1. INTRODUCTION

Salinity is a crucial abiotic environmental factor affecting aquatic organisms, with variations resulting from tidal surges, coastal runoff, and rainfall, each exhibiting diverse durations and ranges (Velasco et al. 2018). Algae inhabiting biotopes with fluctuating salinities have become model organisms for salt tolerance studies, providing valuable insights into salt acclimatization mechanisms that are often more complex in higher plants (Borovkov et al. 2019). Among these, unicellular green algae are particularly notable for their remarkable adaptability to highly saline conditions, making them valuable models for identifying survival mechanisms in extreme environments (Chen and Jiang 2009). Certain microalgae, such as Dunaliella sp., can tolerate a broad spectrum of salinities, surpassing the maximum range that is suitable for most plant species. Their adaptation involves both short- and long-term responses, including osmotic adjustment through intracellular glycerol accumulation and efficient Na⁺ extrusion via plasma-membrane transporters. The absence of a rigid cell wall in Dunaliella enables swift changes in cell volume, allowing rapid responses to fluctuating salt concentrations (Kaçka and Dönmez 2008).

To cope with saline stress, algae and plants assimilate a diverse array of metabolites, including sugars (fructose, sucrose, and trehalose) and charged molecules (proline and glycine betaine), which function as osmoprotectants (Banu et al. 2009). The accumulation of these compatible solutes allows algae to retain water and maintain cellular turgor under hyperosmotic stress. In addition to osmotic regulation, salinity influences various physiological and biochemical processes, including respiration rate, mineral distribution, ion toxicity, photosynthetic rate, and membrane permeability (Sudhir and Murthy 2004). Algal adaptation is often accompanied by dynamic rearrangements in lipid and fatty acid profiles and increased lipid accumulation, particularly under environmental stress conditions, such as nutrient depletion or increased salinity. These lipid reserves serve as both metabolic buffers and energy storage molecules, enabling survival and subsequent recovery.

The study of microalgal lipid productivity in saline media is of growing importance for sustainable biofuel applications. Environmental factors, particularly salinity, play a pivotal role in regulating lipid production in microalgae (Zhila et al. 2011). Numerous reports have shown that increased salinity may stimulate lipid accumulation in species such as Isochrysis sp. and Nannochloropsis oculata (Renaud et al. 1994). Understanding these salt-induced metabolic shifts is vital for optimizing microalgal biofuel production systems, especially as freshwater resources become increasingly scarce (Chokshi et al. 2017;Borovkov et al. 2019). However, the responses of many freshwater microalgae to seawater-level salinities remain poorly characterized.

To address this, the present study investigated the morphological and physiological effects of artificial seawater media on two freshwater species, Chlorella thermophila (a green alga) and Anabaena variabilis (a cyanobacterium). These species were selected because of their contrasting phylogeny and morphology, as well as their recognized bioindustrial relevance. Chlorella thermophila is known for its thermotolerance and robust protein and lipid production, which are desirable qualities for scalable biomass cultivation. Anabaena variabilis combines nitrogen-fixation capacity with high environmental plasticity, further expanding its biotechnological potential (Chokshi et al. 2017). By comparing both, this study aims to clarify the adaptive mechanisms and practical implications for saline microalgaebased biorefinery processes.

2. MATERIALS AND METHODS

2.1. Collection, morphological identification of algal species

At least ten algal samples were collected from each site in July and August, targeting rice fields, ponds, and runoff water from the Johila Dam in the Lalpur region of Amarkantak, Madhya Pradesh, India. The collected samples were examined microscopically using an EVOS XL Core microscope (Thermo Scientific, USA) to assess the algal diversity. For taxonomic identification, standard references were consulted, including Desikachary (1959) and Bellinger and Sigee (2015).

