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Loss of Dictyostelium ATG9 results in a pleiotropic phenotype affecting growth, development, phagocytosis and clearance and replication of Legionella pneumophila

Infection of Dictyostelium discoideum with Legionella pneumophila resulted in a large number of differentially regulated genes among them three core autophagy genes, ATG8, ATG9 and ATG16. Macroautophagy contributes to many physiological and pathological processes and might also constitute an important mechanism in cell‐autonomous immunity. For further studies we selected the highly conserved ATG9. In colocalization studies with GFP‐tagged ATG9 and different organelle marker proteins we neither observed colocalization with mitochondria, the ER nor lysosomes. However, there was partial colocalization with the Golgi apparatus and many ATG9‐GFP‐containing vesicles localized along microtubules and accumulated around the microtubule organizing centre. ATG9‐deficient cells had pleiotropic defects. In addition to growth defects they displayed severe developmental defects, consistent with the known role of autophagy in Dictyostelium development. Unexpectedly, the ATG9 mutant also had a strong phagocytosis defect that was particularly apparent when infecting the cells with L. pneumophila. However, those Legionellae that entered the host could multiply better in mutant than in wild‐type cells, because of a less efficient clearance in the early and a more efficient replication in the late phase of infection. We conclude that ATG9 and hence macroautophagy has a protective role during pathogen infection.


Introduction
The social amoeba Dictyostelium discoideum is a wellestablished model organism for the study of basic biological processes such as signal transduction, cytokinesis, chemotaxis and cell motility as well as basic aspects of development (Noegel and Schleicher, 2000;Chisholm and Firtel, 2004;Robinson and Spudich, 2004;King and Insall, 2009). All stages of the life cycle are motile and upon starvation the amoebae enter a developmental program, in which up to 100 000 cells aggregate by chemotaxis towards cAMP. The aggregate transforms via distinct morphological states into a mature fruiting body, composed of a ball of spores supported by a thin, long stalk made of vacuolized dead cells (Eichinger, 2003). In the vegetative phase the amoebae grow as separate, independent cells and feed on bacteria and yeast. Uptake of the food occurs by phagocytosis, a complex and evolutionarily highly conserved mechanism. The ingested microorganism is trapped in a phagosome and, via the phagolysosomal pathway, is ultimately delivered to a lysosome where it is degraded by an intricate cocktail of hydrolytic enzymes (Duhon and Cardelli, 2002;Cosson and Soldati, 2008). However, despite its impressive armory to kill bacteria, Dictyostelium can be infected with a number of medically relevant pathogenic bacteria (Skriwan et al., 2002;Steinert et al., 2007).
In higher eukaryotes phagocytosis is also a means to fight invading pathogens. This task is performed by specialized cells such as macrophages or neutrophils that have the ability to rapidly and efficiently internalize a variety of microorganisms and particles and degrade them. These cells represent professional phagocytes that are important for innate and adaptive immunity in metazoa. In recent years it became also clear that the basic mechanisms of host pathogen interactions are conserved in lower eukaryotes (Dorer and Isberg, 2006;Bozzaro et al., 2008;Jin et al., 2009). Thus, the professional phagocyte D. discoideum has become an attractive model system for the study of the infection with human pathogens in particular the investigation of bacterial virulence traits and the analysis of the roles of host gene products in phagocytosis and killing (Steinert and Heuner, 2005;Hilbi et al., 2007;Cosson and Soldati, 2008;Isberg et al., 2009). For the study of the host side a large number of D. discoideum mutants are available from the Dictyostelium stock centre, additional genes of interest can be easily disrupted and a complete and well-curated genome sequence is on-hand (Eichinger, 2003;Eichinger et al., 2005).
In order to survive bacterial pathogens manipulate host cell processes to avoid phagolysosomal fusion and to establish a replicative niche (Isberg et al., 2009). The host on the other hand initiates elaborate defence processes of which one appears to be macroautophagy (hereafter autophagy) (Huang and Klionsky, 2007). Autophagy is an ancient cellular pathway that is conserved from yeast to man. It presumably evolved to enable cells to survive periods of starvation. More than 30 autophagy (ATG) genes have been identified, mainly in yeast, of which 18 constitute the core machinery for starvation induced autophagy. Cytosolic material is captured into double membrane-bound vesicles that mature into autophagosomes and then, after fusion with lysosomes, become autophagolysosomes. There the cargo is degraded and then recycled for further use (Nakatogawa et al., 2009). Autophagy contributes to many physiological and pathological processes, including cell differentiation and development, programmed cell death, cancer and neurodegenerative disorders (Xie and Klionsky, 2007). Previous work in Dictyostelium showed that autophagy is required for normal development. Autophagy mutants were generated in five core autophagy genes, and all mutants displayed developmental defects albeit of variable severity (Otto et al., 2003;2004a). There is accumulating evidence that autophagy is also an important mechanism in the complex interactions that occur between host and pathogen. Interestingly, autophagy appears to be a defence mechanism that is able to degrade some pathogens while others apparently modulate and subvert it (Huang and Klionsky, 2007). In the case of Salmonella typhimurium or Mycobacterium tuberculosis cells target bacteria for degradation through autophagy. (Gutierrez et al., 2004). Shi-gella flexneri on the other hand escapes autophagy, while Staphylococcus aureus even utilize the autophagosome for replication (Ogawa et al., 2005;Schnaith et al., 2007). In the case of Dictyostelium it has been reported that Legionella pneumophila is able to replicate in autophagy mutants (Otto et al., 2004b). However, the authors did not examine if disruption of the autophagic machinery might be advantageous for the replication of the pathogen. Such a possibility is supported by data from macrophages where autophagy apparently constitutes a defence mechanism that L. pneumophila must modulate to establish infection (Amer and Swanson, 2005).
Here we describe the subcellular localization of Dictyostelium ATG9 and its function in growth, development and during phagocytosis and infection. ATG9 is so far the only known integral membrane protein of the core autophagic machinery and is thought to deliver membrane lipids to the site of autophagosome formation (Reggiori et al., 2005;Mari and Reggiori, 2007;Webber et al., 2007). We show in immunofluorescence experiments and live cell imaging that ATG9-containing vesicles (ACVs) randomly appeared at the cell periphery, travelled, presumably along microtubules, to the perinuclear region and accumulated around the microtubule organizing centre (MTOC). Deletion of atg9 resulted in a pleiotropic phenotype with defects in growth, development, phagocytosis, and during infection with L. pneumophila. Expression of GFP-tagged ATG9 rescued the mutant phenotype. The strong phagocytosis defect points to an unexpected link between phagocytosis and autophagy. The results of the infection experiment support a protective function of ATG9 during L. pneumophila infection.

