Nitrogen stress induced changes in the marine cyanobacteriumOscillatoria willei BDU 130511

Sushanta Kumar Saha, Lakshmanan Uma, Gopalakrishnan Subramanian �

National Facility for Marine Cyanobacteria, Bharathidasan University, Tiruchirappalli 620 024, India

Received 30 March 2003; received in revised form 20 May 2003; accepted 3 June 2003

First published online 5 July 2003

Abstract

Exclusion of combined nitrogen (NaNO3) from the growth medium caused certain changes in metabolic processes leading to cessationin growth of the non-heterocystous, non nitrogen-fixing marine cyanobacterium Oscillatoria willei BDU 130511. But antioxidativeenzymes, namely superoxide dismutase and peroxidase, helped the organism to survive the nitrogen stress. Prominent effects observedduring nitrogen starvation/limitation were: (i) reduction of major and accessory photosynthetic pigments, (ii) impairment ofphotosynthesis due to loss of one major Rubisco isoenzyme, (iii) reduced synthesis of lipids and fatty acids, (iv) modifications ofprotein synthesis leading to the repression of three polypeptides and synthesis of two new polypeptides, (v) enhanced glutamine synthetaseand reduced nitrate reductase activities, (vi) enhanced production of hydrogen peroxide and (vii) induced appearance of four newperoxidase isoenzymes. The observed metabolic changes were reversible, and the arrested growth under prolonged nitrogen deficiencycould be fully restored upon subculturing in freshly prepared ASN III medium containing nitrogen (NaNO3). The present studydemonstrates the capability of a non-nitrogen-fixer to withstand nitrogen stress making it an ecologically successful organism in themarine environment. The above pleiotropic effects of nitrogen deficiency also demonstrate that nitrogen plays a crucial role in growth andmetabolism of marine cyanobacteria.9 2003 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved.

Keywords: Cyanobacterium; Oscillatoria willei ; Nitrogen; Starvation; Limitation; Oxidative stress ; Active oxygen species; Superoxide dismutase;Hydrogen peroxide; Peroxidase

1. Introduction

Living organisms, especially micro-organisms, are ex-posed to various types of natural stresses, such as nutrientlimitation, light intensity and quality, temperature, pH,salinity, drought, pollution, etc. In photosynthetic organ-isms, the above stress factors cause an imbalance in theelectron transport system of photosystems which is the¢rst indication of an unfavourable condition. Cyanobac-teria, a group of prokaryotic, oxygen-evolving, photosyn-thetic Gram-negative bacteria, survive in a wide variety ofenvironmental extremes ranging from ultra-oligotrophicoceans to nutrient-enriched estuarine, aphotic and anoxic

waters, geothermal and sul¢de-rich conditions (See reviews[1^3]). Of the di¡erent kinds of nutritional stresses, nitro-gen limitation ranks ¢rst in oligotrophic environments [4].Nitrogen is an essential major element required for thesynthesis of primary and secondary amino acids, proteins,nucleic acids, coenzymes, chlorophyll and other accessoryphotosynthetic pigments (phycobilins in cyanobacteria)[5]. Nitrogen comprises about 10% of cell dry weight incyanobacteria [6]. It is an irony that nitrogen is the limit-ing factor in marine environments [7]. The availability ofthis element is a key factor in regulating the productivityand thus in£uencing the species composition of a givenarea and their survival and sustenance in marine habitats[4,8].

Cyanobacteria, though, have specialised biochemicaland ecological mechanisms to access essential nutrients(nitrogen, phosphorus and iron) that most often limitgrowth, only heterocystous and a few non-heterocystousforms ¢x atmospheric nitrogen in nitrogen-limited envi-ronments [9^13]. Several other non-heterocystous forms

0168-6496 / 03 / $22.00 9 2003 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved.doi :10.1016/S0168-6496(03)00162-4

* Corresponding author. Tel. : +91 (431) 2407082;Fax: +91 (431) 2407084.

E-mail address: gs@nfmc.res.in (G. Subramanian).

