential immune elicitors in Tollo mutant and RNAi trachea. This could be due to the presence of higher bacterial load in the trachea. However, as shown in Figure 5B and Figure S4B, the number of Ecc-GFP in Tollo mutant trachea was identical in Tollo mutants and in controls. Alternatively, Tollo mutant trachea could be more permeable to contaminated external fluid. To test this hypothesis, wild type and Tollo mutant larvae were incubated in the presence of a fluorescent dye, bromophenol blue. External fluid penetration inside the trachea lumen was not different in wild-type and Tollo mutant larvae ( Figure 5C and 5D). This indicates that overactivation of Drosomycin in both Tollo mutant and RNAi trachea cannot be attributed to an increase in fluid penetration, and therefore putative immune elicitor load within the tracheal lumen. Altogether, these results demonstrate that the infection-dependent Drosomycin over-activation observed in Tollo mutant, is not secondary to defective trachea but rather suggests a direct implication of the Tollo protein in the regulation of IMDdependent Drosomycin expression.

Ect4/SARM and DNT1/Spätzle 2 loss-of-function mutants phenocopy Tollo mutants
We then tried to identify the intra-and extra-cellular components that may mediate Tollo signalling in trachea. The Drosophila Toll-1 receptor and vertebrate TLR functions have all been shown to be mediated by TIR domain-containing proteins, respectively, the Drosophila dMyd88 and the mammalian Myd88, TRIF, SARM, TRAM and MAL [33][34][35]. However, the Drosophila proteome contains two TIR domain proteins, Ect4/ SARM the Drosophila ortholog of vertebrate SARM and dMyd88, the latter mediating the Toll signalling during dorso-ventral axis specification and immune response [36][37][38]. In order to test whether Tollo acts through a TIR domain-containing protein, we analyzed the tracheal immune response of dMyd88 and Ect4/ SARM mutants. We showed that larvae carrying Ect4/SARM mutations ( Figure S2B) display a strong over-activation of Drosomycin expression (visualized with the Drs-GFP reporter transgene and quantified by q-RT-PCR) upon infection, phenotype that was not observed in dMyd88 mutant larvae ( Figure 6A and 6B and Figure S5A). This suggests that Ect4/SARM is the bone fide TIR domain adaptor transducing Tollo signalling in the tracheal immune response. Whereas TLRs function as Pattern Recognition Receptors by directly binding to microbial motifs, previous work has shown than the Drosophila Toll-1 receptor is activated during both embryonic dorso-ventral axis specification and immune response by its ligand Spä tzle [39][40]. Since spz mutant larvae did not present higher activation of Drs-GFP in the respiratory tract after infection, we believe that Spz is not a functional ligand for Tollo ( Figure 6C). Since Spz-like genes are present in the fly genome, we screened them for a Tollo-like phenotype. We observed that removing Spz2 (known as DNT1) function in trachea phenocopies Tollo mutant as far as Drosomycin over-activation is concerned ( Figure 6C and 6D and Figure S5B). This suggests that DNT1 could be the, or one of the ligand(s), responsible for Tollo activity in the tracheal immune response. Similarly to Tollo, q-RT-PCR data indicate that Ect4 and DNT1 are specifically acting in the tracheal epithelium and do not contribute to IMD pathway regulation in the gut and in the fat body ( Figure 6B and 6D).

Tollo is hypostatic to IMD in the trachea
Taking into account the above results, it appears that the function of Tollo is specifically to down-regulate the IMD pathway in the tracheal cells following infection. In order to genetically place Tollo with respect to known IMD pathway components, we performed epistatic experiments. We showed that the Tollo mutant   Figure 4A). This epistatic relationship was confirmed by q-RT-PCR on Drosomycin mRNA ( Figure 4C). In genetic terms, Tollo is hypostatic, or acts in parallel to PGRP-LC and imd. Consistently, Relish nuclear translocation monitored with an anti-Relish antibody was higher in Tollo RNAi-infected tracheal cells than in controls ( Figure S6). These results suggest that Tollo is not directly involved in IMD pathway activation per se but that, in its absence, IMD pathway activation is more efficient upon infection.