ATG9 is highly conserved and upregulated during Legionella infection
In previous work we studied the global transcriptional regulation of Dictyostelium cells upon infection with L. pneumophila. Comparative analysis of the 24 h time point post infection revealed 240 differentially regulated genes, which are thought to be involved in the pathogenic response (Farbrother et al., 2006). Bioinformatic analysis showed that three of the more than 50 human orthologues among the 240 genes, namely ATG8, ATG9 and ATG16, encoded proteins of the core autophagy machinery (Table 1). Because the role of autophagy in cellautonomous immunity is still a matter of debate, we decided to further study this process with a particular emphasis on phagocytosis and infection. We selected ATG9 for further study because: (i) it was upregulated during infection, (ii) it is highly conserved and (iii) it is so far the only known integral membrane autophagy protein whose study could provide information about the unknown membrane source for the autophagosome.
Dictyostelium atg9 encodes a protein of 699 amino acids. It contains the conserved APG9 (autophagy 9) domain of approximately 300 amino acids. The program TMHMM (http://www.cbs.dtu.dk/services/TMHMM/) predicted six transmembrane domains between amino acids 75 and 500 (Fig. 1A). Pairwise alignments of the APG9 domain of Sacharomyces cerevisiae (yeast), D. discoideum, Arabidopsis thaliana (plant) and Homo sapiens (human) showed highest similarity of the Dictyostelium APG9 domain with the yeast orthologue and slightly lower similarities with plant and with human. However, comparison of the human APG9 domain with those of yeast, plant and Dictyostelium revealed highest similarity with the latter sequence (Fig. 1B). We next performed sequence alignments with orthologous proteins from animals, plants and fungi and generated an evolutionary tree, which showed that the Dictyostelium protein is most closely related to the fungal orthologues and that the ATG9 proteins from animals and plants cluster in two distinct groups (Fig. 1C).

ATG9 is upregulated during development
Autophagy is an important pathway in eukaryotes for recycling cellular material during developmental processes. We therefore investigated the developmental regulation of ATG9. Real-time polymerase chain reaction (PCR) showed an approximately fivefold upregulation at the mRNA level after 4 and 8 h, eightfold after 12 and 16 h and sevenfold after 20 h of Dictyostelium development ( Fig. 2A). On the protein level we observed an approximately eightfold upregulation during the first 6 h of development. The slight decrease at the 8 h time point is probably due to inefficient transfer of the protein (Fig. 2B).
The need to recycle cellular components during starvation is apparently reflected in the strong upregulation of ATG9. A similar developmental regulation was observed for Dictyostelium ATG1 and ATG8 (Otto et al., 2004a).
To investigate the cellular function of ATG9 we generated two independent knockout mutants and two mutants that express ATG9 fused to GFP in the mutant background. AGA expresses GFP fused N-terminally to ATG9 while in the AAG mutant GFP is fused to the C-terminus of ATG9. Southern blot and PCR verification of the knockout mutant is shown in Fig. 3A and B. No ATG9 protein is detected in the ATG9mutant while AGA and AAG express the GFP-ATG9 or, respectively, ATG9-GFP fusion protein of approximately 105 kDa (Fig. 3C). Both strains significantly overexpress the fusion protein in comparison with the expression of ATG9 in wild-type AX2 cells. In the case of the AAG mutant a degradation product is seen with the polyclonal ATG9-specific antibodies but not with the GFP-specific antibody ( Fig. 3C and D).

