Vol. 171, No. 6

Ammonium Inhibition of Nitrogenase Activity inHerbaspirillum seropedicaet

HAIAN FU AND ROBERT H. BURRIS*Department of Biochemistry, College of Agricultural and Life Sciences, University of Wisconsin-Madison,

Madison, Wisconsin 53706

Received 6 December 1988/Accepted 13 February 1989

The effect of oxygen, ammonium ion, and amino acids on nitrogenase activity in the root-associated N2-fixingbacterium Herbaspirillum seropedicae was investigated in comparison with Azospirillum spp. and Rhodospiril-lum rubrum. H. seropedicae is microaerophilic, and its optimal dissolved oxygen level is from 0.04 to 0.2 kPafor dinitrogen fixation but higher when it is supplied with fixed nitrogen. No nitrogenase activity was detectedwhen the dissolved 02 level corresponded to 4.0 kPa. Ammonium, a product of the nitrogenase reaction,reversibly inhibited nitrogenase activity when added to derepressed cell cultures. However, the inhibition ofnitrogenase activity was only partial even with concentrations of ammonium chloride as high as 20 mM. Amidessuch as glutamine and asparagine partially inhibited nitrogenase activity, but glutamate did not. Nitrogenasein crude extracts prepared from ammonium-inhibited cells showed activity as high as in extracts from N2-fixingcells. The pattern of the dinitrogenase and the dinitrogenase reductase revealed by the immunoblottingtechnique did not change upon ammonium chloride treatment of cells in vivo. No homologous sequences were

detected with the draT-draG probe from Azospirillum lipoferum. There is no clear evidence that ADP-ribosylation of the dinitrogenase reductase is involved in the ammonium inhibition of H. seropedicae. Theuncoupler carbonyl cyanide m-chlorophenylhydrazone decreased the intracellular ATP concentration andinhibited the nitrogenase activity of whole cells. The ATP pool was not significantly disturbed when cu!tureswere treated with ammonium in vivo. Possible mechanisms for inhibition by ammonium of whole-cellnitrogenase activity in H. seropedicae are discussed.

Nitrogenase catalyzes the energy-demanding reduction ofN2 to ammonia coupled with H2 production (see reference 33for a review). The nitrogenase complex is composed of twoprotein components, dinitrogenase (MoFe protein) and dini-trogenase reductase (Fe protein). Because of the high energycost of N2 fixation, tight control of nitrogenase synthesis andactivity should be beneficial to the bacteria. The nif(nitrogenfixation) genes are repressed in the presence of combinednitrogen, such as ammonia, the immediate product of thenitrogenase reaction, and 02, which inactivates nitrogenaseproteins (see reference 43 for a review). Besides regulationat the gene expression level, nitrogenase also is regulated atthe enzymatic level in many organisms.The rapid inhibitory effect of the ammonium ion on

N2-fixing cultures ofAzotobacter vinelandii, which could notbe explained by the repression of nitrogenase synthesis, was

documented in 1946 with '5N as a tracer (5). This phenom-enon was reexamined and confirmed by others (13, 25, 29)and extended to diverse N2-fixing bacteria, such as thefree-living Azotobacter chroococcum (6, 7); Azospirillumspp. (17); the methanotrophs Methylosinus trichosporium(36, 50) and Methylococcus capsulatus (36); the sulfate-reducing bacterium Desulfovibrio gigas (40); the sulfide-oxidizing Beggiatoa alba (38); the phototrophs Rhodospiril-lum rubrum (24, 47, 48), Rhodobacter sphaeroides (15, 22,50), Rhodobacter capsulata (19, 23), Rhodopseudomonaspalustris (51), Rhodopseudomonas viridis (21), Chromatiumvinosum (14), Chlorobium limicola (42), Thiocapsa roseop-

ersicina (42), and Ectothiorhodospira sp. (3); the cyanobac-

* Corresponding author.t Unbeknownst to Professor Bums, this paper is dedicated to him

by his student, Haian Fu, on the occasion of his 75th birthday.

terium Anabaena sp. (49); and symbiotic Azorhizobiumcaulinodans ORS571 (28).The biochemical mechanisms accounting for rapid inhibi-

tion by ammonium are not all clearly understood. In Rho-dospirillum rubrum, the molecular basis for ammoniumswitch-off-on (terminology of Zumft and Castillo [51]), is thereversible ADP-ribosylation of arginine 101 of the dinitroge-nase reductase elucidated by Ludden and co-workers (for a

review, see reference 39; P. W. Ludden and G. P. Roberts,Curr. Top. Cell Reg., in press). Introduction of ammoniumto cells of Rhodospirillum rubrum growing on glutarhateresults in the covalent modification of the dinitrogenasereductase catalyzed by dinitrogenase reductase ADP-ribosyltransferase (DRAT) (31) and the rapid loss of whole-cellnitrogenase activity (t112 = 24 min) (24). The ADP-ribosegroup is attached to only one of the two identical subunits ofthe dinitrogenase reductase (24, 39). The inactivated dinitro-genase reductase can be activated in vitro in the presence ofmagnesium ATP, free Mn2+, and dinitrogenase reductase-activating glycohydrolase (DRAG) (32, 45; Ludden andRoberts, in press). The genes coding for DRAT and DRAGhave been cloned recently (11; Ludden and Roberts, inpress), and they will be useful in screening for the presenceof the DRAT-DRAG system in other organisms. Substantialevidence has accumulated to support the operation of similarcontrol mechanisms in Rhodobacter capsulata (23), Chro-matium vinosum (14), and Azospirillum brasiliense andAzospirillum lipoferum (17).

Recently, Yoch et al. (50) examined the dinitrogenasereductase pattern before and after ammonium inhibition wasinduced in Rhodobacter sphaeroides and Methylosinus tri-chosporium and found no evidence for modification of thedinitrogenase reductase. This is consistent with the obser-vation of Haaker et al. (15). Different mechanisms may

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NITROGEN FIXATION IN HERBASPIRILLUM SEROPEDICAE 3169

operate for ammonium inhibition of whole-cell nitrogenasein different microorganisms.

