Lipogenesis and Redox Balance in Nitrogen-Fixing Pea Bacteroids

Jason J. Terpolilli,a,b Shyam K. Masakapalli,c Ramakrishnan Karunakaran,b Isabel U. C. Webb,b,c Rob Green,b Nicholas J. Watmough,d

Nicholas J. Kruger,c R. George Ratcliffe,c Philip S. Pooleb,c

Centre for Rhizobium Studies, Murdoch University, Perth, Australiaa; Department of Molecular Microbiology, John Innes Centre, Norwich, United Kingdomb; Departmentof Plant Sciences, University of Oxford, Oxford, United Kingdomc; Centre for Molecular Structure and Biochemistry, University of East Anglia, Norwich, United Kingdomd

ABSTRACT

Within legume root nodules, rhizobia differentiate into bacteroids that oxidize host-derived dicarboxylic acids, which is as-sumed to occur via the tricarboxylic acid (TCA) cycle to generate NAD(P)H for reduction of N2. Metabolic flux analysis of labo-ratory-grown Rhizobium leguminosarum showed that the flux from [13C]succinate was consistent with respiration of an obligateaerobe growing on a TCA cycle intermediate as the sole carbon source. However, the instability of fragile pea bacteroids pre-vented their steady-state labeling under N2-fixing conditions. Therefore, comparative metabolomic profiling was used to com-pare free-living R. leguminosarum with pea bacteroids. While the TCA cycle was shown to be essential for maximal rates of N2

fixation, levels of pyruvate (5.5-fold reduced), acetyl coenzyme A (acetyl-CoA; 50-fold reduced), free coenzyme A (33-fold re-duced), and citrate (4.5-fold reduced) were much lower in bacteroids. Instead of completely oxidizing acetyl-CoA, pea bacteroidschannel it into both lipid and the lipid-like polymer poly-�-hydroxybutyrate (PHB), the latter via a type III PHB synthase that isactive only in bacteroids. Lipogenesis may be a fundamental requirement of the redox poise of electron donation to N2 in all le-gume nodules. Direct reduction by NAD(P)H of the likely electron donors for nitrogenase, such as ferredoxin, is inconsistentwith their redox potentials. Instead, bacteroids must balance the production of NAD(P)H from oxidation of acetyl-CoA in theTCA cycle with its storage in PHB and lipids.

IMPORTANCE

Biological nitrogen fixation by symbiotic bacteria (rhizobia) in legume root nodules is an energy-expensive process. Within le-gume root nodules, rhizobia differentiate into bacteroids that oxidize host-derived dicarboxylic acids, which is assumed to occurvia the TCA cycle to generate NAD(P)H for reduction of N2. However, direct reduction of the likely electron donors for nitroge-nase, such as ferredoxin, is inconsistent with their redox potentials. Instead, bacteroids must balance oxidation of plant-deriveddicarboxylates in the TCA cycle with lipid synthesis. Pea bacteroids channel acetyl-CoA into both lipid and the lipid-like poly-mer poly-�-hydroxybutyrate, the latter via a type II PHB synthase. Lipogenesis is likely to be a fundamental requirement of theredox poise of electron donation to N2 in all legume nodules.

Biological reduction (or fixation) of atmospheric nitrogen (N2)to ammonia (NH3) provides up to 50% of the biosphere’s

available nitrogen, mostly through symbioses between soil bacte-ria (rhizobia) and legumes (1, 2). These symbioses are initiated byrhizobia infecting legume roots, resulting in the formation of nod-ules. Rhizobia differentiate into N2-fixing bacteroids that expressnitrogenase to reduce N2 to NH3 under microaerobic conditions(3). Bacteroids receive carbon from the legume while secretingNH3 to the plant. The overall stoichiometry of N2 fixation underideal conditions is as follows:

N2 � 8e� � 8H� � 16ATP → 2NH3 � H2 � 16ADP � 16Pi

(1)

Thus, 8 moles of electrons and protons and 16 moles of ATPreduce a single mole of N2, making N2 fixation energetically ex-pensive.

Legumes energize bacteroid N2 fixation by supplying dicar-boxylates, principally malate (4), which must be oxidized to yieldATP and electrons to reduce N2. Bacteroids metabolize malate bythe use of NAD�-dependent malic enzyme (5–7) and pyruvatedehydrogenase to provide acetyl coenzyme A (acetyl-CoA), whichcan be completely oxidized in the tricarboxylic acid (TCA) cycle,yielding FADH2 and NAD(P)H. The standard model is thatNAD(P)H supplies electrons both to nitrogenase via ferredoxin,

or via an equivalent low-potential electron donor, and to an elec-tron transport chain for ATP synthesis (8, 9).

This model is supported by work in Rhizobium leguminosarumand Sinorhizobium meliloti, where TCA cycle mutants are unableto fix N2 in symbiosis with pea (Pisum sativum) and alfalfa (Medi-cago sativa), respectively (10–13). However, the TCA cycle pro-vides both reductant and biosynthetic precursors, so the abolitionof N2 fixation in these mutants could be due to insufficientNAD(P)H to directly power nitrogenase or, equally, could resultfrom biosynthetic deficiencies. In contrast, in soybean (Glycinemax) bacteroids, the TCA cycle either is dispensable for N2 fixa-tion or can be bypassed, with isocitrate dehydrogenase and 2-oxo-

Received 6 June 2016 Accepted 26 July 2016

Accepted manuscript posted online 8 August 2016

Citation Terpolilli JJ, Masakapalli SK, Karunakaran R, Webb IUC, Green R,Watmough NJ, Kruger NJ, Ratcliffe RG, Poole PS. 2016. Lipogenesis and redoxbalance in nitrogen-fixing pea bacteroids. J Bacteriol 198:2864 –2875.doi:10.1128/JB.00451-16.

Editor: I. B. Zhulin, University of Tennessee

Address correspondence to Philip S. Poole, philip.poole@plants.ox.ac.uk.

Supplemental material for this article may be found at http://dx.doi.org/10.1128/JB.00451-16.

Copyright © 2016, American Society for Microbiology. All Rights Reserved.

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glutarate dehydrogenase mutants of Bradyrhizobium japonicumable to fix N2 at wild-type rates (14, 15). Moreover, standard mid-point potentials indicate that NAD(P)H is unlikely to donate elec-trons directly to ferredoxin (the E0= for NAD�/NADH is �320mV, for NADP�/NADPH is �324 mV, and for ferredoxin [Fe3�/Fe2�] is �484 mV) (16, 17). Thus, some other, as-yet-undefinedmechanism must exist to transfer electrons to nitrogenase in rootnodule bacteroids.

Finally, N2-fixing bacteroids in nodules formed by soybeanand common bean (Phaseolus vulgaris) accumulate large quanti-ties of the lipid-like polymer poly-�-hydroxybutyrate (PHB),while bacteroids from pea, alfalfa, and clover (Trifolium spp.) ap-parently do not (18). While abolishing PHB synthesis does notadversely affect N2 fixation rates in soybean and common bean(19–21), in Azorhizobium caulinodans, mutation of PHB synthaseprevents N2 fixation in both free-living and symbiotic forms (22),implying a fundamental role for PHB synthesis in at least someN2-fixing rhizobia.

Determining how N2 is fixed by bacteroids, in what is arguablythe second most important nutrient assimilation cycle after pho-tosynthesis, requires an understanding of bacteroid carbon me-

tabolism. Metabolic profiling and flux analysis as well as muta-tional and N2 fixation studies were used to investigate carbon flowin bacteroids. Remarkably, the results revealed that the TCA cycleis not the only sink for plant-derived carbon in symbiotic N2 fix-ation; rather, pea bacteroids divert appreciable quantities ofacetyl-CoA into the production of lipid or PHB. N2-fixing bacte-roids are therefore inherently lipogenic, and this is probably ametabolic requirement for N2 fixation.

MATERIALS AND METHODSBacterial strains and culture conditions. Bacterial strains and plasmidsused in this study are detailed in Table 1. Rhizobium leguminosarum bv.viciae (Rlv3841) was grown at 28°C on tryptone yeast extract (TY) (28) oracid minimal salts (AMS) medium (29) with succinate (20 mM) andNH4Cl (10 mM) as the sole carbon source and nitrogen source, respec-tively. Where appropriate, antibiotics were used at the following concen-trations: streptomycin, 500 �g/ml; neomycin, 80 �g/ml; spectinomycin,50 �g/ml; gentamicin, 20 �g/ml; and ampicillin, 50 �g/ml.

