Distinct roles of ppGpp and DksA in Legionella pneumophila differentiation

SUMMARY

To transit between hosts, intracellular Legionella pneumophila transform into a motile, infectious, transmissive state. Here we exploit the pathogen’s life cycle to examine how guanosine tetraphosphate (ppGpp) and DksA cooperate to govern bacterial differentiation. Transcriptional profiling revealed that during transmission alarmone accumulation increases the mRNA for flagellar and Type IV-secretion components, secreted host effectors, and regulators, and decreases transcripts for translation, membrane modification and ATP synthesis machinery. DksA is critical for differentiation, since mutants are defective for stationary phase survival, flagellar gene activation, lysosome avoidance, and macrophage cytotoxicity. The roles of ppGpp and DksA depend on the context. For macrophage transmission, ppGpp is essential, whereas DksA is dispensable, indicating ppGpp can act autonomously. In broth, DksA promotes differentiation when ppGpp levels increase, or during fatty acid stress, as judged by flaA expression and evasion of degradation by macrophages. For flagella morphogenesis, DksA is required for basal fliA (σ²⁸) promoter activity. When alarmone levels increase, DksA cooperates with ppGpp to generate a pulse of Class II rod RNA or to amplify the Class III sigma factor and Class IV flagellin RNAs. Thus, DksA responds to the level of ppGpp and other stress signals to coordinate L. pneumophila differentiation.

INTRODUCTION

During infection, pathogens respond to local cues by altering their metabolism and virulence factor production. To coordinate physiological adaptation with virulence mechanisms, many pathogens employ two factors known to be critical for stationary phase resilience of most bacterial species, guanosine tetraphosphate (ppGpp) and DksA. Components of the stringent response regulate processes as diverse as Mycobacterium tuberculosis persistence (Dahl et al, 2003; Stallings et al, 2009), Salmonella enterica invasion (Pizarro-Cerda & Tedin, 2004; Thompson et al, 2006), Shigella flexneri intercellular spread (Sharma & Payne, 2006), and Legionella pneumophila transmission (Hammer & Swanson, 1999; Dalebroux et al, 2009).

L. pneumophila resides in aquatic reservoirs within biofilms or protozoa. When humans inhale contaminated aerosols, L. pneumophila infects alveolar macrophages. In host cells, the bacteria differentiate between two forms, replicative and transmissive (Molofsky & Swanson, 2004; Bruggemann et al, 2006) . Upon phagocytosis, transmissive bacteria utilize the Dot/Icm Type IV secretion system to avoid lysosomes and traffic to a vacuole derived from the endoplasmic reticulum (Isberg et al, 2009). In this compartment, bacteria that sense favorable conditions repress transmissive functions and activate genes needed for protein synthesis and replication (Sauer et al, 2005; Bruggemann et al, 2006). Eventually conditions deteriorate, cueing bacteria to synthesize ppGpp. The alarmone triggers differentiation to the motile, transmissive form, which resist degradation, lyse the exhausted host cell and are equipped to infect naive host cells (Molofsky & Swanson, 2004; Dalebroux et al, 2009). Under certain conditions, transmissive L. pneumophila develop into ‘mature intracellular forms’, which are fit to persist in the environment (Faulkner et al, 2008).

In most gamma-proteobacteria, including L. pneumophila, ppGpp levels are controlled by the synthetase RelA and the bifunctional synthetase/hydrolase SpoT. RelA monitors amino acid availability through its association with the ribosome, whereas SpoT responds to a variety of stimuli including fatty acid starvation, which requires direct interaction with acyl-carrier protein (Potrykus & Cashel, 2008; Battesti & Bouveret, 2009). Synthesis of ppGpp by exponential (E), replicative phase broth-grown L. pneumophila triggers differentiation to the post-exponential (PE), transmissive form (Hammer & Swanson, 1999; Dalebroux et al, 2009). For transmission between macrophages, ppGpp synthesized from SpoT is sufficient (Dalebroux et al, 2009). When conditions are favorable in host cells or in media, transmissive bacteria require SpoT to hydrolyze ppGpp and initiate replication. Thus, L. pneumophila modulates ppGpp levels to coordinate timely differentiation.

Many of the physiological effects of ppGpp are mediated through interactions with RNA polymerase (RNAP) in cooperation with the RNAP secondary channel interacting protein DksA (Haugen et al, 2008; Potrykus & Cashel, 2008). Whether ppGpp and DksA co-exert positive or negative regulation depends upon intrinsic properties of the promoters. While repressing ribosomal RNA operons (rRNA), ppGpp and DksA activate amino acid biosynthetic operons and alternative metabolic pathways. Direct co-positive regulation has also been observed during in vitro studies of promoters of critical virulence regulators (Nakanishi et al, 2006; Sharma & Payne, 2006; Aberg et al, 2008). Recently, DksA and ppGpp were shown to directly inhibit transcription of the E. coli σ⁷⁰-dependent promoters of critical flagellar gene regulators, flhDC and fliA (σ²⁸) to repress flagellar synthesis during starvation (Lemke et al, 2009).

Indirect transcriptional control reflects the impact of ppGpp and DksA on RNAP availability. During E. coli growth, nearly half the cellular RNAP is localized to rRNA operons by the σ⁷⁰ vegetative sigma factor (Bremer, 1996). Upon nutrient limitation, ppGpp and DksA deactivate transcription from these loci, increasing the amount of core RNAP available to alternative sigma factors. These specialized subunits then direct polymerase to promoters of genes involved in particular stress responses (Bernardo et al, 2006; Szalewska-Palasz et al, 2007; Costanzo et al, 2008; Gummesson et al, 2009). Therefore, ppGpp and DksA indirectly control transcription by alternative sigma factors.

Recent evidence from E. coli suggests that DksA is more than a cofactor for ppGpp-dependent transcriptional control. Overproduction of DksA in ppGpp⁰ bacteria can compensate for lack of alarmone, indicating that DksA can act independently of ppGpp (Potrykus & Cashel, 2008). Additionally, phenotypic and in vitro assays show that ppGpp and DksA oppositely regulate some processes and promoters (Magnusson et al, 2007, Lyzen et al, 2009). Therefore, we exploited L. pneumophila differentiation and its flagellar cascade to investigate the functional relationship between ppGpp and DksA.

RESULTS

ppGpp induces rapid accumulation of virulence transcripts

Although ppGpp is known to induce L. pneumophila transmission traits, genes regulated by the alarmone have not been identified. To begin to define the ppGpp regulon, we developed a genetic system to synchronize ppGpp accumulation by L. pneumophila. When treated with IPTG, a relA spoT double mutant strain (ppGpp⁰) that carries an inducible allele of relA is locked in the transmissive state and exhibits heightened virulence (Dalebroux et al, 2009). Although replication is stunted, viability is not compromised. By 60 min after IPTG addition, a pool of ppGpp was evident in ppGpp⁰ prelA^L.p. cell extracts, but not in ppGpp⁰ pempty controls; by 90 min, this pool had increased (Fig. 1).

Figure 1. Kinetics of ppGpp accumulation after prelA^L.p induction.

To evaluate the timing of ppGpp accumulation after relA^L.p. induction, early-E phase AYET broth cultures of ppGpp⁰ pempty and ppGpp⁰ prelA^L.p. labeled with ³²P phosphoric acid were treated with 500 µM IPTG for the periods indicated, then nucleotides in cell extracts were separated by PEI-TLC. An arrow indicates the major ppGpp species, guanosine tetraphosphate; an asterisk denotes the position of guanosine pentaphosphate and an unidentified phosphorylated species. The autoradiogram shown represents one of two independent experiments.

To identify ppGpp-sensitive transcripts, cDNA prepared from ppGpp⁰ prelA^L.p. and ppGpp⁰ pempty bacteria harvested at 45 and 90 min post-induction was hybridized to a comprehensive L. pneumophila microarray (Bruggemann et al, 2006), and relative transcript levels were calculated. This experimental design will identify loci whose expression is affected by cell growth, together with genes for which ppGpp accumulation activates or represses transcription initiation or transcript stability. For brevity, we will use “up-” and “down-regulation” to describe the relative differences between the two cultures, but kinetic studies of the corresponding promoters and transcripts will be required to identify the mechanism(s) that contribute.

Microarray results were validated by qRT-PCR analysis of six loci, and a strong correlation was apparent (R² = 0.98; Fig. S1). Although statistically significant differences were observed at 45 min, most were less than two-fold, our criterion for ppGpp-regulated transcripts (http://www.ebi.ac.uk/microarray-as/ae/). Therefore, the 90 min data set was analyzed further. Many transcripts identified as ppGpp-sensitive at 90 min (Tables 1 and S1) were also differentially expressed at 45 min. By 90 min these differences had increased, suggesting that as time with ppGpp increased, existing differences were amplified.

Table 1.

Select list of transcripts up-regulated 90 min after relA^L.p. induction.

