Defining the Metabolic Pathways and Host-Derived Carbon Substrates Required for Francisella tularensis Intracellular Growth

Gluconeogenesis, but not glycolysis, is essential for F. tularensis intracellular growth and virulence.Unlike most enzymatic reactions of the glycolytic pathway, the conversion between fructose 6-phosphate (F6P) and fructose 1,6-bisphosphate (FBP) is physiologically irreversible, and is catalyzed by enzymes specific to either glycolysis or gluconeogenesis. In F. tularensis, the glycolytic enzyme phosphofructokinase (PfkA) converts F6P to FBP, and the gluconeogenic enzyme fructose 1,6-bisphosphatase (GlpX) performs the reverse reaction (Fig. 1). Deletion of pfkA should prevent F. tularensis from utilizing glucose or glucose 6-phosphate imported from the host, while deletion of glpX should prevent the bacterium from producing F6P during growth on gluconeogenic carbon sources. F6P is a precursor of the pentose phosphate pathway and is used for the de novo synthesis of lipopolysaccharide, peptidoglycan, pentose phosphates, and aromatic amino acids. We hypothesized that if glucose represents a major carbon source for F. tularensis within the host cell, then pfkA would be essential. Alternatively, if glucose is not a major source of carbon utilized by F. tularensis, then the gluconeogenic enzyme glpX would be required in order to synthesize sufficient F6P and glucose-6P from alternate carbon sources.

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FIG 1

An overview of Francisella tularensis subsp. tularensis Schu S4 central carbon metabolism. Labeled enzymes (red) are predicted to be required for the flux of specific carbon substrates (blue) for F. tularensis central carbon metabolism. Orange arrows indicate reactions that are specific to gluconeogenesis. PPP, pentose phosphate pathway; PEP, phosphoenolpyruvate; acetyl-CoA, acetyl coenzyme A; PfkA, phosphofructokinase (FTT_0801); GlpX, fructose 1,6-bisphosphatase (FTT_1631); GlpK, glycerol kinase (FTT_0130); GlpA, glycerol 3-phosphate dehydrogenase (FTT_0132); PckA, phosphoenolpyruvate carboxykinase (FTT_0449); PpdK, pyruvate phosphate dikinase (FTT_0250); MaeA, malic enzyme (FTT_0917); GdhA, glutamate dehydrogenase (FTT_0380).

We first sought to confirm the predicted functions of pfkA and glpX for glycolysis and gluconeogenesis, respectively. Markerless, in-frame deletions were created for pfkA and glpX in Francisella tularensis subsp. tularensis Schu S4, and the deletion strains were grown in defined medium with either glycolytic or gluconeogenic carbon substrates. For all broth cultures, F. tularensis was grown in Chamberlain’s defined medium (CDM) containing a low concentration (∼3mM) of 13 essential and nonessential amino acids and no other major carbon sources (21) (Text S1). In this medium, wild-type (WT) Schu S4 cells grew to low, but detectable levels, presumably by assimilating amino acids for protein synthesis or energy production (Fig. 2A and Fig. S1A). Indeed, Brissac et al. recently demonstrated that supplementation of this medium with 30 mM select amino acids (threonine, proline, methionine, lysine, tyrosine, tryptophan, phenylalanine, asparagine, or serine) permits various degrees of F. novicida growth, suggesting that these amino acids may be utilized as carbon sources (20). Supplementation with either glucose or glutamate supported robust growth of WT Schu S4 cells (Fig. 2A and Fig. S1A). A ΔpfkA mutant grew to WT levels during growth on glutamate, but did not grow on glucose, possibly due to glucose-mediated repression of alternative carbon catabolic pathways (Fig. 2A). Importantly, though terminal optical density at 600 nm (OD₆₀₀) describes the overall capacity of each mutant to grow on different carbon substrates, closer attention to in vitro doubling times reveals subtle nuances in how each deletion affects growth. For instance, a ΔpfkA mutant reaches the same OD₆₀₀ as WT Schu S4, but grows at a much lower rate (Fig. S1B). As expected, the ΔglpX mutant grew on glucose but not on glutamate (Fig. 2A and Fig. S1C). The growth defects of each mutant were restored to WT levels when the deleted genes were complemented in trans (Fig. 2A and Fig. S1A to C).