Discussion
Epithelial responses are first and foremost local responses to prevent the epithelium from unnecessary immune reactions. Since the recognition steps in Drosophila respiratory epithelia involve the transmembrane receptor PGRP-LC and occur within the extracellular space, it is expected that molecular mechanisms must be at work to prevent constitutive or excessive immune response in this tissue, particularly essential for animal growth and viability. In this report, we present data demonstrating that the transmembrane receptor Tollo is part of a signalling network, whose function is to specifically down-regulate AMP production in the trachea. We show that Tollo antagonizes IMD pathway activation in the respiratory epithelium, and that DNT1/Spz2 and Ect4/SARM are putative Tollo ligand and transducer, respectively, in this process. Our data demonstrate that, in addition to the family founder Toll-1, another member of the Leucine-Rich-Repeats family of Toll proteins, is regulating the Drosophila innate immune response. Although it has been abundantly documented that every single mammalian TLR has an immune function [4], the putative implication of Toll family members, other than Toll-1 itself, in the Drosophila immune response has been a subject of controversy [41]. Data showing that Drosophila Toll-9 overexpression was sufficient to induce AMPs expression in vivo has prompted the idea that Toll-9 could maintain significant levels of anti-microbial molecules, thus providing basal protection against microbes [42]. However, our recent analysis of a complete Toll-9 loss-of-function allele has shown that this receptor is neither implicated in basal anti-microbial response nor required to mount an immune response to bacterial infection [43]. The present data are also fully consistent with a recent report showing that Toll-6, Toll-7 and Toll-8 are not implicated in systemic AMP production in flies [44], and demonstrate that a Toll family member, Tollo, is a negative regulator of local airway epithelial immune response upon bacterial infection. In contrast to Toll-1, whose activation is inducible in the fat body, Tollo pathway activation seems to be constitutive in the trachea. Despite these differences, both receptors use a member of the Spz family as ligand. Interestingly, sequence similarities, intron's size and conservation of key structural residues, indicate that Spz2/DNT1 is phylogenetically the closest family member to the Toll ligand Spz [45]. Furthermore, both Spz and Spz2/DNT1 have been shown to have neurotrophic functions in flies [46]. It would be of great interest to test whether Tollo also mediates Spz2 function in the nervous system.
Both during embryonic development and immune response, Spz is activated by proteolytic cleavage [10,[47][48]. This step depends upon the Easter protease that is implicated in D/V axis specification and on SPE for Toll pathway activation by microbes. Since Spz orthologs are also produced as longer precursors, they are likely to be activated by proteolysis. The fact that Tollo and Spz2 loss-of-function phenotypes correspond to excessive AMP production, suggests that in wild-type conditions, the Tollo pathway is constitutively activated by an active form of the Spz2 ligand. This situation is reminiscent to that observed in the embryonic ventral follicle cells, in which a Pipe-mediated signal induces a constitutive activation of the Easter cascade leading to Spz cleavage, Toll activation and, in turn, ventral fate acquisition [49]. It should be noted that Easter and one Pipe isoform are very strongly expressed in the trachea cells (Flyatlas), and are candidate proteins in mediating Tollo activity in the respiratory epithelia.
