sadasivan and neyra- 1985

JOURNAL OF BACTERIOLOGY, Aug. 1985, p. 716-7230021-9193/85/080716-08$02.00/0Copyright C 1985, American Society for Microbiology

Vol. 163, No. 2

Flocculation in Azospirillum brasilense and Azospirillum lipoferum:Exopolysaccharides and Cyst Formation

LAKSHMI SADASIVAN AND CARLOS A. NEYRA*Department ofBiochemistry and Microbiology, New Jersey Agricultural Experiment Station, Cook College, Rutgers

University, New Brunswick, New Jersey 08903

Received 25 February 1985/Accepted 22 May 1985

The phenomena of flocculation and floc formation by Azospirillum brasilense Sp7 (ATCC 29145) andAzospirillum lipoferum Sp59b (ATCC 29707) were studied in aerobic liquid cultures. Carbon sourcesrepresentative of various entry pathways in combination with various nitrogen sources induced flocculation inboth species of azospirilla. Noticeably, the combination of fructose and nitrate was the most effective in termsof floc yields. Phase-contrast microscopic observations revealed a transition in cell morphology from freelymotile, vibrioid cells to nonmotile, highly refractile encysting forms during the formation of flocs. Thenonmotile forms in flocs appeared to be entangled within a fibrillar matrix, and the cells were highly resistantto desiccation. Dried flocs kept for almost 6 months still tnaintained the highly refractile encysting forms, andtheir viability was confirmed by pellicle formation and acetylene reduction in semisolid malate medium.Electron microscopic observations of the desiccated flocs revealed the presence of cell forms containingabundant poly p-hydroxybutyrate granules within a central body and surrounded by a thick layer ofexopolysaccharides. The latter were characterized by alkali and acid digestion, crude cellulase hydrolysis, andcalcofluor staining. It was concluded that the overproduction of exocellular polymers induces the flocculentgrowth and is associated with the concomitant transformation of vegetative cells to the desiccation-resistantencysting forins under limiting cultural conditions.

Azospirillum spp. have been shown to be a commoninhabitant of tropical and temperate soils and have also beenshown to reduce acetylene in culture media and in associa-tion with the roots of forage grasses, cereals, and otherplants (11, 19, 26). A. brasilense and A. lipoferum are knownto grow well in culture media on a variety of carbon sources,including various sugars and organic acids (15, 24, 30, 32).These organisms, under aerobic conditions, also grow wellon a variety of combined nitrogen sources, including ammo-nia, nitrate, or amino acids (20, 21, 22, 24).

Recently, special attention has been given to the use andmetabolism of fructose by strains of A. brasilense and A.lipoferum grown with N2 or NH4Cl as the nitrogen source (4,9, 15, 32). During recent investigations in our laboratory onthe various carbon sources for nitrate-dependent growth byazospirilla, we observed intense flocculation in media con-taining fructose and nitrate. A medium containing KNO3 andglucose has been reported to enhance exopolysaccharidesynthesis in certain soil bacteria (14). Also, flocculation inliquid culture medium has been reported for several gram-negative bacteria, but azospirilla were not included in thosestudies (5, 8, 16). Thus, this research was undertaken tostudy the following: (i) the optimization of cultural condi-tions to achieve maximum flocculation, (ii) the interactionbetWeen carbon and nitrogen sources to bring about floccu-lation, and (iii) the ultrastricture and chemical characteriza-tion of the flocs. (In this paper, "floc formation" refers to theaggregation of cells during growth, and "flocculation" refersto the clumping of cells in a turbid suspension.)

MATERIALS AND METHODSBacterial strains and culture media. A. brasilense Sp7

(ATCC 29145) and A. lipoferum Sp59b (ATCC 29707) wereused throughout this study. These strains were maintained on

* Corresponding author.