Metabolic flux analysis. Rlv3841 cells grown in succinate-NH4ClAMS medium were harvested at mid-log phase (optical density at 600 nm[OD600] of approximately 0.5) and subcultured into fresh AMS media toa reach a starting OD600 of 0.02, with 20 mM [13C4]succinate (20% frac-

TABLE 1 Strains, plasmids, and primers used in this study

Strain, plasmid, or primer Genotype and/or description or sequencea

Reference orsource

StrainsR. leguminosarum Rlv3841 Str derivative of R. leguminosarum bv. viciae strain 300 23R. leguminosarum RU137 Rlv3841 phaC1::Tn5 Nmr 12R. leguminosarum RU116 Rlv3841 sucD::Tn5 Nmr 12R. leguminosarum RU156 Rlv3841 sucA::Tn5 Nmr 12R. leguminosarum RU724 Rlv3841 sucA::Tn5-lacZ Nmr 12R. leguminosarum RU725 Rlv3841 sucC::Tn5-lacZ Nmr 12R. leguminosarum RU726 Rlv3841 sucB::Tn5-lacZ Nmr 12R. leguminosarum RU733 Rlv3841 sucA::Tn5-lacZ Nmr 12R. leguminosarumLMB814

Rlv3841 phaC2::� Str Spr This work

R. leguminosarumLMB816

Rlv3841 phaC1::Tn5 phaC2::� Str Nmr Spr This work

E. coli DH5� Strain used for cloning; F� �80lacZM15 (lacZYA-argF)U169 recA1 endA1hsdR17(rK

� mK�) phoA supE44 thi-1 gyrA96 relA1

Invitrogen

PlasmidspJET1.2/Blunt PCR product cloning vector; Apr Thermo-FisherpHP45-�SmSp pHP derivative with �SmSp cassette; Smr Spr 24pJQ200SK pACYC derivative, P15A origin of replication insertional mutagenesis inactivation

vector; Gmr Sucs

25

pRK2013 Helper plasmid used for mobilizing plasmids; ColE1 replicon with RK2 tra genes; Kmr 26pLMB834 pr1645-1646 PCR product (2.8 kbp) from pRL100105 (phaC2) cloned into pJET1.2/

Blunt; Apr

This work

pLMB835 pLMB834 with �SmSp cassette from pHP45-�SmSp cloned into unique EcoRI site;Apr Smr Spr

This work

pLMB838 pJQ200SK with BglII fragment from pLMB835 containing phaC2::� cloned intoBamH1 site; Smr Spr Gmr Sucs

This work

Primerspr1645 AACGCTACAGCGCAACGCTC This workpr1646 ACTTTCTTCGCTCCCGTCGG This workpr1647 ACCCCGAAGACGCTCGTCAT This workpr1648 ATGATCGTGACGGCATCGGC This workpotfarforward GACCTTTTGAATGACCTTTA 27Tn5-1 ATAGCCTCTCCACCCAAGC This work

a Ap, ampicillin; Gm, gentamicin; Km, kanamycin; Nm, neomycin; Sp, spectinomycin; St, streptomycin; Suc, succinate.

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tional abundance). Cells were harvested at an OD600 of 0.3 and centri-fuged at 8,500 g for 5 min. The resulting pellet was washed with freshAMS medium and centrifuged, and the resulting cell pellet was extractedin 80% (vol/vol) ethanol at 80°C for 5 min prior to centrifugation at12,000 g for 5 min. The supernatant containing the soluble amino acids,organic acids, and sugars was dried by vacuum centrifugation. The insol-uble pellet was rapidly frozen in liquid N2 and freeze-dried. Protein in theinsoluble fraction was hydrolyzed to its component amino acids by incu-bation with 6 M HCl for 24 h at 100°C.

Gas chromatography-mass spectrometry (GC-MS) analysis of deriva-tized amino acids, organic acids, and sugars was performed on an Agilent7890A GC/5975C quadrupole MS system as described elsewhere (30).Amino acids and organic acids were analyzed after derivatization usingN-tert-butyldimethylsilyl-N-methyltrifluoroacetamide (MTBSTFA) orN-methyl-N-(trimethylsilyl)-trifluroacetamide (MSTFA); sugars weretreated with methoxyamine hydrochloride and then derivatized withMSTFA. Protein-derived and soluble amino acids were examined sepa-rately. Mass isotopomer abundances were quantified using Chemstationand corrected for the presence of naturally occurring heavy isotopes in-troduced during derivatization. The chemical fragments used for meta-bolic flux analysis are detailed in Table S1 in the supplemental material.

Metabolic modeling was performed with 13C-FLUX (version20050329) using an iterative procedure described before (30, 31). A com-plete description of the model, which also defines the network carbonatom transitions, is provided in Table S2 in the supplemental material,and net flux data are provided in Table S3 in the supplemental material.During initial parameter fitting, fluxes corresponding to biomass outputswere allowed to vary, and the mean values from the 10 best-fit estimateswere then used to constrain the network output flux values in subsequentsimulations. Malate and oxaloacetate were combined into a single metab-olite pool, as were phosphoenolpyruvate and pyruvate, to improve thedeterminability of fluxes between these intermediates. No adjustmentswere required to compensate for the contribution of preexisting unlabeledpools of metabolites. Molar fluxes are reported relative to a succinateuptake flux value of 1.

Material for metabolite profiling. To prepare samples of free-livingRlv3841, six independent cultures of Rlv3841, derived from six isolatedcolonies of the strain, were grown in AMS medium on a gyratory shaker at250 rpm to an OD600 of 0.4. Cell pellets were collected by centrifugation(5,000 g, 5 min), washed with isolation buffer (8 mM K2HPO4, 2 mMKH2PO4, 2 mM MgCl2), and stored at �80°C for later use in metaboliteprofiling.

To prepare bacteroid and nodule cytosolic samples, seeds of P. sativumcv. Avola were surfaced sterilized with 70% (vol/vol) ethanol for 30 s,rinsed once in sterile water, and then immersed in a 2% (wt/vol) NaOClsolution for 2 min, prior to 10 rinses in sterile water. Seeds were sown into2-liter beakers containing washed and autoclaved fine-grade vermiculite.Six independent cultures of test strain Rlv3841 or test strain RU116, de-rived from six isolated colonies of each strain, were prepared. A 1-mlaliquot of each culture was inoculated into a minimum of six pots, at celldensities of between 5 107 and 9 107 cells ml�1. Seeds were initiallysown in duplicate and thinned to one plant per pot after 7 days. Plantswere watered once with 250 ml nitrogen-free nutrient solution as previ-ously described (29) and were incubated in an illuminated environment-controlled growth room at 22°C on a 16-h day, 8-h night cycle.

Plants were harvested at 28 days postinoculation (dpi) for metabolo-mic profiling. Approximately 1.5 g of nodule tissue was excised fromplants from each set of pots. Nodules were ground in isolation buffer (8mM K2HPO4, 2 mM KH2PO4, 2 mM MgCl2), and the homogenate waspassed through muslin and centrifuged (250 g for 5 min) to removeplant debris before a further round of centrifugation (5,000 g, 10 min)was performed to pellet the bacteroids. The resulting supernatant, repre-senting the nodule cytosol fraction, was freeze-dried; the pellet, represent-ing the bacteroid fraction, was washed twice with isolation buffer and

centrifuged (5,000 g, 10 min); and the pellets were frozen at �80°C forlater use in metabolite profiling.

Metabolite profiling platform. Determinations of metabolomic pro-files of free-living, bacteroid, and nodule cytosol samples were each per-formed using nonbiased, global metabolome profiling technology basedon GC-MS and ultra-high-performance liquid chromatography/massspectrometry/tandem mass spectrometry (UHLC/MS/MS2) platforms(32, 33) developed by Metabolon (Durham, NC). Six replicate samplesfrom each treatment (free-living, bacteroid, and nodule cytosol samples)were extracted using an automated MicroLab Star system (Hamilton).Recovery standards were added prior to the first step in the extractionprocess for quality control purposes. To monitor total process variability,a series of technical replicates were taken from a pool made from smallaliquots of all of the experimental samples. These were spaced evenlyamong the randomly ordered experimental samples, and all consistentlydetected metabolites were monitored for reproducibility. Sample prepa-ration was conducted using methanol extraction to remove the proteinfraction while allowing maximum recovery of small molecules. The re-sulting extract was divided into two fractions, one for analysis by LC andone for analysis by GC. Samples were placed briefly on a TurboVap evap-orator (Zymark) to remove the organic solvent. Each sample was frozenand dried under a vacuum. Samples were then prepared for the appropri-ate instrument (either LC/MS or GC/MS).

The LC/MS portion of the platform was based on a Waters AcquityUHPLC and a Thermo-Finnigan LTQ mass spectrometer, which con-sisted of an electrospray ionization source and a linear ion trap massanalyzer. The sample extract was split into two aliquots, dried, and thenreconstituted in acidic or basic LC-compatible solvents, each of whichcontained 11 or more injection standards at fixed concentrations. Onealiquot was analyzed under acidic positive-ion optimized conditions andthe other under basic negative-ion optimized conditions in two indepen-dent injections using separate dedicated columns. Extracts reconstitutedin acidic conditions were subjected to gradient elution using water andmethanol, with both containing 0.1% (vol/vol) formic acid, while thebasic extracts, which also used water and methanol, contained 6.5 mMNH4HCO3. The MS analysis alternated between MS and data-dependentMS/MS scans using dynamic exclusion.

