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. Author manuscript; available in PMC: 2008 May 5.
Published in final edited form as: Mol Microbiol. 2007 Oct 1;66(3):758–770. doi: 10.1111/j.1365-2958.2007.05956.x

LplA1-dependent utilization of host lipoyl peptides enables Listeria cytosolic growth and virulence

Kristie M Keeney 1, Jeanne A Stuckey 2,3, Mary X D O'Riordan 1,*
PMCID: PMC2367003  NIHMSID: NIHMS45909  PMID: 17908209

Summary

The bacterial pathogen Listeria monocytogenes replicates within the cytosol of mammalian cells. Mechanisms by which the bacterium exploits the host cytosolic environment for essential nutrients are poorly defined. L. monocytogenes is a lipoate auxotroph and must scavenge this critical cofactor, using lipoate ligases to facilitate attachment of the lipoyl moiety to metabolic enzyme complexes. Although the L. monocytogenes genome encodes two putative lipoate ligases, LplA1 and LplA2, intracellular replication and virulence require only LplA1. Here we show that LplA1 enables utilization of host-derived lipoyl peptides by L. monocytogenes. LplA1 is dispensable for growth in the presence of free lipoate, but necessary for growth on low concentrations of mammalian lipoyl peptides. Furthermore, we demonstrate that the intracellular growth defect of the ΔlplA1 mutant is rescued by addition of exogenous lipoic acid to host cells, suggesting that L. monocytogenes dependence on LplA1 is dictated by limiting concentrations of available host lipoyl substrates. Thus, the ability of L. monocytogenes and other intracellular pathogens to efficiently use host lipoyl peptides as a source of lipoate may be a requisite adaptation for life within the mammalian cell.

Introduction

Nutrient scavenging is critical for replication and persistence of intracellular bacterial pathogens. The facultative Gram-positive bacterium Listeria monocytogenes establishes a replicative niche within the host cytosol, where it is protected from humoral immunity. Cytosolic growth of L. monocytogenes is rapid, approximating the doubling time of the bacterium in rich broth culture (Marquis et al., 1993). Upon entry into the host cell, L. monocytogenes escapes from the phagosome by secreting a cholesterol-dependent cytolysin, Listeriolysin O (LLO) (Portnoy et al., 2002). LLO-deficient bacteria remain within the vacuole, do not proliferate and are rapidly cleared by the host immune response (Bouwer et al., 1992). Thus, L. monocytogenes must access the cytosol in order to replicate within an infected host. Although the host cytosol contains many nutrients that support its own growth, nonadapted bacterial species like Bacillus subtilis or Yersinia enterocolitica placed in the cytosolic environment replicate poorly (Portnoy et al., 1992; Goetz et al., 2001). These observations suggest that L. monocytogenes has adaptations for survival and nutrient acquisition that facilitate rapid cytosolic growth; however, relatively little is known about how L. monocytogenes exploits the biochemical environment of the host cell.

The requirements for in vitro replication of L. monocytogenes are well defined. For optimal growth in defined medium, the bacterium requires nine essential amino acids, vitamins (biotin, lipoic acid, riboflavin and thiamine), as well as a carbon source such as glucose (Phan-Thanh and Gormon, 1997; Tsai and Hodgson, 2003). The mechanisms of nutrient acquisition during L. monocytogenes intracellular growth have not been extensively studied. At this time, several processes have been empirically demonstrated to directly promote intracellular replication of L. monocytogenes via synthesis or transport of essential nutrients. These include hexose phosphate transport, aromatic amino acid biosynthesis and activation of pyruvate dehydrogenase (PDH) by the lipoate ligase-like protein, LplA1. The hexose phosphate transporter, hpt, contributes to optimal intracellular growth by L. monocytogenes through uptake of glucose phosphates, which are abundant in the host cytosol (Chico-Calero et al., 2002). Hpt mutant bacteria are approximately 10-fold less virulent than wild-type bacteria in a mouse model of infection. Genes involved in aromatic amino acid biosynthesis (aroA, aroB and aroE) are also important during intracellular growth; deletion of any one of these genes results in a 104-fold decrease in virulence (Stritzker et al., 2004). Lastly, the lipoate ligase-like protein, LplA1, is necessary for intracellular, but not extracellular growth (O'Riordan et al., 2003). The LD50 of LplA1-deficient L. monocytogenes is 250-fold greater than the wild-type parental strain in C57BL/6 mice demonstrating the importance of LplA1 for virulence.

The function of lipoate ligases has been extensively characterized in the model organism Escherichia coli (Morris et al., 1994; 1995; Fujiwara et al., 2005). Lipoate, a thiol-containing cofactor, is essential for the oxidative decarboxylation reactions of aerobic metabolism, and also acts as an antioxidant (Jordan and Cronan, 1997; Bast and Haenen, 2003). Lipoate ligases catalyse formation of an amide linkage between lipoate and a conserved lysine residue within target apoenzymes that include components of the PDH, α-ketoglutarate dehydrogenase (KGDH) and glycine H cleavage complexes (Perham, 2000; Zhao et al., 2003). E. coli can synthesize lipoyl groups de novo, but also scavenges extracellular lipoate by a pathway dependent on the lipoate protein ligase LplA (Morris et al., 1995). L. monocytogenes is a lipoate auxotroph and does not encode the genes necessary for lipoate biosynthesis (Welshimer, 1963; Glaser et al., 2001). However, the L. monocytogenes genome encodes two proteins, termed LplA1 and LplA2, that share 54% and 49% similarity, respectively, with E. coli LplA (Glaser et al., 2001). The presence of two LplA-like enzymes in L. monocytogenes suggests the possibility that the bacterium uses two distinct external sources of lipoic acid. Deletion of the lplA1 gene impairs bacterial replication and lipoylation of L. monocytogenes PDH within host cells, but not in rich medium (O'Riordan et al., 2003). As LplA1 and LplA2 are not redundant during intracellular replication, we hypothesized that LplA1 might be required for utilization of a host-derived form of lipoate. Here we show that LplA1 enables L. monocytogenes to use small host-derived lipoyl peptides, revealing an adaptive mechanism to exploit the host cytosol for essential nutrients.