nutrient agar (Difco Laboratories, Detroit, Mich.) slants at300C.Minimal salts medium, described as medium A by Neyra

and Van Berkum (21), was prepared as follows to avoidprecipitation. Salts other than organic carbon, inorganicnitrogen, or phosphate salts for 1 liter of medium weredissolved in 10 ml of distilled water and autoclaved sepa-rately. After cooling, it was added to 940 ml of sterile KPO4buffer (100 mM; pH 6.8). Organic acids and sugars (at aconcentration range between 8 and 100 mM) as the carbonsource and various inorganic nitrogen sources (viz., NH4Cl,KNO3, and glutamate) at concentration ranges between 0.5and 20 mM were filter sterilized and added individually asrequired for various experiments. The final volume of themedium was then made up to 1 liter.Inoculum for all experiments was harvested from a log-

phase culture grown in nutrient broth (NB; Difco) by cen-trifugation at 5,000 rpm for 10 min at 40C, washed three timeswith equal volumes of 100 mM KPO4 buffer (pH 6.8), andinoculated into test medium to an initial optical density at660 nm (OD6w) of '0.1 for floc formation experiments and>0.2 to 0.4 for flocculation experiments. (An OD6w of 0.8corresponds to 5 x 109 CFU/ml for both strains.)

Experiments were conducted either in 250-ml conicalflasks containing 100 ml of media or in large batch cultureswith 5-liter bottles containing 4 liters of media. Small flaskcultures were incubated on a Gyrotory shaker (NewBrunswick Scientific Co., Inc., New Brunswick, N.J.) at 200rpm and kept in a chamber with temperature control at 34° C.The large batch cultures were incubated with vigorous airsparging at 34° C in a water bath.

Phase-contrast and fluorescence microscopy. Flocs wereallowed to develop by flocculation experiments in minimalsalts medium containing 8 mM fructose and 0.5 mM KNO3.After 72 h, the supematant was decanted, and the settledflocs were washed several times with distilled water and

716

EXOPOLYSACCHARIDES AND CYST FORMATION IN AZOSPIRILLA

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FIG. 1. Growth curves and floc formation ofA. brasilense Sp7 (ATCC 29145) in NB cultures provided with (0) or without (0)5 mM KNO3(A) or minimal salts media containing 5 mM KNO3 and 40 mM malate (0), 4 mM malate (0), 27 mM fructose (A), or 8 mM fructose (A) (B).Floc formation was only observed in the fructose treatments.

photographed either directly on a petri dish or under a Nikonphase-contrast microscope.

Fresh flocs were stained with 0.025% calcofluor (Ameri-can Cyanamid Co., Pearl River, N.Y.) which yields fluores-cence staining of cellulose and other P-linked polysac-charides (16). Stained preparations were examined under along-wavelength UV lamp and Nikon epifluorescence micro-scope with a UV excitation filter at 330 to 380 nm. TheNB-grown cells were pelleted and stained with calcofluor toserve as the control.

Quantification of flocs. The net weight of flocs was ob-tained by filtering the flocs through filter paper (Whatmanno. 1) and weighing after the paper and flocs were air-driedfor 30 min. For the dry weight measurements, the flocs weredried in a desiccator oven at 80° C for 2 h.

Viability test and phase-contrast microscopy of desiccatedflocs. The dried flocs obtained as described above werestored in a closed vial at room temperature (30° C) for up to6 months. A small piece of the dried floc was transferred tothe semisolid nitrogen-free malate medium (6) and incubatedat 34° C for 24 to 48 h to observe growth, pellicle formation,and acetylene reduction. The dry flocs of A. brasilense andA. lipoferum were also photographed under a phase-contrastmicroscope after imbibition of the flocs in KPO4 buffer (pH6.8; 100 mM) for 10 min.

Ultrastructural studies of desiccated flocs. The dried flocswere fixed in 1% glutaraldehyde for 4 h, followed by 1%osmium tetroxide fixation for 10 h, dehydrated in a series ofgraded ethyl alcohol solutions, and finally embedded inSpurr plastic as described by Cole and Popkin (3). Ultrathinsections were cut in an ultramicrotome and observed undera transmission electron microscope (Siemen model 1AElmiscope).Chemical characterization of floes. (i) Alkali and acid treat-

ment. A 500-mg (fresh weight) sample of 72-h-old flocs wastreated with 10 ml of 1 N NaOH for 1 h at 100° C to dissolvethe cellular proteins and nucleic acids (5). The residue(NaOH-insoluble fraction) was further treated with 5 ml of0.66 N HCl for 2 h at 100° C to eliminate noncellulosicglucans (5). The remaining residue was treated with 2 ml of6 N HCl for complete hydrolysis. Later, dry weights of the

insoluble fractions of 1 N NaOH and 0.66 N HCl hydrolysisof the flocs obtained from a 5-liter culture in 8 mM fruc-tose-0.5 mM KNO3 were also recorded.