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www.fems-microbiology.org

are able to withstand a nitrogen-limited environment for alimited period of time, which would be su⁄cient for theirre-establishment. The present study demonstrates the roleof physiological mechanisms and antioxidative enzymes incombating nitrogen stress in the marine ¢lamentous non-heterocystous cyanobacterium Oscillatoria willei BDU130511.

2. Materials and methods

2.1. Organism

Axenic culture of O. willei BDU 130511, a fast-growing,non-heterocystous, ¢lamentous, marine cyanobacterium,was obtained from the culture collection of the NationalFacility for Marine Cyanobacteria, Bharathidasan Univer-sity, Tiruchirappalli, India. The organism was maintainedin ASN III medium [14].

2.2. Culture conditions

The organism was grown in 250-ml Erlenmeyer £askscontaining 100 ml of ASN III medium [14] except in nitro-gen limitation experiments, wherein combined nitrogen(NaNO3) was omitted. Experimental cultures were incuba-ted at 25 O 2‡C, 14/10-h light/dark cycle, with illuminationof 27 WE m32 s31 under cool white £uorescent lamps(Philips). The cultures were mildly shaken by hand onalternate days.

2.3. Microscopy

Microscopic observation in bright ¢eld, dark ¢eld andphotomicrography were done using a Leitz diaplan micro-scope equipped with Photoautomat Wild MPS 46 (Leica,USA).

2.4. Harvesting

All experiments except growth were carried out on day5 after inoculation. Cultures were harvested by centrifug-ing the contents at 6000Ug for 8^10 min at 4‡C. Thepellets were thoroughly washed with distilled water andcentrifuged as above. Fresh weights of the cyanobacterialpellets were noted after blotting them with ¢lter paper.Dry weights were recorded after drying the pellets at60‡C until constant weights were obtained. Results areaverages of triplicates.

2.5. Estimation of pigments

Known amounts (300 mg each) of wet biomass wereused for pigment extraction and all the operations werecarried out exclusively in dim laboratory light to avoidphoto-oxidation.

2.5.1. Chlorophyll aChlorophyll a was extracted in cold methanol (90%)

overnight at 4‡C in the dark. The optical density of thesupernatant was read at 663 nm (Jasco V-550 spectropho-tometer, Japan) and the quantity estimated followingMackinney [15].

2.5.2. CarotenoidsCarotenoids were extracted overnight in 85% acetone at

4‡C in the dark. After centrifugation, the optical densityof the supernatant was read at 450 nm and the amountwas estimated by employing the extinction coe⁄cient ofJensen [16].

2.5.3. PhycobilinsPhycobilins of cyanobacterial biomass were extracted in

cold 0.05 M phosphate bu¡er (pH 6.8) by repeated freeze-thawing at 320‡C until all the phycobilins were extracted.Absorption spectra of supernatants were recorded with theaid of a double-beam UV-visible spectrophotometer andthe absorbances at 615 nm and 652 nm were used to esti-mate the amount of phycocyanin and allophycocyaninfollowing the formula of Siegelman and Kycia [17].

2.6. Lipids and fatty acids

Extraction of lipids was done following the method ofFolch et al. [18]. A known amount of cyanobacterial pelletwas ground in a mortar and pestle by adding pulverisedglass powder (V0.5 mm) and extraction solvent (2:1chloroform:methanol). The extract was ¢ltered throughWhatman No. 1 ¢lter paper where a third volume of dis-tilled water was added to remove water-soluble impurities.Then the ¢ltrate was vortexed and let stand for separationof two layers and the lower lipid layer was transferredcarefully. The moisture content of lipids was eliminatedby the addition of sodium sulfate crystals and dried in avacuum concentrator (Savant Speed Vac, USA) and thelipids were measured gravimetrically.