Results

LplA1 and LplA2 both contribute to PDH lipoylation during extracellular growth

Deletion of lplA1 impairs growth and lipoylation of bacterial protein in the intracellular environment, but not in a rich complex medium, brain–heart infusion (BHI) broth, suggesting that L. monocytogenes encodes a second functional lipoate ligase (O'Riordan et al., 2003). LplA1 shares significant amino acid identity and similarity with LplA2 (lmo0764) and E. coli LplA (Fig. S1) (Glaser et al., 2001). To determine whether lplA2 contributes to lipoylation during extracellular growth, we constructed strains containing an in-frame deletion of lplA2, or a disruption of both lplA1 and lplA2, and examined modification of bacterial proteins after growth in BHI (Fig. 1A). The ΔlplA1 and ΔlplA2 mutant strains exhibited lipoyl modification of a 75 kDa protein, previously identified as the E2 subunit of L. monocytogenes PDH (LmPDH), indicating that either enzyme could function during growth in rich medium (O'Riordan et al., 2003). In contrast, the double mutant (lplA1::Tn917ΔlplA2) showed no detectable lipoylation. Both the ΔlplA1 and ΔlplA2 mutant strains grew similarly to the parental wild-type L. monocytogenes strain in BHI, but the lplA1::Tn917ΔlplA2 strain exhibited slower growth (data not shown). Modification of LmPDH was also measured in bacteria grown in improved minimal medium (IMM) containing free lipoic acid (Fig. 1A). LmPDH lipoylation was decreased in the single mutants compared with the wild-type strain. However, the single mutants grew as well as wild-type L. monocytogenes in IMM, demonstrating functional redundancy of LplA1 and LplA2 in medium containing free lipoic acid (Fig. 1B). While we could observe limited growth of the double mutant in BHI, this strain did not grow in IMM, thus the lplA1::Tn917ΔlplA2 strain was not used for further analysis in this study (data not shown). These data demonstrate that at least one of these enzymes, LplA1 or LplA2, is necessary for lipoyl modification in L. monocytogenes.

Fig. 1. L. monocytogenes has two functional lipoate ligase activities.

Fig. 1

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A. Equivalent numbers of stationary phase wild type (WT), ΔlplA1, ΔlplA2 and lplA1::Tn917ΔlplA2 L. monocytogenes, based on OD600, were pelleted and protein harvested using the FastProtein™ Blue Matrix (MP Biomedicals). Bacterial lysates were analysed by SDS-PAGE, followed by immunoblot with an anti-lipoic acid antibody. The band slightly below 75 kDa has previously been identified by Mass-Spectrometry as lipoyl-E2-PDH (O'Riordan et al., 2003). Relative intensities of the bands to wild type were calculated using ImageJ. Loading of equivalent bacterial lysates was confirmed by stripping the blot and reprobing with an anti-E2 antibody (Stein and Firshein, 2000). Bacteria were grown in both the rich medium BHI and IMM, a medium that is limiting for lipoate; lplA1::Tn917ΔlplA2 was only studied in BHI as it exhibited impaired growth in IMM.

B. Wild-type L. monocytogenes and the ΔlplA1 and ΔlplA2 mutant strains were grown in IMM containing different concentrations of lipoic acid. The OD600 was measured after bacteria had reached stationary phase (30 h) and plotted against lipoic acid concentration. The OD600 was measured in a Bioscreen Growth Curve Analyzer.

LplA1, but not LplA2, contributes to bacterial virulence in a mouse model of infection

As both LplA1 and LplA2 enabled lipoate ligase activity in the extracellular environment, we used an animal model of infection to define their respective contributions to in vivo growth and virulence. C57BL/6 mice were infected intraperitoneally (i.p.) with wild-type and mutant L. monocytogenes, and colony-forming units (cfu) from spleen and liver were enumerated at 72 h (Fig. 2A). The bacterial burden in mice infected with wild-type or ΔlplA2 bacteria was several orders of magnitude higher than mice infected with the ΔlplA1 mutant strain (Fig. 2A). We also assessed the ability of ΔlplA1 and ΔlplA2 mutants to compete against wild-type L. monocytogenes by performing a competitive index (CI) analysis. Mice were co-infected with wild-type and mutant L. monocytogenes and cfu were enumerated at 72 h (Fig. 2B). The CI was calculated by dividing the number of wild-type cfu (anti-biotic sensitive) by the mutant cfu (antibiotic resistant). While the CI of the ΔlplA1 strain was approximately 32 and 603 in liver and spleen respectively, the CI of the ΔlplA2 strain was close to 1 for both liver and spleen. These data indicate that LplA2 does not contribute to bacterial fitness or virulence in vivo. In contrast, successful replication of L. monocytogenes in a mouse model of infection required LplA1, even in the presence of LplA2.

Fig. 2. Bacterial virulence in vivo requires LplA1, but not LplA2.

Fig. 2

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A. 2 × 105 total cfu of exponentially growing cultures of wild-type and mutant L. monocytogenes were injected i.p. into ten 5- to 7-week-old male C57BL/6 mice. After 72 h, spleens and livers were harvested, homogenized and plated onto LB.

B. Exponentially growing cultures of wild-type and mutant L. monocytogenes were mixed at a 1:10 (WT : ΔlplA1) or 1:1 ratio (WT : ΔlplA2) and 2 × 105 total cfu injected i.p. into six 5- to 7-week-old male C57BL/6 mice. After 72 h, spleens and livers were harvested, homogenized and plated onto LB with or without 1 μg ml−1 erm. The competitive index was calculated by dividing the number of wild-type strain cfu (ermS) by the number of mutant cfu (ermR). The horizontal line represents the median value. Statistically significant differences between two groups were determined by the Student's t-test at P < 0.05, as indicated by the symbol ‘*’.