Gene Name / Function	Gene.ID	Annotation	ppGpp0 prelAL.p./pempty
Flagellar Genes
fleQ	lpg0853	Master regulator of flagellar gene transcription (Class I)	2.01
fliH	lpg1758	Export and assembly (Class IIa)	2.41
fliG	lpg1759	Motor switch protein (Class IIa)	3.88
fleR	lpg1762	Two-component response regulator (Class IIa)	3.54
fleS	lpg1763	Two-component sensor histidine kinase (Class IIa)	2.58
fliN	lpg1791	Motor switch protein (Class IIa)	5.25
flgB	lpg1216	Proximal rod protein (Class IIb)	4.32
flgC	lpg1217	Proximal rod protein (Class IIb)	4.03
flgG	lpg1221	Distal rod protein (Class IIb)	4.81
flgH	lpg1222	L-ring protein precursor (Class IIb)	4.42
flgJ	lpg1224	Peptidoglycan-hydrolyzing protein (Class IIb)	2.87
fleN	lpg1783	Regulator of flagellar synthesis (Class IIb)	3.01
flhF	lpg1784	Biosynthesis regulator GTP-binding protein (Class IIb)	3.87
Dot/Icm Structural Genes
icmR	lpg0443	Chaperone for IcmQ; a pore forming molecule	2.46
icmL/dotI	lpg0449	Inner membrane protein	2.23
icmK/dotH	lpg0450	Might be an outer membrane channel	2.18
icmE/dotG	lpg0451	Major component of a channel	1.96
icmG/dotF	lpg0452	Interacts with substrates; predicted channel component	2.14
icmC/dotE	lpg0453	Predicted inner-membrane protein, Similar to DotV	2.31
icmD/dotP	lpg0454	Predicted inner-membrane protein	3.43
icmJ/dotN	lpg0455	Probable ATPase component	2.10
Dot/Icm Secreted Effectors
legC3	lpg1701	Coiled-coil domain, disrupts vacuolar trafficking, IcmSW-dependent	2.11
ylfA	lpg2298	Coiled-coil domain, disrupts vacuolar trafficking,	3.33
sdeD	lpg2509	Substrate of the Dot/Icm system	1.98
lepA	lpg2793	SNAREs and coiled-coil domain; bacterial egress	3.99
Regulatory Factors
lpg0433	lpg0433	Putative transcriptional regulator	1.99
letE	lpg0537	Transmission trait enhancer protein	3.60
rpoS	lpg1284	RNA polymerase sigma factor (sigma-38)	3.11
relA	lpg1457	ppGpp synthetase	16.16
lpg2732	lpg2732	Putative response regulator	2.28
rsmZ		non-coding, CsrA binding, regulatory RNA	4.18
Known Virulence Factors
mip	lpg0789	Macrophage infectivity potentiator	2.06
katB	lpg2389	Catalase-peroxidase	3.03
enhC	lpg2639	Enhanced entry protein	2.73
Fatty Acid / Carbon Metabolism
plsC	lpg0551	similar to 1-acyl-sn-glycerol-3-phosphate acyltransferase	2.51
phaB1	lpg0560	Similar to acetoacetyl-CoA reductase	2.09
yfcX	lpg1596	similar to alpha subunit of fatty-acid oxidation complex	3.25
phbC3	lpg2260	Similar to poly(3-hydroxyalkanoate) synthetase	2.25
pta	lpg2261	Similar to phosphotransacetylase	2.12
pksJ	lpg2186	Similar to polyketide synthase of type I	2.17
lpg2837	lpg2837	phospholipase/lecithinase/hemolysin, lysophospholipase A	6.13

The data were collected from two independent biological replicates. See Fig. S1 for qRT-PCR validation. p < 0.01 for all targets shown.

Consistent with the E. coli stringent response to amino acid starvation (Durfee et al, 2008; Traxler et al, 2008), by 90 min of exposure to ppGpp, the transcripts for several ribosomal proteins were less abundant (Table S1). RNAs encoding components of the ATP synthase complex also decreased. Strikingly, RNAs for enzymes involved in lipopolysaccharide (LPS) modification and phospholipid biosynthesis were less abundant, including plsB, a known mediator of ppGpp-dependent regulation of fatty acid and phospholipid biosynthesis in E. coli (Heath et al, 1994). Thus, L. pneumophila employs ppGpp for stringent control over its membrane machinery and energy intensive processes like protein synthesis.

RNAs for several virulence factor loci accumulated within 90 min of relA induction. Consistent with their motility and toxicity toward macrophages (Dalebroux et al, 2009), ppGpp⁰ prelA^L.p. bacteria contained significantly more transcripts for structural and regulatory elements of the flagellar apparatus (Table 1). Additionally, the amount of RNAs encoding membrane, channel and other components of the Dot/Icm Type IV secretion system increased. Consistent with transcriptional activation and secretion of Type IV effectors in PE phase (Nagai et al, 2002; Tiaden et al, 2007; Rasis & Segal, 2009), we observed an increase in the legC3, ylfA, sdeD and lepA Dot/Icm effector RNAs. Transcripts encoding virulence regulators also accumulated, including the alternative sigma factor rpoS (Bachman & Swanson, 2001; Hovel-Miner et al, 2009), the transmission trait enhancer letE (Bachman & Swanson, 2004), and the regulatory RNA rsmZ (Rasis & Segal, 2009; Sahr et al, 2009). In addition, the macrophage infectivity potentiator mip (Cianciotto & Fields, 1992), the catalase peroxidase katB (Bandyopadhyay et al, 2003), and the enhancer of macrophage uptake enhC (Liu et al, 2008) were up-regulated. Together these data demonstrate that L. pneumophila employs ppGpp to down-regulate protein synthesis machinery and up-regulate factors dedicated to survival and transmission in host cells.

L. pneumophila encodes a DksA homologue required for stationary phase morphogenesis and survival

L. pneumophila encodes a 158 amino acid DksA-like protein (lpg2338) that is 72% identical to that of E. coli K-12 (Fig. 2A). Conserved in DksA_Lpn are two aspartate residues known to comprise the acidic tip of DksA_Eco and other related RNAP secondary interacting proteins. The amino acid sequences in the two α-helical regions immediately adjacent to this pair of aspartates are also highly conserved, suggesting that DksA_Lpn adopts a similar coiled-coil fold (Perederina et al, 2004). Also present in DksA_Lpn are four cysteine residues known in DksA_Eco to comprise a zinc-finger motif. Obvious sequence dissimilarities lie at the extreme N-terminus, where the E. coli protein adopts a globular fold (Perederina et al, 2004). Therefore, L. pneumophila encodes a DksA protein with several but not all features of its E. coli counterpart.

Figure 2. To survive stationary phase stress L. pneumophila requires DksA and ppGpp.

A. L. pneumophila encodes a 158 amino acid protein that is similar to the 151 amino acid DksA protein of E. coli K-12. Residues in bold, black lettering are identical; those in bold and grey are conserved. The underlined aspartate residues comprise the acidic tip, conserved in DksA and other RNAP secondary channel interacting proteins, while the underlined cysteine residues comprise a zinc finger motif. Light grey shading denotes two α-helical domains sandwiching the acidic tip that adopt a coiled-coil structure in the E. coli protein (Perederina et al, 2004). Dark grey shading indicates a C-terminal α-helix. B. Early-E phase AYET broth cultures of WT (triangles) and ppGpp⁰ (circles) carrying empty vector, and dksA mutants transformed with empty vector (squares) or pdksA (diamonds), were diluted to OD₆₀₀ of 0.15, and bacteria were treated with 25 µM IPTG. At the times indicated, culture density and viability were quantified by reading OD₆₀₀ (dashed lines) and enumerating colony forming units (CFU ± SE) ml⁻¹ from duplicate samples on CYET (solid lines). The data represent one of three independent experiments.

To assess the contribution of DksA to differentiation, a dksA deletion insertion mutant was constructed and analyzed. Since L. pneumophila requires the ppGpp alarmone to survive in stationary phase (Dalebroux et al, 2009), we first tested if DksA also contributes to persistence in broth. From 3 – 24 h, both culture density and viability were similar for WT, ppGpp⁰, and dksA mutant bacteria (Fig. 2B). However, beginning in early stationary phase (~ 22 h), ppGpp⁰ and dksA mutant bacteria began to filament; by 48 h, or ~ 28 h into stationary phase, the mutants exhibited several defects. WT bacteria had become coccoid and motile (data not shown), two characteristics of transmissive cells (Molofsky & Swanson, 2004), whereas the ppGpp⁰ and dksA mutants remained amotile and had become more filamentous. Filamentation of dksA mutant bacteria was modest relative to ppGpp⁰ bacteria, which typically elongated to ~ 50 × the length of PE phase WT L. pneumophila (data not shown). After extended culture in stationary phase, ppGpp⁰ and dksA mutant bacteria lost viability: By 72 h, their CFU values had declined ~2 logs below the WT yield (Fig. 2B). The dksA mutant survival, morphogenesis, and motility defects were each restored when expression of plasmid-borne dksA was induced continuously from early-E phase to the late stationary phase with 25 µM isopropyl-beta-D-thiogalactopyranoside (IPTG; Fig. 2B; data not shown). Therefore, DksA and ppGpp each contribute to survival, morphology, and motility of stationary phase L. pneumophila.

