Type II Secretion Is Necessary for Optimal Association of the Legionella-Containing Vacuole with Macrophage Rab1B but Enhances Intracellular Replication Mainly by Rab1B-Independent Mechanisms.

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Type II Secretion Is Necessary for Optimal Association of the Legionella-Containing Vacuole with Macrophage Rab1B but Enhances Intracellular Replication Mainly by Rab1B-Independent Mechanisms Richard C. White, Nicholas P. Cianciotto Department of Microbiology and Immunology, Northwestern University Medical School, Chicago, Illinois, USA

Previously, we documented that type II secretion (T2S) promotes intracellular infection of macrophages by Legionella pneumophila. In the present study, we identified infection events that are modulated by T2S by comparing the behaviors of wild-type and T2S mutant bacteria in murine bone marrow-derived macrophages and human U937 cells. Although the two strains behaved similarly for entry into the host cells and evasion of lysosomal fusion, the mutant was impaired in the ability to initiate replication between 4 and 8 h postentry and to grow to large numbers in the Legionella-containing vacuole (LCV), as evident at 12 h. At 4 h postinoculation, mutant LCVs had a significantly reduced association with Rab1B, a host GTPase that facilitates the tethering of endoplasmic reticulum (ER)-derived vesicles to LCVs. The mutant did not lose expression or translocation of six type IV secretion effectors (e.g., SidM) that are well known for mediating Rab1B association with the LCV, indicating that T2S promotes the interaction between the LCV and Rab1B via a novel mechanism. Interestingly, the mutant’s growth defect was exacerbated in macrophages that had been depleted of Rab1B by short hairpin RNA (shRNA) treatment, indicating that T2S also potentiates events beyond Rab1B association. In support of this, a sidM lspF double mutant had an intracellular growth defect that was more dramatic than that of the lspF mutant (and a sidM mutant) and showed a growth difference of as much as a 400fold compared to the wild type. Together, these data reveal a new role for T2S in intracellular infection that involves both Rab1Bdependent and Rab1B-independent processes.

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egionella pneumophila is widespread in natural and man-made water systems and is best known as the etiologic agent of Legionnaires’ disease (1–3). Recently, the incidence of pneumonias caused by Gram-negative L. pneumophila has roughly tripled in the United States and elsewhere (4–6). In water, L. pneumophila grows primarily as an intracellular parasite of amoebae (7–11). Human infection occurs mainly by the inhalation of contaminated water droplets originating from a wide range of aerosolgenerating devices (12, 13). Yet the first case of person-to-person spread was reported in 2016 (14). In the lungs, L. pneumophila invades and grows in the resident macrophage population and ultimately triggers severe inflammation (3). In both amoebae and macrophages, the bacterium evades the host’s degradative lysosomal pathway and replicates to large numbers within a membrane-bound vacuole, the Legionella-containing vacuole (LCV) (15, 16). The Dot/Icm type IV secretion (T4S) system of L. pneumophila plays a major role in pathogenesis (17–19). Indeed, the Dot/Icm apparatus translocates myriad proteins across the LCV membrane and into the host cell cytoplasm, and then the delivered effectors proceed to modulate a wide variety of host processes. One of the earliest and most notable events that has been ascribed to the Dot/Icm system is the recruitment of host GTPases and endoplasmic reticulum (ER)-derived vesicles to the LCV for the purpose of potentiating bacterial replication (20). L. pneumophila possesses another protein secretion system that also has a major role in the natural history of disease, i.e., the Lsp type II secretion (T2S) system (21–23). In T2S, substrates are first transported across the bacterial inner membrane and then, upon the action of the T2S pilus-like apparatus, exit the cell through a dedicated pore in the outer membrane (24). Based on proteomic

December 2016 Volume 84 Number 12

and enzymatic assays, we determined that the T2S system of L. pneumophila secretes more than 25 proteins, including 18 known enzymes and various novel proteins (25, 26). Within water systems, T2S enhances the survival of L. pneumophila at low temperatures and is essential for infection of amoebae belonging to the genera Acanthamoeba, Naegleria, Vermamoeba (formerly Hartmannella), and Willaertia (26, 27). Among the secreted proteins that promote intracellular infection of amoebae are the novel proteins NttA and NttC as well as the PlaC acyltransferase, ProA metalloprotease, and SrnA RNase (26, 28–30). L. pneumophila T2S also promotes biofilm formation and sliding motility (31, 32). In the lungs, T2S promotes infection of macrophages, suppression of innate immunity, and destruction of tissue (23, 33, 34). Lsp mutants of L. pneumophila that lack T2S, but not their complemented derivatives, are impaired ⬃10-fold for infection of U937 cells, a human macrophage-like cell line, and MH-S cells, a murine mac-

Received 29 August 2016 Accepted 31 August 2016 Accepted manuscript posted online 6 September 2016 Citation White RC, Cianciotto NP. 2016. Type II secretion is necessary for optimal association of the Legionella-containing vacuole with macrophage Rab1B but enhances intracellular replication mainly by Rab1B-independent mechanisms. Infect Immun 84:3313–3327. doi:10.1128/IAI.00750-16. Editor: C. R. Roy, Yale University School of Medicine Address correspondence to Nicholas P. Cianciotto, [email protected]. Supplemental material for this article may be found at http://dx.doi.org/10.1128 /IAI.00750-16. Copyright © 2016, American Society for Microbiology. All Rights Reserved.

Infection and Immunity

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rophage line (33–35). However, the basis for the importance of T2S in macrophage infection has remained undefined. We now report that T2S is required for the optimal association of the host GTPase Rab1B with the LCV, coincident with the initiation of intravacuolar replication between 4 and 8 h after bacterial entry into the macrophage. We further show that T2S continues to promote replication at 12 h and beyond, in a manner that is independent of Rab1B recruitment. MATERIALS AND METHODS Bacterial strains and media. L. pneumophila strain 130b (American Type Culture Collection [ATCC] strain BAA-74) served as our wild-type (WT) strain (36). The lspF mutant NU275, the lspE mutant NU361, and the complemented lspF mutant NU275(pMF) were also described previously (33, 37), as were the chiA mutant NU318, the lapA mutant NU320, the lipA mutant NU262, the map mutant NU254, the rcp mutant NU260, the plaC mutant NU367, the plcA mutant NU268, the proA mutant AA200, and the srnA mutant NU328 (25, 28–30, 38–41). To help monitor the behavior of L. pneumophila in macrophages, we introduced a green fluorescent protein (GFP)-expressing plasmid, pMGFP, into strains 130b and NU275. To that end, a DNA fragment encoding GFP was PCR amplified from pMMB-GRN (42) by use of the primers RW1 and RW2 (see Table S1 in the supplemental material). The EcoRI/HindIII-digested PCR product was then cloned into EcoRI/HindIII-digested pMMB2002 (33), yielding pMGFP. Finally, the plasmid was electroporated into the L. pneumophila strains (43). Legionellae were routinely grown at 37°C on buffered charcoal-yeast extract (BCYE) agar, which, when appropriate, was supplemented with chloramphenicol at 3 ␮g/ml, kanamycin (Kn) at 25 ␮g/ml, or gentamicin at 2.5 ␮g/ml (33). GFP-expressing bacteria were grown on BCYE agar supplemented with 1 mM isopropyl-␤-D-thiogalactopyranoside (IPTG) to induce expression of GFP. To monitor the extracellular growth of L. pneumophila, bacteria were incubated in buffered yeast extract (BYE) broth with shaking at 37°C, and the optical densities of cultures were determined at 660 nm (26). Escherichia coli strain DH5␣ (Life Technologies, Carlsbad, CA) was the host for recombinant plasmids and was grown in Luria-Bertani medium with Kn (50 ␮g/ml), chloramphenicol (30 ␮g/ml), or ampicillin (100 ␮g/ml). Unless otherwise noted, chemicals were obtained from Sigma-Aldrich (St. Louis, MO). Mutant construction. Mutants of L. pneumophila strain 130b containing nonpolar deletions in sidM or dotA were constructed using a form of allelic exchange. To begin, mutagenized alleles were generated using overlap extension PCR (OE-PCR) as previously done (44, 45). The 5= and 3= regions flanking the open reading frames (ORFs) of sidM and dotA were PCR amplified from 130b DNA by use of primers RW3 and RW4 for 5= sidM, RW5 and RW6 for 3= sidM, RW9 and RW10 for 5= dotA, and RW11 and RW12 for 3= dotA (see Table S1 in the supplemental material), as well as HiFi Taq polymerase (Life Technologies), and the Kn resistance cassette flanked by Flp recombination target sites was similarly PCR amplified from pKD4 (44) by use of primers RW7 and RW8 for sidM and RW13 and RW14 for dotA (see Table S1). We then performed two-step OE-PCR to combine the 5= and 3= regions of sidM or dotA with the respective Kn resistance cassette. PCR products corresponding to the correct target sizes were gel purified and ligated into pGEM-T Easy (Promega, Fitchburg, WI) to yield either pGsidM::Kn or pGdotA::Kn. After transforming (46) strain 130b with the newly made plasmids, bacteria containing an inactivated sidM or dotA gene were obtained by plating on BCYE agar containing Kn and verified by PCR using primers RW19 and RW20 or RW21 and RW22 (see Table S1). Next, following electroporation (43) of pBSFLP (44) into the sidM mutant and subsequent plating on BCYE agar containing 1 mM IPTG and gentamicin, the colonies obtained were patched onto ordinary BCYE agar in order to promote the loss of pBSFLP. Clones that were sensitive to gentamicin and Kn were isolated, and the loss of the plasmid and the chromosomal antibiotic cassette (leaving only an unmarked deletion) was confirmed by PCR. The new sidM deletion mutant was designated NU427, and the dotA deletion mutant was designated