Identi¢cation and quanti¢cation of fatty acids weredone by the modi¢ed method of Miller and Berger [19].A known amount of lipid was saponi¢ed by boiling it with1 ml of saponi¢cation reagent (15 g NaOH in 100 ml of1:1 methanol:water) for 30 min. The sample was thenboiled in a water bath at 80‡C for 20 min with 2 ml ofmethylation reagent (1:1.18 methanol:6 N HCl). Aftercooling, 1 ml of extraction solvent (1:1 distilled hexane:anhydrous diethyl ether) was added and mixed thor-oughly. Thereafter the lower aqueous phase was discardedand the remaining upper phase was washed with 3 ml ofbase wash solution (1.2% NaOH w/v). Finally, 2 Wl of theorganic phase was chromatographed in a gas chromato-graph (5890A Hewlett Packard, USA) ¢tted with 10%DEGS column (6 ftU3.12 mm) using a £ame ionisationdetector. The conditions were: oven temperature, 180‡C;detector temperature, 230‡C; carrier gas, nitrogen at

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30 ml min31. Fatty acids were identi¢ed and quanti¢ed bycomparing the retention time and area of the authenticstandards (Sigma, USA).

2.7. Protein estimation

The total soluble protein content of extracts was mea-sured as described by Lowry et al. [20] using bovine serumalbumin as standard.

2.8. Hydrogen peroxide estimation

Production of hydrogen peroxide (H2O2) in nitrogen-free and nitrogen-supplemented medium, with equalamounts of inoculum, was compared from the absorbanceat 485 nm of the chromogen formed by 4-aminoantipyrineand phenol [21].

2.9. Nitrogenase assay

Nitrogenase activity was determined by acetylene reduc-tion assay [22] in aerobic light (AL), microaerobic light(MAL), microaerobic dark (MAD) and microaerobic lightwith 10 WM DCMU (MALD) conditions. Cultures fromthe mid-exponential growth phase were transferred to ni-trogen-free ASN III medium prior to nitrogenase assay.To 5 ml of homogenised culture in 20-ml serum bottles,argon was £ushed thoroughly to provide microaerobicconditions. Assay bottles were sealed with a rubber sep-tum and 1.5 ml of air was withdrawn and replaced withthe same volume of acetylene. Assay bottles of AL, MALand MALD were incubated in an illuminated shaker at27 WE m32 s31 of light intensity, 30‡C and 50 rpm. Fordark estimation, cultures were kept in the dark for 15 minfor dark adaptation immediately before the addition ofacetylene and were incubated in a dark box under theabove condition. At 2, 24, 48 and 72 h of incubation,500 Wl of 10% trichloroacetic acid was added to eachvial to stop the enzyme activity. For the determinationof ethylene concentration, 100 Wl of gaseous phase fromthe assay bottle was injected into a Porapak-T column(oven temperature, 75‡C; detector temperature, 120‡C;carrier gas, nitrogen at 30 ml min31) in a gas chromato-graph (5890A Hewlett Packard, USA) and detected with a£ame ionisation detector. Ethylene from EDT Research,London was used as standard.

2.10. Glutamine synthetase (GS) assay

Washed cyanobacterial pellet was permeabilised by tolu-ene at 4‡C. Then, GS activity was estimated in vitro byincubating the clear supernatant with 1 ml of reactionmixture for 30 min in the dark at 37‡C [23]. After stoppingthe reaction, absorbance was read at 540 nm and theamount of activity was expressed as Wg Q-glutamyl hydrox-amate formed mg Chl a31 min31.

2.11. Nitrate reductase (NR) assay

NR activity was assayed as nitrate reduction (with so-dium dithionite-reduced methyl viologen as the electrondonor) [24]. The amount of activity was expressed as Wgnitrite formed mg Chl a31 min31.

2.12. Protein and enzyme electrophoresis

Samples for protein pro¢le and assay of enzyme activ-ities were prepared from thoroughly washed cyanobacte-rial pellets rewashed with extraction bu¡er (0.0625 MTris^HCl, pH 6.8). Cyanobacterial pellets were homoge-nised using a pre-chilled mortar and pestle in the presenceof glass powder (V0.5 mm) adding ice-cold extractionbu¡er. The samples were centrifuged at 15 000Ug for15 min and the process was repeated twice to obtain clearsupernatants. The amounts of proteins were estimated [20]and used for both sodium dodecyl sulfate^polyacrylamidegel electrophoresis (SDS^PAGE) and activity staining ofgels.