The fact that Ect4, but not dMyd88 mutant, loss-of-function mutant phenocopies Tollo mutant suggest that Ect4 could be the TIR domain adaptor transducing Tollo signal in the tracheal cells. Alternatively, Ect4/SARM could mediate Tollo function by interfering with IMD pathway signalling. In mammals, SARM is under the transcriptional control of TLR and negatively regulates TLR3 signalling by directly interfering with the association between the RHIM domain-containing proteins TRIF and RIP [50]. Since PGRP-LC contains a RHIM domain as TRIF, and IMD is the Drosophila counterpart of RIP, one can envisage that Drosophila SARM could act by interfering with the PGRP-LC/ IMD association required for IMD pathway signalling. Similarly to its function as a negative regulator in fly immunity, SARM is the only TIR domain-containing adaptor that acts as a suppressor of TLR signalling [36,50].
One obvious question relates to the mode of action of Tollo on IMD pathway downregulation. Two mechanisms have been recently described that result in the down-regulation of the IMD pathway. The first one regulates PGRP-LC membrane localization, and is dependent on the PIRK protein [51][52][53]. Upon infection, the intracellular PIRK protein is up-regulated and, in turn, represses PGRP-LC plasma membrane localization leading to the shutdown of the IMD signalling [53]. In infected pirk mutants, IMD-dependent AMPs are overproduced in both the gut and the fat body. In our conditions, however, inactivation of PIRK specifically in the trachea did not influence Drosomycin activation in trachea ( Figure S7A). To verify whether Tollo is acting via a mechanism similar to PIRK, we looked at PGRP-LC membrane localization using a UAS-PGRP-LC::GFP construct. PGRP-LC membrane localization was identical in wild-type and Tollo mutant tracheal cells ( Figure S7B). The second mechanism that modulates IMD activation, acts directly on the promoters of IMD target genes. Ha et al. (2005) have shown that the Caudal transcription factor sits on some of the IMD target promoters preventing their activation by Relish [54]. We thus tested the putative implication of Caudal in Tollo signalling by using Drs-GFP reporter transgenes containing either wild-type Caudal Responsive Elements (CDREs) or mutated versions unresponsive to Caudal activity [55]. Upon weaker in trachea when compare to midgut. Colony forming units (CFUs) per tissue are shown for each condition. n = 6 in triplicates. (C) Example of control larvae (Btl-Gal4) without (top panels) or with (bottom panels) red fluorescent BPB in trachea (arrow). Dorsal views of third instar larvae are shown. Pictures were taken 24 h after Bromophenol Blue (BPB) incubation. (D) Quantification of (BPB) entry in trachea of non-infected larvae. BPB entry is rare (around 5%) and not affected by Tollo mutations (Tollo C5 /Tollo R5A ), Tollo RNAi (Btl-Gal4;UAS-Tollo IR ) or infection (data not shown). Each histogram corresponds to the mean value +/2 SD of 3 experiments. A total number of 90 larvae were counted for each experiment. Values indicated by identical symbols (*) are not significantly different (P.0.05) from one another. doi:10.1371/journal.ppat.1002319.g005 infection, Drs-GFP with mutated CDREs was activated in fat body but not in gut or trachea ( Figure S7C). In conclusion, Caudal acts as a transcriptional activator, rather than a repressor, for the Drs-GFP reporter in trachea. These results indicate that Tollo does not regulate the IMD pathway via PGRP-LC membrane localization or through promoter targeting of Caudal. One challenging task for the future will be to identify the mechanism used by Tollo to counter-balance tracheal PGRP-LC activation. It has been reported that the loss of Tollo function in ectodermal cells during embryogenesis alters glycosylation in nearby differentiating neurons [31,[56][57]. Since the pattern of oligosaccharides expressed in a cell can influence its interactions with others and with pathogens, Tollo could function by modifying glycosylation pattern in response to microbes. It could be envisaged that Tollo mediates PGRP-LC glycosylation, and thereby reduces its ability to respond to bacterial elicitors. Further work will be required to address the above hypothesis, whereby Tollo activity and glycosylation modification could be linked in order to regulate the IMD pathway activation in trachea.