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NU428. To construct a double mutant lacking both sidM and T2S, we introduced pGlspF::Kn (33) into NU427 by transformation (46) and then selected for the acquisition of Kn resistance. The genotype of the sidM lspF double mutant, designated strain NU429, was verified by PCR using primers RW47 and RW48 (see Table S1). A complemented derivative of the sidM lspF mutant was obtained by introducing the lspF-encoding plasmid pMF (33) into NU429 by electroporation. Macrophage cell lines and intracellular infection assays. In order to assess L. pneumophila growth within mammalian cells, we utilized bone marrow-derived (BMD) macrophages that were obtained from A/J mice (Jackson Laboratory, Bar Harbor, ME) and human U937 cells (ATCC CRL-1593.2) terminally differentiated into macrophages by treatment with phorbol myristate acetate (47). To aid in determining the effect of Rab1B on L. pneumophila growth, we generated a stable U937 Rab1Bknockdown cell line. Silencing was achieved by transduction with a vesicular stomatitis virus glycoprotein (VSV-G)-pseudotyped lentivirus that carries a GFP reporter gene and encodes a short hairpin RNA (shRNA) molecule targeting Rab1B, i.e., clone V2LHS_137920 from OpenBioSystems (GE Healthcare, Chalfont, United Kingdom). As a control, we generated a lentivirus encoding a nontargeting shRNA molecule (“scramble”), i.e., clone RHS4346 (GE Healthcare). Lentiviral vectors were prepared from HEK293T cells (ATCC CRL-3216) as previously described (48), and then a 2.5-ml suspension of undifferentiated, replicating U937 cells (approximately 2 ⫻ 106) in RPMI medium (Corning, Corning, NY) with 10% fetal bovine serum (FBS) (Atlanta Biologicals, Flowery Branch, GA) was mixed with 750 ␮l unconcentrated virus in the presence of 4 ␮g/ml Polybrene (Santa Cruz Biotechnology, Dallas, TX) in a 6-well dish (Falcon; Becton Dickinson, Franklin Lakes, NJ). The resulting cell suspension was transduced upon centrifugation at 1,000 ⫻ g for 90 min at 32°C followed by incubation at 37°C. After 16 h, the medium was replaced with fresh RPMI medium containing FBS (RPMI-FBS) to remove the Polybrene. After 3 days of further incubation at 37°C, transduced knockdown cells were selected by incubation in the presence of 1 ␮g/ml puromycin (Corning) for 2 or 3 passages, at which time all cells in the suspension were GFP positive. Silencing of Rab1B was confirmed by immunoblot analysis. To that end, cell pellets were lysed in Laemmli sample buffer and analyzed via SDS-polyacrylamide gel electrophoresis (SDS-PAGE). Proteins were transferred to a polyvinylidene difluoride (PVDF) membrane (Thermo Scientific, Waltham, MA), blocked in Tris-buffered saline with 0.1% Tween 20 (TBS-T) and 5% milk (Bio-Rad Laboratories, Hercules, CA), and probed with a rabbit polyclonal anti-Rab1B antibody (sc-599; Santa Cruz Biotechnology) at 1:250 for 12 h at 4°C. Proteins were finally detected using an anti-rabbit IgG– horseradish peroxidase conjugate (Cell Signaling Technology, Danvers, MA) and an enhanced chemiluminescence (ECL) detection reagent (GE Healthcare). Densitometry analysis of the blots was performed using Image Lab software (Bio-Rad). In the standard infection assay done at 37°C (34), 2.5 ⫻ 105 BMD macrophages or 1 ⫻ 106 differentiated U937 cells were seeded in triplicate into 24-well plates (Falcon) in RPMI-FBS and inoculated with L. pneumophila that had been grown to post-exponential phase on BCYE agar for 3 days. As described before, bacteria were added at a multiplicity of infection (MOI) (i.e., ratio of added bacteria to host cells) equal to 1 for BMD macrophages or 0.5 for U937 cells. Following a 2-h incubation to allow for bacterial uptake, the monolayers were washed extensively to remove remaining extracellular legionellae, and then the infection was continued in the presence of fresh RPMI-FBS for up to 3 days. In infection assays using GFP-expressing bacteria, RPMI-FBS was supplemented with 1 mM IPTG. For CFU determination, the macrophages were lysed with 0.1% saponin, and then dilutions of the lysates were plated onto BCYE medium. As a variation on this protocol, monolayers were inoculated at an MOI of 1 or 50, and then the bacteria were centrifuged onto the macrophages (250 ⫻ g for 5 min) in order to hasten and synchronize the infection event, as done previously (49–51). After 5 min of incubation at 37°C to permit bacterial uptake (49), monolayers were washed with RPMI medium and then exposed to 200 ␮g/ml gentamicin (Corning) in 1 ml RPMI-FBS for 1