LplA1 is required for intracellular growth using host-derived lipoate

We hypothesized that attenuation of virulence and loss of lipoylation in the ΔlplA1 mutant strain was due to inability of the remaining ligase-like protein, LplA2, to use lipoate from the host cytosol. To ensure that intracellular growth was dependent upon host-derived lipoate, we starved wild-type, ΔlplA1 and ΔlplA2 strains in IMM without lipoic acid prior to intracellular infection. LmPDH was extensively lipoylated when the bacteria were grown in the presence of free lipoic acid, but loss of PDH modification was observed in the absence of exogenous lipoate (Fig. 3A). To test the respective contributions of LplA1 and LplA2 towards intracellular growth, bacteria grown with or without lipoic acid were used to infect J774 macrophages. Wild-type and LplA2-deficient L. monocytogenes exhibited similar intracellular growth whether or not the bacteria were lipoate starved (Fig. 3B). However, lipoate-starved ΔlplA1 mutant bacteria no longer replicated within host cells. These results show that LplA2 is dispensable for intracellular growth, but demonstrate an absolute requirement for LplA1 in utilization of host-derived lipoate for proliferation in the macrophage cytosol.

Fig. 3. LplA1, but not LplA2, is essential for intracellular growth.

Fig. 3

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A. A culture of wild-type L. monocytogenes grown in IMM with or without lipoic acid was grown at 37°C to stationary phase, and protein was harvested as in Fig. 1. Bacterial lysates were analysed by immunoblot using an anti-lipoic acid antibody. The 75 kDa band (black arrowhead) corresponds to the L. monocytogenes E2-PDH. Porcine PDH was used as a positive control for the anti-lipoic acid antibody. Loading of equivalent bacterial lysates was confirmed by reprobing the blot with polyclonal anti-Listeria antibody.

B. Wild-type, ΔlplA1 and ΔlplA2 bacterial strains were grown in IMM in the presence (dashed lines) or absence (solid lines) of lipoic acid overnight at 37°C, and used to infect J774 cell cultures. Intracellular growth was quantified by enumerating cfu.

C. Wild-type bacteria were grown in BHI overnight at 37°C and used to infect J774 cells. At 6 h post infection, WT infected J774 cells were lysed and bacteria isolated. Quantitative RT-PCR was performed in triplicate on isolated bacterial RNA to determine template quantities of rpoA, lplA1 and lplA2. Template quantities were normalized against rpoA levels. To control for genomic DNA contamination, a portion of each RNA sample was removed from the reaction prior addition of reverse transcriptase (labelled ‘no RT’) and analysed by QRT-PCR.

D. Starved bacterial strains were used to infect J774 cells with (dashed lines) or without (solid lines) 50 mM DHLA, then cells were lysed and cfu were enumerated. For all growth curves, the mean ± SD was calculated for each time point (n = 3).

Attenuated intracellular growth by the ΔlplA1 mutant could be explained by lack of lplA2 expression or an absence of substrate utilized by LplA2-dependent mechanisms inside the host cell. To examine whether attenuated intracellular growth of the ΔlplA1 mutant strain was due to lack of intracelluar lplA2 expression, we measured levels of lplA1 and lplA2 transcript by quantitative reverse transcriptase polymerase chain reaction (qRT-PCR) in wild-type L. monocytogenes isolated from J774 macrophages at 6 h post infection. lplA2 transcripts were present in intracellular bacteria, albeit at lower levels than lplA1 (Fig. 3C). As there is no detectable free lipoate found in the cytosol, we next investigated if lack of intracellular free lipoate was the limiting factor for growth of the ΔlplA1 strain in the cytosol (Akiba et al., 1998; Podda et al., 1994). Prior to infection, we incubated J774 macrophages with 50 μM dihydrolipoic acid (DHLA) and assessed the effect of supplementation on intracellular growth (Fig. 3D). DHLA supplementation did not further enhance the intracellular growth of wild-type L. monocytogenes but did rescue growth of the ΔlplA1 mutant. Taken together, these results suggest LplA2 is competent to enable intracellular bacterial growth only if free lipoate is added exogenously. As LplA2 does not normally contribute to intracellular growth, we propose that the requirement for LplA1 during cytosolic replication of L. monocytogenes results from the absence of lipoyl substrates used by an LplA2-dependent pathway.

Host-derived lipoyl peptides support LplA1-dependent growth of L. monocytogenes

Lipoyl moieties in mammalian cells are primarily found attached to host proteins rather than as free lipoic acid (Akiba et al., 1998; Perham, 2000). We therefore hypothesized that LplA1, but not LplA2, would permit utilization of lipoyl peptides found in the host cytosol. The predominant lipoyl proteins in J774 macrophages are the E2 subunit of PDH (PDH-E2) and the E2 subunit of α-ketoglutarate dehydrogenase (KGDH-E2), as measured by immunoblot analysis of whole cell lysates using anti-E2/E3 and anti-LA antibodies (data not shown). However, full-length lipoylated PDH-E2 and KGDH-E2 are localized to the mitochondrial matrix and would likely be unavailable to cytosolic bacteria (Margineantu et al., 2002). To determine if lipoylated proteins were present in the cytosol of J774 macrophages, cytosolic and mitochondrial fractions were analysed by SDS-PAGE and immunoblot using an anti-lipoic acid antibody (Fig. 4A). We consistently observed a similar profile of low-molecular-weight lipoylated proteins in the cytosol of uninfected or infected J774 cells. The lipoylated proteins in the cytosolic fraction were unlikely to be the result of mitochondrial rupture during preparation as the abundant mitochondrial E2 PDH and KGDH full-length proteins were not observed in the cytosolic fraction, even though it was concentrated 100-fold. These data suggest that L. monocytogenes has access to lipoyl groups in the host cytosol in the form of lipoylated polypeptides.

Fig. 4. LplA1 enables utilization of degraded host-derived PDH for bacterial growth.

Fig. 4

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A. J774 cell lysate was separated into cytosolic and mitochondrial fractions as described in Experimental procedures. Mitochondrial fractions and concentrated cytosolic fractions (100×) were analysed by immunoblot using a rabbit polyclonal anti-lipoic acid antibody. The 70 kDa band corresponds to mammalian E2-PDH, while the 55 kDa band corresponds to mammalian E2-KGDH.