ACVs localize along microtubules
The membrane source for the growing autophagosome is still unknown. A possible candidate for the unidentified membrane carrier is ATG9, the only integral membrane protein required for autophagosome formation. In yeast, ATG9 is indispensable for pre-autophagosomal structure (PAS) formation and part of its peripheral pool appears to cycle between the mitochondria and the PAS. The remaining part of the peripheral ATG9 pool resides in a hitherto unknown cellular compartment (Reggiori et al., 2005  A. Southern blot of AX2 and two independent ATG9 gene replacement mutants. Genomic DNA was digested with EcoRI and BglII, separated in 0.7% agarose, transferred to nitrocellulose and probed with the 3′-arm of the gene replacement construct. #1, ATG9 -#1; #2, ATG9 -#2. B. Reverse transcriptase PCR of AX2 and ATG9 -#1. No addition of cDNA (blank) was used as a negative control and AX2 genomic DNA as a positive control. C. and D. Western blot of wild-type AX2, the ATG9 -#1 replacement mutant and two strains that either express GFP fused N-terminally (AGA) or C-terminally (AAG) to ATG9 in the gene replacement mutant ATG9 -#1. (C) Polyclonal anti-ATG9 antibody; (D) monoclonal anti-GFP antibody.
and . In mammalian cells part of the peripheral pool localizes to the trans-Golgi network (TGN) and endosomes (Young et al., 2006). We made use of the generated AGA and AAG strains and performed colocalization studies using markers for different subcellular organelles that could supply membrane material for autophagosome formation. We used porin as marker for mitochondria, the lysosomal antigen to label lysosomes, protein disulfide isomerase (PDI) as marker for the ER, vatA for the contractile vacuole and comitin for the Golgi apparatus. ATG9-GFP expressing cells contained a large number of dot-like structures that presumably represented small vesicles (Fig. 4, left panel). We did not detect ATG9-GFP at mitochondria, lysosomes, the ER and at the contractile vacuole ( Fig. 4A-C, middle and right panels; vatA staining not shown). However, for some cells we observed partial colocalization with the Golgi marker comitin (Fig. 4D, arrows), reminiscent of the TGN localization of ATG9 in mammalian cells (Young et al., 2006). Another interesting observation was that ACVs were strongly enriched in the perinuclear region. We therefore performed colocalization studies with the centrosomal marker protein CP250 and microtubules (Fig. 4E). Interestingly, ACVs were enriched around the MTOC, in addition, we frequently observed them in inter-phase cells in close proximity to microtubules, suggesting a possible role for microtubules in their transport (Fig. 4E, inset). Transport along microtubules could also explain the accumulation around the MTOC. This enrichment was lost in dividing cells, and ACVs were evenly distributed throughout the cytoplasm (Fig. 4F, arrow). Immunofluorescence studies with cells that were starved for 3 h showed similar colocalization and subcellular distribution of ACVs (data not shown). We also studied the dynamics of ACVs by life cell imaging. In a number of instances we saw random appearance of ACVs at the cell periphery. The newly generated ACVs usually remained briefly at the 'birth site' and then travelled swiftly in direction of the MTOC, suggesting transport along microtubules (Movie S1).

The ATG9mutant displays severe developmental defects
It has already been shown that autophagy is required for a number of developmental processes (Otto et al., 2003;2004a). We therefore investigated the development of AX2, the ATG9mutant and the AGA and AAG mutants by plating cells on phosphate agar. Development was severely impaired in the ATG9knockout mutant, and we observed four main developmental defects (Movie S2). (i) ATG9aggregates were considerably larger than wildtype AX2 aggregates, and the mutant displayed a multi-tip phenotype. While AX2 aggregates produce only one tip each, mutant aggregates produced generally three tips per aggregate (Fig. 5A). (ii) Slug formation in the mutant was very efficient. Moreover, they were usually thinner and much longer than AX2 slugs (Fig. 5B). (iii) In addition the ATG9slugs frequently broke into several pieces ( Fig. 5B, orange arrows). Furthermore, some slugs migrated normally in the case of the mutant (Fig. 5B, white and yellow arrows); others, however, did not migrate and only elongated (Fig. 5B, green arrow, back of the slug, and blue arrow, front of the slug). This observation prompted us to investigate the phototactic ability of the ATG9mutant. Strikingly, in contrast to AX2 the mutant slugs generally did not move at all into the direction of the light source making it impossible to measure a phototactic index (Fig. S1). Occasionally there appeared to be a very small shift of some slugs in the inoculation spot into direction of the light source, indicating that phototactic ability might still be intact while slug migration is defective (Fig. S1). (iv) Finally, only a minority of the mutant slugs proceeded through terminal differentiation and produced fruiting bodies. The few fruiting bodies that were produced in the ATG9mutant were misshapen. They possessed a thickened stalk, were much smaller than wild-type fruiting bodies and generated only a tiny spore head. Re-expression of ATG9 fused to GFP in the AGA or AAG strains rescued the developmental phenotype (Fig. 5C).
These results clearly indicate that autophagy is required at several steps during Dictyostelium development. Disturbance of the process leads to severe impairments in aggregation, the tipped aggregate stage, the slug stage and fruiting body generation.