The samples destined for GC/MS analysis were redried using vacuumdesiccation for a minimum of 24 h prior to being derivatized using driedN2 and bistrimethyl-silyl-trifluoroacetamide (BSTFA). The GC columnwas 5% phenyl, and the temperature ramp was 40°C to 300°C over a16-min period. Samples were analyzed on a Thermo-Finnigan Trace DSQfast-scanning single-quadrupole gas chromatograph mass spectrometerusing electron impact ionization.

Compound identification, data handling, and statistical analysis.For metabolite profiling, identification of known chemical entities wasbased on comparisons to metabolomic library entries of purified stan-dards as previously described (33, 34). Statistical analysis was performedusing the software packages Array Studio (Omicsoft) and R (http://www.r-project.org/). Where a given metabolite was not detected in a particularsample, the observed minimum detected value for that metabolite fromthe analysis was assigned, under the assumption that the missing valueswere not random but resulted from the amount of the compound beingbelow the limit of detection. Data for free-living and bacteroid sampleswere then normalized to protein content, as determined by Bradford assay(35). For the comparison of the bacteroids to the nodule cytosol, normal-ization was performed by extracting proportional amounts of bacteroidand cytosolic fractions of matched starting samples. That is, the total yieldof bacteroid and cytosolic fractions for each sample was known, and aconstant percentage of each fraction was analyzed in order to compare therelative amounts of metabolites in each fraction. The statistical modelutilized the matched-pair nature of the samples to account for absolutedifferences between the samples. Welch’s two-sample t test was used toidentify metabolites that differed significantly between experimentalgroups (P � 0.05), and the false-discovery rate (FDR) was also calculated

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(36) to account for the multiple comparisons that normally occur inmetabolomic-based studies (Q � 0.1). Thus, metabolites were consideredto be significantly different if they met the statistical significance criteria(P � 0.05 and Q � 0.10).

Assessment of N2 fixation. Plants intended for assessment of N2 fix-ation were grown as described above in “Material for metabolite profil-ing,” with the following exceptions. For measurement of N2 fixation byacetylene reduction assay, plants were grown in 1-liter pots and harvestedat the onset of flowering (21 dpi). Whole plants were removed fromgrowth pots and transferred to 250-ml sealed bottles. After the rates ofacetylene reduction of detached nodules were measured, nodules wereexcised and immediately transferred into a 25-ml bottle and assayed.Rates of N2 fixation were determined by the amount of acetylene reducedafter 1 h in an atmosphere consisting of 95% air–5% acetylene, as previ-ously described (37). Following the acetylene reduction assay, bacteroidprotein was quantified by excising nodules from roots and grinding in 40mM HEPES (pH 7.0). The homogenate was passed through muslin andthe eluate centrifuged (250 g for 5 min) to remove plant debris. Thesupernatant was then centrifuged (5,000 g, 10 min) to pellet the bacte-roids. Bacteroids were lysed by two rounds of ribolysing on a FastPrepFP120 Ribolyser (BIO101/Savant) at a setting of 6.5 for 30 s, with sampleskept on ice for 5 min between rounds. The protein content in the resultingsupernatant was determined by Bradford assay (35) using a Pierce Co-omassie assay kit (Pierce; catalog no. 23200) with bovine serum albumin(BSA) as the protein standard.

For assessment of N2 fixation by plant biomass accumulation, plantswere grown in 2-liter pots and were supplied with 200 ml of additionalsterile water at 28 dpi. Plants were then harvested at 47 dpi by cuttingshoots below the hypocotyl and drying at 60°C for 48 h prior to weighing.

Lipid analysis. Bacteroids were collected from nodules harvestedfrom plants grown for analysis of lipids as described in “Material formetabolite profiling” and harvested at 28 dpi. Nodules were ground in 20mM HEPES buffer (pH 7.0) and purified by Percoll gradient (38). Cells offree-living Rlv3841 were grown in AMS medium with succinate andNH4Cl and harvested at an OD600 of 0.4 to 0.6 by centrifugation (5,000 g for 10 min). The resultant bacteroid and cell pellets were stored at �80°Cfor later use. Bacteroid and cell pellets were lysed by the use of a Ribolyseras described above and centrifuged (10,000 g for 10 min). The super-natant was then centrifuged (20,000 g for 20 min) prior to furtherultracentrifugation (60,000 g for 60 min) to remove cell membranes.The supernatant was concentrated by vacuum centrifugation prior tolipid quantification performed using a triglyceride determination kit (Sig-ma; catalog no. TR0100). Protein determination was performed using theBradford assay as described above.

Mutant construction and phenotyping. To construct the phaC2(pRL100105) mutant of Rlv3841, primers pr1645 and pr1646 (see TableS1 in the supplemental material) were used to amplify 2.8 kb of the regioncontaining the gene and the PCR product was cloned into pJET1.2/blunt,giving plasmid pLMB834. The �-streptomycin/spectinomycin cassettefrom pHP45-�SmSp was cloned into the unique EcoRI site of pLMB834to produce pLMB835. The BglII fragment from pLMB835 was cloned intopJQ200SK to produce pLMB839. Plasmid pLMB839 was then conjugatedinto strain Rlv3841, using pRK2013 as a helper plasmid, to produce phaC2mutants as previously described (5), resulting in LMB814. The mutationwas confirmed by PCR mapping using primer pairs pr1648-potfarforwardand pr1657-potfarforward. Strain LMB816, the phaC1 (RL2098) phaC2(pRL100105) double mutant, was made by using the general transducingphage RL38 to lyse strain RU137. The kanamycin-marked phaC1::Tn5mutation was then back-transduced into LMB814 to generate LMB816, aspreviously described (39), and the mutation was confirmed by PCR map-ping with pr1647-potfarforward, pr1648-potfarforward, and pr1647-Tn5-1 primer pairs. Assessment of N2 fixation of the resulting mutantswas performed as described above. Transmission electron microscopywas performed on nodules harvested from plants at 28 dpi, and the meth-

ods for nodule sectioning, staining, and microscopy were performed asdetailed previously (20).

RESULTSMetabolic flux analysis of free-living rhizobia. Dicarboxylatesare provided to bacteroids by plants to support N2 fixation (3, 4),so the pathways operating in free-living Rhizobium legumino-sarum bv. viciae 3841 (Rlv3841) growing on [13C4]succinate werequantified. The major flux of succinate metabolism in Rlv3841was via fumarate to malate (Fig. 1) and subsequently from malateto pyruvate and oxaloacetate to phosphoenolpyruvate. Thesefluxes would support the major metabolic requirements of cellsgrowing on a TCA cycle intermediate for synthesis of acetyl-CoAto supply the TCA cycle and phosphoenolpyruvate for biosynthe-sis of sugars. Large fluxes were also detected in gluconeogenesisconverting phosphoenolpyruvate to triose phosphates, in the ox-idative decarboxylation of pyruvate to acetyl-CoA, and in the TCAcycle from oxaloacetate to 2-oxoglutarate. Overall, these fluxes areconsistent with respiration of an obligate aerobe growing on aTCA cycle intermediate as the sole carbon source.

Currently, metabolic flux analysis cannot be conducted on no-

FIG 1 Flux map of central carbon metabolism for free-living R. legumino-sarum Rlv3841, grown on succinate and NH4Cl. Net fluxes are expressed on amolar basis relative to succinate uptake. The thickness of each arrow is pro-portional to the net flux with the exception that fluxes representing �1% ofsuccinate uptake are indicated by broken arrows. Biosynthetic outputs areshown in solid boxes, and metabolites treated as a single pool in the model areshown in dashed boxes. Flux identifiers, defined in Table S2 in the supplemen-tal material, are shown in italics. The precise values for the deduced fluxes arepresented in Table S4 in the supplemental material. Standard abbreviations areused for amino acids and metabolic intermediates, and PPP data represent thereversible nonoxidative steps of the pentose phosphate pathway.

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toriously fragile isolated pea bacteroids (40). Nitrogenase activity,as measured by acetylene reduction, in isolated pea nodules col-lapsed 90 min after excision to less than 2% of that in nodules onroots (0.25 � 0.03 versus 18.3 � 2.5 nmol acetylene reduced · mgnodule�1 · h�1). This precludes labeling of nodule metabolites atthe isotopic steady state under physiologically relevant conditionsin an isolated system. Moreover, the likely low rate of proteinturnover in nondividing bacteroids compromises the use of thelabeling patterns of protein-derived amino acids to reflect those oftheir metabolic precursors. We therefore used metabolite profil-ing to examine the differences in levels of metabolic intermediatesbetween cultured cells and bacteroids.