Legionella Type II Secretion and Macrophage Infection

h to kill any remaining extracellular legionellae. After another series of three washes to remove the antibiotic, the infected monolayers (n ⫽ 4) were incubated for another 2 to 72 h and then lysed, and the numbers of CFU were determined by plating. By measuring the levels of lactate dehydrogenase released into the tissue culture supernatants as previously described, using a CytoTox 96 nonradioactive cytotoxicity assay kit (Promega) (52), we confirmed that the infections done using an MOI of 50 did not result in contact-dependent cytotoxicity for the host cells (data not shown). Assessment of bacterial entry into macrophages. To assess the extent of L. pneumophila entry into macrophages, a gentamicin protection assay was done (36, 53). Briefly, the bacterial strains were added to the BMD macrophages at an MOI equal to 1 or 50, and then the infection was synchronized as described above. Following gentamicin treatment to kill residual extracellular legionellae and a final wash step as detailed above, the infected monolayers (n ⫽ 4) were lysed and the numbers of internalized CFU were determined by plating. Entry was measured as the percentage of intracellular CFU recovered from the monolayers compared to the number of CFU in the inoculum. As a second means of monitoring bacterial entry, we utilized a trypan blue quenching-based entry assay which has been described previously for L. pneumophila (54–56). To that end, BMD macrophages (5 ⫻ 104) or differentiated U937 cells (1 ⫻ 105) were seeded into black, clear-bottomed, 96-well microtiter plates (Falcon), and bacteria suspended in RPMI medium lacking phenol red (in order to reduce background fluorescence) were added at an MOI of 50 to six replicate wells. After synchronizing the infections as described above, a fluorescence reading (excitation, 485 nm; emission, 530 nm) was taken using a Synergy H1 hybrid microplate reader (BioTek, Winooski, VT) in order to determine the initial amount of fluorescent legionellae in the infected wells. Following the removal of extracellular bacteria by washing, 50 ␮l of trypan blue solution (54) was added for 1 min to quench any residual extracellular fluorescence. Finally, another fluorescence reading was taken to determine the amount of trypan blue-resistant fluorescence (i.e., intracellular legionellae) remaining in the wells. Entry was defined as the percentage of resistant fluorescence over total fluorescence. Phagosome-lysosome fusion assays. As a first method for assessing the fusion of LCVs with macrophage lysosomes, we examined infected monolayers by using the marker LysoTracker Red, as previously done for the study of L. pneumophila (57–59). BMD macrophages (1 ⫻ 105) or differentiated U937 cells (2 ⫻ 105) were seeded onto sterile circular glass coverslips (Fisher Scientific, Waltham, MA) which had been placed within a 24-well dish and then infected at an MOI of 50 with synchronization. As a positive control, macrophages were also exposed to WT bacteria that had been heat killed by incubation at 65°C for 10 min (60). Following removal of extracellular bacteria by washing, LysoTracker Red (Life Technologies) was added to 75 nM in RPMI-FBS, and after 2 h, the colocalization of bacteria and LysoTracker Red was assessed using immunofluorescence microscopy. To that end, infected macrophages were fixed in 2% paraformaldehyde (Electron Microscopy Sciences, Hatfield, PA), permeabilized in ice-cold methanol, and then blocked in PBS containing 2% goat serum (Life Technologies) as previously described (61). To then detect intracellular bacteria, the coverslips were incubated with the mouse monoclonal anti-Legionella antibody MAB2 (62, 63) at 1:10,000 for 1 h at 37°C, washed, and then treated with goat anti-mouse–Oregon Green (Life Technologies) at 1:10,000 for 1 h at 37°C. After final washes, coverslips were mounted on glass slides (Fisher Scientific) by use of ProLong Diamond antifade mounting reagent with DAPI (4=,6-diamidino-2-phenylindole) (Life Technologies). Finally, the infected cells were inspected with an Evos FL cell imaging system, using the manufacturer’s DAPI, Texas Red, and GFP filter cubes (Advanced Microscopy Group, Life Technologies). As a second method for monitoring phagosome-lysosome fusion, we utilized the Texas Red ovalbumin (TROv) (Life Technologies) detection assay, which has been applied previously to the study of L. pneumophila (64, 65). Following prelabeling with 50 nM TROv for 30 min and a 30-min chase with fresh RPMI-FBS, BMD macrophages were infected at


an MOI of 50 with synchronization, and at 2, 4, and 6 h, the infected cells were examined for colocalization of TROv and bacteria by immunofluorescence microscopy as detailed above. For the LysoTracker Red- and TROv-based assays, at least 100 infected cells were scored per strain per experiment. Microscopic assessment of intravacuolar replication. GFP-expressing bacteria were added to 7.5 ⫻ 104 BMD macrophages on coverslips at an MOI of 5, and then the infection was synchronized as noted above. Following the usual wash step, the infected monolayers were replenished with RPMI-FBS supplemented with 1 mM IPTG in order to induce GFP expression and then incubated for another 4, 8, or 12 h. Finally, the infected macrophages were processed for immunofluorescence microscopy, and the number of GFP-expressing legionellae per LCV was determined by visual inspection (66–68). However, to precisely distinguish intracellular legionellae from any residual extracellular bacteria, the monolayers were treated with anti-L. pneumophila antibody and then with goat antimouse–Alexa Fluor 594 (Life Technologies) at 1:250 for 1 h in the absence of permeabilization. Thus, only LCVs containing bacteria expressing GFP but not labeled with Alexa Fluor 594 were scored. For this assay, we counted the number of bacteria within at least 300 vacuoles per strain per experiment, using an Evos FL cell imaging system with DAPI and GFP filter cubes. Measurement of the association of Rab1B with the LCV. BMD macrophages (1 ⫻ 105) or differentiated U937 cells (2 ⫻ 105) were infected with L. pneumophila strains at an MOI of 50, as detailed above, and then incubated for another 1, 2, 4, 6, and 8 h. The samples were then fixed and permeabilized as described above. In addition to being treated with the anti-L. pneumophila monoclonal antibody to detect intracellular legionellae in LCVs, the coverslips were incubated with rabbit polyclonal antiRab1B at 1:100 for 1 h at 37°C followed by goat anti-rabbit–Alexa Fluor 594 (Life Technologies) at 1:250 for 1 h at 37°C. The colocalization of LCV and Rab1B was then assessed using immunofluorescence microscopy, with at least 100 infected cells scored per strain per experiment, using an Evos FL cell imaging system with DAPI, Texas Red, and GFP light cubes. qRT-PCR analysis. Bacterial RNA was isolated from infected BMD macrophages by use of TRIzol reagent (Life Technologies) according to the manufacturer’s instructions. Briefly, after removal of the culture medium, the monolayers were lysed with TRIzol, and RNA was extracted via phase separation (69). Samples were then treated with RQ1 DNase (Promega) for 45 min at 37°C. The purity and concentration of the RNA were confirmed by spectrophotometry (NanoDrop; Thermo Scientific). cDNA was synthesized from the DNase-treated RNA in a 20-␮l reaction mixture as previously described (47). The PCR primers used to detect transcripts from the L. pneumophila chiA (RW37 and RW38), gyrB (RW39 and RW40), lspF (RW41 and RW42), plaC (RW43 and RW44), and srnA (RW45 and RW46) genes are listed in Table S1 in the supplemental material. Quantitative reverse transcription-PCR (qRT-PCR) was done as described before (47), using a SYBR green probe and a CFX96 Touch real-time PCR detection system (Bio-Rad), with the only modification being the addition of 5 more amplification cycles. The primers used for qRT-PCR analysis were designed using the Primer-BLAST tool on the NCBI website (70) for the sidM (RW23 and RW24), ankX (RW25 and RW26), lidA (RW27 and RW28), lepB (RW29 and RW30), lem-3 (RW31 and RW32), sidD (RW33 and RW34), and dotA (RW35 and RW36) genes, and they are listed in Table S1. The relative RNA abundance was determined using the ⫺⌬CT method (CT for reference ⫺ CT for gene of interest) (71), with gyrB used as the reference gene (47, 72, 73). T4S translocation assay. In order to study the translocation efficiency of the T4S system in WT- versus mutant-infected macrophages, effector protein genes were cloned just downstream of the ␤-lactamase (Bla) gene in pXDC61, a derivative of pMMB2002 (74), creating fusion proteins with the Bla reporter at the N terminus. Plasmids pXlidA, pXlepB, pXralF, pXsidM, and pXankX, encoding Bla fusions with LidA, LepB, RalF, SidM, and AnkX, respectively, were a kind gift from Howard Shuman, University of Chicago (54). To also study the translocation of the T4S effectors


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FIG 1 Intracellular infection of A/J mouse macrophages by WT and T2S mutant L. pneumophila strains. BMD macrophages were inoculated with WT strain 130b (black circles), the lspF mutant NU275 (white squares), or the complemented mutant NU275(pMF) (black triangles) at an MOI of 1 (A) or 50 (B), and the numbers of CFU in the infected wells were determined at the indicated time points. Growth is presented as the fold change in number of CFU over that at time zero. Results are the means and standard deviations for triplicate wells and are representative of three independent experiments. Asterisks indicate significant differences in CFU recovery between the mutant and the WT or the complemented mutant (Student’s t test; *, P ⬍ 0.05; **, P ⬍ 0.01; ***, P ⬍ 0.001).