Herbaspirillum is a newly described genus of nitrogen-fixing bacteria (2). They are gram-negative, vibrioid (some-times helical) organisms. Herbaspirillum seropedicae (thetype species) has been isolated from surface-sterilized rootsof rice, maize, and sorghum in Brazil, and it utilizes N2 as asole source of nitrogen. These bacteria have a broaderoptimal pH range (5.3 to 8.0) for N2 fixation than Azospiril-lum spp. The association of these nitrogen-fixing bacteriawith important cereal crops make them attractive subjectsfor study of their control mechanisms. The Azospirillum spp.constitute a group of well-studied associative N2-fixing bac-teria (see reference 9 for review). Azospirilla includes fourdescribed species: Azospirillum brasiliense, Azospirillumlipoferum, Azospirillum amazonense, and Azospirillum halo-praeferans. Originally, H. seropedicae was identified asanother species of the azospirilla because of its physiologicaland morphological similarities. However, RNA-RNA hy-bridization experiments revealed very low levels (25%) ofRNA complementarity with azospirilla; hence, a new genuswith one species was proposed (2).We compared the mechanisms of ammonium regulation of

nitrogenase activity in three species of azospirilla and foundthat the nitrogenases of Azospirillum brasiliense and Azos-pirillum lipoferum are regulated by the covalent modificationof dinitrogenase reductase, whereas in Azospirillum ama-zonense a different, noncovalent inhibitory mechanism isinvolved (17). In this report, we extend our observations toinclude a newly described microaerobic N2-fixing bacterium,H. seropedicae, and we report the P02 profile for N2 fixationand that ammonium rapidly and reversibly inhibits nitroge-nase activity under conditions optimal for N2 fixation. Thepossible mechanism for inhibition by ammonium ion isdiscussed.

MATERIALS AND METHODS

Bacterial strains. H. seropedicae Z78 (ATCC 35893) andZ176 were kindly provided by J. Dobereiner. Azospirillumbrasiliense Sp7 (ATCC 29145) and Azospirillum lipoferumSpBrl7 (ATCC 29709) were obtained from the AmericanType Culture Collection (Rockville, Md.).Media and growth conditions. H. seropedicae and Azos-

pirillum lipoferum strains were grown in minimal salts me-dium essentially as described by Albrecht and Okon (1) withbiotin added only for Azospirillum lipoferum. To obtainN2-fixing cells, 200 ml of culture on minimal medium with 20mM NH4Cl was grown for 24 h at 30°C and was used toinoculate 2 liters of minimal medium. The initial ammoniumconcentration of the 2-liter batch was about 1 mM derivedfrom the inoculant. The optimal dissolved oxygen concen-tration in equilibrium with the gas phase was kept constantwith an oxystat set at 0.2 kPa of 02 for H. seropedicae and0.3 kPa for Azospirillum lipoferum. The pH was controlled at6.8 with a pH-stat. Nitrogenase was derepressed after theinitially added ammonium was consumed in the constantpresence of N2. For DNA isolation, H. seropedicae andAzospirillum brasiliense were grown in nutrient broth (DifcoLaboratories, Detroit, Mich.) medium.

Nitrogenase assays in vivo. Nitrogenase activity was mea-sured by the acetylene reduction technique (4). To controlthe dissolved 02 level during assays, we used a vigorouslystirred chamber (30 ml) equipped with a Clark-type 02electrode (20). For each assay, 4 ml of the N2-fixing culturewas transferred into the chamber, which had been flushed

with N2 to achieve anaerobiosis. The dissolved 02 concen-trations were adjusted to desired levels by injecting air, andthe dissolved 02 levels were monitored and adjusted duringthe measurements. The reactions were run at 30°C and werestarted by adding acetylene (0.1 atm [10.1 kPa]). The ethyl-ene produced was measured with a gas chromatography unitequipped with a flame ionization detector. The nitrogenaseactivity of whole cells was expressed as nanomoles ofethylene formed per milliliter of cell culture per unit of timewhen the optical density at 580 nm (OD580) was normalizedto 1.0. At an OD580 of 1.0, 1 ml of H. seropedicae culturecontains approximately 500 ,ug of protein. Thus, Fig. 1 to 3are presented with the activity expressed as if the OD580 was1.0 and the cell protein was approximately 500 ,ug/ml.

Preparation of cell-free crude extracts. For H. seropedic ae,500 ml of N2-fixing culture or ammonium-treated N2-fixingculture (see text) was harvested under N2 by centrifugationat 7,000 x g for 10 min and then suspended in 2 to 3 ml of 200mM Tris acetate buffer (pH 8.0, containing 2 mM sodiumdithionite and 1 mM dithiothreitol). After incubation withlysozyme (1 mg/ml) at 4°C for 30 min, cells were disrupted bysonication (2-min pulses, twice with 1-min intervals, on ice;output, 4; 50% duty cycle; ultrasonic cell disruptor model350; Heat Systems Ultrasonics, Inc.). Cell debris was re-moved by centrifugation at 125,000 x g for 2 h, and the clearsupernatant was used as the crude extract.

Nitrogenase assay in vitro. The dithionite-dependent acet-ylene reduction assay was performed anaerobically in 9-mlvials essentially as described by Burris (4). The reactionmixture (1 ml) contained 100 pug of creatine phosphokinase,40 ,umol of creatine phosphate, 15 pumol of magnesiumacetate, 30 p,mol of MOPS (morpholine propanesulfonicacid) buffer (pH 7.0), 5 ,umol of ATP, 5 ,umol of sodiumdithionite, and 0.5 pumol of MnCl2 (when present).SDS-PAGE and enzyme-linked immunoblotting. The so-

dium dodecyl sulfate-polyacrylamide gel electrophoresis(SDS-PAGE) system of Laemmli (30) was used as modifiedby Kanemoto and Ludden (24) with an acrylamide/N,N'-methylenebis(acrylamide) ratio of 30:0.174 to provide satis-factory resolution of the dinitrogenase reductase subunits ina 10% acrylamide gel. Protein samples were boiled in SDS-containing buffer for 1 min before loading. The immunoblot-ting procedures of Hartmann et al. (17) were used. Afterelectrophoresis, the separated proteins were electroblottedonto a nitrocellulose membrane in Tris glycine buffer. Thenitrocellulose membrane was incubated with the antiserumraised against the dinitrogenase or dinitrogenase reductaseof Azotobacter vinelandii. Goat anti-rabbit immunoglobulinG-horseradish peroxidase conjugate (Bio-Rad Laboratories,Richmond, Calif.) was used as a second antibody. Thespecific protein components were visualized on the mem-brane by color development catalyzed by horseradish per-oxidase.ATP extraction and determination. Intracellular ATP was

extracted in perchloric acid (final concentration, 0.3 M) onice as described by Privalle and Burris (41). The ATP levelwas determined by the firefly luciferin-luciferase assay (41).An Aminco Chem-Glow photometer with integrator wasused to monitor the luminescence.