B. Wild-type (WT) L. monocytogenes was grown in IMM without lipoic acid supplemented with proteinase K (ProK)-digested, trypsin (Tryp)-digested or undigested porcine PDH. Growth was measured by OD600 and the mean value ± SD was calculated for each time point (n = 3).

C. Wild-type and ΔlplA1 L. monocytogenes were grown in IMM containing undigested or ProK-digested porcine PDH or KGDH at 5 mg l−1. After 35 h of growth in the Bioscreen Growth Curve Analyzer, bacterial cultures had reached stationary phase, and the OD600 values were plotted for each condition. BSA at 5 mg l−1 was also digested with ProK as a negative control. Mean values ± SD were calculated for each time point (n = 3).

D. Wild-type, ΔlplA1 and ΔlplA1 complemented with a plasmid expressing LplA1 were grown in IMM containing the lipoate sources indicated; ProK-digested BSA at 5 mg l−1 was used as a negative control. Free lipoic acid (206 Da) (5 μg l−1) and ProK-digested porcine PDH (5 mg l−1) were dialysed against a 100 Da or a 500 Da MWCO membrane, and the retentate was used to supplement IMM. After 19.5 h of growth in conical tubes, the bacterial cultures had reached stationary phase; OD600 values for this time point were plotted for each condition. Growth was not determined (ND) for BSA and LA filtration experiments for ΔlplA1 and ΔlplA1 complemented with a plasmid expressing LplA1.

As lipoyl groups in mammalian cells are protein bound, we tested the ability of modified host proteins, such as PDH-E2 and KGDH-E2, to act as a sole source of lipoyl groups for L. monocytogenes in defined minimal medium (IMM). After lipoate pre-starvation, lipoylated porcine PDH did not support L. monocytogenes growth (Fig. 4B). However, it was possible that L. monocytogenes could not transport the large PDH holoenzyme complex. To determine if LplA1 could enable bacterial utilization of smaller lipoylated peptides, we digested porcine PDH with trypsin, a treatment predicted to release a lipoyl peptide of 17 amino acids in length. Trypsin-digested PDH was not able to supplement growth of L. monocytogenes (Fig. 4B). Thus, the lipoyl peptides generated by trypsin digestion either were not transported into the bacteria, or were not suitable substrates for LplA1-dependent ligase activity. In contrast, proteinase K (ProK)-digested porcine PDH or KGDH, which would be predicted to contain lipoylated tripeptides (∼520 Da), supported growth of wild type but not ΔlplA1 (Fig. 4B and C). The increase in bacterial growth upon ProK digestion of PDH or KGDH was not due to the presence of contaminating free lipoate in ProK, or an increase in non-lipoylated peptides, as ProK-digested BSA did not support replication.

We next considered the possibility that LplA1-dependent growth using ProK-digested PDH was due to the release of free lipoamide from PDH. To detect free lipoamide that may have been generated by ProK digestion, we dialysed ProK-digested PDH using a membrane with a 500 Da molecular weight cut-off (MWCO), and supplemented minimal medium with the retentate (> 500 Da) as a potential lipoate source. As lipoic acid (206 Da) and lipoamide (205 Da) are both smaller than 500 Da, retentate from a control filtration of free lipoic acid using a membrane with a 500 Da MWCO failed to stimulate bacterial growth. Similarly, filtered ProK-digested PDH-supplemented growth of the wild-type bacteria to the same extent (Fig. 4D). However, the ΔlplA1 mutant was unable to grow in the presence of the 500 Da retentate, unless complemented by a plasmid containing the lplA1 gene. As a control for the accuracy of the MWCO of the dialysis membranes, we confirmed that retentate of a free lipoic acid solution after dialysis against a 100 Da MWCO was able to supplement growth of wild-type bacteria. These results demonstrate that LplA1 permits L. monocytogenes usage of degraded host lipoyl proteins as a sole source of lipoic acid.

LplA1 is required for utilization of the synthetic lipoyl tripeptide DKLA

The predicted lipoyl tripeptide released from porcine PDH and KGDH after ProK digestion shares amino acid identity with human, murine and rat lipoyl domains (Fig. 5A). To establish that LplA1-mediated growth of L. monocytogenes was dependent upon lipoyl peptide and not other non-modified peptides, we used synthetic lipoyl DKA (DKL.A), the smallest lipoyl peptide predicted from ProK digestion of porcine PDH (Fig. 5A, Fig. S2A and B). At 5 μg l−1, DKL.A supported growth of wild type but not ΔlplA1, demonstrating the specific contribution of LplA1 in utilizing lipoylated peptides for proliferation (Fig. 5B). To test whether replication required the lipoyl moiety, we also supplemented IMM with non-lipoylated tripeptide (DKA), which did not support L. monocytogenes growth in the absence of any lipoyl substrate even at very high concentrations. To determine the optimal concentration of lipoyl peptide for LplA1-mediated multiplication, we performed a dose–response curve by adding DKL.A to lipoate-starved bacterial cultures (Fig. 5C). Lower concentrations of lipoyl tripeptide only supported replication of wild-type bacteria, while high concentrations supported growth of both the wild-type and ΔlplA1 mutant strains. These results imply that LplA2 can also use lipoyl peptides for growth, but only at very high concentrations. Our data suggest that the concentration of available lipoyl peptide in the host cytosol is low, resulting in dependence on LplA1 for intracellular replication of L. monocytogenes.

Fig. 5. LplA1 is required for optimal growth on small lipoyl peptides.

Fig. 5

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A. The amino acid sequences of the dihydrolipoyl transacylase lipoyl domain from Homo sapiens (Accession No. AAA64512), dihydrolipoamide S-acetyltransferase lipoyl domain of Bos taurus (Accession No. XP_588501), dihydrolipoamide branched chain transacylase E2 lipoyl domain of Mus musculus (Accession No. NP_034152), dihydrolipoamide S-acetyltransferase lipoyl domain of Rattus norvegicus (Accession No. AAI07441) and the dihydrolipoamide acetyltransferase lipoyl domain of Sus scrofa (Accession No. NP_999159) aligned.