2.2. Isolation and purification of algal species

Algal strains were isolated and purified under controlled laboratory conditions using sterile equipment. Cultures were maintained in 250 mL Erlenmeyer flasks containing BG11 medium for Chlorella thermophila and BG11 (N^-) medium for Anabaena variabilis. All cultivations were performed using a batch system and incubated at a constant temperature of 25±2°C. Cultures were exposed to a 12 : 12 h light-dark cycle, with illumination provided at 200 μmol photons m^-2 s^-1 using cool-white fluorescent lamps. The pH of the medium was adjusted and maintained at 7.5 for the duration of the experiment. The cultures were agitated daily by gentle swirling to avoid cell sedimentation. The isolation and purification process employed serial dilution, plating, and repeated streaking techniques (Guillard 2005), and all cultures were monitored for two weeks to ensure the growth of unialgal axenic cultures before subsequent experimental treatments.

2.3. Algal species cultivation

For the experiment, Anabaena variabilis and Chlorella thermophila were grown in both the control and treatment groups, with n=3 independent biological replicates per condition (i.e., three separate flasks for each medium and species). The control group was maintained in standard 1/4th strength BG11 medium (for C. thermophila) or BG11 (N^-) medium (for A. variabilis) (Tripathi et al. 2003), whereas the treatment group was cultivated in artificial seawater medium (SWM). Each flask contained 150 mL of culture, the and initial inoculum densities were standardized for all samples. Artificial seawater medium (SWM) was prepared according to the UTEX algal culture collection protocols. The major components per liter include NaCl 18 g L^-1; MgSO₄·7H₂O 2.6 g L^-1; KCl 0.6 g L^-1; NaNO₃ 1.0 g L^-1; CaCl₂·2H₂O 0.3 g L^-1; KH₂PO₄ 0.05 g L^-1; Tricine 4.48 g L^-1; and NH₄Cl 0.027 g L^-1. In addition, trace nutrient solutions were incorporated: P-II Metal Solution at 10 mL L^-1, which contained compounds such as Na₂EDTA·2H₂O, H₃BO₃, FeCl₃·6H₂O, MnSO₄·H₂O, ZnSO₄·7H₂O, and CoCl₂·6H₂O in trace amounts; and Chelated Iron Solution at 1 mL L^-1, derived from a stock containing Na₂EDTA·2H₂O, FeCl₃· 6H₂O, and 0.1 M HCl and Vitamin B₁ at 1 mL L^-1 concentration. The pH of the medium was adjusted to 7.5. All cultures were grown for a total duration of 20 days under a 12 : 12 h light-dark cycle (Tripathi et al. 2004), at 25±2°C and an illumination intensity of 200 μmol photons m^-2 s^-1 (provided by cool white fluorescent lamps). After the initial 10 days of growth, samples were harvested from each flask to analyse cell biomass, lipid and protein content, chlorophyll a concentration, cell volume, and images were collected for morphological assessment. Following this, a subculture from each replicate flask was carefully transferred to the corresponding artificial seawater medium (SWM). Parallel BG11 control cultures were maintained without any transfer. After an additional 10 days (i.e., day 20), all remaining cultures from both the control (BG11/BG11-N^-) and treatment (SWM) groups were harvested, and the same set of analyses was performed. All measurements were performed in triplicate for statistical analysis.

2.4. Cell size and cell volume measurement

The size of the algal cells was measured using the freely available software, ImageJ. Calibrations for cell size measurements were performed using a Stage Micrometer SM-001 (Genex Erma, India). All measurements were performed in triplicates. The cell volume of both algal species was calculated using the method described by Matthews (2016). The following equations were used for Chlorella thermophila (sphere) and Anabaena variabilis (spheroids):

where W is the width of a single cell, and L is the length of a single cell.