To test the simple model that L. pneumophila require DksA to generate ppGpp, we performed thin layer chromatography. Similar to PE phase WT bacteria, dksA mutant L. pneumophila accumulated ppGpp, and the size of the pool increased with the length of the PE phase (data not shown). Therefore, the stationary phase defects of L. pneumophila dksA mutants could not be attributed to a deficiency in alarmone accumulation.

DksA contributes to PE phase virulence

Concomitant with its morphogenesis, stationary phase L. pneumophila activate several virulence phenotypes, including infectivity, lysosome evasion, cytotoxicity, and sensitivity to sodium, a phenotype associated with Type IV secretion (Vogel et al, 1996; Byrne & Swanson, 1998). Since ppGpp activated several loci implicated in particular transmisson traits, we investigated whether DksA also contributes.

L. pneumophila required both DksA and ppGpp for PE phase activation of a panel of virulence phenotypes. Like E phase WT bacteria, PE phase dksA and ppGpp⁰ mutants were poorly infectious (Fig. 3A), frequently degraded within 2 h of infection (Fig. 3B), non-cytotoxic to macrophages (Fig. 3C), and sodium resistant (Fig. 3D). Induction of plasmid-borne dksA restored macrophage infectivity and cytotoxicity of the mutants (Fig. 3A and C). Thus, in broth, L. pneumophila employs both DksA and ppGpp to activate transmission traits.

Figure 3. To activate PE phase transmission traits L. pneumophila requires DksA and ppGpp.

A. To analyze whether DksA contributes to infectivity of PE phase L. pneumophila, macrophages were infected at an MOI of ~ 1 for 2 h with WT pempty, ppGpp⁰ pempty dksA pempty, or dksA pdksA cultured to the growth phase shown. Bacteria carrying plasmids were cultured from early-E phase to PE phase with 25 µM IPTG. Graphed are the mean percent of cell-associated CFU ± SE from duplicate wells in one of two independent experiments. B. The ability of bacteria to resist degradation in macrophage lysosomes was quantified using fluorescence microscopy by scoring the percent of intracellular bacteria that were intact at 2 h post-infection. Shown are the mean percentages from duplicate coverslips ± SE from three independent experiments. C. To determine the contribution of DksA to L. pneumophila cytotoxicity to macrophages, mid-E phase WT pempty (triangles, dashed lines), PE phase WT pempty (triangles, solid lines), PE phase ppGpp⁰ pempty (circles, solid lines), PE phase dksA pempty (squares, dashed lines), or PE phase dksA pdksA (squares, solid lines) bacteria, cultured with IPTG as described in A, were added to triplicate wells of macrophages at the MOI shown. The values plotted represent the mean ± SE for triplicate samples determined in one of three similar experiments. D. To measure sodium resistance, E or PE phase bacteria of the strains depicted were plated onto medium with or without 100 mM NaCl. Shown are the mean percentages from duplicate samples ± SE from three independent experiments.

DksA and ppGpp promote growth in amoebae

In the environment, L. pneumophila parasitizes numerous species of protozoa, including Acanthamoeba castellanii. Optimal growth in A. castellanii required both DksA and ppGpp. During the 4-day incubation, WT bacteria replicated robustly, increasing their CFU ~ 10⁴ fold by 48 h and exhibiting a growth rate constant (µ) of 0.19 h⁻¹ between 1 and 48 h (Fig. 4). The yield of dksA and ppGpp⁰ mutants was less than that of WT at each time analyzed (Fig. 4). Bacteria that lacked dksA were better equipped for intracellular replication than those that could not generate ppGpp (µ = 0.10 h⁻¹ and 0.06 h⁻¹, respectively, between 1 and 48 h). By 96 h, the yield of dksA CFU approached the WT level, while ppGpp⁰ mutant values were 1–2 logs lower (Fig. 4). Thus, for optimal growth in A. castellanii, L. pneumophila requires both DksA and ppGpp, but the alarmone is more critical.

Figure 4. For optimal growth in A. castellanii L. pneumophila requires both DksA and ppGpp.

Thy⁺ derivatives of WT (triangles), dksA (squares) and ppGpp⁰ (circles) bacteria were cultured to PE phase, then added to amoebae in infection buffer at an MOI of ~ 0.05. At the times shown aliquots were suspended and lysed, and lysates plated for CFU enumeration. To obtain relative CFU, viable counts for 24, 48, 72, and 96 were divided by the 1 h CFU value for each strain. Depicted are the relative CFU values from duplicate infections ± SE. The data represent one of two independent experiments.

L. pneumophila requires ppGpp but not DksA for transmission between macrophages

As in broth, L. pneumophila alternates between replicative and transmissive forms in amoebae and macrophages (Byrne & Swanson, 1998; Hammer & Swanson, 1999; Sturgill-Koszycki & Swanson, 2000; Alli et al, 2000; Molofsky & Swanson, 2003; Sauer et al, 2005; Bruggemann et al, 2006). Whereas ppGpp is dispensable for intracellular replication, the alarmone is essential for intracellular progeny to survive transmission from one macrophage to another (Dalebroux et al, 2009). Therefore, we analyzed the contribution of DksA to transmission in murine macrophages.

As reported in Figure 3A, DksA and ppGpp were essential for efficient infection by PE phase L. pneumophila. Both mutants behaved like E phase WT bacteria, yielding less CFU at 2 h than PE phase WT (Fig. 5A). As expected, neither DksA nor ppGpp were required during the primary replication period: From 2 – 24 h, their growth curves were nearly indistinguishable, each exhibiting an ~ 1 log increase in CFU. Between 18 and 24 h, when replicating WT bacteria differentiate to the transmissive form (Byrne & Swanson, 1998; Hammer & Swanson, 1999; Molofsky & Swanson, 2003), DksA was dispensable, but ppGpp was not. The dksA mutant CFU continued to increase until 48 h, when their numbers were similar to WT. In contrast, the yield of ppGpp⁰ mutants failed to increase throughout the remainder of the infection (Fig. 5A).

Figure 5. L. pneumophila requires ppGpp for macrophage transmission; DksA is less critical.

A. Macrophages were infected at an MOI of ~ 1 with E phase WT (triangles, dashed lines), PE phase WT (triangles, solid line), ppGpp⁰ (circles), or dksA (squares) bacteria. The number of viable bacteria was determined at the time points shown. Depicted are the mean CFU ± SE from duplicate samples in one of four independent experiments. * Indicates that the difference in the mean CFU values calculated from four independent experiments for WT and ppGpp⁰ mutant bacteria at 72 h was statistically significant by a paired, two-tailed Student t test (p = 0.047). ** Indicates that the difference between ppGpp⁰ and dksA mutant bacteria at 72 h was also statistically significant (p = 0.047). B. Macrophages were infected at an MOI of ~ 1 with PE phase WT or an MOI ~ 3 of E phase WT, PE phase ppGpp⁰ and PE phase dksA mutant bacteria, then at 18 h coverslips were fixed. A total of 100 infected macrophages from duplicate coverslips were scored as follows: macrophages with a single intact or a single degraded bacterium, and macrophages with multiple intact or multiple degraded bacteria. Shown is the mean percent ± SE of macrophages with degraded bacteria at 18 h from three independent experiments.

At the end of the primary replication period in macrophages, L. pneumophila that lack ppGpp are vulnerable to lysosomal degradation (Dalebroux et al, 2009). Consistent with their lower yield between 48 and 72 h, ppGpp⁰ mutants were more frequently degraded at 18 h than were dksA mutants, as judged by immunofluorescence microscopy (Fig. 5B). To assess the ability of transmitted L. pneumophila to resist lysosomal degradation, macrophages that contained only a single intact or degraded bacterium at 24 h were scored in two independent experiments. By this criterion, ppGpp⁰ mutant L. pneumophila were more susceptible to lysosomal degradation during host-to-host transmission than either dksA mutant or WT L. pneumophila, since 73% ± 8% of infected macrophages harbored a single degraded ppGpp⁰ bacterium, whereas 53% ± 5%, 51% ± 5%, and 49% ± 1% of infected macrophages contained a single degraded WT E, WT PE, or dksA bacterium, respectively. Together, the CFU and morphological data indicate that, whereas DksA is dispensable, L. pneumophila requires the alarmone ppGpp for efficient transmission to a new macrophage.

Constitutive ppGpp synthesis by L. pneumophila can halt replication and stimulate pigmentation independently of DksA

The distinct phenotypes of dksA and ppGpp⁰ mutant bacteria during growth in A. castellanii (Fig. 4) and transmission in macrophages (Fig. 5) motivated us to investigate whether ppGpp or DksA can activate L. pneumophila transmission traits independently of each other. For this purpose, dksA mutant L. pneumophila were transformed with prelA^E.c., which encodes an inducible E. coli K-12 RelA with a C-terminal truncation that prevents its interaction with the ribosome, resulting in constitutive synthetase activity (Schreiber et al, 1991; Hammer & Swanson, 1999). In parallel, ppGpp⁰ bacteria were transformed with pdksA.