Electrophoresis was carried out at 20 O 2‡C with a1.5 mm thick polyacrylamide gel in Tris^glycine bu¡er(pH 8.3). In the case of SDS^PAGE, SDS was includedin the running bu¡er (0.1%) as well as in sample bu¡er(10%). The sample was boiled for 3 min with sample bu¡erand centrifuged brie£y before loading. In both PAGE andSDS^PAGE, a uniform amount (300 Wg) of protein wasloaded to each well. Samples were then electrophoresed at60 V through the stacking gel (6%) and at 120 V throughthe separating gel (10%) [25].

2.13. Denatured gel ^ staining and analysis

After electrophoresis, SDS gels were ¢xed and stainedwith Coomassie brilliant blue R-250 (SRL, India) for 3 hand destained with distilled water containing 5% methanoland 7% glacial acetic acid in a shaker (50 rpm) at roomtemperature. The images were captured with a CCD cam-era and the protein pro¢les were analysed by the gel doc-umentation system (Alpha Imager1 2200, USA).

2.14. Native gel ^ enzyme staining

2.14.1. Rubisco (EC 4.1.1.39)Electrophoresed bands of Rubisco isoenzymes were

¢xed and stained by brie£y soaking the gel in 100 ml of¢xative (5:1:5 methanol:acetic acid:double-distilled water)containing 50 mg of amido black (Loba Chemie, India)for exactly 15 min [26]. Then the gel was quickly destainedusing the same ¢xative.

2.14.2. Superoxide dismutase (EC 1.15.1.1)Activity staining for superoxide dismutase (SOD) on gel

[27] and the identi¢cation of metal prosthetic groups in theactive enzyme were carried out following Chadd et al. [28].

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The gels were soaked in staining solution containing 50 mlof 50 mM Tris^HCl (pH 8), 10 mg NBT (Hi-Media, In-dia), 1 mg EDTA (Sigma, USA) and 2 mg ribo£avin (Sig-ma, USA) for 30 min in the dark at room temperature(25‡C) and then illuminated on a light box with white£uorescent light (80 WE m32 s31) for 30 min or until ach-romatic bands appeared. For the identi¢cation of metalprosthetic groups, enzyme activity was tested by soakingone gel in bu¡er containing 5 mM H2O2, another in bu¡ercontaining 2 mM KCN and one more in bu¡er withoutany inhibitors (H2O2 or KCN) for 20 min in the dark,prior to exposing all three gels to staining solution andillumination.

2.14.3. Peroxidase (EC 1.11.1.7)Activities of peroxidase isoenzymes were detected by

incubating the gel in 100 ml of 0.1 M sodium acetate bu¡-er (pH 4.6) containing 30 mg 3,3P-diaminobenzidine tetra-hydrochloride (SRL, India). The reaction was initiated byadding 250 Wl of 30% hydrogen peroxide (Qualigens, In-dia) and let stand until intense brown bands appeared [29].

3. Results

3.1. Growth

Prolonged starvation for over 5 months caused the ma-jority of cells to turn yellow and fragmentation of ¢la-ments resulted in unicells of varying morphology (Figs. 1and 2). Few hormogonia and live ¢laments with less pig-mentation were also noticed (Fig. 3). These cells returnedto normal (Fig. 5) and started growing when reinoculatedto complete medium (Fig. 4).

Nitrogen limitation reduced growth by 17% of fresh anddry biomass compared to growth in nitrogen-supple-mented medium (Table 1). In N-starved conditions, chlo-

rophyll a and carotenoids were reduced by 31.81% and18.38% compared to their respective controls (Table 1).Among phycobili-pigments, phycocyanin alone was de-creased by 19.67%, while the amount of allophycocyaninremained almost equal to that of cultures grown in nitro-gen-supplemented medium (Table 1). From the absorptionspectrum of cell-free extract, no phycoerythrin peak couldbe detected in this organism (Fig. 6).