Protein assay. Protein concentrations were determined bythe microbiuret method of Goa (12) with bovine serumalbumin as the standard.DNA manipulation and Southern hybridization. Total DNA

was isolated as described by Maniatis et al. (35). Therestriction fragment probe of draG-draT of Azospirillumlipoferum (11) was labeled with 32P by the random hexamer-

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Dissolved Oxygen ( kPa )FIG. 1. P02 profile for nitrogenase activity of H. seropedicae in

comparison with Azospirillum brasiliense. Samples (4 ml) of mi-croaerobically grown cultures were transferred to a controlledoxygen chamber, and acetylene reduction rates were measured atdifferent dissolved oxygen concentrations. The P02 profile of Azos-pirillum brasiliense is from reference 18. As explained in Materialsand Methods, Fig. 1 to 3 are presented with the activity expressed as

if the OD580 was 1.0 and the cell protein was approximately 500,ug/ml.

labeling procedure of Feinberg and Vogelstein (10). ForSouthern hybridization, essentially the procedures of Mani-atis et al. (35) were used, and the following conditions were

used for washing after hybridization at 42°C: 2x SSC (0.3 MNaCl and 0.03 M sodium citrate) at room temperature, 5 mineach time, twice; 2x SSC-1% SDS, 650C, 30 min each time,twice; 0.1x SSC, room temperature, 30 min each time,twice.

Chemicals. All chemicals and gases used were of highpurity or analytical grade and were from Bio-Rad Laborato-ries, Sigma Chemical Co. (St. Louis, Mo.), United StatesBiochemical Corp. (Cleveland, Ohio), Boehringer Mann-heim Biochemicals (Indianapolis, Ind.), Aldrich ChemicalCo., Inc. (Milwaukee, Wis.), Dupont, NEN Research Prod-ucts (Boston, Mass.), or Amersham Corp. (ArlingtonHeights, Ill.).

RESULTS

Effect of dissolved oxygen on nitrogen fixation. H. serope-dicae is a microaerobic N2-fixing bacterium (2). 02 at lowconcentrations serves as an electron acceptor but is toxic athigh concentrations because nitrogenase is 02 labile (27). Todetermine the optimal P02 for N2 fixation by H. seropedicae,we monitored its acetylene reduction rates in a vigorouslystirred chamber at different continuously recorded dissolved02 levels. Figure 1 presents the P02 profile of nitrogenaseactivity for H. seropedicae in comparison with that ofAzospirillum brasiliense. The optimal P02 level is in the0.02- to 0.4-kPa range, similar to that for Azospirillumbrasiliense or Azospirillum lipoferum (18). When the P02was increased, the acetylene reduction rate decreased. H.seropedicae appeared more 02 tolerant than Azospirillumbrasiliense, because a PG2 of 2 kPa completely abolished thenitrogenase activity of Azospirillum brasiliense, whereas H.seropedicae retained about 30% of its activity. Anaerobiosis

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Time (minute)FIG. 2. Inhibition of nitrogenase activity by NH4Cl in H. sero-

pedicae (A) and in Azospirillum lipoferum (B). Exponentially grownN2-fixing cells (4 ml) of H. seropedicae (0.2 kPa of dissolved 02) orAzospirillum lipoferum (0.3 kPa) were transferred to a controlledoxygen chamber, and acetylene reduction was measured at anoptimal P02. At 20 min (arrow) after the introduction of acetylene,NH4Cl was added at final concentrations of 0.2 (0), 1.0 (U), and 20(A) mM. As a control (A), the same volume of H20 was added.

or a high dissolved oxygen concentration (4.0 kPa) totallyinhibited nitrogenase activity. For the following experi-ments, the P02 level was controlled at 0.2 kPa for H.seropedicae.

Effect of combined nitrogen on nitrogenase activity in vivo.We observed the acetylene reduction rate by whole cells ofH. seropedicae and Azospirillum lipoferum under an optimalP02 to determine the effects of combined nitrogen com-pounds on their nitrogenase activity (Fig. 2). Nitrogenaseactivity of H. seropedicae was linear with time withoutammonium treatment (Fig. 2A). The introduction of NH4Cl(e.g., 0.2 mM) immediately decreased the acetylene reduc-tion rate. The inhibition by NH4CI was reversible when lowconcentrations of NH4+ were used because of the exhaus-tion of the added nitrogenous compounds. In Azospirillumlipoferum (Fig. 2B), ammonium inhibited nitrogenase activ-ity and the inhibition was complete after about 15 to 20 min.The inhibition was reversible as reported previously (17), butthe inhibition pattern was distinct in these two organisms. InH. seropedicae, NH4Cl rapidly inhibited nitrogenase activitywithout a transition period detectable under our conditions,and the inhibition was only partial. Even after the addition of20 mM NH4Cl, about 26% of the nitrogenase activity re-

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NITROGEN FIXATION IN HERBASPIRILLUM SEROPEDICAE

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Time (minute)FIG. 3. Effect of NH4CI and amino acids on nitrogenase activity

of H. seropedicae. The experiments were performed as described inthe legend to Fig. 2. At 20 min (arrow) after the introduction ofacetylene, NH4C1 (0), glutamine (a), or asparagine (A) was addedto give a final concentration of 2 mM and glutamate (+) was addedto give a final concentration of 5 mM. As a control (O), the samevolume of H20 was added.

mained. L-Methionine-DL-sulfoximine (1.0 mM), a glutaminesynthetase inhibitor, abolishes NH4+ switch off in Azospiril-lum brasiliense or Azospirillum lipoferum (17), but it did notblock ammonium inhibition of N2 fixation by H. seropedi-cae; L-methionine-DL-sulfoximine did inhibit glutamine syn-thetase activity in vitro (data not shown).

