Library selection and analysis of sequencing data
In order to assess the fitness of insertion mutants en masse, a library of ∼ 50 000 transposon mutants was generated on oxic Shewanella basal medium (SBM) (Covington et al., 2010) agar plates with 40 mM lactate and 80 mM fumarate, and all subsequent growth experiments were performed in this medium unless otherwise noted. Following transposition, the entire mutant library was pooled, aliquoted and frozen. To perform a selection experiment, a library aliquot was thawed and genomic DNA was extracted from a portion of the culture. The remaining culture was back diluted to a final optical density (OD600) of 0.005 and grown shaking at 30°C either with oxygen or anaerobically with fumarate. When culture OD600 reached 0.2, genomic DNA was extracted resulting in populations under selection for ∼ 5–6 generations. We deliberately chose an OD600 of 0.2 because under both oxic and anoxic conditions cultures still exhibit exponential growth and are not limited for carbon source, electron donor, or electron acceptor (Fig. S1). One of the primary advantages of Tn-seq is the semi-quantitative detection of subtle changes in fitness that are commonly overlooked by other methods. Accordingly, we limited the number of doublings during selection to only 5–6 with the hope of maintaining the ability to differentiate subtle and strong phenotypes.
Tn-seq relies on the sequencing of transposon–genome junctions en masse to determine the relative abundance of each transposon mutant in the population, before and after selection. MR-1 genomic DNA from selection experiments was prepared as described in Experimental procedures (Opijnen and Camilli, 2010) and subjected to Illumina sequencing. Genomic DNA flanking each transposon was mapped to the MR-1 genome to determine the site of transposition, insertion sites were parsed to individual genes, and the number of reads found within a given gene were normalized and tabulated. Our data set derives from a library of 26 793 unique insertion mutants and each coding region has on average five insertions within the central 89% portion of the gene. Fitness values were calculated by dividing the number of reads mapped to each gene at the end of a selection by the number mapped to the same gene in the starting pool, followed by a logarithmic transformation. The entire selection experiment was independently performed twice and values derived from sequencing, as well as fitness calculations, are reported for each experiment individually (Table S1). Fitness values are only calculated for genes with at least three independent insertion sites to reduce variability that can result in misleading fitness calculations. Genes with a fitness value of ≤ −0.5 in both experiments for a given condition are considered to cause a fitness defect when inactivated. To validate our results we compared Tn-seq fitness data (Table S1) with studies of genes known to be important for aerobic or anaerobic growth of MR-1 using lactate and fumarate, limiting our comparison to studies in which actual mutants were analysed using growth assays (Table 1). In every case Tn-seq fitness data correctly measured a fitness defect under the conditions previously reported (see Table 1 for specific genes and references).
Table 1. Previously characterized genes important for growth of MR-1 with lactate and fumarate.
|Locus tag||Gene name||Tn-seq fitness values||Reference|
|Replicate 1||Replicate 2|
|Previously reported to have a fitness defect only when grown aerobically on lactate|
|SO_0424|| aceE ||−1.96||0.06||−2.59||−0.33||Flynn et al. (2012)|
|SO_1518|| lldG ||−0.81||−0.28||−1.10||−0.40||Pinchuk et al. (2009)|
|SO_1519|| lldF ||−0.57||0.29||−0.76||−0.27||Pinchuk et al. (2009)|
|Previously reported to have a fitness defect only when grown anaerobically on lactate and fumarate|
|SO_0624|| crp ||−0.42||ND||−0.55||−1.35||Saffarini et al. (2003)|
|SO_1329|| cyaC ||−0.13||−0.99||−0.12||−0.41||Charania et al. (2009)|
|SO_0970|| fccA ||0.08||−4.34||0.11||−1.65||Myers and Myers (1997)|
|SO_2916|| pta ||0.34||−4.64||0.52||−2.12||Hunt et al. (2010)|
|SO_2915|| ackA ||0.19||−5.15||0.05||−3.75||Hunt et al. (2010)|
|SO_2912|| pflB ||−0.01||−2.87||−0.01||−2.48||Flynn et al. (2012)|
|SO_1910|| menA ||0.39||ND||0.04||ND||Myers and Myers (1993)a |
|SO_4713|| menF ||0.08||−3.06||−0.02||−1.70||Myers and Myers (1993)a |
|SO_4739|| menB ||0.12||−3.23||0.43||−2.11||Myers and Myers (1993)a |