Lem3 and SidD, we constructed plasmids pXlem3 and pXsidD by using previously described methods (74). Thus, the lem-3 and sidD ORFs were amplified by use of primer pairs RW15-RW16 and RW17-RW18, respectively (see Table S1 in the supplemental material). The lem-3 PCR product was digested with KpnI and XbaI and then cloned into KpnI/XbaI-digested pXDC61, whereas the sidD product was digested with SmaI and XbaI and then cloned into similarly treated pXDC61. The in-frame insertions of Bla were confirmed by PCR and DNA sequence analysis. Finally, pXlidA, pXlepB, pXralF, pXsidM, pXankX, pXlem3, and pXsidD were introduced into WT strain 130b, the lspF mutant NU275, and the dotA mutant NU428 by electroporation. Assessments of protein translocation out of the LCV and into the host cell cytosol were performed as previously described (54, 74–76). Briefly, BMD macrophages were seeded into black, clear-bottomed microtiter plates at 5 ⫻ 104 cells per well for 72 h prior to infection. L. pneumophila strains carrying the various Bla fusions were grown for 24 h on BCYE agar containing chloramphenicol and 0.5 mM IPTG to induce production of the hybrid proteins. Bacteria were then added to the macrophages at an MOI of 50, and the infection was synchronized by centrifugation (600 ⫻ g, 10 min). Following a 1-h incubation, the infected cells were exposed to the Bla substrate CCF2-AM (Life Technologies) in the presence of 15 mM probenecid (Santa Cruz Biotechnology), which promotes retention of CCF2-AM. Finally, after a further 1 h of incubation, the levels of fluorescence generated in the macrophage cytoplasm due to the cleavage of CCF2-AM by the translocated Bla fusions were quantified on a BioTek fluorescence microplate reader, with excitation at 405 nm and emission at 460 nm and 530 nm (74). Translocation was measured as the 460-nm/530-nm emission ratio adjusted for background fluorescence. To confirm the expression of the SidD fusion protein, which did not show evidence of translocation (see Fig. 6 and below), bacteria containing pXsidD were examined by immunoblot analysis using anti-Bla antibodies (QED Bioscience, San Diego, CA) at 1:200 in TBS-T containing 5% nonfat dry milk. Assessment of bacterial lysis within infected macrophages. The level of lysis of intracellular Legionella was determined essentially as previously described (67). BMD macrophages were infected with GFP-expressing L. pneumophila strains at an MOI of 50 as described above. At 4 h, the RPMI medium was removed from the wells and replaced with 1 ml of fresh medium containing 0.2% saponin in order to lyse the infected macrophages but not the intact bacteria contained within them. After dislodging the macrophages with a cell scraper (Falcon), the entire content of each well was centrifuged in a microcentrifuge tube at 8,000 ⫻ g for 10 min at 4°C. The resultant pellet, containing intact bacteria and large cellular debris, was resuspended and lysed in Laemmli buffer. The soluble proteins in the supernatant were isolated and concentrated via methanol-chloroform extraction (77) and then reconstituted in Laemmli buffer. Cellular and soluble fractions were subjected to SDS-PAGE on a 4 to 20% gradient

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gel (Bio-Rad), and the separated proteins were transferred to a PVDF membrane. After incubating for 12 h at 4°C in TBS-T containing 5% nonfat dry milk, the membranes were treated with rabbit anti-GFP antibody (Cell Signaling Technology) at a dilution of 1:1,000 in TBS-T containing 5% nonfat dry milk for 12 h at 4°C. As a loading control, membranes were also treated with mouse anti-glyceraldehyde-3-phosphate dehydrogenase (anti-GAPDH) (Santa Cruz Biotechnology) at 1:400, which detects a host cytoplasmic protein that is abundant in the soluble fraction. Following washes, the blots were incubated with anti-rabbit or anti-mouse IgG– horseradish peroxidase conjugate (Cell Signaling Technology) at a dilution of 1:1,000 in TBS-T containing 5% dry milk for 1 h at room temperature and then developed using ECL detection reagent, and the immunoblot was analyzed using Image Lab software as noted above.

RESULTS

T2S promotes L. pneumophila infection of A/J mouse macrophages. We previously demonstrated that the L. pneumophila T2S system is required for optimal infection of human macrophages, as an lspF mutant of strain 130b that lacks T2S, but not a complemented lspF mutant, exhibited an approximately 10fold reduction in CFU in infected U937 cell monolayers (33). We also determined that T2S promotes L. pneumophila replication and persistence in the lungs of A/J mice (25, 33, 34). Thus, to begin this study, we compared WT strain 130b, its lspF mutant, and the complemented mutant for the ability to replicate within monolayers of BMD macrophages from A/J mice over a 3-day incubation period. In this assay, initially infected macrophages are lysed by 24 h, and then one or two rounds of reinfection follow over the next 24 to 48 h. After inoculation at an MOI equal to 1, significantly fewer mutant CFU were recovered from the monolayers at 24, 48, and 72 h postinoculation (Fig. 1A). When the MOI was increased to 50, fewer mutant bacteria were again observed (Fig. 1B). Importantly, the complemented lspF mutant, carrying an intact copy of lspF on a plasmid, behaved like the WT strain did (Fig. 1A and B), confirming that the mutant defect was due to the loss of T2S. Together, these data confirmed that T2S promotes L. pneumophila infection of A/J macrophages and does so over a range of infectious doses and during multiple rounds of infection. Since the lspF mutant defect was seen in the first 24 h, as is the case for infected U937 cells (33), we posited that T2S promotes the initial round of infection, helping to mediate bacterial entry, resistance to immediate intracellular killing, and/or replication itself.




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FIG 2 Entry of WT and T2S mutant L. pneumophila strains into macrophages. (A) BMD macrophages (BMDM) were infected with WT strain 130b (black bars) or the lspF mutant NU275 (white bars) at the indicated MOIs, and following treatment with gentamicin, the numbers of intracellular CFU were determined. Entry was calculated as the percentage of gentamicin-resistant (i.e., intracellular) CFU relative to the inoculum CFU, and data are presented as means and standard errors for three independent experiments for each MOI tested. (B) As indicated, BMD macrophages and U937 cells were infected with GFP-expressing WT strain 130b (black bars) or lspF mutant NU275 (white bars) bacteria at an MOI of 50, and following quenching of extracellular fluorescence by the addition of trypan blue solution, the remaining (intracellular) fluorescence was determined. Entry was calculated as the percentage of trypanresistant GFP intensity relative to the initial GFP intensity, and data are presented as means and standard errors for three independent experiments for each cell type.

T2S is dispensable for entry of L. pneumophila into macrophages. As the first step toward explaining the role of T2S in intracellular infection, we assessed the contribution of T2S to L. pneumophila entry into host cells. BMD macrophages were briefly exposed to WT 130b and the lspF mutant at an MOI of 1 or 50, and after the removal of residual extracellular bacteria by treatment with gentamicin, intracellular bacteria were counted by plating. At either MOI, the entry levels of WT and mutant bacteria were comparable (Fig. 2A). The observation that 1 to 3% of the inoculum entered into the macrophages is similar to past observations made with L. pneumophila and this type of gentamicin protection assay (53). To confirm our results, we utilized a trypan blue quenching method that was recently adapted to assess L. pneumophila entry into host cells (54–56). In this case, we infected the BMD macrophages with GFP-expressing bacteria at an MOI equal to 50, and after treatment with trypan blue to quench residual extracellular bacteria, the level of trypan blue-resistant fluorescence remaining associated with the monolayer (i.e., intracellular bacteria) was assessed using a fluorescence microplate reader. By this method, the

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entry levels of WT and mutant bacteria were again comparable (Fig. 2B). Moreover, the overall levels of entry detected were similar to those obtained using the gentamicin protection assay. When the assay was done using U937 cells, there appeared to be an overall increase in bacterial entry, which is compatible with the results of a previous study that compared U937 cells and BMD macrophages (67). However, there was still no difference between the WT and the lspF mutant (Fig. 2B). In sum, these data showed that the lspF mutant is not impaired for entry into A/J macrophages or human U937 cells, indicating that T2S is dispensable for L. pneumophila entry into macrophages. T2S is dispensable for L. pneumophila evasion of the host degradative pathway. Shortly after its entry into macrophages, L. pneumophila evades trafficking to the endolysosomal pathway and thereby avoids acidification and degradation (78, 79). Thus, as the next step toward defining the role of T2S in intracellular infection, we compared the WT and lspF mutant strains for the ability to escape this degradative pathway in BMD macrophages, as measured by bacterial colocalization with LysoTracker Red at 2 h postinoculation. As expected (79, 80), WT 130b evaded the lysosomal compartment (3%), whereas the heat-killed WT strain did not (83%) (Fig. 3A). The lspF mutant displayed a very low level of colocalization that was identical to that of the WT (Fig. 3A). To confirm these results, we monitored colocalization using a different marker, namely, TROv (81). Again, the WT and mutant strains exhibited equivalent abilities to evade lysosomes (4%), whereas heat-killed bacteria readily trafficked to the degradative compartment (89%) (Fig. 3B). The capacity of the lspF mutant to evade degradation was evident not only at 2 h but also at 4 and 6 h postinoculation. Together, these data indicated that the T2S mutant was not preferentially delivered to degradative lysosomes during infection of murine macrophages. The same outcomes were obtained when we assessed the colocalization of bacteria and LysoTracker Red in human U937 cells, i.e., 4% for the WT and mutant strains and 75% for the heat-killed WT (Fig. 3C). Thus, we concluded that T2S is dispensable for the ability of L. pneumophila to evade the macrophage’s degradative pathway, and we suspected that the lspF mutant is defective in the later, replicative phase of intracellular infection. T2S, although dispensable for early intracellular survival, is required for intravacuolar replication. To begin to determine when T2S becomes important during infection, we exposed BMD macrophages to WT 130b and the lspF mutant and then enumer-