Other ammonium salts such as ammonium acetate andammonium sulfate (2 mM tested) showed inhibitory effectssimilar to those of NH4Cl, indicating that the effect was

specific to the ammonium ion rather than to a particularammonium salt (data not shown). Figure 3 shows the effectof some amino acids on nitrogenase activity in H. seropedi-cae. As with azospirilla (17) and Rhodospirillum rubrum(24), glutamine (2 mM) or asparagine (2 mM) rapidly andpartially inhibited nitrogenase activity with a pattern similarto that of ammonium inhibition. Glutamate at a concentra-tion of 5 mM had no effect on the activity of whole-cellnitrogenase during the time tested.

Effect of ammonium inhibition in vivo on subunit pattern ofdinitrogenase reductase. In Rhodospirillum rubrum or Azos-pirillum lipoferum, the inhibition of nitrogenase activity byammonium chloride results from covalent modification onone subunit of dinitrogenase reductase by the addition of anADP-ribose group (17, 24). The modified subunit can beseparated from the unmodified subunit by SDS-PAGE; itexhibits a slower-migrating band, and the protein pattern ofdinitrogenase reductase can be visualized by the immuno-blotting technique. With ADP-ribosylated dinitrogenase re-ductase from Rhodospirillum rubrum as a control, crudeextracts of H. seropedicae prepared from cells before orafter treatment in vivo for 30 min with 10 mM NH4Cl weretested to examine the subunit pattern of dinitrogenase reduc-tase. The protein from Azotobacter vinelandii was used as a

negative control, because there is no evidence for covalentmodification of its dinitrogenase reductase. Dinitrogenasereductase of H. seropedicae formed a closely migratingdoublet on the gel system used, much as did that of Azoto-

LANE 1 2 3 4 5 6 7

FIG. 4. Immunoblots of crude extracts from H. seropedicaeZ176 separated by SDS-PAGE with antiserum against dinitrogenasereductase of Azotobacter vinelandii. Extracts of N2-fixing H. sero-pedicae were prepared from 500-ml culture samples before (lanes 2,3, and 4 carrying 4, 7, and 10 ,ug of protein, respectively) and 30 minafter (lanes 5 and 6 carrying 7 and 10 ,ug of protein, respectively) theaddition of 10 mM NH4C1. As controls, lane 1 carries the highlypurified active dinitrogenase reductase of Azotobacter vinelandiiand lane 7 carries the convalently modified inactive dinitrogenasereductase of Rhodospirillum rubrum.

bacter vinelandii (Fig. 4). No change in the protein patternwas apparent after the addition of ammonium in vivo (lanes5 and 6). The pattern of dinitrogenase was not changed upontreatment with ammonium (data not shown).

Effect of ammonium inhibition in vivo on nitrogenase activ-ity in vitro. In Rhodospirillum rubrum or Azospirillum lipof-erum, the inhibition of nitrogenase activity by in vivoaddition of ammonium ions also caused an in vitro decreaseof nitrogenase activity because of the ADP-ribosylation ofdinitrogenase reductase. To test the possibility that thecovalent modification of the nitrogenase complex upon theaddition of ammonium ions causes its inhibition, we har-vested nitrogen-fixing cells of H. seropedicae before or 30min after the addition of 10 mM ammonium chloride. Crudeextracts were tested for their acetylene reduction activity;sodium dithionite was furnished as an artificial electrondonor, and an ATP-regenerating system was included in the

TABLE 1. Nitrogenase activity in crude extractsof H. seropedicae Z78

Source of crude extractsa Sp actandassayconditions (nmol of ethylene/and assay conditions m pemgbmin per mg)b

Before NH4' treatmentNo MnCl2 ............... ...................... 150.5 mM MnCI2 ..................................... 13

After NH4' treatmentNo MnCl2 ............... ...................... 150.5 mM MnCl2 ..................................... 140.5 mM MnCl2 and DRAG'.............................. 13d" The extracts were prepared from cells harvested before or 30 min after

ammonium (10 mM) treatment.b The data are from one specific experiment. The specific activity in crude

extracts was 14 to 20 nmol of ethylene formed per min per mg of protein.' 12 U of DRAG from Rhodospirillum rubrum were used in the assay. One

unit of DRAG activity is defined as the amount of enzyme required to causethe formation of 1 nmol of ethylene per min. This unit was defined in detail bySaari et al. (45). They used a partially purified nitrogenase preparation fromRhodospirillum rubrum for their assay. They disrupted Rhodospirillum ru-brum cells, centrifuged the suspension, passed the supernatant through aDEAE column, desalted the eluted material with Sephadex G-25, and storedthis. The dinitrogenase was not separated from the dinitrogenase reductase.Reductions of C2H2 were run with additions of the inactive nitrogenasepreparation, activating enzyme, ATP, MgCI2, MnCl2, Na2S204, phosphocre-atine, and creatine phosphokinase at pH 7.8. C2H2 was added, and the C2H4formed was measured.

" It is apparent that DRAG from Rhodospirillum rubrum did not increaseactivity, in contrast to its activation of covalently modified dinitrogenasereductase from Azospirillum brasiliense and Azospirillum lipoferum (34;additional unpublished data).