3.2. Lipids and fatty acids

Nitrogen starvation resulted in a decrease in total lipidcontent (26.08%) and caused qualitative and quantitativevariations in whole cell fatty acids. In nitrogen-starved

Fig. 1. Prolonged starvation for over 5 months caused the majority ofcells to turn yellow.

Fig. 2. Fragmentation of ¢laments resulted in unicells of varying mor-phology.

Fig. 3. Few hormogonia and live ¢laments with less pigmentation.

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cultures, fatty acids such as pentadecanoic acid (C15:0),oleic acid (C18:1 cis), linoleic acid (C18:2 cis), behenicacid (C22:0) and eicosapentaenoic acid (C20:5) disap-peared, whereas eicosenoic acid (C20:1) appeared new.There was a quantitative reduction of other fatty acids(C14:0, C16:0, C16:1 and C17:0) except lauric acid andQ-linolenic acid, which increased in nitrogen-starved con-ditions by 14.53% and 40.55% respectively (Table 2).

3.3. Hydrogen peroxide

Hydrogen peroxide generation increased by 34.35% innitrogen-starved cultures compared to control (Table 1).

3.4. Proteins and enzymes

Nitrogen starvation caused a decrease in protein content

by 38.83% as estimated spectrophotometrically. The pro-tein pro¢les of nitrogen-starved cells di¡ered markedlyfrom those of control cells. In nitrogen-starved cells, threepolypeptides, of 11.2, 16.7 and 55 kDa, found in controlwere missing, while two new polypeptides of molecularmass 52.6 and 90.5 kDa appeared. There was also a quan-titative increase of 59.7-kDa polypeptide in nitrogen-starved cells compared to control (Fig. 7).

Nitrogenase activity could not be detected in any of thefour experimental conditions even after 72 h of incubation.In nitrogen-starved cells the GS activity was nearly 10-foldmore compared to control cells (Table 1). NR activity innitrogen-starved cells decreased by six-fold compared tocontrol (Table 1). A prominent band (Rm 0.484) of Rubis-co isoenzyme found in control cells could not be detectedin nitrogen-starved cells of O. willei BDU 130511 (Fig. 8).

The SOD isoenzyme pro¢les of both control and nitro-gen-starved cultures were essentially similar except for thedisappearance of an isoform of Mn-SOD (Rm 0.855) innitrogen-starved cells. A prominent isoenzyme (Rm

0.596) appearing in both cases was identi¢ed as Fe-SODas it was sensitive to H2O2 incubation and the four lessachromatic bands (Rm 0.690, 0.716, 0.787 and 0.855) were

Fig. 5. Appearance of normal morphology.

Table 1E¡ect of nitrogen limitation/starvation on end-point growth, cellular contents, nitrogen-assimilating enzymes and hydrogen peroxide production ofO. willei BDU 130511

Control N-deprived

End-point biomass (day 8)Fresh weight (g) 3.55 O 0.177 2.94 O 0.212Dry weight (mg) 378.96 O 18.844 313.4 O 22.613

Cellular contentsChlorophyll a (Wg g31 DW) 769.27 O 12.091 524.59 O 0.462Carotenoids (Wg g31 DW) 277.80 O 7.659 228.52 O 20.883Phycocyanin (mg g31 DW) 99.05 O 2.015 79.56 O 6.914Allophycocyanin (mg g31 DW) 19.14 O 0.355 19.43 O 1.066Total protein (mg g31 DW) 370.00 O 11.98 226.3 O 22.34Lipid (mg g31 DW) 88.38 O 1.910 65.33 O 1.109

Nitrogenase activity (2, 24, 48 and 72 h) ND NDGS activitya 0.59 O 0.049 5.79 O 0.674NR activityb 13.91 O 0.229 2.27 O 0.207H2O2 production (OD475) 0.6196 O 0.003 0.9438 O 0.005

ND, not detected.aWg Q-glutamyl hydroxamate formed mg Chl a31 min31.

bWg nitrite formed mg Chl a31 min31.

Fig. 4. Initiation of growth after transfer to complete medium.