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FIG 3 Evasion of host lysosomes by WT and T2S mutant L. pneumophila strains. BMD macrophages were infected with WT strain 130b (black bars), the lspF mutant NU275 (white bars), or heat-killed WT bacteria (gray bars) at an MOI of 50, and at the indicated time points, the percent colocalization of bacteria with either LysoTracker Red (LT) (A) or TROv (B) was determined by immunofluorescence microscopy. (C) U937 cells were infected as described above, and bacterial colocalization with LT was determined at the indicated time point. The data in all panels are presented as means and standard errors for three independent experiments. In all cases, heat-killed WT bacteria displayed a significantly higher level of association with the lysosomal markers than did either WT or lspF mutant bacteria (Student’s t test; ***, P ⬍ 0.001).



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FIG 4 Early survival and replication of WT and T2S mutant L. pneumophila strains in macrophages. (A and B) BMD macrophages were infected with WT strain 130b (black circles), the lspF mutant NU275 (white squares), or the complemented lspF mutant NU275(pMF) (black triangles) at an MOI of 1, and following treatment with gentamicin, the numbers of intracellular CFU were determined by plating at the indicated time points. Growth was measured as the percent CFU relative to the number of CFU at time zero, and data are presented as means and standard errors for four (A) or five (B) independent experiments. #, significant reductions in both WT and mutant CFU numbers relative to those at time zero (one-sample t test; P ⬍ 0.01); asterisks indicate time points at which there were significantly fewer mutant bacteria than WT or complemented lspF mutant bacteria recovered (Student’s t test; *, P ⬍ 0.05; **, P ⬍ 0.01; ***, P ⬍ 0.001; ns, not significant). (C and D) BMD macrophages were infected with WT (black bars) or lspF mutant (white bars) bacteria at an MOI of 5, and at the indicated time points, the numbers of bacteria within LCVs were determined using immunofluorescence microscopy. Growth was judged by recording the percentage of LCVs that had 1, 2, 3 to 9, or ⱖ10 bacteria, and data are presented as means and standard errors for three experiments. Asterisks indicate LCVs for which there were significant differences between the lspF mutant and the WT (Student’s t test; *, P ⬍ 0.05; **, P ⬍ 0.01).

ated intracellular CFU at 0, 2, 4, and 12 h postinoculation. Between 0 and 4 h, WT numbers declined approximately 3-fold (Fig. 4A), indicating killing by the macrophages, which was likely due at least in part to the low level of lysosome association that we observed (Fig. 2B). Compatible with the results obtained using LysoTracker Red and TROv, there were no significant differences between the WT and mutant strains in terms of numbers of CFU recovered at 2 and 4 h (Fig. 4A), supporting the view that T2S is dispensable for early intracellular survival. Between 4 and 12 h, the WT clearly began to replicate, showing a 17-fold increase in the number of CFU recovered (Fig. 4A). The fact that L. pneumophila begins replication at approximately 4 h postentry has been observed previously in studies using various other WT strains and macrophage hosts (67). At 12 h, the lspF mutant had 3-fold fewer CFU than the WT, revealing the emergence of a significant mutant phenotype between 4 and 12 h (Fig. 4A). To more precisely determine when the mutant begins to display its growth defect, we repeated the infection assay but enumerated intracellular bacteria at both 8 and 12 h postinoculation. At the 8-h time point, the mutant displayed 2-fold fewer CFU than the WT (Fig. 4B), implying that T2S begins to promote intracellular replication sometime between 4 and 8 h after entry into the macrophage. Once again, the mutant showed approximately 3-fold fewer CFU at 12 h (Fig. 4B). The complemented lspF mutant behaved as the WT did at both 8 and 12 h (Fig. 4B), confirming that the mutant’s growth defect at these early time points was due to the loss of T2S. To support the results obtained from the CFU recovery assay, we infected BMD

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macrophages with GFP-expressing WT and lspF mutant bacteria and used immunofluorescence microscopy to count the numbers of legionellae within individual LCVs. As a prelude to this analysis, we confirmed that the expression of GFP did not alter the ability of either the WT or the lspF mutant to grow in the macrophage host (see Fig. S1 in the supplemental material). Based on the microscopic counting method, there was little evidence of intracellular replication at 4 h postinoculation, since ⱖ90% of WT LCVs had only a single bacterium, and compatible with the results of the CFU recovery assay, the lspF mutant behaved comparably to the WT at this early time point (Fig. 4C). By 8 h, the WT had obviously begun to replicate, as demonstrated by the marked reduction in the number of LCVs harboring 1 bacterium and by ⬎70% of the LCVs having at least 3 bacteria (Fig. 4C). As in the CFU recovery assay, the mutant phenotype emerged by the 8-h time point, i.e., more mutant LCVs contained only 1 bacterium, and there were also fewer mutant LCVs that had ⱖ10 bacteria, although the latter difference was not quite statistically significant (Fig. 4C). When we repeated the immunofluorescence detection experiment by assaying at both 8 and 12 h postentry, the mutant growth defect was even more apparent at 12 h (Fig. 4D). Whereas the most common WT LCVs (at 56%) were those that contained ⱖ10 bacteria, approximately 40% of mutant LCVs still contained 1 bacterium, and only about 29% of the mutant LCVs had ⱖ10 bacteria. For both the WT and the mutant, the proportion of LCVs containing 1 bacterium did not decrease further between 8 and 12 h (Fig. 4D), suggesting that the replicative capacity of the LCV is defined





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sometime between 4 and 8 h. Together, these data indicated that the T2S mutant was impaired not only in the ability to initiate replication between 4 and 8 h but also in the capacity to grow to large numbers within LCVs at 12 h. Thus, we concluded that L. pneumophila T2S first promotes intravacuolar growth sometime between 4 and 12 h after bacterial entry, which explains, at least in part, its contribution to growth during more extended incubation periods, such as in Fig. 1. T2S promotes the association of host Rab1B with the LCV. A hallmark of the LCV is its association with various smooth vesicles derived from the endoplasmic reticulum (ER)-to-Golgi trafficking pathway (82–85). These host vesicles are thought to deliver membrane material, foodstuffs, and other “cargo” that facilitate intravacuolar growth of L. pneumophila (86). Among the host GTPases that modulate ER-to-Golgi vesicle traffic, Rab1B is well known for being recruited rapidly and substantially to LCVs in A/J BMD macrophages and U937 cells as well as murine RAW264.7 macrophages, human A549 lung epithelial cells, and human HEK293 embryonic kidney cells (61, 66, 67, 87–98). Expression of a dominant negative form of Rab1 restricts L. pneumophila growth in Chinese hamster ovary (CHO) cells and COS1 cells, suggesting that Rab1B recruitment enhances bacterial growth (61, 66). Thus, we explored the possibility that T2S is needed for the association of Rab1B with the LCV. To begin to test this, we infected BMD macrophages with WT 130b and its lspF mutant and then monitored the association of endogenous Rab1B with the LCVs by using immunofluorescence microscopy. At 1 and 2 h postinoculation, WT and mutant LCVs exhibited identical levels of Rab1B association, which reached a maximum of 60% (Fig. 5A, left panel), indicating that T2S is not required for the initial recruitment of Rab1 to the LCV. Yet when we examined the infected cells at 4 h, there was a significant (37%) reduction in Rab1B association with the mutant LCVs but not the WT LCVs (Fig. 5A, left panel; see Fig. S2 in the supplemental material for representative images showing the presence versus absence of colocalization), suggesting that the lspF mutant had an impaired ability to continue Rab1B recruitment between 2 and 4 h or to retain the Rab1B that had already been recruited. To determine if the difference between the WT and mutant strains changed over time, we analyzed the Rab1B association at 4, 6, and 8 h postentry. Compatible with previous observations (89, 92), Rab1B association with LCVs containing the WT declined between 4 h and 8 h (Fig. 5A, right panel). At the 6- and 8-h time points, the lspF mutant LCVs displayed levels of Rab1B association that were now comparable to those with the WT LCVs. Together, these data indicated that the lspF mutant had a transient defect in Rab1B association that was seen 4 h after bacterial entry into BMD macrophages. A second, independently derived T2S mutant (i.e., the lspE mutant NU361) exhibited the same inability to associate with Rab1B (Fig. 5B), indicating that the defect we observed was due to the loss of T2S, not to a spontaneous second-site mutation(s). Further support for a role for T2S in Rab1B association was obtained when reintroduction of an intact copy of lspF into the lspF mutant restored Rab1B association to WT levels (Fig. 5B). The observation that the lspF mutant showed neither increased trafficking to lysosomes nor impaired survival at 4 h postinoculation (Fig. 3B and 4A and C) implied that its reduction of Rab1B association was not an artifact of there being more dead or damaged bacteria within the LCVs. Nonetheless, to directly determine whether the lspF mutant undergoes increased lysis, we infected