Growth of the ATG9mutant is impaired in shaking culture and on bacteria
We were then curious to find out if autophagy is also important during vegetative growth. First we checked growth of the ATG9mutant and the AGA and AAG strains in comparison with wild-type AX2 cells in shaking culture. Log phase cells from a shaken axenic culture were adjusted to a density of 2 ¥ 10 4 cells ml -1 , and the growth followed over 120 h. We observed a strong growth defect in the case of the ATG9mutant. After 3 day growth at 21°C with shaking at 160 r.p.m. the cell titer of the mutant was about sixfold lower than that of AX2 wild-type cells.
The growth defect was rescued through expression of ATG9 fused to GFP in the AGA and AAG strains. ATG9continued to grow from days 4 to 5 while AX2, the AGA and the AAG strains had almost reached stationary phase and the cell titer increased only moderately (Fig. 6A).
Final cell densities of AX2 wild-type and ATG9cells were comparable (data not shown). We calculated the generation time in the logarithmic growth phase from days 1 to 3 and found an approximately 45% longer generation time (14.5 vs.~10 h) of the ATG9cells as compared with AX2 We then compared the growth of wild-type AX2 cells, the ATG9mutant, the AGA and AAG strain on Klebsiella aerogenes and Escherichia coli B/2 bacteria. We quantified growth by measuring the plaque size of the different strains on both bacteria over several days. For both bacteria we found that the ATG9mutant formed considerably smaller plaques in comparison with the other strains. The growth defect was not as dramatic as in shaking culture, and plaque size in case of the ATG9mutant was reduced on average by about 30%. Again, expression of ATG9 fused to GFP in the mutant background (AGA and AAG strains) rescued this defect (Fig. 6B). The reduced plaque size of the ATG9mutant could either be due to a decreased phagocytosis rate or to problems in intracellular digestion of consumed bacteria or to both. Figure 6C depicts examples of single plaques and of fruiting bodies of the four strains grown on Klebsiella. In addition to the smaller plaque size in case of the ATG9mutant it is Fig. 5. Development of the ATG9 mutant is impaired. Cells were developed on phosphate agar and pictures were taken every 10 min. Shown are three developmental stages. A. Tipped mound stage. Wild-type AX2 cells produced only one tip (white arrows) while mutant cells produced generally three tips (white arrows) per mound. Scale bar is 0.2 mm. B. Slug stage: mutant slugs were long and thin and frequently broke (orange arrows). Some migrated normally as was the case for AX2 (white and yellow arrows) while others were immobile and only elongated (blue arrows). Scale bar is 0.2 mm. C. Side view of terminally differentiated fruiting bodies: the ATG9 mutant only occasionally generated fruiting bodies that had thickened stalks and were considerably smaller than wild-type fruiting bodies (please note the different scale). Development and fruiting body generation were rescued in the AGA and AAG strains. Scale bars 0.1 mm.

AX2
ATG9 -AGA AAG evident that only a very small fraction of the mutant cells in the aggregates were able to further develop and to produce fruiting bodies while most cells apparently did not proceed with development (Fig. 6C, top and middle rows). The few fruiting bodies that were generated were very tiny in comparison with AX2 wild-type and the rescued AAG and AGA strains (Fig. 6C, bottom row). In comparison with development on phosphate agar fruiting body generation was even more impaired upon growth on a lawn of Klebsiella. We also found that the viability of the mutant spores was strongly reduced (data not shown).
The ATG9mutant displays a decreased phagocytosis rate The decrease in plaque size on bacterial lawns in case of the ATG9mutant indicated a phagocytosis defect. We therefore performed a number of phagocytosis assays. First we analysed the uptake of pHrodo™ E. coli by fluorescence measurements. The fluorescence of pHrodo™ E. coli strongly increases as the pH of its surrounding becomes more acidic. This is the case when phagosomes fuse with lysosomes, therefore phagolysosomal fusion of ingested particles can be followed. Using this tool we observed an approximately twofold lower fluorescence in the ATG9mutant as compared with AX2 wild-type and the AGA and AAG strains (Fig. 7A). Next we analysed the uptake of fluorescent yeast cells. AX2 wild-type cells showed a strong increase in fluorescence after a lag phase of about 15 min. In the case of the ATG9cells the increase in fluorescence was only moderate and final fluorescence was approximately threefold lower than for AX2 (Fig. 7B).
Both results indicate a phagocytosis defect in the ATG9mutant for microorganisms that are considered as 'food' by wild-type cells. Next we wanted to know whether there would also be a defect in the uptake of pathogenic bacteria.
To analyse this we used S. aureus, a representative of the Gram-positive bacteria, and L. pneumophila, a Gramnegative bacterium. For both pathogens we found a reduction in uptake in case of the ATG9mutant. Fluorescent activated cell sorting (FACS) analysis of Dictyostelium cells that had phagocytosed green fluorescent S. aureus showed an approximately twofold decrease for mutant cells (Fig. 7C). The uptake of L. pneumophila was measured by determining the colony-forming units (cfu) 3 h post infection. At this stage intracellular bacteria have not yet started to replicate. For the ATG9mutant we observed a dramatic decrease in uptake rate that was approximately eightfold lower than for AX2 wild-type cells (Fig. 7D).