Bacteroid central metabolism. The metabolic profiles of free-living and bacteroid forms of Rlv3841 were analyzed using non-biased, untargeted metabolome analysis (32, 33). The metaboliteswhose levels were most highly elevated in bacteroids relative tofree-living Rlv3841 were homoserine and asparagine (105- and58-fold increases, respectively; Fig. 2). Levels of both were alsohigh in the nodule cytosolic fraction relative to bacteroids (33-and 11-fold increases; Table S4 in the supplemental material), inaccordance with previous observations (41, 42) and consistentwith their known plant origin. Asparagine is made in the plantcytosol as the primary nitrogen export product from nodules (40).Furthermore, free asparagine is not made by Rlv3841, which, fromanalysis of its genome, uses the GatCAB pathway to insert aspar-

agine into proteins by charging asparaginyl-tRNA with aspartateand then transamidating aspartate to asparagine (43). In addition,catabolism of asparagine and homoserine is not upregulated inbacteroids (44), nor do catabolic mutants show reduced N2 fixa-tion rates (45, 46), consistent with minor roles in symbiosis.

Our fundamental issue of investigation is whether the TCAcycle is altered during symbiotic N2 fixation. The dicarboxylatesmalate, fumarate, and succinate are the carbon sources for bacte-roids in planta, and levels of all three were increased in bacteroidsrelative to free-living cells (Fig. 2). Moreover, levels of thesemetabolites were also much higher in the plant nodule cytosolfraction relative to bacteroids (malate 14-, fumarate 20-, andsuccinate 2.5-fold increased [see Table S4 in the supplementalmaterial]), consistent with active plant dicarboxylate synthesis.

Metabolism of dicarboxylates by bacteroids is performedvia malic enzyme and phosphoenolpyruvate carboxykinase topyruvate and phosphoenolpyruvate, respectively, with pyru-vate subsequently oxidatively decarboxylated to acetyl-CoA(5–7). The levels of intermediates of sugar metabolism such as3-phosphoglycerate, fructose-6-phosphate, and glucose-6-phos-phate and the pentose phosphate pathway (ribulose-5-phosphateand xylulose-5-phosphate) were greatly reduced (Fig. 2), suggest-ing that little sugar synthesis occurs in bacteroids. Remarkably,levels of pyruvate (5.5-fold reduced), acetyl-CoA (50-fold re-duced), free coenzyme A (33-fold reduced), and citrate (4.5-fold

FIG 2 Metabolite profile of Rlv3841 bacteroids versus Rlv3841 free-living cells showing fold change in metabolite abundance relative to Rlv3841 bacteroids.Bacteroids were isolated from nodules from pea plants 28 days postinoculation (dpi). Cells were harvested from log-phase cultures grown in AMS broth with 20mM succinate and 10 mM NH4Cl as the carbon and nitrogen sources, respectively. Bolded intermediates were detected by metabolite profiling, with a statisticallysignificant fold difference (P � 0.05 by Welch’s t test and Q � 0.1 for the false-discovery rate) denoted with a red (increase) or green (decrease) arrow. A signindicates that the metabolite was undetectable in either the free-living or the bacteroid sample, so the difference reported is therefore a lower-limit estimate of thefold change. Intact arrows indicate single-step enzyme-catalyzed reactions. Broken arrows indicate instances in which two or more enzyme-catalyzed steps areinvolved in a series of reactions. Abbreviations: UD, undetectable; BT, bacteroid; FL, free-living; GABA, �-amino butyric acid; GSH, glutathione (reduced);GSSG, glutathione (oxidized); 2OG, 2-oxoglutarate; OAA, oxaloacetate; PEP, phosphoenolpyruvate.

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reduced) were much lower in bacteroids (Fig. 2). In sharp con-trast, the transcription and enzymatic activity of citrate synthase(RL2234; icdB) were increased 3.2- and 12-fold, respectively, andincreases in the activity and transcription of other enzymes of thedecarboxylating arm of the TCA cycle have been noted (44, 47).While such increased enzyme biosynthesis might indicate increasedflux into the TCA cycle, it is equally consistent with lower feedbackinhibition of the synthesis and activity of enzymes displayed by keyintermediates such as acetyl-CoA and citrate (48, 49).

Carbon in the TCA cycle could also be channeled to glutamate,which is synthesized from 2-oxoglutarate by glutamine synthe-tase/glutamate synthase (GS/GOGAT) (50). However, glutamatelevels were 20-fold lower in bacteroids relative to free-living cells(Fig. 2), consistent with GS/GOGAT activity being both low andnot essential in mature bacteroids (51). Levels of metabolites derivedfrom glutamate, including glutathione and N-acetylglutamate, werealso reduced, while levels of many other amino acids were either al-tered only slightly or unchanged in bacteroids (Fig. 2).

However, steady-state metabolite levels do not represent flux.Low levels of pyruvate, acetyl-CoA, coenzyme A, and citrate inbacteroids may indicate a low rate of synthesis but can equallyresult from rapid turnover. Furthermore, metabolites may dra-matically change concentrations during isolation of bacteroidsfrom nodules. We addressed this by comparing wild-type bacte-roids with mutant bacteroids defective in the TCA cycle, whichshould lead to different metabolite profiles. If low acetyl-CoA lev-els in wild-type bacteroids relative to free-living cells result fromincreased flux through the TCA cycle, then TCA cycle mutantsshould have elevated acetyl-CoA levels.

Metabolite profile of a TCA cycle mutant. We previously iso-lated several Tn5 insertions in Rlv3841 genes encoding TCA cycleenzymes (12). Malate dehydrogenase, succinyl-CoA synthetase,and the E1 and E2 components of the 2-oxoglutarate dehydroge-nase complex are transcribed from the mdh-sucCDAB operon(52). Mutations in sucA (RU156, RU724, and RU733) and sucB(RU726), encoding the E1 and E2 components of the 2-oxogluta-rate dehydrogenase complex, respectively, abolished 2-oxogluta-rate dehydrogenase activity (12), resulting in plants that failed toreduce acetylene (Fix�). Therefore, blocking the TCA cycle inRlv3841 prevents N2 fixation. However, we now show that strainRU116, mutated in sucD (encoding the �-subunit of succinyl-CoA synthetase) and originally scored as Fix� based on yellowingof plants and small nodules but retaining low levels of succinyl-CoA synthetase activity (12), is able to reduce acetylene at 35% ofthe wild-type rate (Fig. 3). This mutation may affect the number ofbacteroids in nodules or total nodule mass or reduce nitrogenaseactivity. However, acetylene reduction per unit bacteroid proteinand levels of shoot dry matter of plants grown under nitrogen-freeconditions inoculated with the sucD mutant were 45% and 51% ofthe wild-type values, respectively (Fig. 3). Therefore, sucD bacte-roids have lowered N2 fixation, presumably due to attenuation,but not complete blockage, of the TCA cycle.

Metabolite profiles of the sucD mutant and wild-type bacte-roids (Fig. 4) show that while succinate levels were similar inRU116 and wild-type bacteroids, levels of fumarate and malatewere considerably lower in the mutant bacteroids, indicating re-duced flux of carbon. Our key investigation concerns the decar-boxylating arm of the TCA cycle. Predictably for a mutant strain

FIG 3 Symbiotic phenotype of sucD mutant (RU116) compared to wild-type Rlv3841. (a and b) N2 fixation as measured by acetylene reduction on whole plantsat dpi 28 expressed on (a) a per-plant basis (n � 6 per treatment) and (b) a per-unit bacteroid protein basis (n � 6), where the significance value (*, P � 0.05)was determined by Welch’s t test. (c) A photograph of pea plants taken at dpi 47 with uninoculated water control (WC), Rlv3841, and sucD (RU116) treatments.(d) Mean shoot dry weights of dpi 42 peas (n � 12 per treatment), where treatments not sharing a letter differ significantly (P � 0.05; analysis of variance[ANOVA] and Tukey’s honest significant difference [HSD]). In all cases, error bars represent standard errors of the means.

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blocked in the TCA cycle at succinyl-CoA synthetase, citrate levelswere 11-fold higher in the sucD mutant than in the wild-typestrain and levels of intermediates derived from 2-oxoglutarate,such as glutamate, glutathione, and 2-hydroxyglutarate, were allincreased markedly (Fig. 4). Therefore, attenuation of succinyl-CoA synthetase activity caused an accumulation of metabolitesprior to the activity of the decarboxylating arm of the TCA cycle.Thus, the TCA cycle operates in bacteroids and reducing its activ-ity also reduced N2 fixation. Crucially, though, while the levels ofpyruvate of the two bacteroid types were similar, no acetyl-CoA orfree coenzyme A was detected in the sucD mutant. If the only majorroute for acetyl-CoA metabolism is the TCA cycle, its levels shouldrise dramatically in strain RU116 (sucD). This suggests that acetyl-CoA has other large sinks that are independent of the TCA cycle. Thepresence of alternative sinks for acetyl-CoA would explain its very lowlevel in bacteroids compared to free-living bacteria. It would also haveprofound implications for our understanding of Rhizobium-legumesymbioses, as it suggests a major rerouting of central metabolismduring N2 fixation in pea bacteroids.