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FIG 5 Colocalization of Rab1B with WT and T2S mutant L. pneumophila strains. BMD macrophages (A and B) or U937 cells (C) were infected at an MOI of 50 with either WT strain 130b (black circles or bars), the lspF mutant NU275 (white squares or bars), the lspE mutant NU361 (gray bar), the complemented lspF mutant NU275(pMF) (checkered bar), or the sidM mutant NU427 (hatched bar), and at the indicated time points, the colocalization of Rab1B with the LCVs was ascertained using immunofluorescence microscopy. Results are presented as percentages of LCVs that stained positive for Rab1B, and the data are presented as means and standard errors for at least three independent experiments. Asterisks indicate cases where a mutant strain was significantly different from the WT (A to C) or the complemented mutant (B) (Student’s t test; *, P ⬍ 0.05; **, P ⬍ 0.01; ***, P ⬍ 0.001).

BMD macrophages with WT and mutant strains expressing cellassociated GFP and then, at 4 h postinoculation, assayed levels of GFP released from the bacteria into the soluble fraction of the host cells. Similarly low levels of released GFP were seen for the lysates obtained from WT- versus mutant-infected cells (see Fig. S3 in the supplemental material), affirming that the T2S mutant does not undergo more lysis than the WT. The T2S mutant defect in Rab1 trafficking was also not simply a nonspecific effect of impaired bacterial growth, since it occurred at 4 h, prior to the onset of the mutant replication defect (Fig. 3). In support of this conclusion, a previously described rcp mutant of strain 130b (40), which has normal T2S but has a replication defect that is comparable to that of the lspF mutant, displayed a level of Rab1B association that was identical to that of the WT (see Fig. S4). Moreover, there are various types of L. pneumophila mutants that are also defective for intracellular growth but do not show any alteration in Rab1 association (67, 99). Thus, we concluded that the T2S mutant defect in Rab1B association is physiologically relevant and worthy of further investigation. As noted above, at least 15 studies have documented the association between Rab1B and LCVs; however, only 3 of these studies used human macrophages, and in no case was there a direct comparison between murine and human host cells (66, 93, 97). Therefore, we infected U937 cells with WT 130b and its lspF mutant and monitored the association of Rab1B with the LCVs, using the same immunofluorescence assay as that used to assess BMD mac-


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rophages. Approximately 60% of WT LCVs had a Rab1B association at 4 h (Fig. 5C; see Fig. S2 in the supplemental material), documenting that Rab1B associations are comparable between murine and human macrophages. The lspF mutant, once again, displayed a reduction in the number of Rab1B⫹ LCVs (Fig. 5C; see Fig. S2), demonstrating that the role of T2S in promoting Rab1B association exists in both murine and human macrophages. The role of T2S in Rab1B association is not due to an effect on the expression or translocation of key T4S effectors. The L. pneumophila Dot/Icm T4S system is very well known for its role in modulating the association of Rab1B with the LCV (20, 100). The T4S apparatus translocates myriad effectors across the LCV and into the host cell cytosol, and a subset of those effectors (i.e., AnkX [LegA8], Lem3, LepB, LidA, SidD, and SidM [DrrA]) mediate the tethering of Rab1B to the LCV membrane or the later release of the Rab1 protein (87–89, 91, 92, 101–108). Therefore, we considered the possibility that the lspF mutant’s impaired association with Rab1B might be due to an artifact of an absent T2S system relevant to the expression or translocation function of the T4S system. To assess effects on gene expression, we infected BMD macrophages with WT and lspF mutant bacteria for 4 h and then, using quantitative RT-PCR analysis, analyzed the levels of ankX, lem-3, lepB, lidA, sidD, and sidM transcripts as well as mRNA levels for dotA, which encodes a key component of the T4S apparatus (79). In all cases, the levels of transcripts were comparable between WT and mutant samples (Fig. 6A), indicating that the lspF mutation and the resultant lack of T2S had neither indiscriminately diminished the expression of T4S nor specifically dampened key effector genes. Incidentally, we did observe that the T4S genes were expressed to differing degrees at the 4-h time point, with lepB and lidA being expressed most highly and ankX being the least expressed gene (Fig. 6A). Next, to investigate a potential effect of the lspF mutation on the translocation function of the Dot/Icm system, we used a ␤-lactamase (Bla) reporter assay that has been used extensively to monitor L. pneumophila T4S within infected macrophages (74, 75, 109, 110). To that end, plasmids expressing a fusion between AnkX, Lem3, LepB, LidA, SidD, or SidM and the Bla reporter were introduced into the WT and its lspF mutant, and after infection of BMD macrophages, we monitored the levels of Bla delivered into the host cell cytosol. As a negative control, we also introduced the plasmids into a dotA mutant of strain 130b. As expected (74), WT 130b, but not the dotA mutant, exhibited translocation of AnkX, Lem3, LepB, LidA, and SidM, albeit to differing degrees (Fig. 6B). We did not detect a significant level of SidD translocation, although immunoblot analysis confirmed that the fusion protein was made by the strains (data not shown). Most importantly, the lspF mutant also exhibited effector translocation (Fig. 6B). The translocation of RalF, a T4S effector that acts on a different host GTPase, was also apparent in both the WT and the mutant (Fig. 6B). Together, these data showed that the loss of T2S in the lspF mutant had not broadly eliminated translocation of T4S effectors nor abolished the translocation of effectors that specifically modulate Rab1B. Thus, the effect of T2S on Rab1B association that we observed likely occurred independently of the expression and translocation of the Dot/Icm effectors that are known to influence Rab1B association with the LCV. Hence, we hypothesize that the L. pneumophila T2S system elaborates a secreted protein(s) that promotes the interaction between the LCV and Rab1B in a novel fashion. Compatible with this scenario, transcripts encoding components of the T2S apparatus as well as var-

10 8 6 4 2 0 LepB LidA SidD Lem3 SidM AnkX RalF

T4S Effector FIG 6 Gene expression and translocation of T4S substrates by the WT and T2S mutant strains during infection of macrophages. (A) BMD macrophages were infected at an MOI of 50 with WT strain 130b (black bars) or the lspF mutant NU275 (white bars), and at 4 h postinoculation, the levels of lepB, lidA, sidD, lem-3, sidM, ankX, and dotA transcripts were determined by qRT-PCR analysis. Data are presented as ⫺⌬CT values, such that values larger than zero indicate a higher abundance than that of the gyrB reference gene, while values lower than zero signify a lower abundance. In all cases, the value obtained for the lspF mutant was not significantly different from that for the WT (HolmSidak multiple-comparison test; P ⬎ 0.05). (B) BMD macrophages were infected at an MOI of 50 with WT (black bars), lspF mutant (white bars), or dotA mutant NU428 (gray bars) bacteria carrying a plasmid encoding a Bla fusion with LepB, LidA, SidD, Lem3, SidM, AnkX, or RalF, and the levels of T4S effector translocation were then determined by measuring the cleavage of a Bla substrate, CCF2-AM, that had been added to the macrophage cultures. Translocation is reported as the emission ratio for 460 nm (blue) over 530 nm (green), and the data are presented as means and standard errors for three independent experiments. The dotA mutant showed lower levels of translocation for all effector fusions (Student’s t test; P ⬍ 0.05), with the only exception being SidD, whose level of translocation was not above the background (dashed line) for any of the strains.