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FIG. 5. Detection of draG-draT homology by Southern hybrid-ization. Total DNAs of H. seropedicae Z78 and Azospirillumbrasiliense Sp7 were digested with EcoRI and hybridized to a

32P-labeled draG-draT-containing probe of Azospirillum lipoferumSpBrl7 (11). (A) DNA on the agarose gel; (B) autoradiogram.Bacteriophage lambda DNA cut with Hindlll furnishes markers anda control.

reaction mixture. Table 1 shows nitrogenase activity incrude extracts of H. seropedicae Z78. The in vitro nitroge-nase activity was not affected by the ammonium treatment(10 mM, 30 min), although the whole-cell nitrogenase activ-ity was inhibited dramatically. The cation Mn2", which isrequired for the activation of inactive dinitrogenase reduc-tase of Rhodospirillum rubrum (32), Azospirilliim brasiliense(34), or Azospirillum lipoferum (H.-A. Fu, unpublisheddata), had no stimulatory effect on H. seropedicae Z78. Italso is interesting that Mn2+ plus DRAG from Rhodospiril-lum rubrum was essentially without influence on the activityof nitrogenase from H. seropedicae, so apparently dinitro-genase reductase (as it has no ADP-ribosylated group) is nota substrate for the DRAG of Rhodospirillum rubrum. Am-monium itself had no inhibitory effect on nitrogenase in vitro(data not shown). These results suggested that the effect ofammonium chloride on nitrogenase activity in H. seropedi-cae was not directly on the nitrogenase complex.

Hybridization of genomic DNA to draT-draG probe. Theabove experimental data suggested the absence of an ADP-ribosylation system for the regulation of nitrogenase activityin H. seropedicae. However, one may argue that the expres-sion of this regulatory system may be physiologically depen-dent (24, 47, 48) and that if the attachment of the modifyinggroup is very labile, the response could be missed under theconditions tested. To confirm the results obtained here, we

searched the homologous sequences in genomic DNA with a

32P-labeled 1.1-kilobase Sall fragment probe which carriesthe draG-draT region of Azospirillum lipoferum (11). Figure5 shows the Southern hybridization pattern. Under theconditions used, no hybridization signal was observed for H.seropedicae, whereas a band of a 12-kb EcoRI fragmentappeared for Azospirillum brasiliense, an organism withdemonstrated covalent modification of its dinitrogenase re-ductase.

Intracellular ATP pool. Nitrogenase reactions are energet-ically expensive, and they theoretically consume at least 16ATP molecules for the reduction of one N2 to 2NH3 + H2(33). The availability of an ATP supply to support nitroge-nase affects its activity. Indeed, upon the addition of theuncoupler carbonyl cyanide m-chlorophenylhydrazone(CCCP), the acetylene reduction rate decreased immediately

TABLE 2. Nitrogenase activity in vivo and ATP levels duringtreatment of H. seropedicae with CCCP or NH4Cl

Time Nitrogenase activity ATP levelInhibitor (min)" (nmol of C2H4IMl (Cpmlp.Ll)bper min)

CCCP (0.05 mM) 0 18.5 9425 5.6 522

10 3.9 51620 3.6 426

NH4Cl (1 mM) 0 18.0 1,2365 4.4 1,16410 3.4 1,09220 3.0 1,278

This is the time after the addition of inhibitors.b The relative ATP level is directly expressed as counts per minute

(luminescence) per milliliter of culture when the OD580 is normalized to 1.0.Note that the CCCP treatment decreased both the nitrogenase activity and theATP level, whereas NH4CI decreased the nitrogenase activity without appre-ciable change in the level of ATP.

and the intracellular ATP pool decreased with time (Table 2).The introduction of ammonium chloride to nitrogen-fixingcells of H. seropedicae decreased the acetylene reductionrate (Table 2; Fig. 2A); however, the intracellular ATP poolwas not disturbed dramatically. Therefore, ammonium inhi-bition of nitrogenase activity is not consistently correlatedwith a change in the ATP pool; this is in agreement with thepreviously reported results for Azotobacter vinelandii (29),Rhodobacter sphaeroides (15), and Rhodospirillum rubrum(37).

DISCUSSION

H. seropedicae, like the azospirilla, fixes N2 only undermicroaerobic conditions, which suggests the absence of an02 protection mechanism for the 02-labile cellular compo-nents of nitrogenase. It therefore is important to conduct allexperiments relating to nitrogenase expression and activityat an optimal P02-

Small amounts of exogenous ammonium (e.g., 0.2 mM)rapidly and reversibly inhibited nitrogenase activity in wholecells of H. seropedicae as is true of Rhodospirillum rubrum(24) or Azospirillum lipoferum (17), in which covalent mod-ification of dinitrogenase reductase by ADP-ribosylation hasbeen demonstrated. However, the mechanisms for the inhi-bitions appear different. (i) The addition of ammoniumchloride to N2-fixing cultures of Azospirillum lipoferum orRhodospirillum rubrum gradually and completely (with atransition period) inhibited nitrogenase activity (17, 24); thissuggested the involvement of an enzymatic action during theprocess, such as the catalysis of ADP-ribosylation of dini-trogenase reductase by DRAT. In H. seropedicae, theaddition of ammonium chloride immediately, without detect-able lag, decreased whole-cell nitrogenase activity, and theinhibition was only partial (Fig. 2A). (ii) The direct covalentmodification of dinitrogenase reductase in Azospirillum li-poferum or Rhodospirillum rubrum triggered by the in vivoaddition of ammonium renders the dinitrogenase reductaseinactive both in vivo and in vitro. In H. seropedicae,nitrogenase activity was inhibited only in vivo. The activitieswere identical regardless of the sources of N2-fixing crudeextracts (i.e., whether they were ammonium treated or not),if they were tested in an in vitro system with dithionite as theartificial reductant and with an ATP-regenerating systemprovided (Table 1). (iii) In this in vitro system, the inclusion