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FIG 2

F. tularensis GlpX is essential for replication on gluconeogenic carbon substrates, within host macrophages, and in a murine model of infection. (A) Terminal OD₆₀₀ of WT Schu S4, ΔpfkA, and ΔglpX strains after 48 h of growth in CDM and CDM supplemented with glucose or glutamate at a final concentration of 0.4%. (B) Intracellular replication of WT Schu S4, ΔpfkA, ΔglpX, and ΔglpX pglpX strains in BMDMs, as indicated via relative light units (RLU) measured every 15 min over a 36-hour period. (C) Growth of WT Schu S4 and (D) ΔglpX strains in BMDMs cultured with or without 150 µM AICAR and/or glucose at a concentration of 4.5 g/liter. Growth was measured via luminescence read every 15 min over 36 h. Each growth curve represents one of three independent experiments, and each data point represents the average of three technical triplicates. (E) Organ burdens of mice 3 days post intranasal inoculation with WT Schu S4, ΔpfkA, and ΔglpX strains. Data are pooled from three independent experiments. **, P < 0.01, and ***, P < 0.001, as determined by Student's t test.

To assess the importance of glycolysis and gluconeogenesis during F. tularensis intracellular growth, we utilized a luminescence reporter to monitor intracellular growth, as previously described (16), where F. tularensis Schu S4 strains harbor a plasmid expressing luciferase enzyme and substrate, as well as an addiction system to maintain the plasmid even in the absence of antibiotic selection. As demonstrated previously, an increase in the bacterial burden within the infected cell is directly proportional to an increase in reporter luminescence (16). Bone marrow-derived macrophages (BMDMs) were infected with WT Schu S4, the ΔpfkA mutant, or the ΔglpX mutant, each strain harboring the luminescence reporter. Twenty-four hours postinoculation, WT Schu S4 and ΔpfkA mutant cells grew to similar levels within the BMDMs, while the growth of the ΔglpX mutant was reduced approximately 10-fold relative to that of the WT and the ΔpfkA mutant (Fig. 2B). These data indicate that gluconeogenesis, but not glycolysis, is necessary for WT levels of F. tularensis intracellular growth and suggests that glucose does not represent a major carbon source within macrophage cells.

We hypothesized that the severe intracellular growth defect observed for the ΔglpX mutant was due to the mutant’s inability to synthesize sufficient levels of F6P and G6P from the catabolism of gluconeogenic carbon substrates. Therefore, supplementation with excess glucose should rescue ΔglpX mutant growth within cells. J774A.1 cells are a transformed macrophage cell line constitutively expressing c-Myc, and therefore import large quantities of glucose to increase glycolytic flux (22, 23). To determine if excess glucose could restore the growth defect of the ΔglpX mutant, we infected J774A.1 cells with WT Schu S4, the ΔpfkA mutant, and the ΔglpX mutant, supplied the infected cells with either high-glucose (4.5 g/liter) or glucose-free Dulbecco minimal essential medium (DMEM), and then measured bacterial growth over a 36-hour period. WT Schu S4 and ΔpfkA mutant cells exhibited significant growth with or without glucose supplementation (Fig. S2A). As expected, the ΔglpX mutant strain did not replicate within J774A.1 cells cultured in glucose-free DMEM; however, intracellular replication was restored to WT levels with excess glucose (Fig. S2B).

The rescue of the ΔglpX mutant did not occur in primary BMDMs, as all BMDM infections were performed in high-glucose (4.5g/liter) DMEM (Fig. 2B). This observation suggests that the reduced level of glucose import and glycolytic flux exhibited by BMDMs, relative to those of J774A.1 cells, is insufficient to permit ΔglpX cells from acquiring adequate glucose from the host to restore WT growth properties, even when glucose is present at high concentrations in the medium. To test this, we attempted to rescue growth of ΔglpX in BMDMs by treating the host cells with 5-amnoimidazole-4-carboxamide ribonucleotide (AICAR). AICAR is an analog of AMP (AMP) that stimulates activation of the major host metabolic regulator, AMP-dependent protein kinase (AMPK) (24). When activated, AMPK stimulates glucose uptake and energy production in part by increasing expression of major glucose transporters GLUT1 and GLUT4, and by increasing overall host glycolytic flux (25). We hypothesized that AICAR treatment of BMDMs cultured in high-glucose DMEM would restore ΔglpX mutant intracellular growth by stimulating glucose import. Indeed, while AICAR had little impact on the growth of WT Schu S4 within BMDMs (Fig. 2C), AICAR treatment significantly increased the intracellular growth of ΔglpX in BMDMs cultured in high-glucose DMEM (Fig. 2D). Altogether, our results support the conclusion that the inability of the ΔglpX mutant to fully assimilate gluconeogenic carbon sources results in attenuated growth during periods of glucose limitation.