Induction of dksA did not impact replication of E phase L. pneumophila, since the culture densities of WT, ppGpp⁰ pempty, and ppGpp⁰ pdksA bacteria were similar after IPTG addition (Fig. 6A). In contrast, replication of dksA prelA^E.c. cells halted shortly after IPTG addition, and their OD₆₀₀ values subsequently remained lower than WT and dksA pempty control bacteria (Fig. 6A). Therefore, when expression of relA^E.c. is highly induced, ppGpp can arrest the L. pneumophila cell cycle independently of DksA.

Figure 6. Constitutive ppGpp synthesis is sufficient to bypass the dksA requirement for cell cycle arrest and pigment production.

A. To test the effect of either dksA induction or constitutive ppGpp synthesis on the growth of L. pneumophila mutant bacteria, WT pempty (triangles), ppGpp⁰ transformed with either pempty (circles, solid lines), or pdksA (circles, dashed lines), and dksA mutants carrying either pempty (squares, solid lines), or prelA^E.c. (squares, dashed lines), a truncated and constitutively active form of E. coli RelA, were induced with IPTG in early-E phase AYET broth cultures, then culture density was monitored by reading OD₆₀₀ over the time period shown. B. At 48 h post-IPTG (indicated in A by **), the OD₅₅₀ of the supernatants was read to quantify extracellular pigment production. To account for differences in cell density, pigmentation values were normalized to culture density by dividing OD₅₅₀ by OD₆₀₀. The graph depicts the mean OD₅₅₀/OD₆₀₀ values ± SE from three independent experiments.

In late stationary phase, L. pneumophila produce a secreted pyomelanin pigment by a pathway induced by RelA (Zusman et al, 2002; Chatfield & Cianciotto, 2007). Pigmentation required both DksA and ppGpp, as pyomelanin was not detected after a ~ 44 h treatment of dksA and ppGpp⁰ mutant bacteria that carried only the vector (Fig. 6B). Nevertheless, DksA was not essential to this response, since ectopic expression of relA^E.c. by dksA mutants was sufficient to generate as much pyomelanin as WT cultures. Nor could DksA function independently of ppGpp to activate this pathway, since ppGpp⁰ pdksA cultures failed to pigment. Therefore, ppGpp can bypass the dksA requirement for pigment production, but the opposite is not true.

DksA controls the propionic acid response cooperatively and independently of ppGpp

When treated with 10 mM propionic acid, E phase L. pneumophila rapidly transition to the transmissive state, a response that is specific to carboxylic acids rather than pH (Edwards et al, 2009). To assess the contribution of DksA to L. pneumophila’s stringent response to fatty acid addition, we used a transcriptional reporter of transmission. The plasmid pflaAgfp encodes the flaA promoter fused to the gene encoding green fluorescent protein (gfp; Hammer & Swanson, 1999).

Neither ppGpp nor DksA is required for growth inhibition by propionic acid, since WT, ppGpp⁰ and dksA mutant cultures responded similarly (Fig. 7A, top). However, unlike WT bacteria, which exhibited heightened fluorescence 3 h post-propionic acid treatment, dksA mutant bacteria failed to fluoresce throughout the time course analyzed (Fig. 7A, bottom). Consistent with their lack of motility in the PE phase, dksA mutant control cultures did not activate flaA expression even when the cells reached stationary phase, while WT bacteria did (9 h; Fig. 7A, bottom). Therefore, L. pneumophila require DksA to induce the flaA promoter when the stringent response is triggered by propionic acid or at the transition to PE phase (Hammer & Swanson, 1999; Edwards et al, 2009).

Figure 7. In response to fatty acid stress, DksA activates transmission traits with and independently of ppGpp.

A. To determine whether DksA is required for flaA promoter activation in response to excess propionic acid (PA), mid-E phase bacteria harboring the pflaAgfp reporter plasmid were normalized to an OD₆₀₀ (0.75–0.85), then 10 mM (PA) (solid lines), or water (dashed lines) were added to WT (triangles), ppGpp⁰ (circles), or dksA (squares) bacteria. At 3 h time intervals, cell density (top panel) and flaA promoter activity was quantified by measuring green fluorescent protein (gfp) accumulation using fluorometry. The data depicted represent one of three independent experiments. B. To determine if DksA contributes to increased resistance to degradation by macrophages after PA treatment, PA (light gray), or water (white) was added to cultures as described in A. After 3 h, bacteria were used to infect macrophages, and resistance to lysosomal degradation was scored by fluorescence microscopy. Shown are the mean percentages from triplicate coverslips ± SE from three independent experiments. In addition to the p values depicted, the mean percent resistance values for WT PA and dksA PA were statistically different by a two-tailed paired Student t-test (p = 0.005).

To verify its role in the L. pneumophila response to fatty acid perturbation, we analyzed the contribution of DksA to evasion of lysosomal degradation. After a 3 h propionic acid treatment, ~ 50% of the infected macrophages contained intact WT bacteria, whereas < 10% harbored intact dksA mutant bacteria (Fig. 7B). Therefore, L. pneumophila also require DksA to induce resistance to macrophage degradation when the stringent response is triggered by propionic acid.

Unlike DksA, ppGpp was only partially required for activation of the flaA-gfp transmission reporter. We repeatedly observed modest promoter activation 6 – 9 h after propionic acid addition to ppGpp⁰ cultures (Fig. 7A, bottom). In addition, ~ 20% of alarmone deficient bacteria avoided degradation, another intermediate response (Fig. 7B). Importantly, DksA accounted for the ppGpp-independent flaA promoter activity and degradation evasion, since dksA mutants treated with propionic acid did not increase their fluorescence (Fig. 7A, bottom) or remain intact in macrophages (Fig. 7B). Thus, L. pneumophila transmission trait activation in response to excess fatty acids is mediated by a DksA activity that is enhanced by ppGpp.

DksA induction leads to ppGpp-independent flagellar biosynthesis

To continue to investigate the capacity of DksA to act independently, we asked whether this transcription factor could control flagellar biogenesis in the absence of ppGpp. Using the strains characterized in Figure 6, we measured whether induction of dksA expression could rescue the flagellar synthesis defect of ppGpp⁰ mutant L. pneumophila.

Once control bacteria entered stationary phase (~ 9 h post-IPTG; Fig. 6A), we assayed levels of flagellin and macrophage cytotoxicity, a flagellin-dependent phenotype (Molofsky et al, 2006). Induction of plasmid-borne dksA increased both the cell-associated flagellin and cytotoxicity of ppGpp⁰ bacteria (Fig. 8). Likewise, dksA induction restored motility of ~ 20 – 35% of the ppGpp⁰ bacteria. Full bypass of ppGpp by DksA was not achieved, as flagellin levels, macrophage killing and motility of ppGpp⁰ mutant bacteria were still lower than PE phase WT (Fig. 8). In contrast, relA^E.c. induction did not restore flagellin synthesis, cytotoxicity or motility to dksA mutants (Fig. 8; data not shown). Therefore, when expressed from a plasmid, dksA can activate flagellin-dependent phenotypes independently of ppGpp. Furthermore, this contribution by DksA cannot be bypassed by constitutive ppGpp synthesis.

Figure 8. Induction of dksA is sufficient to bypass the requirement for ppGpp for flagellin-dependent phenotypes.

In the strains subjected to the induction described in Figure 6A, we assessed levels of flagellin and macrophage cytotoxicity. At 9 h post-IPTG (indicated in Fig. 6A by *), bacterial cell pellets were harvested, and levels of cell-associated flagellin were assessed by Western analysis (upper image). At 9 h post-IPTG, bacteria were also added to triplicate wells of macrophages at the MOIs shown (bottom panel), and cytotoxicity was measured. The values plotted represent the mean percent macrophage killing ± SE for triplicate samples determined in one of three similar experiments.

DksA contributes to the flagellar regulon

As an independent approach to analyze the contribution by DksA to the flagellar gene regulon, the transcriptional profiles of PE phase WT and dksA mutant L. pneumophila were compared (Table 2). Consistent with a role for DksA in the down-regulation of ribosomal genes in PE phase, several ribosomal transcripts were elevated in dksA mutants relative to WT bacteria (http://www.ebi.ac.uk/microarray-as/ae/). As in P. aeruginosa, L. pneumophila flagellar gene transcription occurs in a hierarchy of four classes (Dasgupta et al, 2003; Bruggemann et al, 2006; Heuner, 2007; Albert-Weissenberger et al, 2010). DksA was especially important for PE phase accumulation of late flagellar transcripts, including those encoding structural elements critical for final assembly and motor components essential for flagellar rotation (Table 2). In addition, Class II rod and hook transcripts and the Class III sigma factor fliA as well as each of its downstream flagellar gene targets required DksA for up-regulation (Table 2; Albert-Weissenberger et al, 2010). DksA also contributed to an increase in RNA for RpoN (σ⁵⁴), a factor shown to play a modest role in PE phase up-regulation of a few Class II flagellar genes, including flgB (Albert-Weissenberger et al, 2010). Thus, L. pneumophila employs the DksA protein to increase the level of flagellar transcripts in the stationary phase of growth.

