Nitrogen stress induced changes in the marine cyanobacterium Oscillatoria willei BDU 130511

Authors

  • Sushanta Kumar Saha,

    1. National Facility for Marine Cyanobacteria, Bharathidasan University, Tiruchirappalli 620 024, India
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  • Lakshmanan Uma,

    1. National Facility for Marine Cyanobacteria, Bharathidasan University, Tiruchirappalli 620 024, India
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  • Gopalakrishnan Subramanian

    Corresponding author
    1. National Facility for Marine Cyanobacteria, Bharathidasan University, Tiruchirappalli 620 024, India
      *Corresponding author. Tel.: +91 (431) 2407082; Fax: +91 (431) 2407084. E-mail address: gs@nfmc.res.in
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*Corresponding author. Tel.: +91 (431) 2407082; Fax: +91 (431) 2407084. E-mail address: gs@nfmc.res.in

Abstract

Exclusion of combined nitrogen (NaNO3) from the growth medium caused certain changes in metabolic processes leading to cessation in growth of the non-heterocystous, non nitrogen-fixing marine cyanobacterium Oscillatoria willei BDU 130511. But antioxidative enzymes, namely superoxide dismutase and peroxidase, helped the organism to survive the nitrogen stress. Prominent effects observed during nitrogen starvation/limitation were: (i) reduction of major and accessory photosynthetic pigments, (ii) impairment of photosynthesis due to loss of one major Rubisco isoenzyme, (iii) reduced synthesis of lipids and fatty acids, (iv) modifications of protein synthesis leading to the repression of three polypeptides and synthesis of two new polypeptides, (v) enhanced glutamine synthetase and reduced nitrate reductase activities, (vi) enhanced production of hydrogen peroxide and (vii) induced appearance of four new peroxidase isoenzymes. The observed metabolic changes were reversible, and the arrested growth under prolonged nitrogen deficiency could be fully restored upon subculturing in freshly prepared ASN III medium containing nitrogen (NaNO3). The present study demonstrates the capability of a non-nitrogen-fixer to withstand nitrogen stress making it an ecologically successful organism in the marine environment. The above pleiotropic effects of nitrogen deficiency also demonstrate that nitrogen plays a crucial role in growth and metabolism of marine cyanobacteria.

1Introduction

Living organisms, especially micro-organisms, are exposed to various types of natural stresses, such as nutrient limitation, light intensity and quality, temperature, pH, salinity, drought, pollution, etc. In photosynthetic organisms, the above stress factors cause an imbalance in the electron transport system of photosystems which is the first indication of an unfavourable condition. Cyanobacteria, a group of prokaryotic, oxygen-evolving, photosynthetic Gram-negative bacteria, survive in a wide variety of environmental extremes ranging from ultra-oligotrophic oceans to nutrient-enriched estuarine, aphotic and anoxic waters, geothermal and sulfide-rich conditions (See reviews [1–3]). Of the different kinds of nutritional stresses, nitrogen limitation ranks first in oligotrophic environments [4]. Nitrogen is an essential major element required for the synthesis of primary and secondary amino acids, proteins, nucleic acids, coenzymes, chlorophyll and other accessory photosynthetic pigments (phycobilins in cyanobacteria) [5]. Nitrogen comprises about 10% of cell dry weight in cyanobacteria [6]. It is an irony that nitrogen is the limiting factor in marine environments [7]. The availability of this element is a key factor in regulating the productivity and thus influencing the species composition of a given area and their survival and sustenance in marine habitats [4,8].

Cyanobacteria, though, have specialised biochemical and ecological mechanisms to access essential nutrients (nitrogen, phosphorus and iron) that most often limit growth, only heterocystous and a few non-heterocystous forms fix atmospheric nitrogen in nitrogen-limited environments [9–13]. Several other non-heterocystous forms are able to withstand a nitrogen-limited environment for a limited period of time, which would be sufficient for their re-establishment. The present study demonstrates the role of physiological mechanisms and antioxidative enzymes in combating nitrogen stress in the marine filamentous non-heterocystous cyanobacterium Oscillatoria willei BDU 130511.