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of Mn2+ or Mn2+ plus DRAG gave no enhancement ofnitrogenase activity. As reported previously (34) and as inour unpublished data, the activity of the covalently modifieddinitrogenase reductase of Azospirillum brasiliense andAzospirillum lipoferum could be reactivated by DRAG fromRhodospirillum rubrum in an Mn2'-dependent reaction. (iv)The low-cross-linker SDS-PAGE provides an excellent sys-tem for resolving the modified from the unmodified subunitof dinitrogenase reductase. With polyclonal antibodiesraised against dinitrogenase reductase of Azotobacter vine-landii, the dinitrogenase reductase of H. seropedicae wasvisualized on the gel as closely migrating bands such asfound for Azotobacter vinelandii; the slower-migrating bandcorresponding to the covalently modified subunit of dinitro-genase reductase from Rhodospirillum rubrum was not de-tected. (v) Under conditions exhibiting homology betweenAzospirillum brasiliense and the draG-draT probe of Azos-pirillum lipoferum, no comparable homology was detectedfor H. seropedicae. From this evidence in aggregate, it isapparent that ADP-ribosylation of dinitrogenase reductase isnot responsible for inhibition of nitrogenase activity byammonium in H. seropedicae.Because neither nitrogenase activity in vitro nor the

nitrogenase component patterns were changed after in vivotreatment with ammonium, it is reasonable to suggest thatthe ammonium effect on nitrogenase activity in H. serope-dicae is indirect. This differs from the observation forAzorhizobium caulinodans ORS571 (28). In this microorgan-ism, ammonium chloride inhibited nitrogenase activity invivo and the activity remained low in in vitro assays; noevidence for covalent modification of dinitrogenase reduc-tase has been provided.The nitrogenase reaction is an energetically expensive

process requiring ATP and a reductant. Because ADP is aninhibitor of the enzyme (33), a decrease of ATP concentra-tion or a change in the ATP/ADP ratio may influence theactivity of nitrogenase. As reported previously (15, 29, 37),ammonium inhibits nitrogenase activity in Rhodospirillumrubrum, Azotobacter vinelandii, and Rhodobacter sphaeroi-des, but no correlation has been established for inhibition byammonium and changes in the ATP pool or in the ATP/ADPratio. We found that in H. seropedicae, changes in the ATPpool also were not consistently correlated with the inhibitionof nitrogenase activity by ammonium, although CCCP, anuncoupler, decreased the ATP pool and partially inhibitedthe activity of nitrogenase. The energy charge of the organ-ism was not examined.During the reaction of the nitrogenase complex, reduced

dinitrogenase reductase transfers e- to dinitrogenase, whichcarries the site for substrate(s) reduction (33). Oxidizeddinitrogenase reductase obtains e- from the cellular electrontransport system, and the physiological reductants mostcommonly functional are ferredoxin and flavodoxin. Theactivity of the nitrogenase system is decreased by depletingthe e- supply, and the activity can be restored by supplyinga reductant (33) such as Na2S204 in vitro. The physiologicalelectron transport system to nitrogenase has been mostclearly established for Klebsiella pneumoniae (43, 44, 46).The nifJ gene product (pyruvate:flavodoxin oxidoreductase)oxidizes pyruvate and transfers two e- to two molecules ofthe nifF gene product, a flavodoxin. The flavodoxin in turnpasses e to oxidized dinitrogenase reductase. For aerobessuch as Azotobacter vinelandii, the electron transport sys-tem for nitrogenase appears to be different and is not wellunderstood. Haaker and Veeger (16) have proposed that theflow of electrons to nitrogenase is coupled to the proton

motive force of the cellular membrane. There are threeflavodoxins reported in Azotobacter vinelandii, and one ofthem may be involved in electron donation in the nitrogenasesystem (26).To explain ammonium inhibition of nitrogenase activity in

Azotobacter vinelandii, Laane et al. (29) established a cor-relation between the inhibition of nitrogenase activity and adecrease in membrane potential. They suggested that ammo-nium ions inhibit nitrogenase activity by specifically disrupt-ing the membrane potential and thus decreasing the electronflow to nitrogenase. With Azotobacter chroococcum, Cejudoand Paneque (7) observed that the inhibition of nitrogenaseby ammoniutn depends on the intracellular C/N ratio andthat N starvation decreases the sensitivity of nitrogenase toammonium. They reported that the insensitivity of nitroge-nase to ammonium action is not correlated with a change inrespiration rate. They suggested the possible modification ofdinitrogenase reductase, but no one has examined the pat-tern or activity of dinitrogenase reductase before and afterammonium treatment in vivo and in vitro with these organ-isms. Davis and Kotake (8), on the other hand, hypothesizedthat in aerobes the energized membrane will affect the levelsof bound versus free magnesium and that magnesium is thecontrolling variable for nitrogenase in vivo; they did notspecify the effect of ammonium. For the photosyntheticbacterium Rhodobacter sphaeroides, the involvement ofcovalent modification of dinitrogenase reductase or thelowering of the membrane potential have been discounted asmechanisms for ammonium inhibition (15, 50). It has beenpostulated that ammonium functions via a metabolite gener-ated by glutamine synthetase and that it acts specifically toinhibit electron transport to nitrogenase (15). Hartmann etal. (17) investigated inhibition of nitrogenase activity byammonium in the microaerobic bacterium Azospirillum am-azonense and found no evidence for covalent modification ofthe dinitrogenase reductase. As presented here, the inhibi-tion of nitrogenase by ammonium in H. seropedicae seemscompatible with uncoupling, blocking, or reallocating theelectron flow to nitrogenase. This resembles the phenome-non observed for the phenotype of electron transport mutantstrains of K. pneumoniae (44). The nifJ and nifF mutantstrains of K. pneumoniae have no or little nitrogenaseactivity in vivo but have full activity when assayed in vitrowith dithionite as the artificial electron donor. Dithionitereduces dinitrogenase reductase directly and thus bypassesthe low-potential electron donor required for N2 fixation invivo. Because we lack information on the physiologicalpathway of electron donation to nitrogenase in H. seropedi-cae, it is not justified now to propose a specific target for theammonium effect. However, glutamine or asparagine inhib-ited N2 fixation in a manner similar to that of ammonium;this may argue against a direct effect of ammonium on themembrane potential as proposed for Azotobacter vinelandii.The inability of L-methionine-DL-sulfoximine to preventinhibition by ammonium does not necessarily mean thatammonium is the direct effector, because the poor uptake ofL-methionine-DL-sulfoximine might produce the same result.To understand the mechanism for ammonium inhibition ofnitrogenase in H. seropedicae, it appears necessary to definethe electron transport pathway to nitrogenase in this organ-ism.Ammonium inhibits nitrogenase activity in many N2-fixing

organisms. However, the mechanisms of inhibition appearsto be diverse. Apparently, there are at least two generalcategories of inhibition: (i) direct covalent modification ofdinitrogenase reductase, such as in Rhodospirillum rubrum

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3174 FU AND BURRIS

and Azospirillum lipoferum, and (ii) indirect effects on thenitrogenase reaction, such as found in H. seropedicae,Azotobacter vinelandii, and Azospirillum amazonense. Aclear model of this second type of inhibition remains to beestablished.