Lipids are a sink for acetyl-CoA in bacteroids. Apart from itscomplete oxidation in the TCA cycle, the other major metabolicfate of acetyl-CoA is in lipogenesis. Two possible products of lipo-genesis are poly-�-hydroxybutyrate (PHB) and fatty acids. Con-siderable attention has focused on PHB because it is abundant insoybean and common bean bacteroids, although it is thought tobe absent in mature N2-fixing bacteroids from indeterminate

nodulating plants, including pea, alfalfa, and clover. In contrast,there has been relatively little quantification of bacteroid lipids,which we sought to address.

There was a range of chain lengths and degrees of unsaturationin the free fatty acids in both bacteroids and free-living succinate-grown cells (Table 2). Levels of long-chain free fatty acids (C16 toC20) were higher in bacteroids than in either free-living bacteria ornodule cytosolic fractions. There were also significantly higherlevels of monoacylglycerols, with bacteroids containing highlyelevated levels of 1-linoleoylglycerol ( 57-fold), 1-palmitoyl-glycerol (16-fold), and 2-linoleoylglycerol ( 13-fold) as wellas 1-stearoylglycerol (3.9-fold) and 2-oleoylglycerol (5.8-fold).Moreover, the less efficient N2-fixing sucD mutant strain showedlevels of these lipid species that were significantly lower than thoseseen with wild-type Rlv3841 bacteroids. The presence of thesemolecules at high levels in wild-type Rlv3841 suggests that bacte-roids use fatty acids as a sink for acetyl-CoA.

It was not possible to detect diacyclglycerols or triacylglycerolsin these samples as they fall outside the polarity range and uppersize limit of the GC- and LC-MS techniques used. Therefore,membrane-free extracts were isolated by ultracentrifugationand their glycerolipid level quantified by enzyme assay. Levelsof glycerolipids were 22-fold higher in bacteroids than in free-living cells (62 � 2.66 ng/mg protein versus 2.8 � 1.26 ng/mgprotein, respectively). Bacteroids channel a large proportion ofacetyl-CoA away from the TCA cycle and into lipids, suggesting

FIG 4 Metabolite profile of sucD (RU116) bacteroids versus Rlv3841 bacteroids showing fold change in metabolite abundance relative to sucD bacteroids.Bacteroids were isolated from nodules from pea plants 28 dpi. Bolded intermediates were detected by metabolite profiling, with a statistically significant folddifference (P � 0.05 by Welch’s t test and Q � 0.1 for the false-discovery rate) denoted with a red (increase) or green (decrease) arrow. Intact arrows indicatesingle-step enzyme-catalyzed reactions. Broken arrows indicate instances in which two or more enzyme-catalyzed steps are involved in a series of reactions. sucDbacteroids are attenuated in TCA cycle enzymes post-2-oxoglutarate (2-OG). Abbreviations: UD, undetectable; GABA, �-amino butyric acid; GSH, glutathione(reduced); GSSG, glutathione (oxidized); OAA, oxaloacetate; PEP, phosphoenolpyruvate.

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that related storage mechanisms may be utilized under N2-fixing conditions.

Pea bacteroids of Rlv3841 accumulate PHB. PHB accumula-tion occurs in undifferentiated rhizobia in infection threads of peanodules but is thought to be absent in bacteroids (20). When R. legu-minosarum strain A34 was mutated in phaC, encoding a type I PHBsynthase, it lacked detectable PHB both in infection thread bacteriaand in bacteroids. This is consistent with the paradigm that bacte-roids from indeterminate nodules such as pea and alfalfa do not makePHB in bacteroids. However, the genome of R. leguminosarumstrain Rlv3841 has two PHB synthases: a type I PHB synthase onthe chromosome (phaC1, RL2094) and a phaE (pRL100104)phaC2 (pRL100105) type III PHB synthase on the pRL10 symbioticplasmid. The putative operon containing phaE and phaC2 is pre-ceded by a consensus nifA promoter and was induced 7- to 40-fold inbacteroids, while phaC1 was not upregulated (44). As PHB is anotherlipogenic end product of acetyl-CoA metabolism, we investigated thesymbiotic roles of these two PHB synthases in Rlv3841.

Previous work demonstrated that phaC1 was active in free-living Rlv3841, as mutation of this gene reduced PHB accumula-tion in the RU137 mutant by 93% relative to wild-type results(12), although the symbiotic performance of this phaC1 mutantwas not determined. Therefore, we isolated a phaC2 single mutant(LMB814) and a phaC1 phaC2 double mutant (LMB816) inRlv3841 and assessed their symbiotic phenotype, along with thatof the original phaC1 mutant. While rates of N2 fixation in phaC1and phaC2 single mutants and phaC1 phaC2 double mutants werenot significantly different from the rate seen with wild-typeRlv3841 (Fig. S1 in the supplemental material), examination ofnodule sections by transmission electron microscopy (TEM)showed that PHB accumulation was altered. Pea nodules contain-

ing wild-type Rlv3841 exhibited large PHB droplets in bacteria ininfection threads and smaller bodies in mature bacteroids (Fig. 5).Previously, when small PHB droplets were observed in bacteroids,it was assumed they were synthesized by bacteria in infectionthreads. However, while the phaC1 mutant harbored small PHBdroplets in bacteroids, they were absent in the undifferentiatedbacteria in infection threads. In contrast, PHB was largely absentin phaC2 mutant bacteroids but was abundant in bacteria occu-pying infection threads. Finally, PHB was absent from both bac-teroids and bacteria in infection threads in the phaC1 phaC2 mu-tant. Therefore, Rlv3841 has two functional PHB synthases: oneactive in free-living and undifferentiated bacteria (type I, PhaC1)and the other active in bacteroids (type III, PhaE PhaC2). Al-though most sequenced rhizobia carry a type I PHB synthase,analysis of genome sequences shows that other rhizobia con-tain phaE and phaC2 genes, including strains forming symbi-otic interactions not usually thought to make PHB, such as R.leguminosarum bv. viciae VF39 (pea) and R. leguminosarum bv.trifolii TA1 (clover) (Integrated Microbial Genomes [https://img.jgi.doe.gov/cgi-bin/w/main.cgi]). It is therefore likely that theseother type III-harboring bacteroids also accumulate PHB, as hasbeen demonstrated for Rlv3841.

DISCUSSION

The metabolism of free-living Rlv3841 growing on succinate asthe sole carbon source is dominated by flux through the TCA cycleas well as anaplerotic and biosynthetic reactions. However, whilethe TCA cycle is essential for fully effective N2 fixation in peabacteroids, the accumulation of lipid shows a significant alterna-tive fate for acetyl-CoA. Importantly, this observation is sup-ported by the work of Miller and Tremblay (53), who showed that

TABLE 2 Comparison of fatty acids and monoacylglycerols detected in metabolite profiles

Lipid species

Fold changea in metabolite abundance relative to that in:

Rlv3841 free-living samples Nodule cytosol samples sucD bacteroid samples

Free fatty acidscis-Vaccenate (18:1n7) 1.99 5.91 4.00Palmitoleate (16:1n7) 8.20 4.87 2.94Linolenate (� or � [18:3n3 or 6]) 23.0 4.09 1.32Linoleate (18:2n6) 18.7 3.62 2.13Eicosenoate (20:1n9 or 11) 8.39 2.91 2.5610-Heptadecenoate (17:1n7) 8.22 2.54 1.19Dihomo-linoleate (20:2n6) 3.81 2.50 1.28Stearate (18:0) 2.16 1.90 2.44Palmitate (16:0) 3.72 1.88 2.94Margarate (17:0) 3.94 1.15 1.89Pelargonate (9:0) 0.75 0.48 2.17Heptanoate (7:0) 0.19 0.19 0.46Caproate (6:0) 19.7 0.16 0.71Caprylate (8:0) 1.45 0.16 0.75Isovalerate 1.30 0.05 0.36

Glycerolipids1-Linoleoylglycerol (18:2) 57.2 8.04 9.092-Linoleoylglycerol (18:2) 13.2 5.38 6.252-Oleoylglycerol (18:1) 5.83 3.16 2.781-Stearoylglycerol (18:0) 3.86 0.33 3.851-Palmitoylglycerol (16:0) 15.6 0.27 4.35

a Values are fold changes in metabolite abundance in Rlv3841 bacteroids relative to that in the indicated samples. Values highlighted with boldface indicate significant increases(P � 0.05 and Q � 0.1), and values highlighted with italics indicate significant decreases (P � 0.05 and Q � 0.1); data without highlighting showed no significant differences.

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S. meliloti bacteroids from alfalfa nodules contain 34% of the totalneutral lipid fraction as di- and triglycerides whereas these lipidswere undetected in free-living S. meliloti. Moreover, the extraor-dinary deposition of PHB in bacteroids from common bean andsoybean is an extreme example of carbon storage and redox bal-ancing that has hitherto lacked a coherent explanation, particu-larly since preventing synthesis in these symbioses does not pre-vent N2 fixation (19–21). Here we show that bacteroids of somestrains of R. leguminosarum, such as Rlv3841, make PHB via aputative nifA-dependent type III PHB synthase. Therefore, theparadigm that mature bacteroids from indeterminate nodules(such as those formed on pea, alfalfa, and clover) do not synthesizePHB is incorrect. Most importantly, Rlv3841 bacteroids accumu-late both PHB and lipid, showing that, even with acetyl-CoA in-corporated into lipids, more acetyl-CoA accumulates in PHB.Thus, entry of acetyl-CoA into the TCA cycle must be limited,which implies that symbiotic N2 fixation should be thought of as afundamentally lipogenic process.