ious T2S-dependent proteins were easily detected in infected host cells at the 4-h time point, when the effect of T2S on Rab1B is evident (see Fig. S5 in the supplemental material). As an initial attempt to determine which T2S substrate(s) promotes Rab1B association, we examined eight previously characterized mutants that lack individual T2S substrates. However, mutants of strain 130b defective for either chiA, lapA, lipA, map, plaC, plcA, proA, or srnA displayed the same level of Rab1B association as that of the wild type (see Fig. S6), indicating that the ChiA chitinase, LapA aminopeptidase, LipA lipase, Map phosphatase, PlaC acyltransferase, PlcA phospholipase C, ProA metalloprotease, and SrnA RNase are not required for optimal Rab1B association. Depletion of Rab1B within human macrophages further restricts the growth of an L. pneumophila T2S mutant. Before con-




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sidering further the identification of a T2S-dependent protein(s) that modulates Rab1B, we sought to discern the degree to which the impaired ability to harness the Rab1 protein explains the growth defect of the lspF mutant. The fact that impaired Rab1B association preceded the appearance of fewer mutant bacteria by only a few hours was compatible with the two phenotypes being connected. In order to directly test this hypothesis, we generated a stable U937 cell line expressing shRNA targeted against human Rab1B. We focused on the U937 cell model because it would not only help us to understand the T2S mutant but also address, for the first time, the need for Rab1B recruitment for the growth of WT L. pneumophila in human macrophages. The only past study to assess the impact of depleted Rab1 on L. pneumophila utilized knockdown of Drosophila Kc167 cells, and in that case, loss of the protein did not limit growth (111). A similar conclusion was reached when sidM mutants grew to WT levels within A/J BMD macrophages and amoebae despite their inability to recruit Rab1B (87, 88). Upon differentiation of the treated U937 cells into macrophage-like cells and removal of antibiotic selection, we attained excellent (⬎80%) knockdown of Rab1B at the protein level (Fig. 7A). We then compared Rab1B-depleted U937 cells and a U937 cell line containing a nontargeting shRNA (“scramble”) for the ability to sustain the growth of WT 130b and the lspF mutant. The WT grew comparably in the two cell lines and, if anything, grew slightly better in the Rab1B-depleted cells (Fig. 7B; see Fig. S7 in the supplemental material), demonstrating that full expression of Rab1B is not required for the growth of L. pneumophila in human macrophages. Supporting this conclusion, we observed that our sidM mutant, which lacked the ability to recruit Rab1B (Fig. 5C), grew to WT levels in the depleted U937 cells (Fig. 7C; see Fig. S7). We surmise that in the case of the WT, other host GTPases and their associated vesicles compensate for the lack of Rab1B recruitment, as previously postulated (112). Since the growth of the sidM mutant was also unaffected by depletion of Rab1B in U937 cells (Fig. 7B), we inferred that the sidM mutant can still recruit other vesicles from the human host cell in order to support its growth (112). Most importantly, in contrast to the WT and the sidM mutant, the lspF mutant showed a 21% reduction in growth in Rab1B-depleted U937 cells compared to scramble U937 cells (Fig. 7B; see Fig. S7). The complemented lspF mutant behaved as the WT did in the Rab1B-depleted cells (Fig. 7B), confirming that the mutant’s phenotype was due to the loss of T2S. In sum, our knockdown experiments indicate that the association of Rab1B with the LCV in macrophages is not absolutely necessary for the growth of WT L. pneumophila but can be important for a strain that lacks T2S. The same data also indicate that the lspF mutant lacks a function(s) beyond Rab1 recruitment and that this other defect(s) affects growth. T2S promotes L. pneumophila growth through both Rab1Bdependent and Rab1B-independent processes. In order to validate the data obtained from examining Rab1B-depleted U937 cells, we constructed a sidM lspF double mutant by introducing an lspF mutation into the chromosome of the sidM mutant and then proceeded to test the new mutant for its relative infectivity for macrophages. Because mutation of sidM abolished Rab1B recruitment in both U937 cells and BMD macrophages (Fig. 5D; see Fig. S8 in the supplemental material), we reasoned that the double mutant would recapitulate the larger mutant defect observed in the Rab1B-knockdown cells. As a prelude to the infection assays, we first determined that the double mutant grows normally under

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h post-inoculation FIG 7 Effect of Rab1B depletion on intracellular growth of WT, sidM mutant, and lspF mutant L. pneumophila strains in human macrophages. (A) (Left) U937 cells containing either a nontargeting shRNA (scramble) or an shRNA directed against Rab1B (Rab1B KD) were grown for 0 or 24 h in RPMI-FBS, and cell lysates were then subjected to immunoblot analysis using anti-Rab1B antibodies (top row) or anti-GAPDH antibodies (bottom row). (Right) The levels of Rab1B expression in the scramble versus KD cells at 24 h postincubation were quantitated by performing densitometry on immunoblots obtained from two independent cell cultures. (B) U937 Rab1B KD cells (white bars) or scramble control cells (black bars) were infected with WT strain 130b, the lspF mutant NU275, the sidM mutant NU427, or the complemented lspF mutant NU275(pMF) at an MOI of 0.5. At 0 and 24 h postinoculation, the numbers of CFU within the infected monolayers were determined by plating, and bacterial growth was recorded as the fold change in the number of CFU at 24 h relative to that at 0 h. The growth of each strain in the KD cells is presented as percent growth relative to the growth observed in the scramble control cells, which was set at 100%, and the data presented are means and standard errors for three independent experiments. **, significant reduction in growth of the lspF mutant upon Rab1B silencing (one-sample t test; P ⬍ 0.01). (C) Untransduced U937 cells were infected with the WT or the sidM mutant at an MOI of 0.5, and at the indicated time points, the numbers of CFU within the infected monolayers were determined by plating. Data are presented as mean fold changes in growth and standard deviations for triplicate wells and are representative of three independent experiments.


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FIG 8 Intracellular infection of macrophages by WT and sidM lspF mutant L. pneumophila strains. U937 cells (left) and BMD macrophages (right) were inoculated with WT strain 130b (WT), the sidM mutant NU427 (sidM), the lspF mutant NU275 (lspF), the sidM lspF mutant NU429 (sidM lspF), or the complemented mutant NU429(pMF) (sidM lspF/lspF⫹) at an MOI of 0.5 (left) or 1 (right), and at the indicated times, the numbers of CFU in the infected monolayers were determined by plating. Growth is presented as the fold change (increase) in CFU relative to the level at time zero. Data are means and standard deviations for triplicate wells and are representative of three independent experiments. For both panels, the growth of the sidM lspF double mutant was significantly less than that of all other strains examined (Student’s t test; *, P ⬍ 0.05; **, P ⬍ 0.01). As seen before (Fig. 1), the growth of the lspF mutant was significantly less than that of the WT.

standard extracellular growth conditions (see Fig. S9). As predicted, the sidM lspF double mutant had an intracellular growth defect that was more dramatic than that of the lspF mutant in both the human and murine macrophages, with the difference between the two mutants being as much as 40-fold at 72 h in the BMD macrophages (Fig. 8). Thus, the double mutant was impaired relative to WT strain 130b (and the sidM mutant) at all time points examined, with its defect relative to the WT reaching as much as 400-fold. Providing an intact copy of lspF on a plasmid restored the growth of the double mutant to WT levels (Fig. 8). These results confirmed that T2S promotes intracellular growth and that it does so partly, if not mainly, through a Rab1B-independent process. DISCUSSION