Bacterial load analysis
Bacterial load of surface sterilized individuals was quantified by plating appropriate serial dilutions of lysates obtained from 6 dissected guts or trachea (from larvae) on nutrient agar plates (Luria Bertani + spectinomycin 100 mg/ml). Biological triplicates were collected for each experimental condition at 4h and 24h after Ecc-GFP infection. Homogenization of tissues was performed using the Precellys 24 tissue homogenizer (Bertin technologies, France) and 0,75-1mm glass beads in 500 mL of LB + spectinomycin.

Natural infection of larvae and adults
Cells from overnight bacterial cultures were recovered by centrifugation at 4,000 g for 10 min at 4uC. The supernatant was discarded and the pellet was resuspended in fresh LB media. Cell suspensions were serially diluted in PBS, and the concentration of cells was determined by optical-density (OD) measurement. 200 ml of an overnight bacterial culture of Ecc (OD = 200) were directly added on top of feeding third instar larvae into a standard cornmeal-agar medium at 25uC. A similar volume of LB broth was used in control experiments. Larvae were monitored for Drosomycin and Diptericin transcription by fluorescence analysis using Drs-GFP and Dpt-cherry reporters respectively, and by qRT-PCR, 24h after infection. Septic injuries were performed by pricking adult males with a thin needle contaminated with M. luteus or E. coli. 200 ml of Bromophenol Blue (SIGMA # B8026) at 10 g/l were directly added on top of feeding third instar larvae.

Immunostaining on larvae
Larval tissue were dissected in PBS and fixed for 20 min in 4% paraformaldehyde on ice. After several rinses in PBT (PBS + 0.1% Triton X-100), they were blocked for 1 hr in PBT-3% BSA at 4uC and then incubated with antibody at the appropriate dilution in PBT-BSA 3% overnight at 4uC. Primary antibodies were: rabbit Anti-Relish (1:500) or Mouse Anti-Myc (9E10 Santa Cruz at 1: 500). Several washes in PBT were followed by a 2 hr incubation with secondary antibody at RT (Alexa Fluor 546 goat anti-rabbit IgG and Alexa Fluor 555 goat anti-mouse IgG diluted 1:500, Molecular Probes), then 5 washes in PBT. The tissues were finally mounted in Vectashield (Vector Laboratories) fluorescent mount- ing medium, with DAPI. Images were captured with a LSM 510 Zeiss confocal microscope.

Quantitative real-time PCR
Quantitative real-time PCR and SYBR Green analysis were performed as previously described [58]. Primer information can be obtained upon request. The amount of mRNA detected was normalized to control rp49 mRNA values. Normalized data was used to quantify the relative levels of a given mRNA according to cycling threshold analysis (DCt).

Electronic microscopy
For electron microscopic sections, third instar larvae trachea were dissected and fixed at RT in 4% PFA and 2% glutaraldehyde in 0.12 M sodium cacodylate buffer at pH 7.4 for 1 h. The trachea were then washed 3610 min in 0.12 M sodium cacodylate buffer, post-fixed in 2% OsO4 in 0.12 M sodium cacodylate buffer for 1 h and washed again 3610 min. Samples were subsequently dehydrated through series of ethanol gradients and infiltrated with propylene oxide, embedded in epoxy resin (Fluka, Sigma) and polymerized at 80uC. Ultrathin (80 nm) plastic sections were cut using a Leica UltraCut microtome with a diamond Diatome knife and post-stained with 2% uranyl acetate, followed by treatment with Reynolds'lead citrate, and stabilized for transmission EM by carbon coating. Examination was performed with a Zeiss Leo 912 microscope at 100 kV. Images were captured using a Gatan 792 Bioscan camera using Digital Micrograph software.  The apical localization of PGRP-LC::GFP is un-affected by Tollo mutations. The following genotypes are shown: Tub-Gal4, Tub-Gal80 ts /+; Tollo C5 /+ (control) and Tub-Gal4, Tub-Gal80 ts /+; Tollo C5 /Tollo R5A . A total number of 30 trachea were observed for each genotype. Images were taken 24h after incubation at 29uC. (C) Image representative of Ecc-infected Drs(CDREmut)-GFP larvae is shown with no GFP signal visible in trachea (lower panel). The CDREs of Drosomycin promoter are required for tracheal expression upon Ecc infection. A total number of 50 larvae were analyzed. Images were taken 24h after Ecc infection. (EPS)