The complete oxidation of 1 mole of acetyl-CoA in the TCAcycle yields 4 moles of reducing equivalents [i.e., NAD(P)H orFADH2]. In free-living rhizobia, this reductant can be channeledto the aerobic respiratory chain, driving oxidative phosphoryla-tion, or used as a reductant in biosynthesis to fuel cell growth anddivision. However, mature pea bacteroids are in a metabolicallyactive but nondividing state. In addition, N2 fixation in legumeroot nodules occurs at microaerobic O2 concentrations, estimatedat 3 to 57 nM (54, 55). This low O2 level is likely to restrict bacte-roid respiration and hence TCA cycle activity, thereby forcingacetyl-CoA into lipids. While it is theoretically possible to havelarge rates of electron flux to a high-affinity terminal oxidase suchas cbb3 in bacteroids if O2 flux is also high, the large-scale produc-

tion of lipids and PHB suggests that this route is restricted. In-stead, by channeling acetyl-CoA into lipid and PHB synthesis,bacteroids could overcome this metabolic constraint by consum-ing both carbon and reductant as NAD(P)H. Lipogenesis is a clas-sic response of all domains of life to an excess of carbon and re-ductant that cannot be reoxidized by respiration or fermentation.Thus, free-living bacteria synthesize lipid when growth is nutri-tionally unbalanced, such as under O2- or N2-limited conditions(56, 57). During free-living N2 fixation, both Azotobacter beijer-inckii and A. caulinodans accumulate PHB, and in A. caulinodans,PHB synthesis is essential to both free-living and symbiotic N2

fixation (22, 58). Thus, bacteroids may become lipogenic as aphysiological response to the microaerobic environment insidelegume root nodules.

Although aerobically growing free-living rhizobia and bacte-roids differ with respect to O2 supply and ability to divide, theother obvious metabolic difference is the supply of ATP and re-ductant for bacteroid nitrogenase. Bacteroids must supply reduc-tant and ATP to nitrogenase, requiring 8 moles of electrons and 16moles of ATP to reduce 1 mole of N2 (equation 1). Although theelectron source for nitrogenase in the free-living N2-fixing bacte-rium Klebsiella pneumoniae, where electrons are transferred byNifJ (pyruvate:flavin oxidoreductase) and NifF (flavodoxin) com-plex from pyruvate to nitrogenase (59, 60), is well understood, it isunknown for rhizobia. In the classical model in rhizobia, all re-ductants generated by metabolism, primarily as NAD(P)H, can beallocated to all processes, including N2 reduction or biosynthesiswith excess reductant and ATP consumed by lipogenesis (Fig. 6).However, the standard redox potentials of NADH and ferredoxin(the E0= value for NAD�/NADH is �320 mV and the ferredoxinFe3�/Fe2� value is �484 mV [16, 17]) suggest that it is unlikely

FIG 5 Transmission electron micrographs of pea nodules at 28 dpi. (a and b) Wild-type Rlv3841 bacteroids (a) and bacteroids in an infection thread (b), bothshowing PHB droplets. (c and d) Mutant phaC1 (RU137) bacteroids showing PHB accumulation (c); PHB accumulation was absent from infection threads (d).(e and f) Mutant phaC2 (LMB814) bacteroids where PHB droplets were largely absent (e); the bacteroids were abundant in infection threads (f). (g and h) Doublemutant phaC1 phaC2 (LMB816) bacteroids (g) and bacteroids in infection threads (h), with PHB absent from both. Scale bars are 2 �m in panels a, c, e, and gand 1 �m in panels b, d, f, and h. Arrows point to PHB droplets which appear white.

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that NADH donates electrons directly to ferredoxin and then tonitrogenase. An alternative explanation would be that a specificmolecule acts as the low-potential electron donor to nitrogenase,an example being pyruvate oxidation by the NifJ-NifF complex inK. pneumoniae (59, 60). This process consumes four pyruvatemolecules and produces four acetyl-CoA molecules to generatethe eight electrons needed by nitrogenase. Since this complex isnot present in rhizobia, an alternative pathway is required. Onepossibility is that the electron-transferring flavoprotein (ETF)complex, FixABCX, interacts with pyruvate dehydrogenase, asshown by genetic suppressor analysis in A. caulinodans (61). ETFcomplexes use electron bifurcation in anaerobic bacteria (62, 63),which might enable FixABCX to generate low-potential electronsfor reduction of ferredoxin and then N2 (Fig. 6). While unproven,such a mechanism would require 8 moles of pyruvate to reduce 1mole of N2 and would exacerbate the reductant problem becauseacetyl-CoA oxidized by the TCA cycle would generate excessNAD(P)H. In the absence of convincing experimental evidencefor the pathway of electron donation to nitrogenase, we cannotcomplete a formal model to represent the electron and reductantbalance. However, Fig. 6 illustrates how dramatically redox bal-

ance in bacteroids can be altered by the need for low-potentialelectrons for N2 reduction.

While A. caulinodans must synthesize PHB during N2 fixation(22), synthesis can be blocked in bacteroids of peas, alfalfa, com-mon bean, and soybean (19–21, 64, 65). The ability to preventPHB synthesis and still have a functioning bacteroid may be ex-plained by the existence of multiple storage sinks for acetyl-CoA,including PHB, free fatty acids, glycerolipids, and membranephospholipids, with PHB itself being less important in these sym-bioses. Overall, bacteroids are highly lipogenic, with multiple lipidsinks for excess reductant. This applies to both determinate andindeterminate nodules and is likely to be an essential part of theenergization of nitrogenase and associated redox balance in allN2-fixing symbioses.

ACKNOWLEDGMENTS

This work was supported by the Biotechnology and Biological SciencesResearch Council (grant number BB/F013159/1). The collaboration be-tween J.J.T. and P.S.P. is supported by Murdoch University through theSir Walter Murdoch Adjunct Scheme.

FIG 6 Two possible pathways of electron allocation in N2-fixing bacteroids. (a) In the first scenario, NADH supplies electrons directly to nitrogenase aswell as providing ATP from oxidative phosphorylation. A minimum of 2 moles of malate is required to be oxidized to acetyl-CoA to yield sufficient ATPand electrons to reduce 1 mole of N2. (b) In the second scenario, electrons are supplied to nitrogenase via tight coupling with PDH and electronbifurcation through FixABCX, requiring 8 moles of malate to reduce 1 mole of N2. The 16 moles of electrons liberated from the oxidation of 8 moles ofpyruvate could undergo electron bifurcation at FixABCX, resulting in 8 (high-potential) electrons reducing CoQ via the Fix complex, while 8 (low-potential) electrons are channeled to nitrogenase and N2 fixation. The 16 moles of ATP for N2 fixation could be supplied from oxidative phosphorylation,for example, using the 8 (high-potential) electrons from FixABCX that enter the electron transport chain (ETC) at the level of CoQ (producing 6 molesof ATP), plus the reoxidation of 4 moles of NADH (of a total 8 moles of NADH) produced by malic enzyme (producing 10 moles of ATP). In this schemethere would be no need for all 8 moles of acetyl-CoA produced to enter the TCA cycle and undergo further oxidation to yield 24 moles of NADH and 8moles of FADH2, as that would be far in excess of the requirement to make the additional ATP required for N2 fixation, leading to overreduction of theelectron carrier pool. Instead, only a fraction of the acetyl-CoA might be further oxidized in the TCA cycle and the NADH produced used to supportlipogenesis. The two models are not mutually exclusive, as, in panel a, free NADH might also interact with FixABCX, enabling low-potential electrons tobe generated by bifurcation for reduction of ferredoxin. Note that a P:2e� ratio of 2.5 is assumed for NADH and a ratio of 1.5 is assumed for electronsentering the ETC at the level of CoQ. For simplicity, we have not distinguished between NAD� and NADP� in this model. Abbreviations: CoQ, coenzymeQ; ME, malic enzyme; N2ase, nitrogenase; PDH, pyruvate dehydrogenase.

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We thank Graham O’Hara and Garth Maker (Murdoch University,Australia) for valuable input in preparing the manuscript.

FUNDING INFORMATIONThis work, including the efforts of Philip S. Poole, was funded by Bio-technology and Biological Sciences Research Council (BBSRC)(BB/F013159/1).

The funders had no role in study design, data collection and interpreta-tion, or the decision to submit the work for publication.

REFERENCES1. Olivares J, Bedmar EJ, Sanjuan J. 2013. Biological nitrogen fixation in the

context of global change. Mol Plant Microbe Interact 26:486 – 494. http://dx.doi.org/10.1094/MPMI-12-12-0293-CR.