(ii) Cellulase digestion of cellulosic material. The NaOH-insoluble residue was suspended in 2 ml of 0.05 M sodiumcitrate buffer (pH 4.5), to which was added 20 U of crudecellulase (EC 3.2.1.4) from Trichoderma viridie (type V,Sigma Chemical Co., St. Louis, Mo.) and incubated at 45° Cfor 12 h. The reducing sugars released were measured by thecolorimetric method of Nelson-Somogyi (18).

RESULTSCultural conditions for floc formation and flocculation. (i)

Floc formation. Cultures of A. brasilense (ATCC 29145),growing in NB with or without nitrate, showed normalgrowth kinetics. The turbidity (absorbance at 660 nm) in-creased in a sigmoidal fashion as the time of incubationprogressed (Fig. 1A). Flocs were never observed in NBcultures. The doubling time during the log phase of thecultures was between 1.5 and 2 h. Similar growth patternswere also obtained after transferring washed NB-grown cellsinto a medium containing 40 mM malate as the sole carbonsource and 5 mM KNO3 as the nitrogen source to an initialODw0 of <0.1 (Fig. 1B). However, the cells did not grow tothe log phase in a medium containing fructose and nitrate.Instead, the inoculum grew only up to a period of 6 h andafter reaching an OD6w of 0.2, the cells started aggregating,and no further increase in turbidity could be measured due tointerference by the flocs (Fig. 1B). In the case of malate andKNO3 medium, trace quantities of flocs began to appearafter 10 to 20 h of incubation.

(ii) Flocculation. The phenomenon of flocculation alsooccurs in the case of Azospirillum cultures. When thewashed cells of a log-phase culture grown in NB wastransferred to a minimal medium containing (8 mM) fructoseand (0.5 mM) KNO3 to an initial turbidity of >0.2, theinoculum, instead of growing, flocculated immediatelywithin 2 to 4 h, leaving behind a clear supernatant (Fig. 2A).

Optimization of media condition for floc formation andflocculation. The conditions for floc formation and floccula-tion were initially optimized by visual assessment of the flocs

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A B

FIG. 2. (A) Flocculation of A. lipoferum (ATCC 29707) in minimal salts medium containing fructose (8 mM) and KNO3 (0.5 mM). (B) Themacroscopic appearance of the flocs is more clearly presented after separation from the rest of culture contents and placement on a petriplate.

appearing in the media containing various concentrations ofcarbon and nitrogen sources and by measuring the pH of themedium. Sugars (fructose and glucose) were found to yieldmore flocs than the organic acids (Table 1). In a secondexperiment, a representative from the organic acids (malate)and one from sugars (fructose) were chosen to study theeffects of various concentrations of carbon sources on flocformation and flocculation. Carbon source concentrations of8 mM yielded better flocs in fructose medium than in malatemedium (Table 2). There were no significant pattern differ-ences in floc formation or flocculation by changing theconcentrations of carbon sources. However, in a mediumlacking carbon, the inoculum remained suspended freely,and the cells were vibrioid with impaired motility andreceding cytoplasm. Table 3 gives the effects of nitrogensources on floc formation and flocculation. A medium con-taining KNO3 yielded more flocs than did any other nitrogensource. The formation of flocs did not appear to be pHdependent, since a medium supplemented with NH4Cl de-creased in pH to between 5.9 and 6.4 (acidic) and yieldedless flocs than did a medium with NH4NO3 whose pHremained between 6.9 and 7.1, which is about the optimumpH at which azospirilla grow.

Floc formation and flocculations were also compared in amedium with 8 mM fructose and increasing concentrationsof KNO3 ranging between 0.5 and 20 mM. It was observedthat all concentrations (.s5 mM, 10 mM, 15 mM, and 20 mM)induced flocculation within 2 h of incubation at 34° C. In a

period of 6 to 8 h, maximum settling of the flocs occurred at<5 mM; at 10, 15, and 20 mM, however, there was turbidityalong with flocs. In the floc formation experiments, flocsbegan to appear after the OD660 or absorbance at 660 nm ofthe culture medium increased to ca. 0.4 in a ca. 6- to 8-hperiod. Increasing the fructose and KNO3 concentrations to40 and 20 mM, respectively, resulted in more turbidity in thesupernatant, indicating growth, although flocs were alsopresent. Flocs also appeared when the concentration offructose was increased to 100 mM.