Clearance of L. pneumophila was less efficient and replication was better in ATG9cells
Recently it was reported that autophagy was dispensable for intracellular L. pneumophila replication (Otto et al., 2004b). However, the authors did not address the question if autophagy could also be a means to fight the pathogen. In this case it would be expected that disruption of autophagy would be advantageous for the pathogen and its survival and/or replication would be favoured. We therefore performed time-course experiments with L. pneumophila infecting AX2 wild-type cells and the ATG9mutant. To compensate for the reduced uptake rate of the mutant we used a multiplicity of infection (MOI) of 8 for the mutant and an MOI of 1 for the wild-type cells. This way similar numbers of cfu were obtained for both strains 3 h post infection (data not shown). We carried out 10 independent infection experiments and relative changes in intracellular bacteria at the different time points were calculated based on the cfu of the 3 h time point, which was considered one for both strains. In the early phase of infection, where L. pneumophila has to establish its replicative niche in the host, we observed a stronger decrease in intracellular bacteria in wild-type cells as compared with the ATG9mutant [ Fig. 8 inset and Table 2, 8 and 18 hpi (hours post infection)]. This result indicates a reduced clearance of ingested L. pneumophila in the mutant. In addition, we found that in the late phase of infection the bacteria also replicated better in the ATG9mutant than in AX2 cells (Fig. 8). Both effects together, the reduced clearance and the increased replication of the pathogen, resulted in an approximately twofold higher bacterial load in the case of the ATG9mutant (Table 2, 48 hpi). The data support a model where autophagy has a protective role during infection of D. discoideum with L. pneumophila.