B. Wild-type (WT) L. monocytogenes and the ΔlplA1 mutant strain were grown in IMM containing 5 μg l−1 DKL.A, or 5 μg l−1 non-lipoylated DKA. The OD600 was measured over time in a Bioscreen Growth Curve Analyzer and plotted as a function of time.

C. Wild-type L. monocytogenes and the ΔlplA1 mutant strain were grown as in (B), but in IMM containing different concentrations of DKL.A. The OD600 was measured after bacteria had reached stationary phase (25 h) and plotted against lipoyl peptide concentration.

D. Bacterial growth curves were performed as in (B), but 0.5 μg l−1 tripeptide (lipoylated or non-lipoylated) was added with or without prior aminopeptidase M digestion as indicated.

E. Bacterial growth curves were performed as described in (B), but in IMM containing the concentrations of lipoamide indicated. After 30 h of growth, bacteria had reached stationary phase, and OD600 values were plotted against lipoamide concentration. The mean value ± SD was calculated for each time point (n = 3) in (B–E).

To determine if LplA1 could support growth with smaller lipoyl peptides than lipoylated tripeptides, we digested lipoylated DKL.A with the protease aminopeptidase M (Fig. S2C). Unlike the preferential growth of wild-type bacteria mediated by undigested lipoyl DKL.A, peptide digestion products allowed growth of both the wild-type and ΔlplA1 strains, suggesting that lipoyl dipeptide or lipoyl lysine does not dictate LplA1-dependent bacterial growth (Fig. 5D). Thus, LplA1 allows bacterial proliferation in low concentrations of lipoyl peptides, and the length of the modified peptide may determine preferential usage by LplA1 over LplA2. We also investigated if the free amide derivation of lipoate, lipoamide, supported LplA1-dependent replication of L. monocytogenes. Although there was a slight difference in growth by the ΔlplA1 mutant strain as compared with wild-type bacteria at suboptimal lipoamide concentrations, both wild type and ΔlplA1 replicated in lipoamide-containing medium, demonstrating that lipoamide did not define the host-specific requirement for LplA1 (Fig. 5E). Overall, these data indicate that LplA1 facilitates growth of L. monocytogenes on lipoyl peptides, the predominant intracellular form of lipoate.

L. monocytogenes LplA1 exhibits lipoate ligase activity

Our results suggest a model in which intracellular growth of L. monocytogenes using small host-derived lipoyl peptides is dependent upon LplA1. If LplA1 is able to directly use lipoyl substrates, we predicted that the lplA1 gene would complement an E. coli strain deficient in lipoate utilization, TM131, which cannot grow in minimal medium without a functional lipoate ligase (Morris et al., 1995). As E. coli LplA has been shown biochemically to have lipoate ligase enzymatic activity, this complementation strategy has been used to demonstrate functionality of lipoate ligases from organisms as diverse as the protozoan parasite Plasmodium falciparum and Arabidopsis thaliana (Wada et al., 2001; Allary et al., 2007). We grew TM131 in metabolic bypass medium (+succinate +acetate) to allow transformation with an IPTG-inducible plasmid expressing E. coli lplA, L. monocytogenes lplA1 or the vector alone. All transformants grew on bypass medium (Fig. 6A), but only E. coli lplA and L. monocytogenes lplA1 complemented growth of TM131 when acetate and succinate were removed (Fig. 6B and C). However, TM131 expressing lplA1 grew to a lesser extent, indicating that the L. monocytogenes lipoate ligase only partially compensated for the loss of endogenous E. coli lipoate ligase activity. We were not able to confirm expression of LplA2 in the TM131 strain transformed with an IPTG-inducible lplA2 plasmid although the plasmid could direct LplA2 expression in a wild-type E. coli strain, thus it is still unknown whether LplA2 can act directly as a functional lipoate ligase (Fig. S3A and data not shown). As expected, expression of both L. monocytogenes lplA1 and E. coli lplA resulted in lipoylation of the E2 subunit of PDH (Fig. S3B). Thus, complementation of the E. coli TM131 mutant by L. monocytogenes lplA1 suggests that LplA1 can act enzymatically as a lipoate ligase.

Fig. 6.

Fig. 6

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Complementation of an E. coli strain deficient in lipoate utilization by L. monocytogenes (L.m.) LplA1. E.coli TM131 (lplA−lipA−) transformed with an empty IPTG-inducible vector, or the same vector expressing E. coli LplA or L. monocytogenes LplA1. TM131 is deficient in lipoate biosynthesis as well as endogenous LplA; growth requires either exogenous expression of a lipoate ligase, or supplementation with acetate and succinate. Clones expressing either empty vector, E.coli LplA or L. monocytogenes LplA1 were streaked on M9 minimal medium plates containing IPTG and free lipoic acid. Acetate and succinate were included in (A), but not (B) and (C). The boxes illustrated in (B) are magnified in (C) for viewing of single colonies. Some putative E. coli revertants were observed (large white colony observed in the insert of the LplA1-expressing E.coli strain).

To explore if there might be a structural basis for the difference in lipoate ligase activity between L. monocytogenes LplA1 and LplA2, we modelled the structure of L. monocytogenes LplA1 and LplA2 using previously published crystal structures of the E. coli and Streptococcus pneumoniae LplA proteins (Fujiwara et al., 2005). Every residue in the lipoyl AMP-binding pocket predicted to interact with substrate or residues identified as being important for binding of the target apo-domain was conserved between the E. coli and L. monocytogenes enzymes (Fig. S1) (Kim et al., 2005). However, the electrostatic surface topology was notably different between the three enzymes, with an overall electrostatic potential for LplA1 of −13, while LplA2 and E. coli LplA exhibit electrostatic potentials of −5 and −8 respectively (Fig. 7). Moreover, LplA1 exhibited several regions of clustered negatively charged residues, which might modulate interaction of LplA1 with other proteins or cofactors. These predicted structural differences are consistent with our observations that L. monocytogenes LplA1 and LplA2 have overlapping but distinct functions.