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considered Mn-SODs as they were resistant to both H2O2

and KCN (Fig. 9).Nitrogen starvation induced four new isoforms of per-

oxidase (at Rm 0.107, 0.124, 0.454 and 0.538). Of these, theexpression of the isoform with Rm 0.538 was high; how-ever, the isoform with Rm 0.378 present in control wasabsent under nitrogen starvation (Fig. 10).

4. Discussion

It is well known that a large part of the world’s oceansis nutritionally depleted especially in nitrogen and marinecyanobacteria are able to persist and adapt their metabo-lism to this distinct environmental stress. De¢ciency ofcombined nitrogen in nature is a threat to non-nitrogen-¢xing cyanobacteria. The present laboratory study hasshown the high degree of physiological adaptability ofO. willei BDU 130511 to nitrogen stress. To overcomethe stress, organisms generally have two types of defencestrategies: (i) mechanisms that prevent interaction with thestress factors and (ii) those that counteract the stress-in-

duced damages. The results of this study indicate thatO. willei BDU 130511 has opted for the second strategy.

O. willei BDU 130511 could not grow in N-free mediumdue to the inability of this non-heterocystous cyanobacte-rium to scavenge atmospheric N2. No nitrogenase activitycould be detected by acetylene reduction assay in MAD,MALD, MAL and AL conditions (Table 1). Acetylenereduction activity has been accepted as a simple and e⁄-cient way of estimating nitrogenase activity [9^13,30^36],although 15N ¢xation, molecular and antibody assays willprove useful in identifying the presence or absence of theenzyme per se. The fact that the cyanobacterium could notgrow in the absence of combined nitrogen in the medium

Fig. 6. Relative absorption spectra of phycobilins of O. willei BDU130511 extracted in 0.05 M phosphate bu¡er by repeated freeze-thaw-ing; C, control ; T, nitrogen-starved cells.

Table 2Fatty acid content (mg g lipid31) of O. willei BDU 130511 grown underdi¡erent nitrogen conditions

Fatty acid Nþ N3

Lauric acid (C12:0) 0.553 0.647Myristic acid (C14:0) 1.258 0.580Pentadecanoic acid (C15:0) 0.031 NDPalmitic acid (C16:0) 4.304 1.981Palmitolic acid (C16:1) 12.967 5.756Heptadecanoic acid (C17:0) 4.902 1.458Oleic acid (C18:1 cis) 0.150 NDLinoleic acid (C18:2 cis) 0.114 NDQ-Linolenic acid (C18:3Y) 0.107 0.180Eicosenoic acid (C20:1) ND 2.401Eicosapentaenoic acid (C20:5) 1.226 NDBehenic acid (C22:0) 1.510 ND

ND, not detected.

M C T

94

67

43

30

20.1

14.4

11.2

16.7

52.6

55

59.7

90.5

Fig. 7. Modi¢cation of protein synthesis in O. willei BDU 130511 dur-ing nitrogen (NaNO3) starvation. Protein samples (300 Wg), from cellsgrown for 4 days in complete ASN III medium (lane C) and ASN IIImedium devoid of NaNO3 (lane T), were resolved by SDS^PAGE(10%) and visualised by Coomassie staining. To determine the molecularmasses of prominent polypeptides either missing, appearing new or atenhanced levels, a mixture of six standard proteins (lane M) fromAmersham Biosciences was run simultaneously.

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and the lack of acetylene reduction activity indicate thatthe organism is unable to use atmospheric N2 in requiredamounts.

Nitrogen limitation in O. willei BDU 130511 resulted inthe reduction of photosynthetic pigments such as chloro-phyll a, carotenoids and phycocyanin resulting in chloro-sis. It is known that in photosynthetic organisms N limi-tation triggers ordered degradation of phycobilisomes,ribosomes and thylakoid membranes [37]. This explainsthe lowered pigment content, lipid content and change inpattern of fatty acids observed in O. willei BDU 130511 inthis study as well as in Aphanocapsa 6308 [38], Agmenellumquadruplicatum [37] and Synechococcus 6301 [39] byothers. Furthermore, nitrogen limitation leading to catab-