|SO_4573|| menD ||0.10||−2.98||0.35||−2.84||Myers and Myers (1993)a |
|SO_4574|| menH ||0.02||−1.15||0.01||−0.56||Myers and Myers (1993)a |
|SO_4575|| menC ||0.15||−3.05||0.16||−2.57||Myers and Myers (1993)a |
|SO_4576|| menE ||0.16||−3.05||0.29||−1.76||Myers and Myers (1993)a |
|SO_0259|| ccmE ||−0.23||ND||−0.39||−2.78||Bouhenni et al. (2005)a |
|SO_0261|| ccmC ||−0.49||−4.29||−0.45||ND||Bouhenni et al. (2005)|
|SO_0263|| ccmA ||−0.47||−3.63||−0.26||ND||Bouhenni et al. (2005)a |
|SO_0266|| ccmF-1||−0.51||−3.70||−1.05||ND||Bouhenni et al. (2005)|
|SO_0268|| ccmH ||−0.44||−1.01||0.32||−0.10||Bouhenni et al. (2005)a |
|Previously reported to have a fitness defect when grown aerobically or anaerobically on lactate and fumarate|
|SO_0274|| ppc ||−2.88||−1.38||−2.95||−1.24||Flynn et al. (2012)|
In the current study we use Tn-seq analysis to gain new insights into the central metabolic pathways of MR-1. Here we report fitness values for 3030 of the 4467 genes encoded by the MR-1 genome and megaplasmid. Figure 1 shows a map of the central carbon metabolism of MR-1, with genes colour coded based on fitness data provided by Tn-seq analysis. The effect of inactivation of most genes was consistent with recent metabolic models for MR-1 (Tang et al., 2007b; Pinchuk et al., 2010; Flynn et al., 2012). Inactivation of a few genes, however, resulted in an unexpected fitness defect, or lack thereof, and we chose these genes to study further.
Figure 1. Graphic representation of fitness values for the genes of central carbon metabolism in MR-1 determined by Tn-seq. Solid arrows denote single reactions while dotted arrows denote multiple reactions. The inactivation of a gene is determined to have a fitness cost for a condition if the average fitness value falls below −0.5 (Table S1) and given the corresponding colour (see inset). Genes which did not contain insertions in the parent library (†) or did not meet the filtering requirements outlined in Experimental procedures (*) are indicated. A white box denotes genes for which a fitness value was not determined.
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Lactate metabolism and regulation of lactate utilization
MR-1 is thought to form syntrophic partnerships with fermentative microbes, oxidizing fermentation end-products (e.g. lactate, propionate, formate, hydrogen, and some amino acids) for anaerobic respiration (Nealson and Scott, 2006). Lactate is of particular interest as it is used as a carbon and energy source for the vast majority of studies involving Shewanella sp. grown in a minimal medium, and Fig. 1 shows a model for lactate metabolism in MR-1. Following uptake, lactate is oxidized to pyruvate by membrane associated flavin adenine dinucleotide (FAD)-dependent lactate dehydrogenases specific for the l- or d-isomer (LldEFG or Dld respectively) (Pinchuk et al., 2009). MR-1 grows somewhat faster on l-lactate than d-lactate in the presence of oxygen (Pinchuk et al., 2009), and accordingly, insertions in lldF and lldG resulted in a mild fitness defect in oxic SBM (Table 1 and Fig. 1). Interestingly, insertions in dld had a fitness defect under both oxic and anoxic conditions. These data are consistent with d-lactate inhibition of l-lactate utilization in MR-1, a hypothesis confirmed by phenotypic analysis of deletion mutants and lactate dehydrogenase activity assays (E.D. Brutinel and J.A. Gralnick, manuscript in preparation).
Subsequently, the fate of pyruvate is dependent on the availability of oxygen. In the presence of oxygen pyruvate is oxidized to acetyl coenzyme A (acetyl-CoA), which then enters the TCA cycle, by the pyruvate dehydrogenase complex (AceEF/LpdA) (Scott and Nealson, 1994; Tang et al., 2007a). Under anoxic conditions pyruvate is converted to acetate and formate in a stepwise fashion by the action of pyruvate–formate lyase (PflAB), phosphate acetyltransferase (Pta), and acetate kinase (AckA) (Scott and Nealson, 1994; Tang et al., 2007c; Hunt et al., 2010). In agreement with the above model, transposon insertions in aceE result in a fitness defect in the presence of oxygen while insertions in pflA, pflB, pta, and ackA result in a fitness defect under anoxic conditions (Table 1 and Fig. 1). Insertions in aceF had a fitness defect under both conditions (Table S1), possibly due to polar effects on the expression of the downstream lpdA gene that we infer to be essential based on the absence of insertion mutants in lpdA in our Tn-seq data set.