We next tested whether F. tularensis similarly requires glpX and not pfkA for replication in a murine model of F. tularensis pulmonary infection. Groups of C57BL6/J female mice were infected intranasally with 100 CFU of WT, ΔpfkA, or ΔglpX Schu S4 strains. Three days postinfection, the lungs, livers, and spleens of the infected mice were harvested, homogenized, and plated for bacterial enumeration. Organ burdens for the ΔpfkA mutant strain were similar to those of WT Schu S4 (Fig. 2E). However, the number of CFU recovered from the lungs of mice infected with the ΔglpX mutant was similar to that of the original inoculum and below the limit of detection in the liver and spleen (Fig. 2E). These data align with our observations that glpX, and therefore gluconeogenesis, is necessary for F. tularensis replication in host cells, whereas pfkA and glycolysis are dispensable.

F. tularensis possesses multiple pathways that supply gluconeogenic substrates to support intracellular growth.Our data suggest that a ΔglpX mutant does not produce essential biosynthetic precursors from the nutrients available within the host cell. Since the deletion of glpX precludes the utilization of a large number of gluconeogenic carbon sources, such as glycerol, pentose sugars, amino acids, lactate, pyruvate, and tricarboxylic acid (TCA) cycle intermediates, we generated F. tularensis mutant strains unable to utilize some of these specific carbon sources. Like the conversion of F6P to FBP, the enzymatic conversion of pyruvate to phosphoenol pyruvate (PEP) during glycolysis is physiologically irreversible and must be bypassed during gluconeogenesis. The ATP-dependent decarboxylation of oxaloacetate to PEP is catalyzed by PEP carboxykinase (pckA) and is important for growth on carboxylic or amino acids. Alternatively, pyruvate-phosphate dikinase (ppdK) converts pyruvate to PEP and is required for growth on pyruvate, lactate, and some amino acids. The reactions catalyzed by each enzyme can independently fuel the gluconeogenic pathway to generate essential metabolic precursors necessary for growth. (Fig. 1).

We generated markerless, in-frame deletions of ppdK and pckA. Growth characteristics of each mutant were analyzed in defined medium with specific glycolytic or gluconeogenic substrates to confirm the metabolic function of each enzyme. Both ΔppdK and ΔpckA mutants grew to levels similar to that of WT Schu S4 in CDM supplemented with glucose (Fig. 3A and B). However, while the ΔpckA mutant grew to WT levels in CDM with or without excess glutamate, the ΔppdK mutant had a severe growth defect in CDM and in CDM supplemented with glutamate, similar to that of the ΔglpX mutant (Fig. 3A and B). These data suggest that during growth in defined medium, F. tularensis preferentially synthesizes PEP from pyruvate (PpdK) and not from oxaloacetate (PckA).

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FIG 3

Growth of ΔpckA and ΔppdK mutants in defined medium, host cells, and a murine model of infection. Terminal OD₆₀₀ of (A) ΔpckA and (B) ΔppdK and ΔppdK pppdK strains after 48 h of growth in CDM and CDM supplemented with glucose or glutamate at a final concentration of 0.4%. Data are pooled from three triplicate wells from three independent experiments (mean ± standard deviation [SD]). (C) The intracellular growth kinetics of ΔpckA, ΔppdK, and ΔppdK ΔpckA mutants cultured in high-glucose (4.5 g/liter) DMEM, as indicated via RLU measured every 15 min over a 36-hour period. The data shown represent three independent experiments, and each data point represents the average of three technical replicates. (D) Organ burdens of mice 3 days post intranasal inoculation with WT Schu S4, ΔppdK, ΔpckA, or ΔppdK ΔpckA mutants. Data are pooled from three independent experiments. *, P < 0.05; **, P < 0.01; and ***, P < 0.001, as determined by Student's t test.