Discussion
Infection of D. discoideum with L. pneumophila resulted in the identification of 240 differentially regulated genes that were specific for the pathogenic response (Farbrother  , 2006). Of these more than 50 had clear orthologues in human including ATG8, ATG9 and ATG16 that encode core autophagy proteins (Table 1). We reasoned that their differential expression might either be part of the Dictyostelium defense mechanism or might be triggered by Legionella to support the establishment of a replicative niche or even both players might try to modulate autophagy during the infection process. To gain a better understanding of the highly conserved ATG9 protein we have investigated its cellular function in D. discoideum through the analysis of a gene deletion strain and the expression of GFP-tagged ATG9 in the mutant background. ATG9 is essential for autophagy in yeast, it is so far the only known integral membrane protein of the core autophagy machin-ery and a possible candidate for the unidentified membrane carrier of the growing autophagosome (Reggiori et al., 2005;He and Klionsky, 2007).
In immunofluorescence studies of GFP-tagged ATG9 with markers for different cellular organelles we found partial colocalization with the Golgi marker comitin (Fig. 4D). It is possible that this result may reflect close neighbourhood because we observed a strong enrichment of ACVs in a juxtanuclear position at the MTOC, the same region where the Golgi apparatus is found (Fig. 4E). A connection with the Golgi apparatus is however not excluded as a colocalization with the Golgi was observed in mammalian cells where ATG9 cycles between the TGN and endosomes (Young et al., 2006). In contrast in yeast, part of the ATG9 peripheral pool appears to cycle between the mitochondria and the PAS while the remaining part resides in a hitherto unknown cellular compartment (Reggiori et al., 2005;He and Klionsky, 2007). Interestingly, no colocalization with mitochondria has been observed in mammalian cells and in Dictyostelium. In addition, we found that ACVs often localize along microtubules and that this localization is lost in dividing cells (Fig. 4E inset and F). Live cell imaging showed many instances of ACVs that randomly appeared at the cell periphery and then travelled swiftly to a position close to the MTOC, as revealed by costaining with the centrosomal marker CP250 (Movie S1 and Fig. 4E). Thus, generation and transport of Dictyostelium autophagosomes appear to be similar to that of mammalian cells, where they are generated in the cell periphery, followed by transport along microtubules in a dynein-dependent manner and accumulation at the MTOC (Webber et al., 2007;Jahreiss et al., 2008;Kimura et al., 2008). Interestingly, Lu and Clarke reported that the Legionella containing phagosome is also transported along microtubules in Dictyostelium (Lu and Clarke, 2005). This aspect is worth to be investigated in future infection studies with ATG9-GFP expressing cells.
In recent years the role of autophagy in uptake, establishment and replication of intracellular pathogens has been increasingly studied. And it is now clear that autophagy is an important component of cell-autonomous immunity. Some pathogens are sequestered by autophagosomes and degraded in the autolysosome; others manage to escape from autophagosomes and replicate in the cytoplasm and still others block or delay autophagosome maturation and replicate in the autophagosome (Swanson, 2006;Huang and Klionsky, 2007). For Dicty-ostelium it was shown that L. pneumophila could replicate successfully in ATG1, ATG5, ATG6, ATG7 and ATG8 autophagy mutants and produce replication vacuoles that were morphologically indistinguishable from those in wild type. The authors concluded that macroautophagy is dispensable for intracellular replication of L. pneumophila (Otto et al., 2004b). However, the first measurement of intracellular bacteria was performed only 2 days post infection, and the early phase of infection was not analysed. In addition, it is not clear if the tested autophagy mutants had, similar to the ATG9mutant, a phagocytosis defect (see below). If this were the case, the reported results would imply that L. pneumophila would indeed replicate better in the mutants. We found that clearance of L. pneumophila was less efficient in the ATG9mutant during the first 18 h of infection and intracellular replication in the following 30 h slightly more efficient (Fig. 8 and Table 2). These results strongly support findings that are consistent with ATG9 and hence autophagy having a protective role during infection with L. pneumophila. In studies with Caenorhabditis elegans and Dictyostelium it was shown that autophagy genes protect against S. typhimurium infection (Jia et al., 2009). In macrophages autophagy appears to be a defence mechanism that L. pneumophila and other pathogens must modulate to establish infection (Gutierrez et al., 2004;Amer and Swanson, 2005;Huang and Klionsky, 2007). Surprisingly, the ATG9mutant also had a strong phagocytosis defect in the uptake of yeast, non-pathogenic and pathogenic bacteria (Figs 6C and 7). The defect was strongest with L. pneumophila where we observed an eightfold reduced uptake rate. To our knowledge this is the first report describing a phagocytosis defect upon deletion of a core autophagy gene. Interestingly, there appears to be a link between autophagy and phagocytosis in murine macrophages where it was shown that activation of toll-like receptors triggered recruitment of LC3/ATG8 and Beclin/ ATG6, elements of the classical autophagy pathway, to the phagosome (Sanjuan et al., 2007). The authors also found that engaging the autophagy pathway via TLR signalling enhanced phagosome maturation and destruction of engulfed yeast cells. At present it is not clear how autophagy is linked to phagocytosis in Dictyostelium. Apart from a possible direct link similar to macrophages it could also be that disruption of cellular recycling processes lead to a shortage of membranes and other components that are needed for efficient phagocytosis.
The latter possibility could also be the basis for the severe developmental defect that is seen in Dictyostelium autophagy mutants. Starving Dictyostelium cells need to recycle cellular components to be able to proceed through development. When developed on phosphate agar, aggregates of the atg9 mutant were larger than from wild type, they produced multiple tips and practically all of them entered the slug stage. Slugs were very long and thin and frequently broke. Finally, only a few, tiny and misshapen fruiting bodies were produced in the mutant ( Fig. 5 and Movie S2). Previously it was reported that Dictyostelium mutants in ATG1, ATG6, ATG7, ATG8 and the ATG12-ATG5 conjugation system displayed developmental defects of varying severity (Otto et al., 2003;Otto et al., 2004a). The developmental phenotype of the ATG9mutant was similar to the ATG5, ATG6, ATG7 and ATG8 mutants but clearly not identical. Its phenotype was stronger than in ATG6and ATG8and slightly weaker than in ATG5and ATG7 -. As with other Dictyostelium autophagy mutants, development of ATG9was more aberrant on bacterial lawns than on phosphate agar (Figs 5 and 6C). The fact that autophagy mutants considerably differ in the severity of their developmental defects, indicate additional and as yet largely unknown functions of these proteins.
In conclusion, disruption of ATG9 function in Dictyostelium cells resulted in a pleiotropic phenotype. Mutant cells not only displayed significant growth and severe developmental defects but, surprisingly, also had a strong phagocytosis defect. The latter phenotype indicates a hitherto unknown link between autophagy and phagocytosis. Whether the connection is direct or indirect remains to be determined. Last but not least infection experiments with L. pneumophila resulted in higher intracellular titers in mutant cells, supporting a model in which autophagy is part of a cell-autonomous defence system. Based on opposing differential regulation of three autophagy genes in response to infection we would like to postulate that both players, the host and the pathogen, try to modulate this ancient cellular process.
Dictyostelium strains and growth conditions D. discoideum strain AX2 was used as wild-type strain. The ATG9 mutant (ATG9 -) was generated by gene inactivation of the atg9 gene in AX2 through homologous recombination with the Cre-loxP system (Faix et al., 2004). AGA and AAG were generated by transformation of ATG9after transient expression of the Cre recombinase with expression constructs encoding GFP fused either N-or C-terminally to ATG9. Wild-type and mutant strains were grown at 21°C in liquid nutrient medium with shaking at 160 r.p.m. (Brink et al., 1990) or on SM agar plates with K. aerogenes (Williams and Newell, 1976). Generation times were calculated using n = [logN t -logN0]/log2 where n is the number of generations, N0 is the cell titer at time 0 and Nt is the cell titer at time t. For development in suspension, axenically grown cells were washed twice with Soerensen buffer (17 mM Na-K-phosphate, pH 6.0) and resuspended in the same buffer at a concentration of 1 ¥ 10 7 cells ml -1 and shaken at 160 r.p.m. at 21°C.