ACKNOWLEDGMENTS

This work was supported by the College of Agricultural and LifeSciences, University of Wisconsin-Madison, and by Department ofEnergy grant DE-FG02-87ER13707.We thank P. W. Ludden, G. P. Roberts, A. Hartmann, and L. L.

Saari for enlightening discussions and generous support, W. P.Fitzmaurice for help in DNA manipulation, and J.-H. Liang forpurified nitrogenase components from Azotobacter vinelandii.

LITERATURE CITED1. Albrecht, S. L., and Y. Okon. 1980. Culture of Azospirillum.

Methods Enzymol. 69:740-749.2. Baldani, J. I., V. L. D. Baldani, L. Seldin, and J. Dibereiner.

1986. Characterization of Herbaspirillum seropedicae gen. nov,sp. nov., a root-associated nitrogen-fixing bacterium. Int. J.Syst. Bacteriol. 36:86-93.

3. Bognar, A., D. M. Libman, and E. B. Newman. 1982. Control ofnitrogenase in a photosynthetic autotrophic bacterium. Ec-tothiorhodospira sp. J. Bacteriol. 152:706-713.

4. Burris, R. H. 1972. Nitrogen fixation-assay methods andtechniques. Methods Enzymol. 24B:415-431.

5. Burris, R. H., and P. W. Wilson. 1946. Ammonia as an inter-mediate in nitrogen fixation by Azotobacter. J. Bacteriol. 53:505-512.

6. Cejudo, F. J., A. de la Torre, and A. Paneque. 1984. Short-termammonium inhibition of nitrogen fixation in Azotobacter. Bio-chem. Biophys. Res. Commun. 123:431-437.

7. Cejudo, F. J., and A. Paneque. 1988. Effect of nitrogen starva-tion on ammonium-inhibition of nitrogenase activity in Azoto-bacter chroococcum. Arch. Microbiol. 149:481-484.

8. Davis, L. C., and S. Kotake. 1980. Regulation of nitrogenaseactivity in aerobes by Mg++ availability: an hypothesis. Bio-chem. Biophys. Res. Commun. 93:934-940.

9. D6bereiner, J., and F. 0. Pedrosa. 1987. Nitrogen-fixing bacteriain non-leguminous crop plants. Science Tech Publishers/Spring-er-Verlag, Madison, Wis.

10. Feinberg, A. P., and B. Vogelstein. 1983. A technique forradiolabeling DNA restriction endonuclease fragments to highspecific activity. Anal. Biochem. 137:266-267.

11. Fu, A.-H., W. P. Fitzmaurice, L. J. Lehman, G. P. Roberts, andR. H. Burris. 1988. Regulation of nitrogenase activity in azos-pirilla, herbaspirilla and acetobacter; and cloning of draG- anddraT-homologous genes of A. lipoferum SpBr17; p. 336. In H.Bothe, F. J. de Bruijn, and W. E. Newton (ed.), Nitrogenfixation: hundred years after. Gustav Fischer, Stuttgart.

12. Goa, J. 1953. A micro biuret method for protein determination.Scand. J. Clin. Lab. Invest. 5:218-222.

13. Gordon, J. K., V. K. Shah, and W. J. Brill. 1981. Feedbackinhibition of nitrogenase. J. Bacteriol. 148:884-888.

14. Gotto, J. W., and D. C. Yoch. 1985. Regulation of nitrogenaseactivity by covalent modification in Chromatium vinosum.Arch. Microbiol. 141:40-43.

15. Haaker, H., C. Laane, K. Hellingwerf, B. Houwer, W. N.Konings, and C. Veeger. 1982. Short-term regulation of thenitrogenase activity in Rhodopseudomonas sphaeroides. Eur. J.Biochem. 127:639-645.

16. Haaker, H., and C. Veeger. 1977. Involvement of the cytoplas-mic membrane in nitrogen fixation by Azotobacter vinelandii.Eur. J. Biochem. 77:1-10.

17. Hartmann, A., H.-A. Fu, and R. H. Burris. 1986. Regulation ofnitrogenase activity by ammonium chloride in Azospirillum spp.J. Bacteriol. 165:864-870.

18. Hartmann, A., H.-A. Fu, S.-D. Song, and R. H. Burris. 1985.Comparison of nitrogenase regulation in A. brasiliense, A.lipoferum and A. amazonense, p. 116-126. In W. Klingmuller(ed.), Azospirillum III: genetics, physiology, ecology. Springer-

Verlag, Berlin.19. Hillmer, P., and K. Fahlbusch. 1979. Evidence for an involve-

ment of glutamine synthetase in regulation of nitrogenase activ-ity in Rhodopseudomonas capsulata. Arch. Microbiol. 122:213-218.

20. Hochman, A., and R. H. Burris. 1981. Effect of oxygen onacetylene reduction by photosynthetic bacteria. J. Bacteriol.147:492-499.

21. Howard, K. S., B. J. Hales, and M. D. Socolofsky. 1983.Nitrogen fixation and ammonia switch-off in the photosyntheticbacterium Rhodopseudomonas viridis. J. Bacteriol. 155:107-112.

22. Jones, B. L., and K. J. Monty. 1979. Glutamine as a feedbackinhibitor of the Rhodopseudomonas sphaeroides nitrogenasesystem. J. Bacteriol. 139:1007-1013.

23. Jouannealu, Y., C. M. Meyer, and P. M. Vignais. 1983. Regula-tion of nitrogenase activity through iron protein interconversioninto an active and an inactive form in Rhodopseudomonascapsulata. Biochim. Biophys. Acta 749:318-328.

24. Kanemoto, R. H., and P. W. Ludden. 1984. Effect of ammonia,darkness, and phenazine methosulfate on whole-cell nitroge-nase activity and Fe protein modification in Rhodospirillumrubrum. J. Bacteriol. 158:714-720.