2.5. Estimation of biomass of selected algal species

The biomass of the selected algal species was collected after 10 and 20 d of inoculation. The samples were centrifuged using a Hermle Z366 (Hermle, USA) centrifuge at 4,100×g for 10 min at room temperature. The pellets obtained by centrifugation were washed with distilled water and then dried. The biomass percentage was determined using the following equation:

2.6. Estimation of photosynthetic pigment

Chlorophyll a and carotenoids were extracted from algal samples following the method described by Arnon (1949). Five milliliters of algal culture was harvested and centrifuged using a Hermle Z366 (Hermle, USA) centrifuge at 5,000 rpm for 10 min at RT. The pellet was suspended in 80% acetone and stored at 4°C for 24 h. The sample was centrifuged again, and the absorbance of the supernatant was recorded at 663, 645, and 480 nm using an Orion^TM AquaMate 8100 UV visible spectrophotometer (Thermo Scientific, USA). The chlorophyll a and carotenoid contents were estimated using the following equation:

where A₆₆₃, A₆₄₅, and A₄₈₀ refer to the absorbance values of the sample measured at 663 nm, 645 nm, and 480 nm wavelengths, respectively, as determined by spectrophotometry.

2.7. Determination of protein

Protein content was quantitatively measured as described by Lowry et al. (1951). Absorbance was measured using an Orion^TM AquaMate 8100 UV Visible spectrophotometer (Thermo Scientific, USA).

2.8. Extraction of lipids

Lipids were extracted from the cells using the method outlined by Bligh and Dyer (1959), with minor modifications. Algal samples (20 mg) were first reconstituted in 0.8 mL of chloroform/methanol (in a ratio of 2 : 1, v/v), followed by the addition and agitation of 100 μL of 0.9% KCl solution. After vortexing, the sample mixture was centrifuged at 3,000×g for 5 min at room temperature to facilitate phase separation. The lower chloroform phase, which was enriched with lipids, was then isolated. Lipid extraction was performed three times, and the upper layer was collected each time. The pooled lower chloroform phase was then evaporated to dryness, yielding a lipid-rich residue. These lipids were subsequently dissolved in 20 μL of chloroform and stored at -20°C for further analysis.

2.9. Lipid productivity using FTIR analysis

Total lipid production was calculated using the following equation:

where Y_L is the total lipid production, β_t is the lipid content at time t (demonstrated by FTIR spectra at 2,926 cm^-1), and X_t is the dry cell weight at time t. Here, t is 10 and 20 days (Liu et al. 2013).

2.10. Fatty acid analysis

Fatty acid profiling of algal oil samples was performed by GC-MS using a Shimadzu GC-2010 Plus (Japan), following the Bureau of Indian Standards methods (Kisan et al. 1976). Specifically, 20 mg of algal oil was saponified with 1 mL of a saturated KOH-CH₃OH solution at 50°C for 10 min, followed by methanolysis with 5% HCl in methanol at 60°C for another 10 min in screw-capped test tubes. The methyl fatty acids were separated by adding 2 mL of water, and the fatty acid phase was recovered using a separating funnel. Exactly 1 μL of the solvent-free methyl fatty acid sample was injected into the GC column (Restek-Stabilwax; Polyethylene glycol, 30 m length, 0.25 mm internal diameter). The flow rates of the carrier gases were adjusted to 30 mL min^-1 for nitrogen and 40 mL min^-1 for hydrogen. Chromatographic data were recorded and compared with the fatty acid standard mixtures.

2.11. Transesterification of algal oil

Approximately 40 mg of algal oil was placed in a round-bottom flask and mixed with 1.5 mL of methanolic sulfuric acid containing 2% sulfuric acid in methanol (v/v) and refluxed at 60°C for 4 h with continuous shaking.