Floc yields were later quantified by measuring the freshand dry weights of the flocs obtained through flocculationwith 8 mM malate and fructose as carbon sources and 5 mMNH4Cl lacking nitrate and KNO3 as nitrogen sources (Table4). The floc yield was greatly reduced when NH4Cl was usedin combination with malate, whereas fructose and nitrateyielded maximum flocculation.

Macroscopic appearance of flocs. The massive flocculationof A. lipoferum (ATCC 29707) and its tendency to settledown is clearly evident in Fig. 2A. For this experiment,bacterial cells were inoculated into a large batch aerobicculture containing 8 mM fructose and 0.5 mM KNO3. Figure2B is a close-up of the flocs, showing the slimy and curdlikeaggregation. Both A. brasilense and A. lipoferum yieldedsimilar types of flocs.

Phase-contrast microscopy of developing flocs. The micro-scopic appearance of the vegetative cells was found to

TABLE 1. Floc formation and flocculation in A. brasilense andA. lipoferum in different carbon sources and KNO3 (0.5 mM) as

the nitrogen sourceFloc formation and flocculation in b:

Carbon source"A. brasilense A. lipoferum

NBMalate + +Gluconate + + + + + +a-ketoglutarate + +Fructose + + + + + + + +Glucose + + +

a Filter-sterilized carbon sources were added at 20 mM C. in minimal saltsmedium.

b Symbols: (-) no growth, (+) little floc, (+ +) fair, (+ + +) good, and(+ + + +) excellent.

TABLE 2. Effect of various concentrations of carbona sources inminimal salts medium on floc formation and flocculation in A.

brasilense and A. lipoferum with KNO3 (0.5 mM) asthe nitrogen source

Effect of:

Concn (mM) Malate Fructose

A. A. A. A.brasilense lipoferum brasilense lipoferum

0 - - _ -

8 ++ +++ ++++ ++++20 + ++ +++ +++40 + ++ +++100 ++ +++ +++ +++

aA representative from the organic acids (malate) and one from sugars(fructose) were chosen for this study. Symbols: (-) no flocs, (+) little floc,(+ +) fair, (+ + +) good, and (+ + + +) excellent.

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FIG. 3. Microscopic characteristics of flocs as seen under a phase-contrast microscope. Flocs of A. lipoferum (ATCC 29707) were takenat different stages during the flocculation process. (A) Vegetative cells from 24-h-old NB cultures. (B) Aggregation of cells 24 h after transferto minimal salts medium containing fructose (8 mM) and KNO3 (0.5 mM). The vegetative cells look entrapped within a fibrillar mesh. (C)Forty-eight-hour-old flocs showing a more compact mesh containing polymorphic cells. (D) One-week-old flocs showing highly refractile cells.Similar observations in A. brasilense were also photographed. Bar, 10 ,um.

change upon flocculation (Fig. 3). Cells grown in NB ap-peared as actively motile curved rods and did not form anykind of aggregation (Fig. 3A). The cells (absorbance at 660nm, 0.2 to 0.4), when transferred from NB to minimalmedium containing fructose (8 mM) and KNO3 (0.5 mM),lost their characteristic motility and assumed a more oval tospherical shape within a period of 2 to 4 h (Fig. 3B and 3C).These nonmotile cells appeared to be entangled in a mesh offibrillar material which was not easily separable, even afterhard pressing between slide and cover slip. Flocs observedafter 1 week of culture were arranged as a very compactaggregate of refractile cells, a very different configurationfrom that observed with fresh flocs 24 h after inoculation ortransfer (Fig. 3D). Thus, cell types, metabolic activity, andpatterns of aggregation also appear to change with cultureage under conditions conducive to flocculation.

TABLE 3. Effect of various nitrogen sources on floc formationand flocculation with fructose (8 mM) as the carbon source in

minimal salts mediumFloc formation and flocculation inb:

Nitrogen sourceaA. brasilense A. lipoferum

NH4Cl + (5.9) + (6.4)KNO3 + + + (8.2) + + + (7.5)NH4NO3 + + (6.9) + + (7.1)Glutamate + (8.6) + + (7.9)a Nitrogen sources were added at 5 mM N.b Numbers in parentheses are the pHs of the medium after 24 h at 34° C.

Symbols: (+) little flock, (+ +) good, and (+ + +) excellent.