Growth on lactate necessitates gluconeogenesis to generate essential precursor metabolites. Accordingly, we did not detect transposon insertions in genes required for gluconeogenesis with the exception of pgi (encoding phosphoglucose isomerase) and gapA1 [encoding glyceraldehyde-3-phosphate dehydrogenase (GAPDH)]. Transposon insertions in pgi do not appear to be lethal but result in a reduced fitness under both oxic and anoxic conditions (Fig. 1 and Table S1). Transposon insertions in the gapA1 gene (SO_0538) appeared to have no effect on fitness under either condition, in disagreement with a model predicting that gapA1 is essential when MR-1 is grown on lactate (Flynn et al., 2012). Genomic sequence analysis previously predicted two paralogues of gapA1 in the genome of MR-1, gapA2 (SO_2345) and gapA3 (SO_2347) (Serres and Riley, 2006), whose protein products share 43% and 34% identity respectively with that of gapA1. While transposon insertions in gapA2 did not appear to effect fitness, insertions in gapA3 had a pronounced effect on fitness under both oxic and anoxic conditions (Fig. 1 and Table S1). This result was unexpected and we wanted to determine if each of the predicted gapA paralogues encode an enzyme with GAPDH activity. To this end we performed NAD- and NADP-dependent GAPDH activity assays on cell extracts derived from MR-1 strains expressing gapA1, gapA2, or gapA3 from a plasmid and the results are reported in Table 2. GAPDH activity in cells expressing gapA1 was not significantly different from a strain carrying a vector control, consistent with the lack of a phenotype for insertions in gapA1, and suggesting that gapA1 was either not expressed or inactive under the conditions tested. We detected a significant increase in NAD- or NADP-dependent GAPDH activity in cells expressing gapA2 or gapA3 respectively. It appears that, when grown in minimal medium supplemented with lactate and fumarate, MR-1 primarily generates glyceraldehydes-3-phosphate in an NADP-dependent reaction catalysed by gapA3. This result highlights the utility of Tn-seq in evaluating the contribution of predicted isoenzymes to an essential biochemical reaction under a given condition. The roles played by gapA1 and gapA2 in the physiology of MR-1 remain unclear, though we speculate that these paralogues must be important under conditions not tested in this manuscript.
Table 2. Glyceraldehyde-3-phosphate dehydrogenase activity in cell lysates from MR-1 expressing gapA1, gapA2 or gapA3 from a plasmid.
| ||OD340 min−1 mg−1 |
|MCS2||3.36 ± 0.10||0.31 ± 0.02|
| gapA1 ||3.79 ± 0.75||0.24 ± 0.09|
| gapA2 ||10.52 ± 0.34||0.57 ± 0.09|
| gapA3 ||4.93 ± 0.21||2.00 ± 0.20|
MR-1 uses the non-oxidative branch of the pentose phosphate pathway for anaerobic metabolism
The pentose phosphate pathway is required for the generation of three key metabolites (ribulose-5-phospphate, erythrose-4-phosphate, and sedoheptulose-7-phosphate) and has an oxidative and a non-oxidative branch. Transposon insertions in a number of genes in the pentose phosphate pathway did not affect the fitness of MR-1 under oxic or anoxic conditions (Fig. 1). Two of the dispensable genes were in the oxidative branch (zwf and gnd), essential for the conversion of glucose 6-phoshate to ribulose-5-phosphate coupled to the generation of NAD(P)H (Fig. 1). In contrast, the genes of the non-oxidative branch are critical for growth of MR-1 on lactate, with the notable exception of the gene coding for transaldolase (tal). Transposon insertions in the gene encoding ribulose-5-phosphate 3-epimerase (rpe) drastically reduced fitness under aerobic and anaerobic conditions, and transposon insertions were not detected in the remaining genes involved in the non-oxidative branch of the pentose phosphate pathway (rpiA and tkt), consistent with the essential role of generating key metabolic intermediates. Under the conditions tested, MR-1 is able to generate ribulose-5-phosphate, erythrose-4-phosphate, and sedoheptulose-7-phosphate via the non-oxidative branch of the pentose phosphate pathway with the reactions catalysed by Tkt, RpiA, and Rpe. This conclusion is supported by the fact that the growth rate of a strain lacking gnd is identical to that of MR-1 grown in oxic or anoxic SBM (Flynn et al., 2012). Our data demonstrate that the pentose phosphate pathway of MR-1 functions in a similar manner as that of Escherichia coli, not with genomics and modelling based predictions, but with phenotypic data.