Fig. 7.

Fig. 7

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Structural Modelling of L. monocytogenes (L.m.) LplA1 and LplA2. Electrostatic surface potentials for the crystallographic structure E. coli (E.c.) LplA (PDB ID 1X2H) and the modelled structures of LplA1 and LplA2 were calculated using APBS (Baker et al., 2001) and mapped onto their respective solvent accessible surfaces using Pymol (DeLano, 2002). Negative potentials (−10 kT e−1) are shown in red, positive potentials (10 kT e−1) in blue. The views for individual molecules are separated by a 90° rotation about the x-axis. The protein structures are shown at the same magnification and orientation for each view.

Discussion

Intracellular pathogens such as L. monocytogenes have evolved mechanisms to take advantage of the biochemical environment of the host cell; the study of these mechanisms can reveal critical parameters of the host–pathogen interaction. The essential nutrient lipoic acid is scarce in the host cell in its free form, but our data demonstrate lipoylated polypeptides are present in low abundance in the cytosol where L. monocytogenes replicates. Although L. monocytogenes has two lipoate ligases, LplA1 and LplA2, only LplA1 was required for intracellular growth and virulence. LplA2 was sufficient for lipoylation of target proteins and growth in rich broth medium, which contains free lipoate, but was insufficient during intracellular infection by L. monocytogenes. We showed that LplA1 was necessary for L. monocytogenes growth in low concentrations of lipoyl peptide, yet was dispensable for growth in medium containing free lipoic acid or growth in host cells supplemented with free lipoic acid. These data suggest that optimal replication in the intracellular environment by L. monocytogenes requires LplA1-dependent utilization of host-derived lipoyl peptides.

Local concentrations of host lipoyl peptide vary widely depending primarily on the concentration of mitochondria in that tissue (Baker et al., 1998). As LplA1 and LplA2 both support growth with high concentrations of lipoyl peptide, tissues that contain high amounts of lipoylated proteins, such as the liver, may be able to support more extensive bacterial growth (Baker et al., 1998). The ΔlplA1 strain was substantially less able to compete with the wild-type strain in spleen when compared with liver; this tissue preference could be due to a higher concentration of lipoyl peptide in the liver supporting more robust replication in the absence of LplA1. However, the spleen also contains more immune effector cells that use oxidative stress as a host defence, which might also contribute to decreased fitness of the ΔlplA1 mutant. Lipoic acid is a potent antioxidant that may help bacteria survive oxidative stress; by allowing efficient utilization of host lipoyl peptides, LplA1 may promote L. monocytogenes survival in the spleen (Bryk et al., 2002). Scavenging lipoate is a known growth requirement for several pathogens, such as the auxotrophic bacteria Enterococcus faecalis and L. monocytogenes (Reed et al., 1951). Even pathogenic organisms that synthesize lipoate, such as the protozoan parasites Toxoplasma gondii and Plasmodium falciparum, still scavenge lipoate from the host environment, as the parasite biosynthetic pathways supply only the apicoplast and not the mitochondrion (Wrenger and Muller, 2004; Crawford et al., 2006; Allary et al., 2007). Thus, the use of host-derived lipoate may be more widespread among intracellular pathogens than previously appreciated. The ability to utilize nutrients from diverse host pools is a common theme in pathogenesis, as other essential compounds like iron are scavenged in multiple forms to enable optimal replication (Skaar et al., 2004).

Our structural models of LplA1 and LplA2 suggest that these enzymes share with E. coli LplA the basic catalytic mechanism by which free lipoate is activated and transferred to target apoproteins (Morris et al., 1994; 1995). The amino acid residues involved in the binding of the lipoyl group and the target domain appear to be conserved in both E. coli LplA and the L. monocytogenes lipoate ligases (Fig. S1). However, the enzymatic mechanism by which LplA1 can enable use of low concentrations of lipoyl peptide is still unknown. Utilization of lipoyl peptide by LplA1 would mandate a distinct enzymatic mechanism from ligation of free lipoic acid as synthesis of the activated lipoyl AMP intermediate requires the carboxyl group of lipoate, lacking in lipoyl peptides. The simplest model would predict that, in addition to its LplA-like activity, LplA1 could bind directly to lipoyl peptides and transfer the lipoyl group from the peptide to the target domain, by a mechanism more analogous to LipB-mediated transfer of octanoyl groups from an acyl-carrier protein to the target apoprotein (Zhao et al., 2003; Ma et al., 2006). Although LplA contains no cysteines, which would be required for the lysine/cysteine catalytic dyad characterized in LipB, a cysteine could conceivably be provided by a hybrid interface with an interacting protein (Ma et al., 2006). In this model, LplA1 would bind to lipoyl peptides with higher affinity than LplA2, allowing LplA1 to function efficiently at lower substrate concentrations. As an alternative model, the clustered negatively charged regions on the surface of LplA1 could represent sites of interaction with a critical cofactor. For example, LplA1 may have intrinsic lipoamidase activity or associate with a protein with lipoamidase activity, thus liberating lipoamide from lipoyl peptides and increasing the local concentration of free lipoamide. Such lipoamidase activity would have to be strictly regulated, as it would oppose ligase function (Jiang and Cronan, 2005). Future structure–function studies will elucidate which model more accurately describes how LplA1 contributes to L. monocytogenes pathogenesis.