Viability and phase-contrast microscopy of desiccated flocs.Desiccated flocs maintained for up to 6 months were foundto be viable after transfer into semisolid malate medium andincubation for 48 h at 37° C. The typical subsurface pellicleformed was found to reduce acetylene to ethylene.The dry flocs under a phase-contrast microscope revealed

the presence of encysting forms which were still highlyrefractile (Fig. 4A and B). The refractile forms were ob-

TABLE 4. Effect of carbona and nitrogen sources on floc yieldsafter flocculation by A. brasilense and A. lipoferum

Floc yield (g/liter)bTreatment Culture pH

Fresh wt Dry wt

A. brasilenseFructose +NH4Cl 2.5 0.15 3.2Fructose + KNO3 4.7 0.64 6.2

Malate + NH4Cl TR TR 8.2Malate + KNO3 0.025 TR 7.7

A. lipoferumFructose + NH4Cl 2.0 0.2 5.5Fructose + KNO3 6.0 1.18 6.2

Malate + NH4Cl TR TR 7.5Malate + KNO3 TR TR 7.5a Fructose or malate was added as the carbon source at 8 mM, and KNO3

or NH4C1 was the nitrogen source at 5 mM in minimal salts medium.b Flocs were harvested for weight determinations ater 24 h. TR, Traces.

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FIG. 4. Phase-contrast microscopic observation of the dried flocs after 6 months of desiccation. Highly refractile encysting forms of A.brasilense (A) and A. lipoferum (B). The encysting forms of A. brasilense (C) are seen embedded within a matrix (arrows). (D) Mold of theextracellular material after the encysting forms were gently tapped out. [A. lipoferum revealed similar structures to those shown in (C) and(D)]. Bar, 10 ,um.

served to be embedded inside a honeycomb-like network ofextracellular matrix (Fig. 4C, arrows). Figure 4D shows thecaste of the material after the central resting cell wasremoved by gentle tapping.

Ultrastructure of desiccated flocs. The ultrastructures of A.brasilense are shown in Fig. 5a, c, and e, and the ultra-structures ofA. lipoferum are shown in Fig. 5b, d, and f. Thevegetative cells (Fig. 5a and b) are vibrioid, with minimalquantities of refractile granules which are reported to be polyf-hydroxybutyrate (PHB) (23). The highly refractile centralbodies and the transformed vegetative cells in the floc areshown in Fig. 5e and d, respectively. These forms are verydifferent from vegetative cells in the properties of nonmotil-ity and enlargement into a round, central body with abun-dant PHB granules. The central body is surrounded by thickcapsular material. Ultrathin sections of the dry flocs (Fig. 5eand f) show a network of extracellular matrix tightly holdingtogether the encysting cell forms.

Chemical characteristics of flocs. (i) Alkali and acid hydro-lysis. Treatment of flocs with 1 N NaOH yielded twofractions, one NaOH soluble and one NaOH insoluble. Thesoluble fraction was presumed to contain proteins and nu-cleic acids, and the insoluble fractions, which were 300 and600 mg (dry weight)/5 liter of A. brasilense and A. lipoferum,respectively, were expected to contain the exocellularpolysaccharides. The treatment of NaOH-insoluble fractionswith 0.66 N HCI at 100° C for 2 h possibly removed alla-glucans and left granular and fibrillar residues whose dryweights were 2 and 4 mg/5-liter culture of A. brasilense and

A. lipoferum, respectively. This residue was completelyhydrolyzed by treatment with 6 N HCI.

(ii) Cellulase digestion of alkali-insoluble fraction. Table 5gives the quantities of reducing sugars released during thecellulase digestion as measured by the Nelson-Somogyimethod (18). It was also observed that crude cellulase failedto disperse untreated flocs, indicating that cellulose was notthe sole component involved in flocculation but that otherpolysaccharides were involved. However, the insolubleNaOH residue did disperse significantly but not completely,leaving behind reducing sugars and trace amounts of precip-itates which could be completely hydrolyzed in 6 N HCl.

(iii) Calcofluor staining of wet flocs. To complement ourchemical characterization of flocs, 72-h-old floc, whenstained with 0.025% calcofluor dye, exhibited fluorescencein a UV lamp, and the same was later confirmed with anepifluorescence microscope. The NB-grown cells did notfluoresce after staining with calcofluor (data not shown).