Branching of the MR-1 anaerobic TCA cycle is dynamic and utilizes an additional citrate synthase
MR-1 preferentially uses oxygen as a terminal electron acceptor and, in the presence of oxygen, acetyl-CoA enters the TCA cycle and is completely oxidized to CO2 generating both energy and reducing equivalents (Tang et al., 2007a). Accordingly, transposon insertions that disrupt TCA cycle genes should result in a decreased growth rate in the presence of oxygen, and fitness values measured by Tn-seq are in agreement with this assertion (Fig. 1). Transposon insertions in the genes encoding aconitate hydratase (SO_0432; acnB) and the α-ketoglutarate dehydrogenase complex (SO_0930, SO_0931; sucAB) were not present in the parent library suggesting disruption of these genes is extremely deleterious in the presence of oxygen, consistent with published E. coli and S. oneidensis mutants lacking aconitate hydratase or α-ketoglutarate dehydrogenase activity (Creaghan and Guest, 1978; Gruer et al., 1997; Pinchuk et al., 2011). An acnB deletion strain was unable to grow in the presence of oxygen and required glutamate for growth in SBM without casamino acids under anoxic conditions demonstrating that the acnB gene product, a predicted bifunctional aconitate hydratase II/2-methylisocitrate dehydratase, is the predominant source of aconitate hydratase activity under either condition tested. Significant fitness defects are not observed for insertions in either of the genes annotated to encoding an isocitrate dehydrogenase (icd or SO1538) suggesting that both genes are expressed and functional under oxic and anoxic conditions. Our data are consistent with a complete suite of TCA cycle reactions in MR-1 grown under aerobic conditions.
Under anaerobic conditions many organisms do not complete the TCA cycle, instead operating with an oxidative and reductive branch, primarily for anaplerotic reactions to replenish α-ketoglutarate, oxaloacetate, and succinyl-CoA pools. Numerous reports have suggested that this is the case for MR-1 using transcriptomics, metabolic labelling, enzymatic assays, and computer modelling to infer the expression/functionality of key enzymes in the TCA cycle in the absence of oxygen (Scott and Nealson, 1994; Beliaev et al., 2002; Tang et al., 2007c,c; Pinchuk et al., 2010). The effect of an insertion mutation in each gene of the anaerobic TCA cycle was measured by Tn-seq and a number of unexpected phenotypes were observed.
Citrate synthase (GltA) catalyses the first reaction of the oxidative arm of a branched anaerobic TCA cycle required to generate α-ketoglutarate for biosynthesis of glutamate. Accordingly, E. coli strains lacking a functional gltA gene require exogenous glutamate for growth in minimal medium (Lakshmi and Helling, 1976). Our parent library contained transposon insertions in the gene that encodes citrate synthase (SO_1926; gltA), despite being grown in SBM, and under anoxic conditions the fitness of cells with transposon insertions in gltA was the same as MR-1 (Table S1 and Fig. 1). The growth rate of a gltA deletion strain was substantially slower than wild type in the presence of oxygen (Fig. 2A), and indistinguishable from wild type under anoxic conditions, even in the absence of the 0.01% casamino acids normally present in SBM (Fig. 2A). The following two possibilities could explain the observed phenotype of the gltA mutant: (i) MR-1 uses an alternative pathway to synthesize α-ketoglutarate/glutamate, or (ii) another protein provides sufficient citrate synthase activity under anaerobic conditions. The former possibility seems unlikely because inactivation of the genes encoding the α-ketoglutarate dependent glutamate synthase (SO_1325, SO_1324; gltBD) has a drastic effect on fitness under oxic and anoxic conditions (Table S1 and Fig. 1). A candidate for the latter possibility is 2-methylcitrate synthase (encoded by prpC; SO_0344), which, while predominantly catalysing the reaction of oxaloacetate and propionyl-CoA to form 2-methylcitrate, has been shown to possess citrate synthase activity in E. coli and S. enterica (Gerike et al., 1998; Horswill and Escalante-Semerena, 1999). To evaluate the contribution of 2-methylcitrate synthase to the lactate metabolism of MR-1 we generated a prpC deletion strain as well as a gltA prpC double mutant. While growth of the prpC mutant on lactate was indistinguishable from wild type, the gltA prpC double mutant required exogenous glutamate for growth in SBM without casamino acids (Fig. 2B). The glutamate requirement of the gltA prpC double mutant was abated by expression of either gltA or prpC from a plasmid. Citrate synthase activity assays were performed on cell extracts derived from MR-1 strains grown in the presence and absence of oxygen and the results are reported in Table 3. While citrate synthase activity in the gltA mutant grown under oxic or anoxic conditions was significantly lower than MR-1, the activity measured was well above that of the double mutant, demonstrating a significant contribution by 2-methylcitrate synthase to the formation of citrate in MR-1.