olisation of lipids and fatty acids resulting in their reduc-tion as found in Spirulina platensis and Anacystis nidulans[40] was also observed in this cyanobacterium (Table 2). Adecreased level of total lipid content is believed to indicatea reduced carbon storage mechanism and leads to theavailability of carbon skeletons to synthesise proteins orenzymes required for the defence [41]. The enhanced levelsof C12:0 and C18:3Y fatty acids as well as the newlysynthesised C20:1 (18.47% of total fatty acids) under ni-trogen starvation in O. willei BDU 130511 (Table 2) mayhelp the organism to maintain membrane £uidity to over-come stress and retain cellular integrity as suggested bySmith et al. [42] and Floreto et al. [43]. Reduction in levelsof some fatty acids and disappearance of certain others(Table 2) could lead to reutilisation of these fatty acidsfor other metabolic purposes including the increased pro-duction of certain other fatty acids mentioned above.

Most photosynthetic micro-organisms depend on eitherammonium (NHþ

4 ) or nitrate (NO33 ) as their sole source of

combined nitrogen. However, cyanobacteria have the abil-ity to use a wide variety of nitrogen sources such as am-monia, nitrate, nitrite, urea, amino acids, nucleosides orbases [44,45], atmospheric nitrogen (some cyanobacteriaonly) and even synthetic compounds like ampicillin [46].In general, the preferential order of nitrogenous com-pounds for cyanobacteria and other algae is ammo-nias nitrate or ureas other organic compounds [45].

GS is an e⁄cient ammonia scavenger in cyanobacteria[47]. Enhanced GS activity of O. willei BDU 130511 undernitrogen starvation (Table 1) may probably be due to thechanneling of available nitrogen through breakdown oforganic nitrogenous compounds from dead cells as re-vealed by microscopic observation (Fig. 3) and breakdownof its own phycocyanin [48,49]. In addition, several otherreasons suggested by others such as increased expression

0.484

Rm

Fig. 8. Native PAGE of control (lane C) and nitrogen-starved (lane T)cells of O. willei BDU 130511 stained with amido black for Rubisco iso-enzymes.

Rm

0.596

0.6900.716

0.787

0.855

Fig. 9. Activity staining for SOD and identi¢cation of metal prostheticgroups in control (lane C) and nitrogen-starved (lane T) cells of O. wil-lei BDU 130511.

Fig. 10. Activity staining for peroxidase isoenzymes in control (lane C)and nitrogen-starved (lane T) cells of O. willei BDU 130511, showing in-duced bands in the stress condition.

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of ammonium transport (amt) genes as in Synechocystissp. [50], possession of multiple GS enzymes [45] and de-creased activity of NR as a process of saving energy re-ported in Prochlorococcus sp. strain PCC 9511 [51] (wherethe genes encoding the nitrate-assimilating system werelacking in extremely nitrogen (nitrate or nitrite)-scarce en-vironments) could also contribute to the enhancement ofGS activity. In this study, a signi¢cant reduction of NRactivity during nitrogen starvation was observed (Table 1)which indicates the control of expression of genes for ni-trate-assimilating enzymes, which may help the cyanobac-terium O. willei BDU 130511 to survive successfully inoligotrophic or ultra-oligotrophic environments.

Nitrogen starvation had caused profound alterations inproteins, both qualitatively and quantitatively, in O. willeiBDU 130511. Prominent among the impaired proteins arephycobiliproteins (Fig. 6, Table 1). In Synechococcus sp.also, severe nitrogen limitation was reported to cause pro-teolytic degradation of phycobiliproteins [45]. Degradedproducts of phycobiliproteins could provide C and N ele-ments for the synthesis of other cellular constituents re-quired during nutrient deprivation [48] and reduce absorp-tion of excitation energy, making cells less susceptible tophotodamage [52].

Two of the three newly synthesised proteins (Fig. 7)based on their molecular masses might be consideredstress proteins (52.6 and 90.5 kDa) or hydrolytic enzymeswhile the 59.7-kDa protein whose level is increased duringN starvation (Fig. 7) could be a chaperonin, which helpsin protein protection and repair mechanism under stress asreported by Sanders [53].