DISCUSSIONThe results presented throughout this report have shown

conclusively the phenomena of floc formation and floccula-tion in cultures of A. brasilense (ATCC 29145) and A.lipoferum (ATCC 29707). We have provided evidence for thecultural conditions required to consistently bring aboutmassive flocculation yielding exocellular polymers in liquidcultures of these bacteria.

Malate has been the preferred carbon source for nitrate-dependent growth of these bacteria (2, 13, 21). Our attempts

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..a mmb

FIG. 5. Ultrastructure of vegetative cells and desiccated flocs ofA. brasilense (a, c, and e) and A. lipoferum (b, d, and f). (a and b) Vibrioidrods with few PHB granules. (c and d) Single encysting cells from the desiccated flocs after 6 months, showing a change in cell morphologyto round forms with abundant PHB granules and surrounded by a thick layer of polysaccharide. (e and f) Ultrastructures of the desiccatedflocs, showing the compact aggregations of encysting forms within a mesh of exopolysaccharides. Bar, 0.5 p.m.

to use fructose instead of malate failed to produce a normalgrowth proffle in Azospirillurn cultures in a nitrate-containingmedium (see above). A minimal medium containing fructoseand KNO3 favors floc formation and flocculation in A.brasilense and A. lipoferum cultures and is better thanmalate and NH4Cl. The differential effects on flocculation bydiverse carbon and nitrogen sources are not fully understoodat present, but they are believed to be connected with theproduction of exocellular polymers, particularly n-linkedpolysaccharides.

TABLE 5. Hydrolysis of the NaOH (1 N)-insoluble fraction bycrude cellulase

Cellulose substrate (mg) Amt of Reducing sugarsenzymes (U) produced (mg)aPure cellulose (10)b 0 0Pure cellulose (10)b 20 0.51 N NaOH residue (500) 0 01 N NaOH residue (500) 20 1.38

a After 12.h at 45C.b Whatman cellulose powder (W. & R. Balston, Ltd.).

The capsule formation and the production of exocellularpolysaccharides are the prerequisites for the development ofa mature cyst (27). The exines of the Azotobacter cysts areknown to be composed of basic units of polysaccharidescomplexed with divalent cations (7, 28). The capsular forms(C forms) of azospirilla reported by Berg et al. (1) wereconsidered to be active, nitrogen-fixing cells modified toprotect the 02-sensitive nitrogenase enzyme to facilitatenitrogen fixation (1). These C forms were obtained on alimiting carbon and nitrogen medium (2 g/liter concentrationof malate and 0.5 mM KNO3) and ultrastructurally aresimilar to the dry flocs reported in this paper. The C formswere not believed to be resting structures due to the lack ofcharacteristic exines and intines of a mature cyst as de-scribed in Azobacter spp.; hence, no desiccation experi-ments were conducted (1). However, we have shown thatthe flocs were resistant to desiccation for up to 6 months andare still viable. By definition, the cysts are the transformedvegetative cells containing abundant PHB granules with ahigh degree of resistance to desiccation (27, 28). LikeAzotobacter spp., azospirilla have also been isolated from15-year-old dry soil samples (10, 31). Encysted forms of

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azospirilla obtained by various cultural conditions are foundto be resistant to desiccation from several hours to 1 month(11, 25). There are reports on Azotobacter spp. on theinduction of cyst formation, depending upon the carbon andnitrogen source (27). In a liquid culture medium, the lack ofdevelopment of exine and intine had been reported inAzotobacter spp. (12). However, the induction of exineformation was achieved by the addition of the flocculatingdivalent cations such as CaNO3 (29). Thus, the lack ofobservation of a definite exine and intine in Azospirillum sp.may be a matter of identifying the specific chemical agentsand cultural conditions for their induction. The report hereof the desiccation-resistant forms in flocs suggests that thebacteria, upon transfer from complete nutrient medium to anutritionally minimal medium, immediately undergo meta-bolic changes and synthesize the polysaccharides which arenecessary for encystation during desiccating conditions.