25. Klugkist, J., and H. Haaker. 1984. Inhibition of nitrogenaseactivity by ammonium chloride in Azotobacter vinelandii. J.Bacteriol. 157:148-151.

26. Klugkist, J., J. Voorberg, H. Haaker, and C. Veeger. 1986.Characterization of three different flavodoxins from Azotobac-ter vinelandii. Eur. J. Biochem. 155:33-40.

27. Krieg, N. R., and P. S. Hoffman. 1986. Microaerophily andoxygen toxicity. Annu. Rev. Microbiol. 40:107-130.

28. Kush, A., C. Elmerich, and J.-P. Aubert. 1985. Nitrogenase ofSesbania rhizobium strain ORS571: purification, properties and"switch-off' by ammonia. J. Gen. Microbiol. 131:1765-1777.

29. Laane, C., W. Krone, W. Konings, H. Haaker, and C. Veeger.1980. Short-term effect of ammonium chloride on nitrogenfixation by Azotobacter vinelandii and of bacteroids of Rhizo-bium leguminosarum. Eur. J. Biochem. 103:39-46.

30. Laemmli, U. K. 1970. Cleavage of structural proteins during theassembly of the head of bacteriophage T4. Nature (London)227:680-685.

31. Lowery, R. G., L. L. Saari, and P. W. Ludden. 1986. Reversibleregulation of the nitrogenase iron protein from Rhodospirillumrubrum by ADP-ribosylation in vitro. J. Bacteriol. 166:513-518.

32. Ludden, P. W., and R. H. Burris. 1976. Activating factor for theiron protein of nitrogenase from Rhodospirillum rubrum. Sci-ence 194:424-426.

33. Ludden, P. W., and R. H. Burris. 1986. Nitrogenase: propertiesand regulation, p. 41-58. In C. A. Neyra (ed.), Biochemicalbasis of plant breeding, vol. 2, CRC Press, Inc., Boca Raton,Fla.

34. Ludden, P. W., Y. Okon, and R. H. Burris. 1978. The nitroge-nase system of Spirillum lipoferum. Biochem. J. 173:1001-1003.

35. Maniatis, T., E. F. Fritsch, and J. Sambrook. 1982. Molecularcloning: a laboratory manual. Cold Spring Harbor Laboratory,Cold Spring Harbor, N.Y.

36. Murrell, J. C. 1988. The rapid switch-off of nitrogenase activityin obligate methane-oxidizing bacteria. Arch. Microbiol. 150:489-495.

37. Paul, T. D., and P. W. Ludden. 1984. Adenine nucleotide levelsin Rhodospirillum rubrum during switch-off of whole-cell nitro-genase activity. Biochem. J. 224:961-969.

38. Polman, J. K., and J. M. Larkin. 1988. Properties of in vivonitrogenase activity in Beggiatoa alba. Arch. Microbiol. 150:126-130.

39. Pope, M. R., S. A. Murrell, and P. W. Ludden. 1985. Covalentmodification of the iron protein of nitrogenase from Rhodospiril-lum rubrum by adenosine diphosphoribosylation of a specificarginyl residue. Proc. Natl. Acad. Sci. USA 82:3173-3177.

40. Postgate, J. R., H. M. Kent, S. Hill, and H. Blackburn. 1984.Nitrogen fixation by Desulfovibrio gigas and other species ofDesulfovibrio, p. 225-234. In P. W. Ludden and J. E. Burfis(ed.), Nitrogen fixation and CO2 metabolism. Elsevier Science

J. BACTERIOL.

on February 26, 2020 by guest

http://jb.asm.org/

Dow

nloaded from

NITROGEN FIXATION IN HERBASPIRILLUM SEROPEDICAE 3175

Publishing, Inc., New York.41. Privalle, L. S., and R. H. Burris. 1983. Adenine nucleotide

levels in and nitrogen fixation by the cyanobacterium Anabaenasp. strain 7120. J. Bacteriol. 154:351-355.

42. Radionov, Y. V., N. V. Lebedeva, and E. N. Kondratieva. 1986.Ammonia inhibition of nitrogenase activity in purple and green

bacteria. Arch. Microbiol. 143:345-347.43. Roberts, G. P., and W. J. Brill. 1981. Genetics and regulation of

nitrogen fixation. Annu. Rev. Microbiol. 35:207-235.44. Roberts, G. P., T. MacNeil, D. MacNeil, and W. J. Brill. 1978.

Regulation and characterization of protein products coded bythe nif (nitrogen fixation) genes of Klebsiella pneumoniae. J.Bacteriol. 136:267-279.

45. Saari, L. L., E. W. Triplett, and P. W. Ludden. 1984. Purifica-tion and properties of the activating enzyme for iron protein ofnitrogenase from the photosynthetic bacterium Rhodospirillumrubrum. J. Biol. Chem. 259:15502-15508.

46. Shah, V. K., G. Stacey, and W. J. Brill. 1983. Electron transport

to nitrogenase. J. Biol. Chem. 258:12064-12068.47. Sweet, W. J., and R. H. Burris. 1981. Inhibition of nitrogenase

activity by NH4' in Rhodospirillum rubrum. J. Bacteriol. 145:824-831.

48. Sweet, W. J., and R. H. Burris. 1982. Effects of in vivotreatments on the activity of nitrogenase isolated from Rhodos-pirillum rubrum. Biochim. Biophys. Acta 680:17-21.

49. Yoch, D. C., and J. W. Gotto. 1982. Effect of light intensity andinhibitors of nitrogen assimilation on NH4' inhibition of nitro-genase activity in Rhodospirillum rubrum and Anabaena sp. J.Bacteriol. 151:800-806.

50. Yoch, D. C., J.-D. Li, C.-Z. Hu, and C. Scholin. 1988. Ammoniaswitch-off of nitrogenase from Rhodobacter sphaeroides andMethylosinus trichosporium: no evidence for Fe protein modi-fication. Arch. Microbiol. 150:1-5.

51. Zumft, W. G., and F. Castillo. 1978. Regulatory properties ofthe nitrogenase from Rhodopseudomonas palustris. Arch. Mi-crobiol. 117:53-60.

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