2Materials and methods

2.1Organism

Axenic culture of O. willei BDU 130511, a fast-growing, non-heterocystous, filamentous, marine cyanobacterium, was obtained from the culture collection of the National Facility for Marine Cyanobacteria, Bharathidasan University, Tiruchirappalli, India. The organism was maintained in ASN III medium [14].

2.2Culture conditions

The organism was grown in 250-ml Erlenmeyer flasks containing 100 ml of ASN III medium [14] except in nitrogen limitation experiments, wherein combined nitrogen (NaNO3) was omitted. Experimental cultures were incubated at 25±2°C, 14/10-h light/dark cycle, with illumination of 27 μE m−2 s−1 under cool white fluorescent lamps (Philips). The cultures were mildly shaken by hand on alternate days.

2.3Microscopy

Microscopic observation in bright field, dark field and photomicrography were done using a Leitz diaplan microscope equipped with Photoautomat Wild MPS 46 (Leica, USA).

2.4Harvesting

All experiments except growth were carried out on day 5 after inoculation. Cultures were harvested by centrifuging the contents at 6000×g for 8–10 min at 4°C. The pellets were thoroughly washed with distilled water and centrifuged as above. Fresh weights of the cyanobacterial pellets were noted after blotting them with filter paper. Dry weights were recorded after drying the pellets at 60°C until constant weights were obtained. Results are averages of triplicates.

2.5Estimation of pigments

Known amounts (300 mg each) of wet biomass were used for pigment extraction and all the operations were carried out exclusively in dim laboratory light to avoid photo-oxidation.

2.5.1Chlorophyll a

Chlorophyll a was extracted in cold methanol (90%) overnight at 4°C in the dark. The optical density of the supernatant was read at 663 nm (Jasco V-550 spectrophotometer, Japan) and the quantity estimated following Mackinney [15].

2.5.2Carotenoids

Carotenoids were extracted overnight in 85% acetone at 4°C in the dark. After centrifugation, the optical density of the supernatant was read at 450 nm and the amount was estimated by employing the extinction coefficient of Jensen [16].

2.5.3Phycobilins

Phycobilins of cyanobacterial biomass were extracted in cold 0.05 M phosphate buffer (pH 6.8) by repeated freeze-thawing at −20°C until all the phycobilins were extracted. Absorption spectra of supernatants were recorded with the aid of a double-beam UV-visible spectrophotometer and the absorbances at 615 nm and 652 nm were used to estimate the amount of phycocyanin and allophycocyanin following the formula of Siegelman and Kycia [17].

2.6Lipids and fatty acids

Extraction of lipids was done following the method of Folch et al. [18]. A known amount of cyanobacterial pellet was ground in a mortar and pestle by adding pulverised glass powder (∼0.5 mm) and extraction solvent (2:1 chloroform:methanol). The extract was filtered through Whatman No. 1 filter paper where a third volume of distilled water was added to remove water-soluble impurities. Then the filtrate was vortexed and let stand for separation of two layers and the lower lipid layer was transferred carefully. The moisture content of lipids was eliminated by the addition of sodium sulfate crystals and dried in a vacuum concentrator (Savant Speed Vac, USA) and the lipids were measured gravimetrically.

Identification and quantification of fatty acids were done by the modified method of Miller and Berger [19]. A known amount of lipid was saponified by boiling it with 1 ml of saponification reagent (15 g NaOH in 100 ml of 1:1 methanol:water) for 30 min. The sample was then boiled in a water bath at 80°C for 20 min with 2 ml of methylation reagent (1:1.18 methanol:6 N HCl). After cooling, 1 ml of extraction solvent (1:1 distilled hexane:anhydrous diethyl ether) was added and mixed thoroughly. Thereafter the lower aqueous phase was discarded and the remaining upper phase was washed with 3 ml of base wash solution (1.2% NaOH w/v). Finally, 2 μl of the organic phase was chromatographed in a gas chromatograph (5890A Hewlett Packard, USA) fitted with 10% DEGS column (6 ft×3.12 mm) using a flame ionisation detector. The conditions were: oven temperature, 180°C; detector temperature, 230°C; carrier gas, nitrogen at 30 ml min−1. Fatty acids were identified and quantified by comparing the retention time and area of the authentic standards (Sigma, USA).