Also, extracellular polymers such as polysaccharides andcellulose microfibrils in flocs have been reported to play apossible role in the infection process in the rhizosphere andcell suspension cultures of carrots by causing aggregation ofthe infecting bacterial cells (16, 17). In this report, we haveshown that the polysaccharides may also be playing a role inthe survival mechanism of these bacteria under stress con-ditions such as desiccation and nutritional limitation. It maybe possible that if the conditions in nature in the rhizospherewere favorable for infection, e.g., a microaerophilic condi-tion and presence of suitable carbon and nitrogen sources inthe form of root exudates, then the aggregated cells mightshift their metabolic activity from encystation to germinationand adsorption of the roots. This, however, is yet to beconfirmed.

This paper also demonstrates that azospirilla can be in-duced to produce extracellular polysaccharides by manipu-lating the culture medium, and thus it will enable futureresearchers to identify and screen for strains varying in theirexopolysaccharides and, thereby, the nature of aggregationand adsorption.

ACKNOWLEDGMENTSWe thank Robert L. Tate III, Department of Soils and Crops,

Rutgers University, for allowing us the use of microscopic facilitiesfor the phase-contrast and epifluorescence microscopy. We alsothank R. Triemer and Laura Wood for their collaboration andtechnical assistance in the electron microscopic studies at NelsonBiological Laboratories, Rutgers University. We thank AliceMontana, Madeline Mendoza, and Sylvia Taylor for assistance in thepreparation of this manuscript.

This work, publication no. D-01204-01-85, was supported by theNew Jersey Agricultural Experiment Station, which is supported bystate and U.S. Hatch Act funds. Partial support was also providedby the New Jersey Agricultural Experiment Station project no.04400 on rhizosphere research. L.S. is a visiting scientist andpostdoctoral fellow supported partially by funds provided by theNew Jersey Agricultural Experiment Station, Department of Bio-chemistry and Microbiology, and the International Agriculture andFood Programs at Cook College, Rutgers University.

LITERATURE CITED1. Berg, R. H., M. E. Tyler, N. J. Novick, V. Vasil, and I. K. Vasil.

1980. Biology of Azospirillum-sugarcane association: enhance-ment of nitrogenase activity. Appl. Environ. Microbiol.39:642-649.

2. Bothe, H., B. Klein, M. P. Stephan, and J. Dobereiner. 1981.Transformations of inorganic nitrogen by Azospirillum spp.Arch. Microbiol. 130:96-100.

3. Cole, R. M., and T. J. Popkin. 1981. Electron microscopy, p.

34-51. In P. Gerhardt, R. G. E. Murray, R. N. Costilow, E. W.Nester, W. A. Wood, N. R. Kreig, and G. B. Phillips (ed.),Manual of methods for general bacteriology. American Societyfor Microbiology, Washington, D.C.

4. Das, A., and A. K. Mishra. 1983. Utilization of fructose byAzospirillum brasilense. Can. J. Microbiol. 29:1213-1217.

5. Deinema, M. M., and L. P. T. M. Zevenhuizen. 1971. Formationof cellulose fibrils by gram-negative bacteria and their role inbacterial flocculation. Arch. Microbiol. 78:42-52.

6. Dobereiner, J., and J. M. Day. 1976. Associative symbiosis intropical grasses: characterization of microorganisms anddinitrogen-fixing sites, p. 518-538. In W. E. Newton and C. J.Nyman (ed.), Proceedings of the First International Symposiumon Nitrogen Fixation. Washington State University Press, Pull-man.

7. Eklund, C., L. M. Pope, and 0. Wyss. 1966. Relationship ofencapsulation and encystment in Azotobacter. J. Bacteriol.92:1828-1830.

8. Friedman, B. A., P. R. Dugan, R. M. Pfister, and C. C. Remsen.1969. Structure of exocellular polymers and their relationship tobacterial flocculation. J. Bacteriol. 98:1328-1334.

9 Goebel, E. M., and N. R. Kreig. 1984. Fructose catabolism inAzospirillum brasilense and Azospirillum lipoferum. J. Bacte-riol. 159:86-92.

10. Lakshiui, V., A. Satyanarayana Rao, K. V"ayalakshmi, M.Lakshmi Kumari, K. V. B. R. Tilak, and N. S. Subba Rao.1977. Establishment and survival of Azospirillum lipoferum.Proc. Indian Acad. Sci. Sect. B 86:397-404.

11. Lam'm, R. B., and C. A. Neyra. 1981. Characterization and cystproduction of azospirilla isolated from selected grasses growingin New Jersey and New York. Can. J. Microbiol. 27:1320-1325.