Table 2.

Flagellar gene expression of PE phase dksA mutant L. pneumophila.

Gene Name	Gene.ID	Annotation	dksA/WT
Class I
rpoN	lpg0477	RNA polymerase sigma factor (sigma-54)	−2.17
fleQ	lpg0853	Master regulator of flagellar gene transcription	---
Class IIa
fliJ	lpg1756	Export and assembly	---
fliI	lpg1757	Flagellum-specific ATP synthase	---
fliH	lpg1758	Export and assembly	---
fliG	lpg1759	Motor switch protein	---
fliF	lpg1760	M-ring protein	−2.17
fliE	lpg1761	Hook-basal body complex protein	---
fleR	lpg1762	Two-component response regulator	---
fleS	lpg1763	Two-component sensor histidine kinase	---
flhA	lpg1785	Export and assembly	---
flhB	lpg1786	Export and assembly	---
fliR	lpg1787	Export and assembly	---
fliQ	lpg1788	Export and assembly	---
fliP	lpg1789	Export and assembly	---
fliO	lpg1790	Export and assembly	---
fliN	lpg1791	Motor switch protein	−2.85
fliM	lpg1792	Motor switch protein	---
Class IIb
flgA	lpg0908	P-ring biosynthesis	---
flgB	lpg1216	Proximal rod protein	−2.38
flgC	lpg1217	Proximal rod protein	−4.55
flgD	lpg1218	Rod modification protein	−4.00
flgE	lpg1219	Flagellar hook protein	−11.11
flgF	lpg1220	Proximal rod protein	−4.34
flgG	lpg1221	Distal rod protein	---
flgH	lpg1222	L-ring protein precursor	---
flgI	lpg1223	P-ring protein precursor	---
flgJ	lpg1224	Peptidoglycan-hydrolyzing protein	---
flgK	lpg1225	Hook-associated protein	−2.70
flgL*	lpg1226	Flagellar hook-associated protein	−12.50
fliK	lpg1688	Hook-length control protein	−3.57
fleN	lpg1783	Regulator of flagellar synthesis	−7.14
flhF	lpg1784	Biosynthesis regulator GTP-binding protein	−6.25
Class III
flgN*	lpg0906	Potential chaperone	−8.33
flgM*	lpg0907	Anti-sigma-28 factor	−20.00
motB*	lpg1780	Sodium-type motor protein	−16.67
motA*	lpg1781	Sodium-type motor protein	−20.00
fliA	lpg1782	RNA polymerase sigma factor (sigma-28)	−20.00
motA2	lpg2318	Proton conductor component of motor	−10.00
motB2	lpg2319	Motor protein	−16.67
flhB’*	lpg2583	Unknown	−10.00
Class IV
fliS*	lpg1337	Potential chaperone	−20.00
fliD*	lpg1338	Filament cap	−25.00
flaG*	lpg1339	Unknown	−33.33
flaA*	lpg1340	Flagellin	−50.00
motY*	lpg2962	Sodium-type motor protein	−7.14

The data were collected from two independent biological replicates. See Fig. S2 for RT-PCR validation.

p < 0.01 for all targets shown.

^*indicates a gene regulated by fliA (σ²⁸) (Albert-Weissenberger et al, 2010)

ppGpp modulates DksA-dependent expression of flagellar genes

To investigate how DksA contributes to expression of flagellar genes stimulated by alarmone, we induced synchronous ppGpp synthesis by WT and dksA mutant bacteria, and then analyzed expression of genes representing three tiers of the flagellar regulon 0, 90 and 300 min later. By 300 min, 95–100% of WT cells were motile, verifying functional flagella. Furthermore, the growth of both WT and dksA mutant L. pneumophila was rapidly inhibited (Fig. S3), suggesting that the strains accumulated ppGpp similarly (Schreiber et al, 1991). Therefore, the fold-increase relative to t = 0 was calculated for three representative transcripts: flgB, a Class II gene encoding a flagellar basal-body rod protein; fliA, a putative a Class III gene encoding σ²⁸, the sigma factor that activates flaA (Heuner et al, 2002); and flaA, a Class IV gene encoding flagellin.

By 90 min, flgB, fliA and flaA RNA levels had increased in WT bacteria, revealing that soon after ppGpp synthesis, each tier of the flagellar gene cascade is active (Fig. 9A). Between 90 and 300 min, fliA and flaA levels increased further in WT bacteria, whereas flgB levels declined. Therefore, as alarmone accumulates (Hammer & Swanson, 1999), expression of the Class III fliA and Class IV flaA loci is continuously induced, whereas expression of the Class II flgB gene is induced, and then repressed.

Figure 9. L. pneumophila uses ppGpp to modulate DksA-dependent gene expression.

A. To assess the contribution of DksA to gene expression by ppGpp, qRT-PCR was performed on transcripts isolated from mid-E phase AYET broth cultures of WT and dksA mutant bacteria induced to express prelA^E.c. with 500 µM IPTG. RNA was harvested at 0, 90 and 300 minutes post-IPTG, and the relative transcript levels were assessed by dividing 90 min (dark grey) and 300 min (light grey) values by the 0 min value to give the fold increase for each target. Depicted are the mean log₂ (fold increase) values from triplicate wells in three independent experiments ± SE. Dashed lines delineate the two-fold change cutoff. B. To monitor fliA promoter activity at the E to PE phase transition, mid-E phase WT (triangles), ppGpp⁰ (circles), or dksA (squares) bacteria harboring the pfliAgfp reporter plasmid were diluted to an OD₆₀₀ of 1.0. Cell density (dashed lines) and fliA promoter activity (solid lines) were quantified until bacteria entered stationary phase. Actual fluorescence values were 1000× greater than those shown. The data represent one of three independent experiments, and the differences between WT and dksA, and ppGpp⁰ and dksA at 0* and 3 h** were statistically significant by a two-tailed paired Student t-test (p = 0.01*, 0.04** and 0.03*, 0.006**).

Rapid ppGpp-mediated up-regulation required DksA, as no increase was observed for these loci at 90 min post-induction in dksA mutants (Fig. 9A). The DksA contribution to flaA RNA accumulation could be partially bypassed by relA^E.c. induction, as dksA mutants showed modest ~ 3-fold increase in flaA transcript at 300 min. However, when their flaA levels were compared directly, dksA mutants contained ~ 140 fold less flaA mRNA than WT bacteria (data not shown).

Consistent with fliA mRNA accumulation patterns, ppGpp and DksA were both required for activation of the fliA promoter at the E to PE phase transition, as judged by a fliA-gfp reporter (Fig. 9B). Additionally, DksA contributed to basal fliA promoter activity in E phase, since WT and ppGpp⁰ mutant fluorescence values were consistently 2–3 fold higher than those of dksA bacteria. Likewise, in the uninduced qRT-PCR samples, WT bacteria contained ~ 3 fold more fliA mRNA than dksA mutants (data not shown). Therefore, L. pneumophila requires the DksA transcription factor to activate the flagellar regulon, whereas the alarmone modulates this activation, by either amplifying or repressing flagellar gene expression over time.

DISCUSSION

When their capacity to synthesize either proteins or fatty acids is compromised, replicating L. pneumophila rely on the signaling molecule ppGpp to orchestrate differentiation to a transmissive, infectious form (Edwards et al, 2009; Dalebroux et al, 2009). In the process, intracellular L. pneumophila alter the level of ~ 800 transcripts (Bruggemann et al, 2006). Here we demonstrate that ppGpp and the transcription factor DksA can either cooperate or act independently to coordinate this developmental program.

Composed of over 40 genes whose expression occurs in four distinct phases, the flagellar transcriptional cascade provides a sensitive read-out for analyzing the impact of ppGpp and DksA during L. pneumophila differentiation. Several factors regulate the hierarchy, including the master regulator FleQ (Class II and III), the response regulator FleR and the alternative sigma factors σ⁵⁴ (Class II) and σ²⁸ (Class IV; Bruggemann et al, 2006; Heuner, 2007; Albert-Weissenberger et al, 2010). The alarmone rapidly increases the level of fleQ, fleR and several Class II flagellar transcripts, including flgB (Table 1).

Kinetic analysis of flgB expression revealed the versatility of ppGpp and DksA control. In response to ppGpp, DksA is essential for flgB expression, but the amount and/or duration of the ppGpp stimulus governs the level of flgB RNA. As ppGpp accumulates, RNA for the Class II, σ⁵⁴-dependent gene target flgB first accumulates and then decreases (Fig. 9A). In S. enterica, only six subunits of FlgB polymerize with other rod proteins prior to hook formation (Chevance & Hughes, 2008). Thus, the pulse of flgB RNA orchestrated by ppGpp and DksA may contribute to proper stoichiometry of rod and hook subunits.