2.7Protein estimation

The total soluble protein content of extracts was measured as described by Lowry et al. [20] using bovine serum albumin as standard.

2.8Hydrogen peroxide estimation

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

2.9Nitrogenase assay

Nitrogenase activity was determined by acetylene reduction assay [22] in aerobic light (AL), microaerobic light (MAL), microaerobic dark (MAD) and microaerobic light with 10 μM DCMU (MALD) conditions. Cultures from the mid-exponential growth phase were transferred to nitrogen-free ASN III medium prior to nitrogenase assay. To 5 ml of homogenised culture in 20-ml serum bottles, argon was flushed thoroughly to provide microaerobic conditions. Assay bottles were sealed with a rubber septum and 1.5 ml of air was withdrawn and replaced with the same volume of acetylene. Assay bottles of AL, MAL and MALD were incubated in an illuminated shaker at 27 μE m−2 s−1 of light intensity, 30°C and 50 rpm. For dark estimation, cultures were kept in the dark for 15 min for dark adaptation immediately before the addition of acetylene and were incubated in a dark box under the above condition. At 2, 24, 48 and 72 h of incubation, 500 μl of 10% trichloroacetic acid was added to each vial to stop the enzyme activity. For the determination of ethylene concentration, 100 μl of gaseous phase from the assay bottle was injected into a Porapak-T column (oven temperature, 75°C; detector temperature, 120°C; carrier gas, nitrogen at 30 ml min−1) in a gas chromatograph (5890A Hewlett Packard, USA) and detected with a flame ionisation detector. Ethylene from EDT Research, London was used as standard.

2.10Glutamine synthetase (GS) assay

Washed cyanobacterial pellet was permeabilised by toluene at 4°C. Then, GS activity was estimated in vitro by incubating the clear supernatant with 1 ml of reaction mixture for 30 min in the dark at 37°C [23]. After stopping the reaction, absorbance was read at 540 nm and the amount of activity was expressed as μg γ-glutamyl hydroxamate formed mg Chl a−1 min−1.

2.11Nitrate reductase (NR) assay

NR activity was assayed as nitrate reduction (with sodium dithionite-reduced methyl viologen as the electron donor) [24]. The amount of activity was expressed as μg nitrite formed mg Chl a−1 min−1.

2.12Protein and enzyme electrophoresis

Samples for protein profile and assay of enzyme activities were prepared from thoroughly washed cyanobacterial pellets rewashed with extraction buffer (0.0625 M Tris–HCl, pH 6.8). Cyanobacterial pellets were homogenised using a pre-chilled mortar and pestle in the presence of glass powder (∼0.5 mm) adding ice-cold extraction buffer. The samples were centrifuged at 15 000×g for 15 min and the process was repeated twice to obtain clear supernatants. The amounts of proteins were estimated [20] and used for both sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) and activity staining of gels.

Electrophoresis was carried out at 20±2°C with a 1.5 mm thick polyacrylamide gel in Tris–glycine buffer (pH 8.3). In the case of SDS–PAGE, SDS was included in the running buffer (0.1%) as well as in sample buffer (10%). The sample was boiled for 3 min with sample buffer and centrifuged briefly before loading. In both PAGE and SDS–PAGE, a uniform amount (300 μg) of protein was loaded to each well. Samples were then electrophoresed at 60 V through the stacking gel (6%) and at 120 V through the separating gel (10%) [25].

2.13Denatured gel – staining and analysis

After electrophoresis, SDS gels were fixed and stained with Coomassie brilliant blue R-250 (SRL, India) for 3 h and destained with distilled water containing 5% methanol and 7% glacial acetic acid in a shaker (50 rpm) at room temperature. The images were captured with a CCD camera and the protein profiles were analysed by the gel documentation system (Alpha Imager™ 2200, USA).

2.14Native gel – enzyme staining

2.14.1Rubisco (EC 4.1.1.39)

Electrophoresed bands of Rubisco isoenzymes were fixed and stained by briefly soaking the gel in 100 ml of fixative (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 destained using the same fixative.