The results presented here significantly advance our understanding of the role of T2S in intracellular infection of macrophages by L. pneumophila. Although the T2S system was dispensable for early infection events, including bacterial entry into the host cell, evasion of phagosome-lysosome fusion, and resistance to initial intracellular killing, it was necessary for optimal initiation of intravacuolar replication between 4 and 8 h postentry as well as for the capacity to grow to large numbers within LCVs between 8 and 12 h and beyond. Based upon our further analysis of multiple lsp mutants and a complemented derivative in human and murine macrophages, we determined that the role of T2S in early replication involves at least the promotion of Rab1B association with the LCV. Because the loss of T2S in the lspF mutant did not simply eliminate the transcription or translocation of T4S effectors, including ones known to modulate Rab1B recruitment, we posit that T2S elaborates a secreted protein(s) that promotes the interaction between the LCV and host Rab1B. In support of our data, a past study detected lspE and lspL mutants, which, like our lspF mutant, lack the type II secretion apparatus, in a screen of transposon-mutagenized L. pneumophila mutants for strains with impaired Rab1 association (88). Because it was focused on T4S, that study did not attempt complementation of the lsp mutants or investigate, to any degree, the lsp mutants’ defects in intracellular infection. Thus, our study represents the first documentation of a role for T2S in the association of the LCV with Rab1B. Although T2S was recently shown to be important for the intracellular survival or growth of Burkholderia cenocepacia, Chlamydia trachoma-

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tis, and Yersinia enterocolitica (113–116), the current study is also the first demonstration of a link between T2S and Rab1 proteins associated with vesicular trafficking. The recruitment of the Rab1B GTPase to the LCV and the role that T4S effectors play in that process have been the focus of an extraordinary amount of research, revealing a remarkable array of enzymatic reactions. Rab1B and the other Rab proteins are master regulators of intracellular trafficking, controlling the specificity, direction, timing, and coordination of vesicle transport (117). Rab1B is first activated and recruited to the LCV via T4S effectors, and then activated Rab1B facilitates the tethering of ER-derived vesicles at the LCV (61). In the first step of Rab1B activation, the T4S effector SidM (DrrA) promotes recruitment by acting as a guanine dissociation inhibitor (GDI) displacement factor, guanine nucleotide exchange factor (GEF), and AMPylation factor (87, 88, 101). The effector LidA, with its high affinity for active Rab1B, further enhances the recruitment of Rab1 to the LCV, while AnkX, by virtue of its phosphocholination activity, blocks possible inactivation of Rab1B by other factors (87, 102). Ultimately, SidD antagonizes SidM function with a deAMPylation activity, and Lem3 reverses the effect of AnkX, thus allowing Rab1B to be inactivated by LepB, a GTPase activation protein (GAP) (89, 91, 92, 103). With the hydrolysis of GTP to GDP, the Rab1 protein is released from the LCV and extracted back into the cytoplasmic inactive pool. Looking beyond the well-characterized SidM, LidA, AnkX, SidD, Lem3, and LepB proteins, recent reports implicate additional T4S effectors as interactors/modifiers of Rab1B (68, 118, 119). The fact that so many T4S effectors are dedicated to Rab1B recruitment indicates that the association of this Rab protein and ER-derived vesicles with the LCV is important for L. pneumophila growth. Without diminishing the role of T4S in Rab1B recruitment, our study signals that the recruitment and retention of this host GTPase at the LCV are not strictly the purview of T4S. Because the lspF mutant’s defect in Rab1B association was manifest at 4 h postentry, but not at 1 and 2 h postentry, we suspect that T2S does not act at an early stage of Rab1B recruitment but is needed for ongoing recruitment or retention. Indeed, T2S may promote L. pneumophila’s association with Rab1B by enhancing later stages of Rab1B recruitment and tethering to the LCV or by inhibiting the release of recruited Rab1B. In considering further how T2S facilitates the associa-




tion of Rab1B with the LCV, several intriguing scenarios can be envisioned. In one general scenario, a protein(s) secreted via T2S is translocated out of the LCV and gains access to the macrophage cytoplasm, where it acts directly on Rab1B so as to promote or prolong its association with the LCV. Alternatively, a translocated type II secreted protein indirectly influences Rab1B, either by modulating another factor that acts on Rab1B, such as a host protein or a T4S effector, or by influencing the nature of the cytoplasmic face of the LCV which serves to tether Rab1B. Compatible with this scenario is the fact that the L. pneumophila T2S system secretes a wide range of enzyme activities as well as many novel proteins of undefined function (23, 26, 30). In a second type of situation, the protein(s) secreted by the T2S system remains localized within the lumen of the LCV but proceeds to modify (“from within”) the nature of the LCV and its membrane such that it is more receptive to association with Rab1B. Clearly, future investigations need to determine the subcellular location of type II secreted proteins in infected macrophages as well as to characterize the secreted protein(s) that influences Rab1B. The simple fact that the lspF mutant has a growth defect in both human and murine macrophages yet a sidM mutant that completely lacks Rab1B recruitment grows similarly to the WT indicates that T2S must promote growth beyond facilitating Rab1B association with the LCV. By demonstrating that the magnitude of the lspF mutant defect is increased upon depletion of Rab1B in human macrophages, we confirmed that T2S also promotes intracellular growth by means that are independent of Rab1B. Once again, there are many potential explanations for this additional role of T2S. The magnitude of the defect linked to the loss of T2S was greatly exacerbated by the additional loss of SidM (i.e., the sidM lspF mutant had a larger defect than the lspF mutant), suggesting that T2S facilitates a process whose function is complementary to that of Rab1Bmediated vesicle recruitment. Indeed, simultaneously targeting components of multiple early secretory pathways via small interfering RNA (siRNA)-mediated depletion can result in dramatic inhibition of L. pneumophila growth (111). In a similar vein, a bacterial mutant that lacks the ability to harness a single vesicular pathway and thus still grows to WT levels in normal host cells (e.g., a lidA mutant lacking Rab1 recruitment) exhibits a growth defect in host cells that are defective (via siRNA depletion) for another aspect of vesicle trafficking (120). Thus, we posit that T2S promotes the recruitment of various types of host vesicles to the LCV. On the other hand, it is also possible that T2S-dependent proteins which come to reside in the host cell cytoplasm promote intracellular growth by means that are unrelated to vesicular trafficking, such as promoting the association of the LCV with mitochondria or the regulation of protein turnover and the ubiquitin pathway, among other things (15, 16). Finally, it is also possible that the Rab1B-independent effect of T2S is due to events that occur strictly within the LCV, such as the acquisition of nutrients and the stabilization of the LCV membrane. None of the L. pneumophila mutants lacking individual type II secreted proteins that have been tested in macrophages have exhibited a growth defect (23), suggesting that an as yet undefined secreted protein(s) is responsible for the role of T2S in intramacrophage growth. Compatible with this hypothesis, our proteomic and in silico analyses indicate


that the L. pneumophila T2S system may secrete as many as 60 proteins (25). In summary, we have shown that the T2S system of L. pneumophila promotes intravacuolar growth and enhances the association of Rab1B with the LCV, a process that we have shown can promote growth in human and murine macrophages. Our Rab1B depletion experiments and analysis of various bacterial mutants further indicate that T2S also promotes intramacrophage replication through Rab1B-independent mechanisms that likely involve a novel secreted protein(s). Finally, we strongly suspect that the L. pneumophila T2S system evolved in part to facilitate similar events in infected amoebae, as the intracellular life cycles of L. pneumophila within macrophages and environmental protozoa are remarkably similar (121). Indeed, in both hosts, the LCV associates with secretory vesicles, rough ER, mitochondria, and the ubiquitination machinery, among other things, and proteomic analyses of LCVs purified from amoebae revealed the presence of small GTPases, including Rab1 homologs (122, 123). ACKNOWLEDGMENTS We thank past and present members of the Cianciotto lab for their helpful advice. We acknowledge Ombeline Rossier for construction of pMGFP and Meghan Pearce for introducing the plasmid into L. pneumophila strains. We thank Howard Shuman for providing pXlidA, pXlepB, pXralF, pXsidM, and pXankX, Michele Swanson for pMMB-GRN, and Richard Kohler for anti-L. pneumophila MAB2. We also thank Laimonis Laimins for providing lentiviral packaging plasmids and the nontargeting shRNA control plasmid and Eva Gottwein for providing HEK293T cells. R.C.W. was supported in part by NIH training grant T32 AI0007476. This study was funded by NIH grant R01 AI043987 awarded to N.P.C.

FUNDING INFORMATION This work, including the efforts of Nicholas P. Cianciotto, was funded by NIH-NIAID (R01 AI043987). This work, including the efforts of Richard C. White, was funded by NIH-NIAID (T32 AI0007476).

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