In contrast to flgB, ppGpp and DksA continuously activate fliA (σ²⁸) and flaA during differentiation (Fig. 9A), consistent with induction of fliA in the transmissive phase (Fig. 9B; Bruggemann et al, 2006). As alarmone increases (Hammer & Swanson, 1999), so do fliA and flaA RNA levels, suggesting functional FliA accumulates. FliA activates Class IV flagellar genes such as flaA and fliD, in addition to genes unrelated to flagellar biogenesis (Heuner et al, 2002; Molofsky et al, 2005; Bruggemann et al, 2006; Albert-Weissenberger et al, 2010). Continuous elevated activation of flaA by FliA is fitting, since the typical bacterial flagellum is comprised of ~ 20,000 filament subunits (Chevance & Hughes, 2008).

Phenotypic analyses also indicate that ppGpp modulates activation initiated by DksA. In response to fatty acid stress, DksA controls L. pneumophila flaA promoter activation and evasion of macrophage degradation independently of ppGpp (Fig. 7). When plasmid-borne dksA expression is induced, ppGpp⁰ bacteria initiate flagellar biosynthesis, motility and cytotoxicity to macrophages (Fig. 8). Furthermore, DksA mediates basal fliA promoter activity during E phase (Fig. 9B). Likewise, in both exponential and stationary phase E.coli, DksA regulates flagellar biogenesis directly, in this case by inhibiting transcription from a σ⁷⁰-dependent fliA promoter (Lemke et al, 2009). Furthermore, when over-expressed, DksA suppresses several E. coli ppGpp⁰ mutant phenotypes, including amino acid auxotrophy, autoaggregation, motility and RpoS accumulation (Potrykus & Cashel, 2008). DksA can also regulate E. coli gene expression independently and even oppositely of ppGpp (Lyzen et al, 2009; Merrikh et al, 2009). Perhaps at the promoters of certain flagellar genes or their regulators (FleQ, σ⁵⁴, FleR, and σ²⁸), DksA controls RNAP activity independently of ppGpp.

It is noteworthy that E. coli and L. pneumophila each enlist DksA and ppGpp to regulate flagellar genes, but to the opposite effect. Commensal enteric E. coli utilize DksA and ppGpp to repress ribosome and flagella synthesis simultaneously (Lemke et al, 2009), whereas aquatic intracellular L. pneumophila have co-opted the stringent response to down-regulate ribosomal transcripts while activating flagellar genes, which promote transmission to a new replication niche. Judging from BLASTP analyses, L. pneumophila also lack the canonical chemotaxis components that equip E. coli to swim toward nutrients (data not shown). Because L. pneumophila is constrained within a host vacuole during replication, this pathogen presumably exploits whatever nutrients are available. Also, E. coli is equipped with numerous peritrichous flagella, whereas L. pneumophila synthesize a monopolar flagellum. Regardless of the distinct costs and benefits that motility confers to E. coli and L. pneumophila in their natural reservoirs, mechanistic studies can now exploit this dichotomy. Perhaps sequence differences observed in the N-terminal globular domain of the two DksA proteins (Fig. 2A) account for their distinct mode of regulation. Alternatively, the promoter architecture of key regulators like fliA may dictate the impact of each stress response.

To express certain transmission traits fully, L. pneumophila requires that DksA cooperate with ppGpp. For example, DksA only partially activates the flaA promoter and evasion of lysosomal degradation by ppGpp⁰ mutants in response to propionic acid (Fig. 7). The alarmone is also essential for activation of transmission phenotypes in response to acetate (data not shown) or entry into stationary phase (Figs. 3 and 7A). Moreover, dksA induction fails to rescue pigmentation (Fig. 6B), sodium sensitivity, infectivity and stationary phase survival defects of ppGpp⁰ mutants (data not shown). Whether ppGpp affects DksA protein levels has not been tested in L. pneumophila, but dksA RNA levels are similar during the replicative and transmissive phases in broth and amoebae (Bruggemann et al, 2006). The genetic studies presented here establish that the capacity of DksA and ppGpp to act independently or cooperatively during L. pneumophila differentiation is context-dependent.

Analysis of DksA and ppGpp also revealed that the requirements for L. pneumophila to transit from broth to macrophage and from macrophage-to-macrophage are distinct. Like DksA, the regulators LetA, LetS, LetE and FliA each activate transmission phenotypes in broth, yet are dispensable for transmission from one macrophage to another (Fig. 3; Hammer et al, 2002). In contrast, ppGpp-dependent factors are critical for L. pneumophila transmission from lysosomal vacuoles of permissive mouse macrophages (Sturgill-Koszycki & Swanson, 2000; Dalebroux et al, 2009). Perhaps in host cells, ppGpp induces L. pneumophila pathways that promote infection of A. castellanii (Fig. 4) and spread between macrophages (Fig. 5) but are poorly expressed in broth culture. In fact, of the cohort of genes upregulated during transmission in amoebae, only ~ 77% are also elevated during PE phase in broth (Bruggemann et al, 2006). It is also possible that L. pneumophila regulators control parallel, or perhaps even redundant pathways in particular host environments. Indeed, although dispensable for macrophage transmission, the FliA sigma factor and the LetA response regulator are required for bacterial growth in particular amoebae (Heuner et al, 2002; Gal Mor & Segal, 2003). Also, some bacteria use ppGpp to control cellular processes through physical interactions with factors other than RNAP (Wang et al, 2007; Zhao et al, 2008).

Synthesis of ppGpp by L. pneumophila rapidly alters the level of numerous transcripts. As in E. coli (Durfee et al, 2008; Traxler et al, 2008), ppGpp immediately down-regulates the protein synthesis machinery of L. pneumophila. Indeed, of the targets showing decreased RNA levels, nearly half encode ribosomal proteins and translation machinery (Table S1). While the impact of ppGpp on RNAP has been the focus of much research, the alarmone also induces growth arrest and may alter transcript stability, factors which will also affect the level of transcripts in a cell. Mechanistic studies of particular promoters and transcripts can now identify how ppGpp alters the L. pneumophila transcriptional profile during its life cycle.

In the transmissive state, L. pneumophila activates its Dot/Icm Type IV secretion system; this virulence mechanism is also coordinated in part by ppGpp. For example, genes encoding secreted substrates are activated in the PE phase (Nagai et al, 2002; Bruggemann et al, 2006; Tiaden et al, 2007; Rasis & Segal, 2009), and effectors such as LepA and LepB contribute to non-lytic release from amoebae (Chen et al, 2004). Consistent with these findings, RNAs for three secreted coiled-coil domain containing host cell effectors rapidly accumulate, including LepA (Table 1). Transcripts encoding membrane and channel components of the Dot/Icm system are also responsive to ppGpp. However, since many of these proteins are constitutively expressed during E and PE phase in broth (J. P. Vogel, personal communication), basal levels of ppGpp may mediate their expression, or more complex post-transcriptional mechanisms contribute.

L. pneumophila differentiation also entails a variety of metabolic changes. In the transmissive form, the pathogen activates several phospholipases, some of which are secreted into host cells via the Type II secretion (Lsp) and Type IV Dot/Icm secretion systems (Banerji et al, 2008). The level of a phospholipase (lpg2837) RNA immediately increases in response to ppGpp (Table 1). Additionally, transcripts for central metabolic and fatty acid activating enzymes like pta are up-regulated by ppGpp. Alarmone accumulation also leads to accumulation of an RNA encoding an uncharacterized type I polyketide synthase (Gokhale et al, 2007). During differentiation to the transmissive form, L. pneumophila modifies its surface and sheds vesicles that inhibit phagosome-lysosome fusion (Fernandez-Moreira et al, 2006). Concomitant with this process, several LPS modification and phospholipid biosynthesis enzymes are down-regulated by ppGpp (Table S1). These alterations in LPS structure and phospholipid content may be critical for transmission of progeny, since mutants that lack ppGpp are degraded at the end of the replication period (Fig 5B). Therefore, regulation of a number of lipid pathways by ppGpp is another component of L. pneumophila differentiation.

L. pneumophila also employs ppGpp and DksA to recruit other regulatory factors to orchestrate its cellular differentiation. In response to alarmone accumulation, L. pneumophila up-regulates transcripts for the stationary phase sigma factor RpoS (σ³⁸) and the non-coding regulatory RNA RsmZ (Table 1). On the other hand, the amount of RsmY RNA did not increase in response to ppGpp. Therefore, our study supports the prediction that the RsmY and RsmZ regulatory RNAs exhibit varying degrees of sensitivity to ppGpp, possibly at the level of transcription initiation or RNA stability.

The L. pneumophila flagellar cascade illustrates how stringent response factors can cooperate to govern bacterial life cycles. The interplay between DksA and ppGpp is complex, as their roles are distinct at different loci. With the identification of candidate genes by microarray analysis, detailed kinetic studies in which ppGpp synthesis by WT and dksA bacteria is synchronized can reveal how ppGpp and DksA coordinate complex developmental processes.