2.14.2Superoxide dismutase (EC 1.15.1.1)

Activity staining for superoxide dismutase (SOD) on gel [27] and the identification of metal prosthetic groups in the active enzyme were carried out following Chadd et al. [28]. The gels were soaked in staining solution containing 50 ml of 50 mM Tris–HCl (pH 8), 10 mg NBT (Hi-Media, India), 1 mg EDTA (Sigma, USA) and 2 mg riboflavin (Sigma, USA) for 30 min in the dark at room temperature (25°C) and then illuminated on a light box with white fluorescent light (80 μE m−2 s−1) for 30 min or until achromatic bands appeared. For the identification of metal prosthetic groups, enzyme activity was tested by soaking one gel in buffer containing 5 mM H2O2, another in buffer containing 2 mM KCN and one more in buffer without any inhibitors (H2O2 or KCN) for 20 min in the dark, prior to exposing all three gels to staining solution and illumination.

2.14.3Peroxidase (EC 1.11.1.7)

Activities of peroxidase isoenzymes were detected by incubating the gel in 100 ml of 0.1 M sodium acetate buffer (pH 4.6) containing 30 mg 3,3′-diaminobenzidine tetrahydrochloride (SRL, India). The reaction was initiated by adding 250 μl of 30% hydrogen peroxide (Qualigens, India) and let stand until intense brown bands appeared [29].

3Results

3.1Growth

Prolonged starvation for over 5 months caused the majority of cells to turn yellow and fragmentation of filaments resulted in unicells of varying morphology (Figs. 1 and 2). Few hormogonia and live filaments with less pigmentation were also noticed (Fig. 3). These cells returned to normal (Fig. 5) and started growing when reinoculated to complete medium (Fig. 4).

Figure 1.

Prolonged starvation for over 5 months caused the majority of cells to turn yellow.

Figure 2.

Fragmentation of filaments resulted in unicells of varying morphology.

Figure 3.

Few hormogonia and live filaments with less pigmentation.

Figure 5.

Appearance of normal morphology.

Figure 4.

Initiation of growth after transfer to complete medium.

Nitrogen limitation reduced growth by 17% of fresh and dry biomass compared to growth in nitrogen-supplemented medium (Table 1). In N-starved conditions, chlorophyll a and carotenoids were reduced by 31.81% and 18.38% compared to their respective controls (Table 1). Among phycobili-pigments, phycocyanin alone was decreased by 19.67%, while the amount of allophycocyanin remained almost equal to that of cultures grown in nitrogen-supplemented medium (Table 1). From the absorption spectrum of cell-free extract, no phycoerythrin peak could be detected in this organism (Fig. 6).

Table 1.  Effect of nitrogen limitation/starvation on end-point growth, cellular contents, nitrogen-assimilating enzymes and hydrogen peroxide production of O. willei BDU 130511
  1. ND, not detected.

  2. aμg γ-glutamyl hydroxamate formed mg Chl a−1 min−1.

  3. bμg nitrite formed mg Chl a−1 min−1.

 ControlN-deprived
End-point biomass (day 8)  
Fresh weight (g)3.55±0.1772.94±0.212
Dry weight (mg)378.96±18.844313.4±22.613
Cellular contents  
Chlorophyll a (μg g−1 DW)769.27±12.091524.59±0.462
Carotenoids (μg g−1 DW)277.80±7.659228.52±20.883
Phycocyanin (mg g−1 DW)99.05±2.01579.56±6.914
Allophycocyanin (mg g−1 DW)19.14±0.35519.43±1.066
Total protein (mg g−1 DW)370.00±11.98226.3±22.34
Lipid (mg g−1 DW)88.38±1.91065.33±1.109
Nitrogenase activity (2, 24, 48 and 72 h)NDND
GS activitya0.59±0.0495.79±0.674
NR activityb13.91±0.2292.27±0.207
H2O2 production (OD475)0.6196±0.0030.9438±0.005
Figure 6.

Relative absorption spectra of phycobilins of O. willei BDU 130511 extracted in 0.05 M phosphate buffer by repeated freeze-thawing; C, control; T, nitrogen-starved cells.