Only a meagre amount of information is available onthe e¡ect of N limitation and starvation on photosynthesisand respiration. In the present study, N starvation hasbeen shown to result in reduced chlorophyll a synthesis(Table 1) as well as alteration in Rubisco isoenzyme pro-¢le, due to the disappearance of a prominent band in theRubisco pro¢le of nitrogen-starved cells (Fig. 8). Thiscould be due to the interaction of active oxygen specieswith essential sulfhydryl groups of the enzyme as reportedin several chloroplast enzymes of the carbon di-oxide ¢x-ation pathway [54]. Photosynthesis was thus a¡ected andreduced the growth of O. willei BDU 130511. Similar ob-servations were reported in Synechocystis PCC 6803 cells,where the rate of both O2 evolution and CO2 ¢xationdeclined under nitrogen starvation [55].

De¢ciency of combined nitrogen could also have createdoxidative stress. In N-starved cultures of Spirulina platen-sis, the rate of respiration was reported to have increasedboth at normal and at high CO2 concentration [49]. Ahigh rate of respiration might be one of the reasons forinduction of oxidative stress through the generation ofactive oxygen species [56]. Hence, e⁄cient scavenging isimportant to prevent oxidative damage to the organismas active oxygen species are highly reactive with the pro-teins and membranes. SOD is the ¢rst line of defence

against active oxygen species. In cyanobacteria, three iso-forms of SOD are reported, viz. Mn-SOD, Fe-SOD andCu/Zn-SOD (see [28]). It has been found that Mn-SODprotects thylakoid membranes against superoxides, whileFe-SOD protects cytosolic contents [57]. In O. willei BDU130511, under nitrogen starvation, Fe-SOD could play amajor role in dismutating the superoxide ions, since Fe-SOD was considered to be the sole superoxide-scavengingenzyme in Synechocystis sp. strain PCC 6803 [56]. Billi andCaiola [58] also presumed that Fe-SOD protects the cya-nobacterium against free oxygen radicals generated duringoxidative stress due to nitrogen starvation.

The presence of SOD indicates dismutation of superox-ides to hydrogen peroxide, a powerful oxidant which pro-vides the cellular signal for induction of defence mecha-nisms [59,60]. It also disturbs the cellular redox state andits uncontrolled accumulation may cause damage of mem-brane lipids, DNA, and ¢nally death of the cell [61]. In thepresent study an increased level of H2O2 was observed innitrogen-starved cells (Table 1). Hydrogen peroxide couldbe useful in degrading organic matter of surrounding deadcells and make nutrients available for the surviving cells.However, since H2O2 is the precursor for the most reactiveand destructive active oxygen species in the presence ofO3

2 and Fe2þ [62], the removal of H2O2 from the vicinityof living cells is essential. Probably for this reason an in-creased number of isoforms of peroxidase were observedin this cyanobacterium (Fig. 10). Of the four newly syn-thesised peroxidase isoenzymes, observed under nitrogenstress, three were intensely stained. This induced synthesisof new isoforms could be to play a special role in neutral-ising the lethal e¡ect of H2O2 in living cells favouring theorganism to survive in nitrogen-limited environments.

5. Conclusion

Cyanobacteria (blue-green algae) are of evolutionary in-terest, as they are highly adaptable to various environmen-tal extremes. However, the mechanisms involved in envi-ronmental tolerance by cyanobacteria are not fullyunderstood, especially in a nitrogen-limited environment.From the results of the present study, it could be inferredthat the marine cyanobacterium O. willei BDU 130511,although adversely a¡ected by nitrogen starvation, isable to save some cells or ¢laments from the stress withthe help of antioxidative enzymes, and these cells uponavailability of nitrogen return to normal growth. Thusthe study helps to understand the survival strategy of anon-heterocystous, non-nitrogen-¢xing marine cyanobac-terium in the oligotrophic environment.

Acknowledgements

The authors thank the Department of Biotechnology

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(DBT), Government of India, for ¢nancing the facility andfellowship to S.K.S. Dr D. Prabaharan and Dr A. Kalibare thanked for their valuable suggestions in improvingthe manuscript.

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