Experimental procedures

Bacterial culture

Strains used in this study are described in Table 1. For intra-cellular growth curves, all L. monocytogenes strains were grown to mid-log (OD600 0.2–0.6) in IMM at 37°C unless otherwise specified. IMM was prepared as previously described (Phan-Thanh and Gormon, 1997), using the concentrations listed in Table 1 of the referenced paper. Specifically, lipoic acid was used to supplement IMM at 5 μg l−1 except when otherwise specified in the figure legends, or substituted where indicated with various concentrations of lipoamide (0–5 μg l−1), DKL.A (0–25 μg l−1), digested lipoyl PDH (5 mg l−1) or digested lipoyl KGDH (5 mg l−1). For growth curves in IMM, single colonies from freshly streaked BHI plates were inoculated into IMM without lipoic acid (IMM−L) for 10–14 h at 37°C shaking to OD600 0.2–0.4, back-diluted to an OD600 of 0.02 in IMM and cultured at 37°C shaking. Bacterial growth was determined by measuring changes in OD600 over time. All IMM growth curves, except the experiment shown in Fig. 4D, were performed in a Bioscreen Growth Curve Analyzer (Growth Curves, USA).

Table 1.

Strains used.

Strains Genotype/description Reference
10403S L. monocytogenes serotype 1/2a, parental strain used as wild type Freitag et al. (1993)
MOR125 10403S lplA1::Tn917ΔlplA2 This study
MOR129 10403S ΔlplA2 This study
DP-L3903 L. monocytogenes 10403S::Tn917, unknown site of insertion, wild-type phenotype Auerbuch et al. (2001)
DP-L4263 10403S ΔlplA1 O'Riordan et al. (2003)
MOR142 10403S ΔlplA1 (DP-L4263) containing the Tn917 insertion transduced from DP-L3903 This study
JK1 E. coli rpsL Morris et al. (1995)
TM131 E. coli rpsL lipA182::Tn1000dKn lplA148::Tn10dTc Morris et al. (1995)
MOR226 E. coli rpsL lipA182::Tn1000dKn lplA148::Tn10dTc + ptac85 This study
MOR229 E. coli rpsL lipA182::Tn1000dKn lplA148::Tn10dTc + ptac85-ECLplA This study
MOR227 E. coli rpsL lipA182::Tn1000dKn lplA148::Tn10dTc + ptac85-LMLplA1 This study
MOR228 E. coli rpsL lipA182::Tn1000dKn lplA148::Tn10dTc + ptac85-LMLplA2 This study
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Allelic exchange

A strain containing an in-frame deletion of lplA2 was generated using homologous recombination. The deletion allele was obtained by amplifying 3′ and 5′ genomic sequences flanking lplA2 that were then fused by splice overlap extension-PCR (Horton et al., 1990). The 0.8 kb fragments were amplified from the 10403S bacterial genome with Platinum Pfx polymerase (Invitrogen) using primer sequences described in Table S1. The PCR product was subcloned into the allelic exchange vector, pKSV7 (Smith and Youngman, 1992). Allelic exchange was performed as previously described and confirmed by sequencing (O'Riordan et al., 2003). The in-frame deletion resulted in removal of amino acids 30–300 out of 330 amino acids (strain MOR129). The transposon insertion that disrupted lplA1 in DP-L2214 (O'Riordan et al., 2003) was transduced as previously described (Hodgson, 2000; O'Riordan et al., 2003) into MOR129 to generate a strain deficient in both putative lipoyl ligases (lplA1Tn::917ΔlplA2; MOR125). For complementation studies, pAM401 and pAM401lplA1 were transformed individually into DP-L4263 and 10403S as previously described (O'Riordan et al., 2003).

Mouse infections

Monotypic and CI infections were performed as previously described with the following modifications (Auerbuch et al., 2001). Exponential phase bacterial cultures were diluted in Ca2+- and Mg2+-free Dulbecco's phosphate-buffered saline, and 2 × 105 bacteria in 200 μl were injected i.p. into 4- to 6-week-old C57BL/6 J mice (Jackson Laboratories, Bar Harbor, ME). At 72 h post infection, the animals were sacrificed, and livers and spleens homogenized in 0.1% NP-40, serially diluted, and plated onto Luria–Bertani (LB) agar. For the CI infections, the homogenization buffer also contained 0.1 μg ml−1 erythromycin (erm), and the lysate was plated on LB agar with or without 1 μg ml−1 erm. The CI was calculated by dividing the number of wild-type cfu (erm sensitive) by the number of mutant cfu (erm resistant).

Quantitative real-time polymerase chain reaction

After infection of J774 macrophages at a multiplicity of infection (moi) of 1:1 for 6 h, host cells were lysed with 50 mM Tris, pH 8, 150 mM NaCl and 0.05% NP-40 for 10 min on ice. After vortexing and pelleting the host nuclei at 3000 g for 30 s, the remaining supernatant was used to isolate bacterial RNA with the Pro-Blue Fast RNA kit (MP Biomedicals) followed by DNase digestion on Qiagen RNeasy minicolumns. cDNA was generated from 2.5 μg of RNA using random primers and M-MLV reverse transcriptase (Invitrogen). Primers for lplA1, lplA2 and rpoA (Table S1) were used to amplify cDNA, which was quantified by SYBR Green fluorescence (Stratagene Mx3000p).

Lipoyl substrates

Lipoamide (T5875), PDH (P-7032) and KGDH (K1502) were purchased from Sigma-Aldrich. Prior to using commercial PDH and KGDH, the enzymes were precipitated and washed with Bio-Rad ReadyPrep 2-D Cleanup Kit. Trypsin digestion was performed at 30°C for 4 h in 0.02% Tween-20 and 200 mM ammonium bicarbonate (pH 8.9). The proteins were digested with a ratio of 1 mg of protein to 8 mg of proteomics grade trypsin in a total volume of 500 μl; the reaction was stopped with 0.0003% trifluoroacetic acid. ProK digestion was performed at a ratio of 5 mg of protein to 20 μg of enzyme (Sigma P-4850) in 20 mM Tris-HCl, 1 mM CaCl2 at pH 7.4 at 25°C for 4 h and heat inactivated for 10 min at 90°C. Enzymatic digestion was confirmed by SDS-PAGE and immunoblotting analysis for lipoylated peptides. For dialysis experiments, ProK-digested samples were loaded in 100 or 500 Da Spectra/Por Micro DispoDialyzer membranes and dialysed against 20 mM Tris-HCl, 1 mM CaCl2 at pH 7.4. Synthetic lipoyl tripeptide DKL.A was synthesized by Anaspec using a previously published protocol (Fig. S2A and B) (Konishi et al., 1996). Digestion of DKL.A with immobilized aminopeptidase M was performed at a ratio of 0.5 U enzyme to 250 μg of peptide for 18 h at 37°C (Pierce; #20238); digestion was confirmed by LC-MS on a nanoAcuity/Qtof premier instrument. The non-digested and digested samples were injected on a C18 column and analysed in positive ion mode (Fig. S2C). Non-lipoylated DKA was synthesized and LC-MS was performed by the Protein Structure Facility of the University of Michigan Medical School.