3.2Lipids and fatty acids

Nitrogen starvation resulted in a decrease in total lipid content (26.08%) and caused qualitative and quantitative variations in whole cell fatty acids. In nitrogen-starved cultures, fatty acids such as pentadecanoic acid (C15:0), oleic acid (C18:1 cis), linoleic acid (C18:2 cis), behenic acid (C22:0) and eicosapentaenoic acid (C20:5) disappeared, 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 and γ-linolenic acid, which increased in nitrogen-starved conditions by 14.53% and 40.55% respectively (Table 2).

Table 2.  Fatty acid content (mg g lipid−1) of O. willei BDU 130511 grown under different nitrogen conditions
  1. ND, not detected.

Fatty acidN+N
Lauric acid (C12:0)0.5530.647
Myristic acid (C14:0)1.2580.580
Pentadecanoic acid (C15:0)0.031ND
Palmitic acid (C16:0)4.3041.981
Palmitolic acid (C16:1)12.9675.756
Heptadecanoic acid (C17:0)4.9021.458
Oleic acid (C18:1 cis)0.150ND
Linoleic acid (C18:2 cis)0.114ND
γ-Linolenic acid (C18:3Y)0.1070.180
Eicosenoic acid (C20:1)ND2.401
Eicosapentaenoic acid (C20:5)1.226ND
Behenic acid (C22:0)1.510ND

3.3Hydrogen peroxide

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

3.4Proteins and enzymes

Nitrogen starvation caused a decrease in protein content by 38.83% as estimated spectrophotometrically. The protein profiles of nitrogen-starved cells differed markedly from those of control cells. In nitrogen-starved cells, three polypeptides, of 11.2, 16.7 and 55 kDa, found in control were missing, while two new polypeptides of molecular mass 52.6 and 90.5 kDa appeared. There was also a quantitative increase of 59.7-kDa polypeptide in nitrogen-starved cells compared to control (Fig. 7).

Figure 7.

Modification of protein synthesis in O. willei BDU 130511 during nitrogen (NaNO3) starvation. Protein samples (300 μg), from cells grown for 4 days in complete ASN III medium (lane C) and ASN III medium devoid of NaNO3 (lane T), were resolved by SDS–PAGE (10%) and visualised by Coomassie staining. To determine the molecular masses of prominent polypeptides either missing, appearing new or at enhanced levels, a mixture of six standard proteins (lane M) from Amersham Biosciences was run simultaneously.

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

Figure 8.

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

The SOD isoenzyme profiles of both control and nitrogen-starved cultures were essentially similar except for the disappearance of an isoform of Mn-SOD (Rm 0.855) in nitrogen-starved cells. A prominent isoenzyme (Rm 0.596) appearing in both cases was identified as Fe-SOD as it was sensitive to H2O2 incubation and the four less achromatic bands (Rm 0.690, 0.716, 0.787 and 0.855) were considered Mn-SODs as they were resistant to both H2O2 and KCN (Fig. 9).

Figure 9.

Activity staining for SOD and identification of metal prosthetic groups in control (lane C) and nitrogen-starved (lane T) cells of O. willei BDU 130511.

Nitrogen starvation induced four new isoforms of peroxidase (at Rm 0.107, 0.124, 0.454 and 0.538). Of these, the expression of the isoform with Rm 0.538 was high; however, the isoform with Rm 0.378 present in control was absent under nitrogen starvation (Fig. 10).

Figure 10.

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

4Discussion

It is well known that a large part of the world's oceans is nutritionally depleted especially in nitrogen and marine cyanobacteria are able to persist and adapt their metabolism to this distinct environmental stress. Deficiency of combined nitrogen in nature is a threat to non-nitrogen-fixing cyanobacteria. The present laboratory study has shown the high degree of physiological adaptability of O. willei BDU 130511 to nitrogen stress. To overcome the stress, organisms generally have two types of defence strategies: (i) mechanisms that prevent interaction with the stress factors and (ii) those that counteract the stress-induced damages. The results of this study indicate that O. willei BDU 130511 has opted for the second strategy.