EXPERIMENTAL PROCEDURES

Bacterial strains and culture

L. pneumophila strain Lp02 (thyA hsdR rpsL; MB110), a virulent thymine auxotroph derived from Philadelphia 1 (Berger & Isberg, 1993), was the parental strain for all the strains analyzed. L. pneumophila was cultured at 37°C with agitation in 5 ml of N-(2-acetamido)-2-aminoethanesulfonic acid (ACES; Sigma)-buffered yeast extract (AYE) broth or on ACES-buffered charcoal yeast extract (CYE), supplemented with 100 µg/ml thymidine (AYET, CYET) when necessary. Bacteria from colonies <5 days old were cultured in broth overnight, then subcultured into fresh AYET prior to experiments. Exponential (E) cultures had an optical density at 600 nm (OD₆₀₀) of 0.3 to 2.0; post-exponential (PE) cultures had an OD₆₀₀ of 3.0 to 4.5, a period when the viability of the strains was similar (Fig. 2). Where indicated, ampicillin (amp; Fisher) was added to a final concentration of 100 µg ml⁻¹; gentamycin (gent; Fisher) to 10 µg ml⁻¹; chloramphenicol (cam; Roche) to 5 µg ml⁻¹ for L. pneumophila and 10 ug ml⁻¹ for E. coli; propionic acid to 10 mM; and, isopropyl-beta-D-thiogalactopyranoside (IPTG) to the concentrations specified. To determine colony forming units (CFU), serial dilutions of L. pneumophila were plated on CYET and incubated at 37°C for 4–5 days.

dksA mutant construction

To construct the dksA mutant, a deletion insertion allele was first generated by recombineering in E. coli (Datsenko & Wanner, 2000; Yu et al, 2000). pGEM-dksA was generated by amplifying the dksA region (lpg2338) using primers dksA1 and 2 (Table S2). The FRT::cat::FRT cassette was amplified from pKD3 using primers dksA-pKD3a and dksA-pKD3b. To generate pGEM-ΔdksA::FRT::cat::FRT, pGEM-dksA and the linear PCR product were co-electroporated into DY330, prepared as described (Yu et al, 2000), recombinants were selected on LB-cam, and the insertion verified by PCR. Plasmid DNA from DY330 recombinants was used to transform DH5α, and transformants were verified by PCR. Lp02 was transformed with the ΔdksA::FRT::cat::FRT allele by natural competence (Stone & Kwaik, 1999). Chromosomal recombination was confirmed by PCR, and the resulting ΔdksA::FRT::cat::FRT mutant L. pneumophila was designated MB699 (Table 3).

Table 3.

Bacterial strains and plasmids

Strain	Relevant genotype/phenotype	Reference
E.coli
DH5α	F-endA1 hsdR17 (r- m+) supE44 thi-1 recA1 gyrA/ (Nalr) relA1 Δ(lacZYA-argF)U169Ф80dLacZΔM15λpirRK6	Laboratory collection
DY330	W3110 ΔlacU169 gal490 λcl1857 Δ(cro-bioA)	(Yu et al, 2000)
L. pneumophila
MB110	Lp02 wild-type, StrR, Thy-, HsdR-	(Berger & Isberg, 1993)
MB699	Lp02 ΔdksA::FRT::cat::FRT mutant	This work
MB697	Lp02 relA::gent spoT::kan double mutant	(Dalebroux et al, 2009)
MB698	Lp02 p206-gent, vector control strain	This work
MB701	Lp02 ΔdksA::FRT::cat::FRT mutant p206-gent	This work
MB731	Lp02 ΔdksA::FRT::cat::FRT pdksAi-gent	This work
MB698	Lp02 p206-cat, vector control strain	(Dalebroux et al, 2009)
MB686	Lp02 relA::gent spoT::kan double mutant p206-cat	(Dalebroux et al, 2009)
MB687	Lp02 relA::gent spoT::kan double mutant prelAL.p.	(Dalebroux et al, 2009)
MB700	Lp02 relA::gent spoT::kan double mutant pdksAi-cat.	This work
MB359	Lp02 prelAE.c.	(Hammer & Swanson, 1999)
MB702	Lp02 ΔdksA::FRT::cat::FRT mutant prelAE.c.	This work
MB355	Lp02 pflaAgfp td(Δ)I	(Hammer & Swanson, 1999)
MB732	Lp02 ΔdksA::FRT::cat::FRT mutant pflaAgfp td(Δ)I	This work
MB685	Lp02 relA::gent spoT::kan double mutant pflaAgfp td(Δ)I	(Dalebroux et al, 2009)
MB733	Lp02 pfliAgfp td(Δ)I	This work
MB735	Lp02 ΔdksA::FRT::cat::FRT mutant pfliAgfp td(Δ)I	This work
MB734	Lp02 relA::gent spoT::kan double mutant pfliAgfp td(Δ)I	This work
Plasmids
pGEMT-Easy	MCS within coding region of B-lactamase α-fragment linearized with single-T overhanges, AmpR	Promega
pGEM-Gent	pGEMT-Easy with 1.7 kb gentamycin cassette cloned into the MCS, source of 1.7 kb gentamycin resistance cassette, AmpR, GentR	(Molofsky et al, 2005)
pKD3	Template plasmid for λ Red system, bla FRT::cat::FRT, ori R6K	(Datsenko & Wanner, 2000)
pGEM-dksA	pGEMT-Easy with 1.4 kb PCR amplified dksA chromosomal region ligated into T overhangs, AmpR	This work
pGEM-ΔdksA::FRT::cat::FRT	pGEM-dksA with 1.1 kb FRT::cat::FRT cassette from pKD3 recombineered using strain DY330 to delete and replace the dksA coding sequence leaving ~500 bp homology on either side for Lp02 chromosomal recombination, AmpR , CamR	This work
pGEM-dksAi	pGEMT-Easy with 540 bp PCR amplified dksA chromosomal region ligated into T overhangs, AmpR	This work
p206-cat	pMMB66EH derivative, Δmob lacIq,PtaclacUV5, CamR	(Morales et al, 1991)
pdksAi-cat	p206-cat with 540 bp SalI/HindIII fragment from pGEM-dksAi ligated between the SalI/HindIII sites in the MCS, colinear with the PtaclacUV5 promoter, lacIq, Inducible dksA expression, CamR.	This work
p206-gent	p206-cat with the cat cassette replaced by the gentamycin resistance cassette from pGEM-Gent, GentR.	This work
pdksAi-gent	p206-gent with 540 bp SalI/HindIII fragment from pGEM-dksAi ligated between the SalI/HindIII sites in the MCS, colinear with the PtaclacUV5 promoter, lacIq, Inducible dksA expression, GentR.	This work
prelAE.c.	pMMB66EH derivative, lacIq,PtaclacUV5, GentR with a relA allele from E. coli encoding a truncated, metabolically active RelA, cloned into the MCS, colinear with the PtaclacUV5 promoter, lacIq Inducible relAE.c. expression, GentR.	(Hammer & Swanson, 1999)
pflaAgfp	150 bp flaA promoter fragment fused to GFPmut3 in pKB5 with Ptac and lacIq removed, td(Δ)i	(Hammer & Swanson, 1999)
pfliAgfp	304 bp fliA promoter fragment fused to GFPmut3 in pKB5 with Ptac and lacIq removed, td(Δ)i	This work

Inducible dksA expression

To generate strains in which expression of dksA could be induced, a promoterless fragment of dksA was cloned into either pMMB206-Δmob, a broad host range vector containing a P_taclacUV5 IPTG-inducible promoter with a cam cassette (p206-cat), or the same vector with a gent cassette (p206-gent). To construct pGEM-dksAi, the dksA locus was amplified from Lp02 using primers dksAi1 and i2. The fragment was excised from pGEM-dksAi and ligated into p206-cat or p206-gent immediately 3’ of the P_taclacUV5 promoter, generating pdksAi-cat and pdksAi-gent, respectively. Insertion was confirmed by PCR. For complementation experiments, MB699 transformed with pdksAi-gent were selected on gent, creating MB701 for inducible dksA expression (Table 3). To induce dksA expression by ppGpp⁰ mutant L. pneumophila, MB697 transformed with pdksAi-cat were selected on cam, creating MB700 for inducible dksA expression in the absence of ppGpp.

fliA-gfp promoter fusion

A fragment containing 304 bp 5’ of the fliA RBS (K. Heuner, personal communication) and encoding the putative σ⁷⁰ promoter and transcriptional start was amplified using primers fliAP1 and P2. The fragment was ligated into pKB5 directly 5’of gfp as described (Hammer & Swanson, 1999). This plasmid was used to transform MB110, MB697 and MB699 (Table 3), generating MB733, MB734 and MB735, respectively.

Fluorometry

To monitor expression of the flagellin promoter, E phase cultures of MB355, MB685, and MB732 were diluted to OD₆₀₀ (0.75–0.85) and treated with 10 mM propionic acid or water (t = 0; Fig. 7A). At the times indicated, the cell density of each culture was measured as OD₆₀₀ and fluorescence was quantified as described (Edwards et al, 2009). To quantify fliA promoter activity, E phase cultures of MB733, MB734 and MB735 were diluted to OD₆₀₀ = 1.0 (t = 0) and cultured to stationary phase. Fluorescence was detected as described above except that cultures were normalized to OD₆₀₀ = 3.0.