We observed that both the ΔppdK and ΔpckA mutants grew to WT levels within the BMDMs (Fig. 3C). Furthermore, a ΔppdK ΔpckA double mutant replicated to significant levels within these cells, albeit at a lower rate, suggesting that these gluconeogenic pathways are not essential for F. tularensis growth within BMDMs (Fig. 3C). When we infected J774A.1 cells with the ΔppdK and ΔpckA mutants, we found that the ΔpckA mutant grew to WT levels within J774A.1 cells cultured with or without glucose supplementation (Fig. S2C). The ΔppdK mutant, however, exhibited significantly reduced growth within J774A.1 cells cultured without glucose (Fig. S2D). Intracellular proliferation of the ΔppdK mutant was restored to WT levels upon high glucose supplementation, indicating that ppdK may contribute to the assimilation of host-derived carbon in J774A.1 cells.

While we found that the organ burdens of the ΔppdK mutant were similar to those of WT Schu S4 in our murine model, we recovered significantly reduced numbers of the ΔpckA mutant from the lungs, livers, and spleens of infected mice (Fig. 3D). Furthermore, we recovered similar numbers of the ΔppdK ΔpckA double mutant relative to that of the ΔpckA single mutant. This indicates that pckA is required for optimal replication in a murine model of F. tularensis infection but that ppdK is dispensable (Fig. 3D).

Amino acids feed the gluconeogenic pathway through the TCA cycle.The attenuation of the ΔpckA mutant in mice suggests that F. tularensis relies on the metabolic pathway catalyzed by PckA during infection. Potential nutrients that can fuel the gluconeogenic pathway through PckA include TCA cycle intermediates or amino acids that feed into the TCA cycle. To discern between these possibilities, we evaluated the importance of glutamate dehydrogenase (gdhA) for F. tularensis intracellular growth. GdhA catalyzes the reversible oxidative deamination of glutamate to α-ketoglutarate, a TCA cycle intermediate (Fig. 1). F. tularensis is predicted to require GdhA to shuttle several amino acids into the TCA cycle, including glutamate, glutamine, proline, arginine, and potentially aspartate and asparagine. Therefore, if F. tularensis preferentially catabolizes amino acids and not TCA cycle intermediates, then a ΔgdhA mutant would likely be similarly attenuated relative to ΔpckA during in vivo growth.

To validate the predicted function of gdhA, we tested a ΔgdhA mutant for growth on glycolytic and gluconeogenic carbon sources in defined medium. As expected, we found that gdhA was required for growth in CDM or CDM supplemented with glutamate, but not in CDM supplemented with glucose (Fig. 4A). Because the ΔgdhA mutant grew to significant levels on glucose in defined media lacking glutamate, we reasoned that gdhA was dispensable for glutamate synthesis but was required for glutamate assimilation.

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FIG 4

GdhA fuels gluconeogenesis by shuttling carbon into the TCA cycle. (A) Terminal OD₆₀₀ of ΔgdhA cells after 48 h of growth in CDM and CDM supplemented with glucose or glutamate at a final concentration of 0.4%. Data are pooled from three triplicate wells from three independent experiments (mean ± SD). (B) The intracellular growth kinetics of WT Schu S4, ΔgdhA, and ΔgdhA pgdhA strains within BMDMs, as indicated via RLU measured every 15 min over a 36-h period. (C) ΔgdhA strains expressing the luciferase (Lux) reporter of intracellular growth in BMDMs cultured with or without 150 µM AICAR and/or glucose at a concentration of 4.5 g/liter. Each growth curve represents one of three independent experiments, and each data point represents the average of three technical triplicates. (D) Organ burdens of mice 3 days post intranasal inoculation with WT Schu S4 or the ΔgdhA mutant. Data are pooled from three independent experiments. *, P < 0.05; **, P < 0.01; and ***, P < 0.001, as determined by Student's t test.

The ΔgdhA deletion mutant exhibited reduced growth in BMDMs that was restored upon expression of gdhA in trans (Fig. 4B), suggesting that GdhA-mediated carbon assimilation represents an important metabolic pathway during F. tularensis replication in BMDMs. When we infected J774A.1 macrophage cells with the ΔgdhA mutant, we found that the defect in bacterial intracellular replication during culture in glucose-free DMEM could be partially rescued with excess glucose, similar to that of the ΔglpX and ΔppdK mutants (Fig. S2E). Similarly, BMDMs cultured in high-glucose DMEM and treated with AICAR permitted significant growth of the ΔgdhA mutant relative to that in untreated BMDMs (Fig. 4C). These data suggest that the intracellular growth defect observed for the ΔgdhA mutant is at least in part due to the ability of this mutant to assimilate sufficient host-derived carbon.