O. willei BDU 130511 could not grow in N-free medium due to the inability of this non-heterocystous cyanobacterium to scavenge atmospheric N2. No nitrogenase activity could be detected by acetylene reduction assay in MAD, MALD, MAL and AL conditions (Table 1). Acetylene reduction activity has been accepted as a simple and efficient way of estimating nitrogenase activity [9–13,30–36], although 15N fixation, molecular and antibody assays will prove useful in identifying the presence or absence of the enzyme per se. The fact that the cyanobacterium could not grow in the absence of combined nitrogen in the medium and the lack of acetylene reduction activity indicate that the organism is unable to use atmospheric N2 in required amounts.

Nitrogen limitation in O. willei BDU 130511 resulted in the reduction of photosynthetic pigments such as chlorophyll a, carotenoids and phycocyanin resulting in chlorosis. It is known that in photosynthetic organisms N limitation triggers ordered degradation of phycobilisomes, ribosomes and thylakoid membranes [37]. This explains the lowered pigment content, lipid content and change in pattern of fatty acids observed in O. willei BDU 130511 in this study as well as in Aphanocapsa 6308 [38], Agmenellum quadruplicatum[37] and Synechococcus 6301 [39] by others. Furthermore, nitrogen limitation leading to catabolisation of lipids and fatty acids resulting in their reduction as found in Spirulina platensis and Anacystis nidulans[40] was also observed in this cyanobacterium (Table 2). A decreased level of total lipid content is believed to indicate a reduced carbon storage mechanism and leads to the availability of carbon skeletons to synthesise proteins or enzymes required for the defence [41]. The enhanced levels of C12:0 and C18:3Y fatty acids as well as the newly synthesised C20:1 (18.47% of total fatty acids) under nitrogen starvation in O. willei BDU 130511 (Table 2) may help the organism to maintain membrane fluidity to overcome stress and retain cellular integrity as suggested by Smith et al. [42] and Floreto et al. [43]. Reduction in levels of some fatty acids and disappearance of certain others (Table 2) could lead to reutilisation of these fatty acids for other metabolic purposes including the increased production of certain other fatty acids mentioned above.

Most photosynthetic micro-organisms depend on either ammonium (NH4+) or nitrate (NO3) as their sole source of combined nitrogen. However, cyanobacteria have the ability to use a wide variety of nitrogen sources such as ammonia, nitrate, nitrite, urea, amino acids, nucleosides or bases [44,45], atmospheric nitrogen (some cyanobacteria only) and even synthetic compounds like ampicillin [46]. In general, the preferential order of nitrogenous compounds for cyanobacteria and other algae is ammonia>nitrate or urea>other organic compounds [45].

GS is an efficient ammonia scavenger in cyanobacteria [47]. Enhanced GS activity of O. willei BDU 130511 under nitrogen starvation (Table 1) may probably be due to the channeling of available nitrogen through breakdown of organic nitrogenous compounds from dead cells as revealed by microscopic observation (Fig. 3) and breakdown of its own phycocyanin [48,49]. In addition, several other reasons suggested by others such as increased expression of ammonium transport (amt) genes as in Synechocystis sp. [50], possession of multiple GS enzymes [45] and decreased activity of NR as a process of saving energy reported in Prochlorococcus sp. strain PCC 9511 [51] (where the genes encoding the nitrate-assimilating system were lacking in extremely nitrogen (nitrate or nitrite)-scarce environments) could also contribute to the enhancement of GS activity. In this study, a significant reduction of NR activity during nitrogen starvation was observed (Table 1) which indicates the control of expression of genes for nitrate-assimilating enzymes, which may help the cyanobacterium O. willei BDU 130511 to survive successfully in oligotrophic or ultra-oligotrophic environments.

Nitrogen starvation had caused profound alterations in proteins, both qualitatively and quantitatively, in O. willei BDU 130511. Prominent among the impaired proteins are phycobiliproteins (Fig. 6, Table 1). In Synechococcus sp. also, severe nitrogen limitation was reported to cause proteolytic degradation of phycobiliproteins [45]. Degraded products of phycobiliproteins could provide C and N elements for the synthesis of other cellular constituents required during nutrient deprivation [48] and reduce absorption of excitation energy, making cells less susceptible to photodamage [52].