Detection of ppGpp

Accumulation of ppGpp was detected by thin-layer chromatography (TLC) as described (Dalebroux et al, 2009). Briefly, E phase AYET broth cultures of ppGpp⁰ mutant bacteria carrying either pempty or prelA^L.p were diluted to OD₆₀₀ = 0.25 and labelled with ~ 100 µCi/ml of carrier-free [³²P]-phosphoric acid at 37°C for 6 h (~ 2 generations). After labelling, the E phase cultures were treated with 500 µM IPTG and sampled at 30, 60, and 90 min post-induction.

Flagellin western analysis

After culture to OD₆₀₀ = 1.0 in 50 ml, mutant bacteria were treated with 200 µM IPTG; untreated WT bacteria were the positive control. After culture to PE phase (~ 9 h indicated by * in Fig. 6A), cell pellets were harvested. Since induction of prelA^E.c. immediately inhibited growth of dksA mutants, dksA pempty cultures were normalized to the dksA prelA^E.c. culture density prior to pelleting. Bacteria were lysed with Qproteome™ kit (Qiagen), and equivalent volumes of ppGpp⁰ and dksA mutant L. pneumophila lysates were denatured and separated on a SDS-10% polyacrylamide gel. To avoid signal overload, WT cell lysates were diluted 1:100 in PBS prior to denaturing. Flagellin was detected using a 1:50 dilution of monoclonal antibody 2A5 (Molofsky et al, 2005).

Microarrays

To study the effect of ppGpp induction on L. pneumophila gene expression (Table 1), ppGpp⁰ mutants carrying either pempty, or prelA^L.p., were grown in AYET to E phase OD₆₀₀ = 1.4 before treatment with 500 µM IPTG. After 45 min (when the pempty and prelA^L.p. cultures had increased to OD₆₀₀ = 1.90 and 1.70 , respectively) and 90 min (OD₆₀₀ = 2.45 and 2.10 for pempty and prelA^L.p., respectively), cells were harvested, and total RNA was extracted as described (Milohanic et al, 2003). To test the contribution of DksA to PE phase expression of L. pneumophila flagellar genes (Table 2), WT and dksA mutant bacteria were normalized to an OD₆₀₀ = 1.6 and cultured until 95–100% of the WT culture was motile. RNA was prepared from two independent cultures, and each RNA sample was hybridized twice with dye swap to the microarrays. The design of microarrays was based on all predicted genes of the genomes of L. pneumophila Paris, Lens and Philadelphia (Bruggemann et al, 2006) in addition to the ncRNAs rsmY and rsmZ (Sahr et al, 2009). Reverse transcription, labeling and hybridization were carried out as described (Sahr et al, 2009).

Data normalization and differential analysis were conducted as described (Sahr et al, 2009). If not stated otherwise, only differentially expressed genes with 2-fold changes were taken into consideration. Complete data sets are available at http://genoscript.pasteur.fr in a MIAME compliance public database maintained at the Institut Pasteur and were submitted to the ArrayExpress database maintained at http://www.ebi.ac.uk/microarray-as/ae/ under the Acc. No. pending.

Quantitative Real-time PCR

To validate the microarray data sets (Tables 1, 2 and S1), bacteria were cultured as described for Microarray analysis. To assess whether dksA contributes to ppGpp-dependent transcript accumulation (Fig. 9A), WT and dksA mutant bacteria carrying prelA^E.c.were cultured to E phase OD₆₀₀ (0.7–0.8) and treated with 500 µM IPTG. Bacteria were harvested at 0, 90 and 300 min post-IPTG. By 300 min, WT prelA^E.c. bacteria had transformed to the transmissive state and were fully motile. To isolate RNA for Real-time PCR (RT-PCR), L. pneumophila cell pellets were lysed in TRIzol (Invitrogen), extracted with chloroform/isoamyl alchohol (24/1) and precipitated. Total nucleic acid concentration was assessed, and total nucleic acid was treated with Turbo DNA Free DNase (Ambion) to digest residual genomic DNA (gDNA). RNA integrity was assessed with an Agilent 2100 Bioanalyzer, and cDNA was synthesized using Superscript® II Reverse Transcriptase (RT) (Invitrogen) and random hexamers (New England Biolabs). To control for residual gDNA contamination, a no-RT cDNA synthesis reaction was run for each sample. Prior to qRT-PCR, cDNA was diluted to ~ 300 pg/ul (assuming a 100% RT yield) and ~ 1.5 ng of cDNA was added to the reaction mixture containing Brilliant II SYBR® Green Q-PCR Master Mix (Stratagene), a reference dye, and primers for each target at a final concentration of 300 nM, and reactions were run in triplicate. Non-template control reactions and dissociation curves were run for each primer pair. The lpg2096 locus served as an internal reference, as it showed no change in expression by microarray under any conditions or strains tested in these or other studies (data not shown). qRT-PCR was performed using an MX3000P instrument (Stratagene). For validation of the prelA^E.c. (Table 1 and Table S1) and dksA/WT PE phase (Table 2) microarray data sets, letE, rpoS, flgB, fliA, flaA, fliD, ndk, lag-1 and lpg0260 (Table S2) were targeted (Figs. S1 and S2). To test the contribution of dksA to ppGpp dependent expression of flagellar genes (Fig. 9), flgB, fliA and flaA were targeted. All primers were designed to amplify 100–200 bp segments of cDNA.

Comparisons of the transcriptional profiles of PE phase WT and ppGpp⁰ mutants and of dksA and ppGpp⁰ mutants L. pneumophila were also sought. However, since these ppGpp⁰ mutant microarray data sets failed a series of independent qRT-PCR and promoter fusion validation tests, the data were not analyzed further.

Intracellular growth in A. castellanii

Three-day-old cultures of amoebae were washed in infection buffer (PYG 712 medium without tryptone, glucose, or yeast extract) and adjusted to 5 × 10⁵ to 1.0 × 10⁶ cells ml⁻¹. PE phase AYE broth grown L. pneumophila harboring a plasmid conferring thymine prototrophy (pflaAgfp) were diluted in buffer and mixed with A. castellanii at an MOI of ~ 0.05. After allowing invasion for 1 h at 37°C, adherent amoebae were washed three times with infection buffer, and flasks were returned to 37°C. At the times shown, 300 µl aliquots from each flasks were centrifuged and vortexed to lyse intact amoebae, and CFU were enumerated by plating dilutions on CYE agar. Each infection was carried out in duplicate.

Macrophage cultures

Macrophages were isolated from femurs of female A/J mice (Jackson Laboratory) and cultured in RPMI-1640 containing 10% heat-inactivated fetal bovine serum (RPMI/FBS; Gibco BRL) as described (Swanson & Isberg, 1995). Following a 7-day incubation in L-cell supernatant-conditioned media, macrophages were plated at 5 × 10⁴ per well for cytotoxicity assays or 2.5 × 10⁵ per well for lysosomal degradation assays, infectivity assays and intracellular growth curves.

Infection and growth in macrophages

L. pneumophila binding, entry, and survival inside macrophages during a 2 h incubation was measured as described (Dalebroux et al, 2009). To complement the dksA mutant infectivity defect, plasmid carrying bacteria were induced with 25 µM IPTG in early E phase and cultured to PE phase prior to infection (Fig. 3A). To quantify intracellular growth, each pooled macrophage supernatant and lysate was plated for CFU at various times post-infection as described (Bachman & Swanson, 2001).

Degradation in macrophages

The percentage of intracellular L. pneumophila that remain intact after a 2 h infection was quantified by fluorescence microscopy (Bachman & Swanson, 2001). Except for longer incubation times (18 h) and MOI adjustments, identical procedures were used to image and score infected macrophages at later time points of the primary infection period (Fig. 5B).

Cytotoxicity

To measure contact-dependent cytotoxicity, L. pneumophila were added to macrophages at the indicated MOI, and cytotoxicity was measured spectrophoretically as described (Molofsky et al, 2005). To determine the contribution of dksA to PE phase activation of L. pneumophila cytotoxicity (Fig. 3C), WT pempty was cultured to E or PE phase, ppGpp⁰ pempty to PE phase, and dksA mutants from E to PE phase with 25 µM IPTG prior to infection. To determine if induction of dksA expression or constituitive ppGpp synthesis could restore ppGpp⁰ and dksA mutant cytotoxicity (Fig. 8), WT, ppGpp⁰, and dksA bacteria carrying pempty, pdksA, or prelA^E.c. were cultured from E to PE phase (~ 9 h, indicated by * in Fig. 6A) in the presence of 200 µM IPTG prior to infection.

Sodium sensitivity

The percentage of L. pneumophila that were sensitive to sodium was determined by enumerating colony formation on CYET and CYET containing 100 mM NaCl as described (Byrne & Swanson, 1998).