2. Herridge DF, Peoples MB, Boddey RM. 2008. Global inputs of biologicalnitrogen fixation in agricultural systems. Plant Soil 311:1–18. http://dx.doi.org/10.1007/s11104-008-9668-3.

3. Udvardi M, Poole PS. 2013. Transport and metabolism in legume-rhizobia symbioses. Annu Rev Plant Biol 64:781– 805. http://dx.doi.org/10.1146/annurev-arplant-050312-120235.

4. Terpolilli JJ, Hood GA, Poole PS. 2012. What determines the efficiencyof N2-fixing Rhizobium-legume symbioses? Adv Microb Physiol 60:325–389. http://dx.doi.org/10.1016/B978-0-12-398264-3.00005-X.

5. Mulley G, Lopez-Gomez M, Zhang Y, Terpolilli J, Prell J, Finan T,Poole P. 2010. Pyruvate is synthesized by two pathways in pea bacteroidswith different efficiencies for nitrogen fixation. J Bacteriol 192:4944 –4953. http://dx.doi.org/10.1128/JB.00294-10.

6. Driscoll BT, Finan TM. 1993. NAD�-dependent malic enzyme of Rhizo-bium meliloti is required for symbiotic nitrogen fixation. Mol Microbiol7:865– 873. http://dx.doi.org/10.1111/j.1365-2958.1993.tb01177.x.

7. Driscoll BT, Finan TM. 1997. Properties of NAD�- and NADP�-dependent malic enzymes of Rhizobium (Sinorhizobium) meliloti and dif-ferential expression of their genes in nitrogen-fixing bacteroids. Microbi-ology 143:489 – 498. http://dx.doi.org/10.1099/00221287-143-2-489.

8. Preisig O, Anthamatten D, Hennecke H. 1993. Genes for a microaero-bically induced oxidase complex in Bradyrhizobium japonicum are essen-tial for a nitrogen-fixing endosymbiosis. Proc Natl Acad Sci U S A 90:3309 –3313. http://dx.doi.org/10.1073/pnas.90.8.3309.

9. Delgado MJ, Bedmar EJ, Downie JA. 1998. Genes involved in the for-mation and assembly of rhizobial cytochromes and their role in symbioticnitrogen fixation. Adv Microb Physiol 40:191–231. http://dx.doi.org/10.1016/S0065-2911(08)60132-0.

10. Mortimer MW, McDermott TR, York GM, Walker GC, Kahn ML. 1999.Citrate synthase mutants of Sinorhizobium meliloti are ineffective and havealtered cell surface polysaccharides. J Bacteriol 181:7608 –7613.

11. Koziol U, Hannibal L, Rodriguez MC, Fabiano E, Kahn ML, Noya F.2009. Deletion of citrate synthase restores growth of Sinorhizobium meli-loti 1021 aconitase mutants. J Bacteriol 191:7581–7586. http://dx.doi.org/10.1128/JB.00777-09.

12. Walshaw DL, Wilkinson A, Mundy M, Smith M, Poole PS. 1997.Regulation of the TCA cycle and the general amino acid permease byoverflow metabolism in Rhizobium leguminosarum. Microbiology 143:2209 –2221. http://dx.doi.org/10.1099/00221287-143-7-2209.

13. Pardo MA, Lagunez J, Miranda J, Martinez E. 1994. Nodulating abilityof Rhizobium tropici is conditioned by a plasmid-encoded citrate synthase.Mol Microbiol 11:315–321. http://dx.doi.org/10.1111/j.1365-2958.1994.tb00311.x.

14. Green LS, Emerich DW. 1997. The formation of nitrogen-fixing bacte-roids is delayed but not abolished in soybean infected by an �-ketogluta-rate dehydrogenase-deficient mutant of Bradyrhizobium japonicum. PlantPhysiol 114:1359 –1368.

15. Shah R, Emerich DW. 2006. Isocitrate dehydrogenase of Bradyrhizobiumjaponicum is not required for symbiotic nitrogen fixation with soybean. JBacteriol 188:7600 –7608. http://dx.doi.org/10.1128/JB.00671-06.

16. Carter KR, Rawlings J, Orme-Johnson WH, Becker RR, Evans HJ. 1980.Purification and characterization of a ferredoxin from Rhizobium japoni-cum bacteroids. J Biol Chem 255:4213– 4223.

17. Loach PA. 2010. Oxidation-reduction potentials, absorbance bands andmolar absorbance of compounds used in biochemical studies, p 557–578.In Lundblad RL, Macdonald F (ed), Handbook of biochemistry and mo-lecular biology. CRC Press, Boca Raton, FL.

18. Sprent JI. 2009. Legume nodulation: a global perspective. Wiley-Blackwell, Oxford, United Kingdom.

19. Cevallos MA, Encarnacion S, Leija A, Mora Y, Mora J. 1996. Genetic andphysiological characterization of a Rhizobium etli mutant strain unable tosynthesize poly-beta-hydroxybutyrate. J Bacteriol 178:1646 –1654.

20. Lodwig EM, Leonard M, Marroqui S, Wheeler TR, Findlay K, DownieJA, Poole PS. 2005. Role of polyhydroxybutyrate and glycogen as carbonstorage compounds in pea and bean bacteroids. Mol Plant Microbe Inter-act 18:67–74. http://dx.doi.org/10.1094/MPMI-18-0067.

21. Quelas JI, Mongiardini EJ, Perez-Gimenez J, Parisi G, Lodeiro AR.2013. Analysis of two polyhydroxyalkanoate synthases in Bradyrhizobiumjaponicum USDA 110. J Bacteriol 195:3145–3155. http://dx.doi.org/10.1128/JB.02203-12.

22. Mandon K, Michel-Reydellet N, Encarnacion S, Kaminski PA, Leija A,Cevallos MA, Elmerich C, Mora J. 1998. Poly-�-hydroxybutyrate turn-over in Azorhizobium caulinodans is required for growth and affects nifAexpression. J Bacteriol 180:5070 –5076.

23. Johnston AWB, Beringer JE. 1975. Identification of the Rhizobium strainsin pea root nodules using genetic markers. J Gen Microbiol 87:343–350.http://dx.doi.org/10.1099/00221287-87-2-343.

24. Prentki P, Krisch HM. 1984. In vitro insertional mutagenesis with aselectable DNA fragment. Gene 29:303–313. http://dx.doi.org/10.1016/0378-1119(84)90059-3.

25. Quandt J, Hynes MF. 1993. Versatile suicide vectors which allow directselection for gene replacement in Gram-negative bacteria. Gene 127:15–21. http://dx.doi.org/10.1016/0378-1119(93)90611-6.

26. Figurski DH, Helinski DR. 1979. Replication of an origin-containingderivative of plasmid RK2 dependent on a plasmid function provided intrans. Proc Natl Acad Sci U S A 76:1648 –1652. http://dx.doi.org/10.1073/pnas.76.4.1648.

27. Karunakaran R, Haag AF, East AK, Ramachandran VK, Prell J,James EK, Scocchi M, Ferguson GP, Poole PS. 2010. BacA is essentialfor bacteroid development in nodules of galegoid, but not phaseoloid,legumes. J Bacteriol 192:2920 –2928. http://dx.doi.org/10.1128/JB.00020-10.

28. Beringer JE. 1974. R factor transfer in Rhizobium leguminosarum. J GenMicrobiol 84:188 –198.

29. Poole PS, Schofield NA, Reid CJ, Drew EM, Walshaw DL. 1994. Iden-tification of chromosomal genes located downstream of dctD that affectthe requirement for calcium and the lipopolysaccharide layer of Rhizo-bium leguminosarum. Microbiology 140:2797–2809. http://dx.doi.org/10.1099/00221287-140-10-2797.

30. Masakapalli SK, Kruger NJ, Ratcliffe RG. 2013. The metabolic fluxphenotype of heterotrophic Arabidopsis cells reveals a complex responseto changes in nitrogen supply. Plant J 74:569 –582. http://dx.doi.org/10.1111/tpj.12142.

31. Kruger NJ, Masakapalli SK, Ratcliffe RG. 2012. Strategies for investigat-ing the plant metabolic network with steady-state metabolic flux analysis:lessons from an Arabidopsis cell culture and other systems. J Exp Bot 63:2309 –2323. http://dx.doi.org/10.1093/jxb/err382.

32. Lawton KA, Berger A, Mitchell M, Milgram KE, Evans AM, Guo L,Hanson RW, Kalhan SC, Ryals JA, Milburn MV. 2008. Analysis of theadult human plasma metabolome. Pharmacogenomics 9:383–397. http://dx.doi.org/10.2217/14622416.9.4.383.

33. Evans AM, DeHaven CD, Barrett T, Mitchell M, Milgram E. 2009.Integrated, nontargeted ultrahigh performance liquid chromatography/electrospray ionization tandem mass spectrometry platform for the iden-tification and relative quantification of the small-molecule complement ofbiological systems. Anal Chem 81:6656 – 6667. http://dx.doi.org/10.1021/ac901536h.