Two of the three newly synthesised proteins (Fig. 7) based on their molecular masses might be considered stress proteins (52.6 and 90.5 kDa) or hydrolytic enzymes while the 59.7-kDa protein whose level is increased during N starvation (Fig. 7) could be a chaperonin, which helps in protein protection and repair mechanism under stress as reported by Sanders [53].

Only a meagre amount of information is available on the effect of N limitation and starvation on photosynthesis and respiration. In the present study, N starvation has been shown to result in reduced chlorophyll a synthesis (Table 1) as well as alteration in Rubisco isoenzyme profile, due to the disappearance of a prominent band in the Rubisco profile of nitrogen-starved cells (Fig. 8). This could be due to the interaction of active oxygen species with essential sulfhydryl groups of the enzyme as reported in several chloroplast enzymes of the carbon di-oxide fixation pathway [54]. Photosynthesis was thus affected and reduced the growth of O. willei BDU 130511. Similar observations were reported in Synechocystis PCC 6803 cells, where the rate of both O2 evolution and CO2 fixation declined under nitrogen starvation [55].

Deficiency of combined nitrogen could also have created oxidative stress. In N-starved cultures of Spirulina platensis, the rate of respiration was reported to have increased both at normal and at high CO2 concentration [49]. A high rate of respiration might be one of the reasons for induction of oxidative stress through the generation of active oxygen species [56]. Hence, efficient scavenging is important to prevent oxidative damage to the organism as active oxygen species are highly reactive with the proteins and membranes. SOD is the first line of defence against active oxygen species. In cyanobacteria, three isoforms of SOD are reported, viz. Mn-SOD, Fe-SOD and Cu/Zn-SOD (see [28]). It has been found that Mn-SOD protects thylakoid membranes against superoxides, while Fe-SOD protects cytosolic contents [57]. In O. willei BDU 130511, under nitrogen starvation, Fe-SOD could play a major role in dismutating the superoxide ions, since Fe-SOD was considered to be the sole superoxide-scavenging enzyme in Synechocystis sp. strain PCC 6803 [56]. Billi and Caiola [58] also presumed that Fe-SOD protects the cyanobacterium against free oxygen radicals generated during oxidative stress due to nitrogen starvation.

The presence of SOD indicates dismutation of superoxides to hydrogen peroxide, a powerful oxidant which provides the cellular signal for induction of defence mechanisms [59,60]. It also disturbs the cellular redox state and its uncontrolled accumulation may cause damage of membrane lipids, DNA, and finally death of the cell [61]. In the present study an increased level of H2O2 was observed in nitrogen-starved cells (Table 1). Hydrogen peroxide could be useful in degrading organic matter of surrounding dead cells and make nutrients available for the surviving cells. However, since H2O2 is the precursor for the most reactive and destructive active oxygen species in the presence of O2 and Fe2+[62], the removal of H2O2 from the vicinity of living cells is essential. Probably for this reason an increased number of isoforms of peroxidase were observed in this cyanobacterium (Fig. 10). Of the four newly synthesised peroxidase isoenzymes, observed under nitrogen stress, three were intensely stained. This induced synthesis of new isoforms could be to play a special role in neutralising the lethal effect of H2O2 in living cells favouring the organism to survive in nitrogen-limited environments.

5Conclusion

Cyanobacteria (blue-green algae) are of evolutionary interest, as they are highly adaptable to various environmental extremes. However, the mechanisms involved in environmental tolerance by cyanobacteria are not fully understood, especially in a nitrogen-limited environment. From the results of the present study, it could be inferred that the marine cyanobacterium O. willei BDU 130511, although adversely affected by nitrogen starvation, is able to save some cells or filaments from the stress with the help of antioxidative enzymes, and these cells upon availability of nitrogen return to normal growth. Thus the study helps to understand the survival strategy of a non-heterocystous, non-nitrogen-fixing marine cyanobacterium in the oligotrophic environment.

Acknowledgements

The authors thank the Department of Biotechnology (DBT), Government of India, for financing the facility and fellowship to S.K.S. Dr D. Prabaharan and Dr A. Kalib are thanked for their valuable suggestions in improving the manuscript.

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