Subcellular fractionation

Mitochondria were isolated by lysis of 1 × 107 macrophages in 10 mM HEPES pH 7.9, 1.5 mM MgCl2, 10 mM KCl, 0.5 mM DTT and 0.2 mM PMSF, incubation on ice for 10 min and vortexing for 1 min. After lysis, salts were added to a final concentration of 30 mM HEPES (pH 7.9), 140 mM KCl and 3 mM MgCl2. After removing nuclei by centrifuging at 3000 g for 30 s and washing three times, mitochondria were spun at 80 000 g for 4 h at 4°C. The remaining supernatant was retained and concentrated in a 3 kDa Ultracel YM-3 Centricon Centrifugal Filter Device (Amicon).

Protein analysis

Bacteria were prepared and analysed by immunoblot with an anti-lipoic acid antibody (Calbiochem) as previously described (O'Riordan et al., 2003) except washed exponential phase cultures (OD600 0.5) were used after growth in IMM−L or BHI at 37°C. Protein lysate was prepared with FastProtein Blue Lysing Matrix (MP Biomedicals) by processing in a FastPrep machine for 40 s at a setting of 6.0 in Lysing Matrix B. For cytosolic and mitochondrial peptide analysis by immunoblot, 0.75 M Tris-HCl was used in the resolving gel, running buffer and loading buffer.

Bacterial infections

J774 macrophages were infected as previously described (O'Riordan et al., 2003) except washed exponential phase cultures grown at 37°C in IMM−L pre-starved bacteria were used at a moi of 3, resulting in ∼10% macrophage infection. In DHLA (Sigma-Aldrich T-8260) supplementation experiments, 50 μM DHLA was added to the J774 macrophage growth medium prior to infection and maintained throughout the infection. Addition of high concentrations of non-reduced lipoic acid resulted in precipitation, as well as non-specific host cell effects. Data points represent the mean with standard deviation.

E. coli complementation

TM131 (rpsL lipA182::Tn1000dKn lplA148::Tn10dTc) and its parental wild-type strain JK1 (rpsL) were transformed with E. coli lplA, L. monocytogenes lplA1 or lplA2 in the IPTG-inducible ptac85 plasmid and plated onto LB agar containing 0.4% glucose, 5 mM sodium acetate, 5 mM sodium succinate and 100 μg ml−1 ampicillin. Primers used to amplify the lipoate ligase genes with Pfx platinum polymerase are listed in Table S1. TM131-transformed strains were also grown in the presence of 50 μg ml−1 kanamycin and 125 μg ml−1 tetracycline. Transformants were restreaked onto M9 minimal medium plates containing 0.4% glucose, 1 mM IPTG, thiamine (10 μg ml−1), 2 mM MgSO4, 0.1 mM CaCl2 and the appropriate antibiotics with or without 5 mM sodium acetate and 5 mM sodium succinate as indicated. Lysates for Western blotting were grown in LB containing 0.4% glucose, 5 mM sodium acetate, 5 mM sodium succinate, 1 mM IPTG and the appropriate antibiotics.

Structural modelling of LplA1 and LplA2

Because of its high homology with LplA1 (65.8%) and LplA2 (66.6%), the structure of LplA from S. pneumoniae (PDB ID: 1VQZ; Joint Center for Structural Genomics) was used as the foundation for modelling the LplA1 and LplA2 structures. To model the bound forms of LplA1 and LplA2, the N- and C-terminal domains of 1VQZ were first aligned onto the respective domains of the lipoate-bound form of E. coli LplA (PDB ID: 1X2H) using the graphics program O, then the amino acid sequence of 1VQZ was mutated into that of LplA1 (Jones et al., 1991; Fujiwara et al., 2005). Amino acid insertions and deletions were fit using the lego-loop option in O. The resulting LplA1 model was then placed into a box of waters containing a minimum of two shells of water, minimized and put through simulated annealing using torsion angle dynamics in Crystal-lography & NMR System (CNS; Brunger et al., 1998). The lipoate structure from 1X2H was attached to the NZ atom of a modelled lysine residue using O. CNS parameter and topology files for the lipoate-lysine residue were created via the HIC-Up server (http://xray.bmc.uu.se/cgi-bin/gerard/hicup_server.pl). The tripeptide (DKL.A) was then created via CNS and modelled into the refined LplA1 model using O. Simulated annealing was then performed on the DKL.A-bound LplA1 model. To create the LplA2 model, the amino acid sequence of LplA2 was overlaid onto the bound form of LplA1. The resulting DKL.A-bound LplA2 model was placed into a box of waters and put through simulated annealing.

Supplementary Material

supplemental

Acknowledgements

We gratefully acknowledge members of the O'Riordan laboratory, especially Maggie Evans, and Dr K. Carroll for many helpful discussions. We also thank Dr M. Swanson and Dr V. Carruthers for critical review of the manuscript. This work was funded by NIH R01 AI064540 (M.X.D.O.) and by the University of Michigan Center for Structural Biology (J.A.S.). K.M.K. was a trainee of the Molecular Mechanisms of Pathogenesis Training Grant (T32AI007528).

Footnotes

Supplementary material

This material is available as part of the online article from: http://www.blackwell-synergy.com/doi/abs/10.1111/j.1365-2958.2007.05956.x (This link will take you to the article abstract).

Please note: Blackwell Publishing is not responsible for the content or functionality of any supplementary materials supplied by the authors. Any queries (other than missing material) should be directed to the corresponding author for the article.

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