Vector construction and transformation
The gene replacement construct was generated in the pLBLP vector where the Bsr resistance cassette is flanked by loxP sites (Faix et al., 2004). A PCR-amplified fragment of 434 bp (bases 34-468 of the coding sequence) was cloned into the BamHI and PstI sites and a 560 bp fragment (bases 1586-2146 of the coding sequence) into the HindIII and SalI sites of the vector. Vectors for expression of full-length ATG as GFP fusion proteins in D. discoideum under the control of the actin-15 promoter and actin-8 terminator were constructed using the pBsr-N-GFP N1 and pBsr-C-GFP C1 vectors (Blau-wasser et al., 2009). Proteins composed of either N-or C-terminal GFP fused to ATG9 were expressed in the ATG9mutant. The constructs were generated by PCR and verified by sequencing. In all fusion proteins a linker of six amino acids with the sequence GGSGGS separated the GFP moiety from the full-length ATG9. The plasmids were introduced into AX2 wild-type cells and the ATG9mutant by electroporation; transformants were selected in the presence of 5 mg ml -1 Blasticidin S (ICN Biochemicals). The transformants were either identified by PCR screening of bsr resistant clones followed by Southern and Western blotting and/or by visual inspection under a fluorescence microscope followed by immunological detection of the expressed proteins in Western blots.

Development and phototaxis assays
Cells at a density of 2-3 ¥ 10 6 cells ml -1 were washed twice with Soerensen buffer. A total of 1 ¥ 10 8 cells were then resuspended in 1 ml of Soerensen buffer and 500 ml evenly distributed onto a phosphate-buffered agar plate (9 cm in diameter) and incubated at 21°C. Different stages of development were observed, and the images were either captured manually at defined time points or automatically at 6 or 10 min intervals using a stereomicroscope or an inverted microscope. Movies were processed with Image J. Phototaxis assays were essentially performed as described (Khaire et al., 2007). Approximately 48 h after inoculation, slime trails and cellular material were blotted onto nitrocellulose filter (BA85, Ø 82 mm, Schleicher and Schuell) by keeping the filter on the plate for 1 h. Filters were stained with 0.1% amido black in 25% isopropanol and 10% acidic acid (staining solution) for 10 min. To remove the excess stain two incubations in destaining solution (25% isopropanol, 10% acetic acid) were performed for 10-15 min, then the filters were washed with water and air-dried. The filters exhibiting the stained slime trails were used to determine the distance travelled by the slugs towards the source of light from the point of application.

Phagocytosis analysis
We analysed the phagocytosis of E. coli, S. aureus, L. pneumophila Corby (see below) and yeast cells in the different Dictyo-stelium strains. Phagocytosis of E. coli was analysed with pHrodo™ E. coli Bioparticles (Invitrogen GmbH, Germany; http// www.invitrogen.com). The pH-sensitive pHrodo™ dye is a specific sensor of intracellular bacteria. The dye is non-fluorescent at neutral pH and fluoresces bright red in acidic environments. The assay was carried out essentially according to the instructions of the manufacturer (Invitrogen GmbH, Germany). Briefly, 2 ¥ 10 5 AX2 wild-type, ATG9 -, AGA or AAG cells per well were transferred into the wells of a 96-well black microplate. The Dictyostelium cells were allowed to settle down, after 1 h the medium was removed and 100 ml of the pHrodo™ E. coli Bioparticles in phagocytosis buffer (20 mM Hepes pH 7.0, 2 mM MgCl2, 10 mM NaCl) were added, corresponding to an approximately 150:1 ratio relative to Dictyostelium cells. Fluorescence measurements were performed with excitation at 544 nm and emission at 590 nm. Data were corrected by the fluorescence of bacteria alone. Each measurement was carried out in quadruplicate, and three independent experiments were performed. Data represent mean values with the fluorescence value of AX2 being set to 100%.
For quantifying the uptake of S. aureus by D. discoideum via FACS analysis, Dictyostelium cells were harvested and resuspended in a 1:1 mixture of Soerensen buffer and AX2 medium (AX/2) to a final concentration of 1 ¥ 10 6 cells ml -1 . From this suspension 0.3 ml were transferred into 12-well microtiter plates and incubated for 60 min at room temperature. S. aureus NRS151 GFP were harvested from a preculture and resuspended in AX/2 medium to a concentration of 1 ¥ 10 8 cells ml -1 . The incubation medium in the plate wells was then replaced by 0.3 ml of the bacterial suspension, resulting in an MOI of 100. Dictyostelium and bacterial cells were incubated together for 3 h at room temperature, then washed once with Soerensen buffer and incubated with lysostaphin (0.2 mg ml -1 ) for 15 min on ice in order to kill extracellular bacteria. Cells were then washed twice with ice-cold Soerensen buffer to remove excess lysostaphin and dead bacteria before being transferred into polystyrene FACS tubes. Using the CellQuest FACS software (BD Biosciences, Heidelberg, Germany) viable Dictyostelium cells were gated. Fluorescence intensity generated by intracellular GFPexpressing S. aureus NRS151 was measured, and the acquired data were quantified in a histogram.
Quantitative phagocytosis of TRITC-labelled heat-killed yeast cells was performed as described (Rivero and Maniak, 2006). Data represent the mean values and standard errors of the mean of four independent experiments.