We expected the ΔgdhA mutant to be similarly attenuated relative to a ΔpckA mutant during growth in mice. Strikingly, when we assessed the requirement of gdhA for growth in our murine model of F. tularensis pulmonary infection, we found that the CFU recovered from the lungs, livers, and spleens of gdhA-infected mice were greatly reduced compared to those from the WT, and that the CFU recovered from the liver and spleen of the mice were reduced approximately 3-fold relative to those from the ΔpckA mutant (Fig. 4D). In addition to fueling the gluconeogenic pathway, ΔgdhA-mediated anaplerosis of the TCA cycle may be essential during infection to supply other essential metabolic precursors (e.g., oxaloacetate and/or acetyl coenzyme A [acetyl-CoA]) or to generate reducing power via the use of malic enzyme. This conclusion is consistent with our observation that excess glucose supplementation during growth in J774A.1 or AICAR-treated BMDMs only partially rescued bacterial proliferation of this mutant.

Glycerol catabolism is required for F. tularensis in vivo growth.Since the ΔglpX mutant was more severely attenuated in mice relative to the ΔppdK ΔpckA mutant, we reasoned that F. tularensis may assimilate additional carbon substrates besides those supplied through the gluconeogenic pathways catalyzed by Ppdk and PckA. In F. tularensis, glpA (G3P dehydrogenase) is predicted to be required for the catabolism of glycerol and G3P. We used the Targetron gene knockout system modified for use in Francisella to disrupt glpA in F. tularensis Schu S4 (26). Interestingly, we found that the generation of a ΔglpA mutant strain was only possible through the simultaneous introduction of a secondary mutation in glpK (Fig. S3A). In F. tularensis, glpK is located upstream of glpA and is predicted to encode a kinase responsible for the phosphorylation of glycerol forming G3P during glycerol catabolism (Fig. S3A). The disruption of G3P dehydrogenase in the presence of a fully functional glycerol kinase can lead to increased concentration of intracellular G3P. In Escherichia coli, excess G3P within the cell stimulates the synthesis of the toxic metabolite methylglyoxal (27). We suspect that a similar phenomenon may be responsible for the requirement of a secondary glpK mutation in an F. tularensis ΔglpA background.

We analyzed the growth properties of the glpKA disruption mutant in defined medium supplemented with glucose, glycerol, or G3P to confirm that glpA and glpK are required for growth on glycerol and G3P. As expected, the glpKA mutant grew to WT levels when cultured with glucose but not with glycerol or G3P (Fig. 5A). In fact, supplying the glpKA mutant with G3P led to significantly lower levels of bacterial replication relative to growth on glycerol or just CDM, possibly due to the toxic buildup of intracellular G3P. We found that growth of the glpKA mutant was restored on glycerol and G3P only when these two genes were expressed with the downstream gene, glpF, despite the fact that sequencing of the surrounding genes in our glpKA mutant revealed no additional mutations in or around the coding sequence for glpF. glpF is predicted to encode a glycerol uptake facilitator and may be cotranscribed with glpA (Fig. S3A). As expected, we observed WT levels of growth on G3P but not on glycerol when glpA and glpF, but not glpK, were expressed in trans in the glpKA mutant (Fig. S3B), and growth on both glycerol and G3P was restored when the glpKA mutant was complemented with glpKAF in trans (Fig. 5A). These findings are summarized in Table S1.

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FIG 5

Glycerol metabolism is essential for F. tularensis intracellular replication. (A) Terminal OD₆₀₀ of WT Schu S4, the glpKA insertion mutant, and the corresponding pglpKAF complemented strain grown in CDM and CDM supplemented with glucose, glycerol, or G3P after 48 h of growth. Data are pooled from three triplicate wells from three independent experiments (mean ± SD). (B) Growth curves of WT Schu S4, the glpKA mutant, and the glpKAF complemented strain harboring the LUX reporter within BMDMs. Intracellular bacterial growth was measured via luminescence (RLU), read every 15 min over a 36-hour period. (C) Growth of ΔglpA mutant expressing the LUX reporter of intracellular growth in BMDMs cultured with or without 150 µM AICAR and/or glucose at a concentration of 4.5 g/liter. Each growth curve represents one of three independent experiments, and each data point represents the average of three technical triplicates. (D) Organ burdens of mice 3 days post intranasal inoculation with WT Schu S4 and the glpKA insertional mutant. Data are pooled from three independent experiments. *, P < 0.05; **, P < 0.01; and ***, P < 0.001, as determined by Student's t test.