34. Yobi A, Wone BW, Xu W, Alexander DC, Guo L, Ryals JA, Oliver MJ,Cushman JC. 2012. Comparative metabolic profiling between desicca-tion-sensitive and desiccation-tolerant species of Selaginella reveals in-sights into the resurrection trait. Plant J 72:983–999. http://dx.doi.org/10.1111/tpj.12008.

35. Bradford MM. 1976. A rapid and sensitive method for the quantitation ofmicrogram quantities of protein utilizing the principle of protein-dyebinding. Anal Biochem 72:248–254. http://dx.doi.org/10.1016/0003-2697(76)90527-3.

36. Storey JD, Tibshirani R. 2003. Statistical significance for genomewidestudies. Proc Natl Acad Sci U S A 100:9440 –9445. http://dx.doi.org/10.1073/pnas.1530509100.

37. Yates RJ, Howieson JG, Hungria M, Bala A, O’Hara GW, Terpolilli JJ.

Terpolilli et al.

2874 jb.asm.org October 2016 Volume 198 Number 20Journal of Bacteriology

on February 2, 2020 by guest

http://jb.asm.org/

Dow

nloaded from

2016. Authentication of rhizobia and assessment of the legume symbiosisin controlled plant growth systems, p 73–108. In Howieson JG, DilworthMJ (ed), Working with rhizobia. Australian Centre for International Ag-ricultural Research, Canberra, Australia.

38. Allaway D, Lodwig EM, Crompton LA, Wood M, Parsons R,Wheeler TR, Poole PS. 2000. Identification of alanine dehydrogenaseand its role in mixed secretion of ammonium and alanine by pea bac-teroids. Mol Microbiol 36:508 –515. http://dx.doi.org/10.1046/j.1365-2958.2000.01884.x.

39. Buchanan-Wollaston V. 1979. Generalized transduction in Rhizobiumleguminosarum. J Gen Microbiol 112:135–142.

40. Lodwig E, Poole P. 2003. Metabolism of Rhizobium bacteroids. Crit RevPlant Sci 22:37–78. http://dx.doi.org/10.1080/713610850.

41. Joy KW, Prabha C. 1986. The role of transamination in the synthesis ofhomoserine in peas. Plant Physiol 82:99 –102. http://dx.doi.org/10.1104/pp.82.1.99.

42. Prell J, Bourdes A, Karunakaran R, Lopez-Gomez M, Poole P. 2009.Pathway of �-aminobutyrate metabolism in Rhizobium leguminosarum3841 and its role in symbiosis. J Bacteriol 191:2177–2186. http://dx.doi.org/10.1128/JB.01714-08.

43. Min B, Pelaschier JT, Graham DE, Tumbula-Hansen D, Soll D. 2002.Transfer RNA-dependent amino acid biosynthesis: an essential route toasparagine formation. Proc Natl Acad Sci U S A 99:2678 –2683. http://dx.doi.org/10.1073/pnas.012027399.

44. Karunakaran R, Ramachandran VK, Seaman JC, East AK, Mouhsine B,Mauchline TH, Prell J, Skeffington A, Poole PS. 2009. Transcriptomicanalysis of Rhizobium leguminosarum biovar viciae in symbiosis with hostplants Pisum sativum and Vicia cracca. J Bacteriol 191:4002– 4014. http://dx.doi.org/10.1128/JB.00165-09.

45. Vanderlinde EM, Hynes MF, Yost CK. 2014. Homoserine catabolism byRhizobium leguminosarum bv. viciae 3841 requires a plasmid-borne genecluster that also affects competitiveness for nodulation. Environ Micro-biol 16:205–217. http://dx.doi.org/10.1111/1462-2920.12196.

46. Huerta-Zepeda A, Ortuno L, Du Pont G, Duran S, Lloret A, Merchant-Larios H, Calderon J. 1997. Isolation and characterization of Rhizobiumetli mutants altered in degradation of asparagine. J Bacteriol 179:2068 –2072.

47. McKay IA, Dilworth MJ, Glenn AR. 1989. Carbon catabolism in contin-uous cultures and bacteroids of Rhizobium leguminosarum MNF 3841.Arch Microbiol 152:606 – 610. http://dx.doi.org/10.1007/BF00425495.

48. Tabrett CA, Copeland L. 2000. Biochemical controls of citrate synthase inchickpea bacteroids. Arch Microbiol 173:42– 48. http://dx.doi.org/10.1007/s002030050006.

49. Dunn MF. 1998. Tricarboxylic acid cycle and anaplerotic enzymes inrhizobia. FEMS Microbiol Rev 22:105–123. http://dx.doi.org/10.1111/j.1574-6976.1998.tb00363.x.

50. Patriarca EJ, Tate R, Iaccarino M. 2002. Key role of bacterial NH4�

metabolism in Rhizobium-plant symbiosis. Microbiol Mol Biol Rev 66:203–222. http://dx.doi.org/10.1128/MMBR.66.2.203-222.2002.

51. Mulley G, White JP, Karunakaran R, Prell J, Bourdes A, Bunnewell S,Hill L, Poole PS. 2011. Mutation of GOGAT prevents pea bacteroidformation and N2 fixation by globally downregulating transport of or-

ganic nitrogen sources. Mol Microbiol 80:149 –167. http://dx.doi.org/10.1111/j.1365-2958.2011.07565.x.

52. Poole P, Reid C, East AK, Allaway D, Day M, Leonard M. 1999.Regulation of the mdh-sucCDAB operon in Rhizobium leguminosarum.FEMS Microbiol Lett 176:247–255. http://dx.doi.org/10.1111/j.1574-6968.1999.tb13669.x.

53. Miller RW, Tremblay PA. 1983. Cytoplasmic membrane of Rhizobiummeliloti bacteroids. I. Alterations in lipid composition, physical properties,and respiratory proteins. Can J Biochem Cell Biol 61:1334 –1340. http://dx.doi.org/10.1139/o83-170.

54. Kuzma MM, Hunt S, Layzell DB. 1993. Role of oxygen in the limitationand inhibition of nitrogenase activity and respiration rate in individualsoybean nodules. Plant Physiol 101:161–169.

55. Appleby CA. 1984. Leghemoglobin and Rhizobium respiration. AnnuRev Plant Physiol 35:443– 478. http://dx.doi.org/10.1146/annurev.pp.35.060184.002303.

56. Rehm BH. 2010. Bacterial polymers: biosynthesis, modifications andapplications. Nat Rev Microbiol 8:578 –592. http://dx.doi.org/10.1038/nrmicro2354.

57. Wältermann M, Steinbüchel A. 2005. Neutral lipid bodies in pro-karyotes: recent insights into structure, formation, and relationship toeukaryotic lipid depots. J Bacteriol 187:3607–3619. http://dx.doi.org/10.1128/JB.187.11.3607-3619.2005.

58. Senior PJ, Beech GA, Ritchie GA, Dawes EA. 1972. The role of oxygenlimitation in the formation of poly-�-hydroxybutyrate during batch andcontinuous culture of Azotobacter beijerinckii. Biochem J 128:1193–1201.http://dx.doi.org/10.1042/bj1281193.

59. Shah VK, Stacey G, Brill WJ. 1983. Electron transport to nitrogenase.Purification and characterization of pyruvate:flavodoxin oxidoreductase.The nifJ gene product. J Biol Chem 258:12064 –12068.

60. Nieva-Gómez D, Roberts GP, Klevickis S, Brill WJ. 1980. Electrontransport to nitrogenase in Klebsiella pneumoniae. Proc Natl Acad SciU S A 77:2555–2558. http://dx.doi.org/10.1073/pnas.77.5.2555.

61. Scott JD, Ludwig RA. 2004. Azorhizobium caulinodans electron-transferring flavoprotein N electrochemically couples pyruvate dehydro-genase complex activity to N2 fixation. Microbiology 150:117–126. http://dx.doi.org/10.1099/mic.0.26603-0.

62. Bertsch J, Parthasarathy A, Buckel W, Muller V. 2013. An electron-bifurcating caffeyl-CoA reductase. J Biol Chem 288:11304 –11311. http://dx.doi.org/10.1074/jbc.M112.444919.

63. Hess V, Gonzalez JM, Parthasarathy A, Buckel W, Muller V. 2013.Caffeate respiration in the acetogenic bacterium Acetobacterium woodii: acoenzyme A loop saves energy for caffeate activation. Appl Environ Mi-crobiol 79:1942–1947. http://dx.doi.org/10.1128/AEM.03604-12.

64. Povolo S, Tombolini R, Morea A, Anderson AJ, Casella S, Nuti MP.1994. Isolation and characterization of mutants of Rhizobium meliloti un-able to synthesize poly-�-hydroxybutyrate. Can J Microbiol 40:823– 829.http://dx.doi.org/10.1139/m94-131.

65. Aneja P, Charles TC. 1999. Poly-3-Hydroxybutyrate degradation in Rhi-zobium (Sinorhizobium) meliloti: isolation and characterization of a geneencoding 3-hydroxybutyrate dehydrogenase. J Bacteriol 181:849 – 857.

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