Uptake and infection assays with L. pneumophila Corby
For the uptake and infection assays Dictyostelium strains and L. pneumophila Corby were cultured and prepared as previously described (Hägele et al., 2000). Infections were performed in a 24-well set-up. Briefly, Dictyostelium cells were adjusted to 1 ¥ 10 6 cells ml -1 in infection medium (a 1:1 mixture of HL5 medium and 1¥ Soerensen buffer; HL5 medium: 5 g of yeast, 10 g of glucose, 10 g of proteose peptone, 0.64 g of Na2HPO4, 0.48 g of KH2PO4 dissolved in 1 l of H2O, pH 7.5). A total of 300 ml of this suspension was transferred into the wells of a 24-well plate, and cells were left 30 min at 25.5°C so that they could settle down. Bacteria were adjusted to 10 7 cfu ml -1 in infection medium as determined by OD550. The supernatant and nonadhering Dictyostelium cells were removed from the wells and replaced by 300 ml of this bacterial suspension (MOI 10). After 3 h of infection extracellular bacteria were removed by two washing steps, first with 1¥ Soerensen buffer followed by infection medium containing 100 mg ml -1 gentamicin. In order to kill remaining extracellular bacteria cells were incubated for additional 45 min with 300 ml of infection medium containing 100 mg ml -1 gentamicin. Dead bacteria and gentamicin were removed by two washing steps with infection medium, and the cells were covered with 1 ml of infection medium for the remaining time of infection. At indicated time points cells were collected by resuspending, transferred into 1.5 ml of reaction tubes and mechanically lysed by centrifugation (16 000 g, 6 min) at room temperature and strong vortexing. Serial dilutions in ddH2O were plated out on BCYE-agar for cfu determination. The time point 3 h corresponds to the cfu of intracellular bacteria after gentamicin treatment and was used to determine the uptake of L. pneumophila. To compensate for the differences in bacterial uptake rates between wild-type AX2 and the Datg9 mutant time-course experiments were performed with MOI 8 for the mutant and MOI 1 for the wild type. In these infections comparable numbers of intracellular bacteria were detected 3 h post infection in both strains, and intracellular replication was monitored. Ten independent infections were performed, and cfu values were determined at the different time points post infection. The cfu value 3 h post infection was considered as 1 and used as a reference for calculating the relative changes at later infection time points. Mean values, standard errors of the mean and ratios of the mean values of the relative bacterial titers in AX2 and ATG9cells were calculated. The combined error DR of the ratio R was calculated based on the Gaussian error propagation using the formula: (DR/R)2 = (DA/A)2 + (DB/B)2 with A and B being the mean values of the infections with AX2 and ATG9 -, respectively, and DA and DB being the corresponding standard errors of the mean. A paired t-test was used to calculate the P-values of the different time points.

Antibody generation and Western blotting
For generation of ATG9-specific polyclonal antibodies (pAbs), sequences encompassing the APG9 domain (aa 188-567) were amplified and cloned into a pGEX-4T GST expression vector. The fusion protein was expressed in E. coli DH5a, purified using GST-Sepharose beads and used for the immunization of rabbits (BioGENES GmbH, Berlin, Germany). The resulting ATG9 pAbs were used for Western blotting at a 1:10 000 dilution. GFP was detected with monoclonal antibody K3-184-2 (Noegel et al., 2004). Secondary antibodies used were anti-mouse and antirabbit IgG POD (Sigma, Deisenhofen, Germany).

Miscellaneous methods
For SDS gel electrophoresis the proteins of 2 ¥ 10 5 cells were separated per lane. RNA isolation, northern transfer and real-time PCR were essentially done as described (Farbrother et al., 2006). For real-time PCR the amplified gene fragment from cDNA had a size of 1975 bp. Genomic DNA contains two short introns of 64 and 75 bp respectively. Control amplifications of the generated cDNAs of the different developmental time points with standard PCR resulted in a single band of the expected size after agarose gel electrophoresis. No cross contamination with genomic DNA was observed. The following primers were used. Forward primer: 5′-CGCGGATCCCCAATGATGGACGATCCAG AGG-3′. Reverse primer: 5′-CGCGGTCGACTCTTCCAGTTTT AAATCTTG-3′.