Figure 2. The gltA and prpC gene products contribute to the production of citrate in MR-1.
A. Growth curves of MR-1, the gltA mutant, and the prpC mutant were performed in SBM under oxic or anoxic conditions in the presence and absence of casamino acids. The growth curve of the prpC mutant did not differ significantly from that of MR-1 and is represented by the same line for clarity.
B. Growth curves of the gltA prpC double mutant in SBM without casamino acids under oxic or anoxic conditions in the presence and absence of glutamate. The reported values are the averages of three independent experiments and error bars represent the standard error of the mean (SEM).
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Table 3. Citrate synthase acticity in cell lysates from cultures grown under oxic and anoxic conditions.
| ||OD412 min−1 mg−1 |
|MR-1||4.92 ± 0.13||1.76 ± 0.34|
|ΔgltA ||0.34 ± 0.02||0.19 ± 0.05|
|ΔprpC ||4.76 ± 0.15||1.60 ± 0.23|
|ΔgltA ΔprpC ||0.03 ± 0.01||0.06 ± 0.04|
In the canonical branched anaerobic TCA cycle α-ketoglutarate is derived from citrate and the oxidative branch while oxaloacetate and succinyl-CoA are derived from phosphoenolpyruvate (PEP) and the reductive branch. It has been suggested that MR-1 uses the above pathways based on transcriptional data and enzyme activity measured in cell extracts (Scott and Nealson, 1994; Beliaev et al., 2002); however, definitive genetic analysis has not been performed. Our Tn-seq experiments suggest that under anoxic conditions inactivation of the genes encoding the succinyl-CoA synthase (sucCD) and fumarate reductase (frdABCD) complexes did not affect the fitness of MR-1 (Table S1 and Fig. 1), a surprising result considering the requirement of succinyl-CoA for biosynthesis. E. coli strains lacking succinyl-CoA synthase activity are prototrophic in the presence of oxygen but require exogenous lysine and methionine for anaerobic growth on glucose (Herbert and Guest, 1968; Mat-Jan et al., 1989). In SBM without casamino acids the growth rate of a strain derived from MR-1 with a sucCD deletion was measurably slower than MR-1 in the presence of oxygen, and indistinguishable from MR-1 under anoxic conditions (Fig. 3A). These results indicate that under anoxic conditions succinyl-CoA is not solely derived from succinate and instead derives either partially or entirely from α-ketoglutarate and the oxidative branch in MR-1. Unfortunately, our Tn-seq experiments did not detect any insertions in the sucAB genes (encoding α-ketoglutarate dehydrogenase). In agreement, E. coil strains lacking α-ketoglutarate dehydrogenase activity are unable to grow in the presence of oxygen (our library was generated on oxic SBM agar), but exhibit prototrophic growth in the absence of oxygen (Herbert and Guest, 1969; Creaghan and Guest, 1978). A strain derived from MR-1 with a sucB deletion was unable to grow in the presence of oxygen, even in rich medium, but exhibited the same growth rate as MR-1 in SBM without casamino acids under anoxic conditions (Fig. 3B).
Figure 3. The MR-1 anaerobic TCA cycle can produce succinyl-CoA via succinyl-CoA synthase (SucCD) or α-ketoglutarate dehydrogenase (SucAB). Growth assays under aerobic and anaerobic conditions comparing MR-1 to (A) the sucCD mutant or (B) the sucB mutant were performed in SBM without casamino acids. The reported values are the averages of three independent experiments and error bars represent the standard error of the mean (SEM).
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