12. Layne, J. S., and E. J. Johnson. 1964. Natural factors involvedin the induction of cyst formation in Azotobacter. J. Bacteriol.87:684-689.

13. Magalhaes, L. M. S., C. A. Neyra, and J. Dobereiner. 1978.Nitrate and nitrite reductase negative mutants of N2-fixingAzospirillum sp. Arch. Microbiol. 117:247-252.

14. Maltseva, N. N., and Y. Laski. 1982. Synthesis and monosac-charide composition of exopolysaccharides in certain soil bac-teria. Mikrobiol. Zh. 44:33-36.

15. Martinez-Drets, G., M. D. Gallo, C. Burpee, and R. H. Burris.1984. Catabolism of carbohydrates and organic acids and ex-pression of nitrogenase by azospirilla. J. Bacteriol. 159:80-85.

16. Matthysse, A. G., K. V. Holm'es, and R. H. G. Gurlitz. 1981.Elaboration of cellulose fibrils by Agrobacterium tumefaciensduring attachment to carrot cells. J. Bacteriol. 145:583-595.

17. Napoli, C., F. Dazzo, and D. Hubbel. 1975. Production ofcellulose microfibrils by Rhizobium. Appl. Microbiol. 30:123-131.

18. Nelson, H. 1944. A photometric adaptation of the Somogyimethod for the determination of glucose. J. Biol. Chem.153:375-380.

19. Neyra, C. A., and J. Dobereiner. 1977. Nitrogen fixation ingrasses. Adv. Agron. 29:1-38.

20. Neyra, C. A., J. Dobereiner, R. Lalande, and R. Knowles. 1977.Denitrification by N2-fixing Spirillum lipoferum. Can. J. Micro-biol. 23:300-305.

21. Neyra, C. A., and P. Van Berkum. 1977. Nitrate reduction andnitrogenase activity in Spirillum lipoferum. Can. J. Microbiol.23:306-310.

22. Nur, I., Y. Steinitz, Y. Okon, and Y. Henis. 1981. Carotenoidcomposition and function in nitrogen-fixing bacteria of the genusAzospirillum. J. Gen. Microbiol. 122:27-32.

23. Okon, Y. 1984. The physiology of Azospirillum in relation to itsutilization as inoculum for promoting growth in plants, p.165-174. In L. W. Ludden and J. E. Burris (ed.), Proceedings ofthe Fourteenth Steinbock Symposium at the University ofWisconsin, Madison. Elsevier Science Publishing Inc., NewYork.

24. Okon, Y., S. L. Albrecht, and R. H. Burris. 1976. Factorsaffecting growth and nitrogen fixation of Spirillum lipoferum. J.Bacteriol. 127:1248-1254.

25. Papen, M., and D. Werner. 1982. Organic acid utilization,

J. BACTERIOL.


succinate excretion, encystation and oscillating nitrogenaseactivity in Azospirillum brasilense, under microaerobic condi-tions. Arch. Microbiol. 132:57-61.

26. Patriquin, D. G., J. Dobereiner, and D. K. Jain. 1983. Sites andprocesses of association between diazotrophs and grasses. Can.J. Microbiol. 29:900-916.

27. Sadoff, H. L. 1975. Encystment and germination in Azotobactervinelandii. Bacteriol Rev. 39:516-539.

28. Stevenson, L. H., and M. D. Socolofskey. 1966. Cyst-formationand Poly-B-hydroxybutyric acid accumulation in Azotobacter.J. Bacteriol. 91:304-310.

29. Stevenson, L. H., and M. D. Socolofskey. 1972. Encystation of

Azotobacter vinelandii in liquid cultures. Antonie vanLeeuwenhoek J. Microbiol. Serol. 38:605-610.

30. Tarrand, J. J., N. R. Kreig, and J. Dobereiner. 1978. Ataxonomic study of the Spirillum lipoferum group, with descrip-tions of a new genus, Azospirillum gen. nov. and two species,Azospirillum lipoferum (Beijerinck) comb. nov. and Azospiril-lum brasilense sp. nov. Can. J. Microbiol. 24:967-980.

31. Vela, G. R. 1974. Survival of Azotobacter in dry soil. Appl.Microbiol. 28:77-79.

32. Westby, C. A., D. S. Cutshall, and G. V. Vigil. 1983. Metabolismof various carbon sources by Azospirillum brasilense. J. Bacte-riol. 156:1369-1372.

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