The glpKA mutant replicated in J774A.1 cells to intermediate levels with or without supplemented glucose, indicating that this mutant can replicate within this cell line, but growth was not fully restored by the addition of excess glucose (Fig. S2F). We found that the glpKA disruption mutant did not replicate within BMDMs (Fig. 5B). Interestingly, growth within BMDMs was restored to WT levels upon in trans expression of glpA and glpF without glpK, suggesting that F. tularensis Schu S4 may assimilate G3P and not glycerol during intracellular growth (Fig. S3C). Growth of the mutant within BMDMs was similarly restored to WT levels upon complementation of glpKAF (Fig. 5B). Finally, replication of the glpKA mutant was significantly increased within BMDMs cultured with AICAR and excess glucose, demonstrating that, similar to the glpX mutant, the glpKA disruption mutant could be rescued by supplying an alternative carbon source (Fig. 5C).

We then assessed the importance of glycerol catabolism for F. tularensis during growth in mice and found that the number of CFU recovered from the lungs of mice infected with the glpKA disruption mutant was similar to the original inoculum and below the limit of detection in the livers and spleens of mice (Fig. 5D). These data suggest that a glpKA mutant colonized, but did not proliferate or disseminate, in a murine model of F. tularensis infection.

Data from our mutational analysis suggest that glycerol represents an essential host-derived source of carbon during F. tularensis intracellular growth. However, we could not exclude the alternative possibility that F. tularensis attenuation may be due to a toxic buildup of metabolites or disruption of proper metabolic regulatory mechanisms in our mutant strains. To delineate these possibilities, we sought to reduce the concentration of available glycerol within BMDMs and to examine the impact on WT F. tularensis intracellular proliferation. A significant bulk of host glycerol stores are sequestered as triglycerides in host lipid droplets (28). During lipolysis, a series of enzymatic reactions free glycerol from cellular triglyceride stores and release it into the cytosol of the host (28, 29). We hypothesized that F. tularensis may exploit this process to establish a source of glycerol during intracellular growth. Atglistatin is a selective inhibitor of adipose triglyceride lipase (ATGL), an enzyme responsible for the first catalytic step of lipolysis (30). When we infected Atglistatin-treated BMDMs with WT F. tularensis, we observed that Atglistatin treatment significantly reduced F. tularensis intracellular burden in a dose-dependent manner (Fig. 6A). Importantly, these concentrations were not cytotoxic to BMDMs (Fig. S4A). To verify these findings, we used Cre-Lox recombination to generate ATGL deficient BMDMs. BMDMs derived from C57Bl6/J or ATGL-flox mice were treated with Cre recombinase gesicles during differentiation. Cre-treated BMDMs isolated from ATGL-flox mice demonstrated approximately 60% knockdown of ATGL expression based on reverse transcription-quantitative PCR (qRT-PCR) (Fig. S4B). This was associated with a significant reduction in F. tularensis replication within ATGL knockdown BMDMs (Fig. 6B). From these data, we conclude that host lipolysis is important for sustaining F. tularensis growth, and that host-derived glycerol represents a primary source of carbon necessary for fueling F. tularensis in vivo replication.

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FIG 6

Active host cell lipolysis is required for efficient F. tularensis intracellular replication. (A) Growth curve of WT Schu S4 cells harboring a LUX reporter within BMDMs cultured in high-glucose (4.5 g/liter) DMEM with or without Atglistatin at indicated concentrations. Intracellular bacterial growth was measured via luminescence (RLU), read every 15 min over a 24-h period. Data represent the mean pooled from 3 replicates in 3 independent experiments. (B) Fold change in WT Schu S4 burden between 24 and 4 h postinfection of WT and ATGL knockdown BMDMs. Data are pooled from three independent experiments. *, P < 0.05; **, P < 0.01; and ***, P < 0.001, as determined by Student's t test).