2.12. Fourier transform infrared spectra measurement

FTIR analysis was performed on the dried algal biomass to characterize its biochemical composition and quantitatively assess major cellular macromolecules, especially proteins and lipids. Dried algal samples were finely ground and analyzed using a Nicolet iS5 Fourier Transformation Infrared Spectrophotometer (Thermo Scientific, USA) equipped with an ATR accessory. Spectra were recorded over the wavenumber range of 4,000- 400 cm^-1, with a minimum of 32 scans per sample and a resolution of 4 cm^-1. Each sample was measured in duplicate to ensure its reproducibility. Spectral data were processed using OMNIC^TM Series Software (Thermo Fisher Scientific, USA) for baseline correction and normalization. Specific absorption peaks were used as biomarkers for macromolecular groups: the region of 3,000-2,800 cm^-1 (centered at ~2,925 cm^-1) was assigned to C-H stretching vibrations of aliphatic chains, indicative of total lipid content; the amide I band (~1,650 cm^-1) and amide II band (~1,545 cm^-1) were assigned to proteins, while the region near 1,740 cm^-1 (C=O stretching) was ascribed to ester linkages in lipids. Peaks between 1,200 and 900 cm^-1 corresponded to carbohydrates (C-O-C and C-O stretching). Quantitative analysis was performed by calculating the relative intensities and integrated areas of the characteristic absorption bands, particularly at 2,925 cm^-1 for lipids and 1,650 cm^-1 for proteins, following established protocols (Liu et al. 2013). For comparison, the intensities of these bands were normalized to the total absorbance in the spectrum. An increase in the intensity or area under the respective peaks was interpreted as an increase in the concentration of the corresponding biochemical constituents in algal biomass. These FTIR-derived biochemical estimations were further used to complement gravimetric lipid estimations and to support the interpretation of physiological changes under different culture conditions.

3. RESULTS AND DISCUSSION

3.1. Isolation, purification, and cultivation of algal species

Microscopic examination and morphological identification of water samples collected from rice fields, ponds, and runoff in Amarkantak, Madhya Pradesh, led to the selection of two taxonomically and physiologically distinct freshwater algal species: C. thermophila and A. variabilis. Both species were successfully isolated and purified, as detailed in Section 2.2. Both C. thermophila and A. variabilis were molecularly identified by sequencing and their nucleotide sequences are available in the NCBI GenBank database under accession numbers PP968571 and PX339966, respectively. Cultivation experiments revealed notable differences in salinity responses. C. thermophila exhibited consistent and vigorous growth in both BG11 and artificial seawater medium, demonstrating pronounced adaptability likely attributable to its thermophilic nature and inherent metabolic flexibility. In contrast, A. variabilis showed optimal development in BG11 (N^-) medium and a comparatively slower acclimation to saline conditions, occasionally manifesting filament aggregation, consistent with its requirement for gradual osmotic adjustment and reliance on nitrogen fixation (Banu et al. 2009;Chokshi et al. 2017). These distinct physiological responses underscore species-specific tolerance mechanisms to salinity stress and highlight how environmental factors, such as medium composition, temperature, and illumination, interact to influence algal growth and biomass potential (Zhila et al. 2011;Borovkov et al. 2019;Almutairi et al. 2020). The specific cultivation parameters used for both C. thermophila and A. variabilis in this study, including temperature, light intensity, photoperiod, pH, and culture system, are summarized in Table 1. Notably, the ability of C. thermophila to grow successfully in artificial seawater medium aligns with previous research demonstrating enhanced lipid production in microalgae cultivated under saline or seawater-based conditions, as observed in species such as Synechocystis sp. and Nannochloropsis (Bartley et al. 2013;Cai et al. 2013;Sheets et al. 2014;Zhao et al. 2015). Similarly, Chen et al. (2013) reported significantly higher lipid yields in Chlorella sorokiniana when cultivated with deep-sea water supplementation. Collectively, these findings confirm that microalgae sourced from freshwater environments can exhibit substantial adaptive plasticity and favourable biochemical profiles when exposed to saline media, supporting their potential application in sustainable biofuel production and saline water-based cultivation systems for biofuel production.

3.2. Effects of seawater medium on cell size, volume and biomass percentage

Both C. thermophila and A. variabilis exhibited marked increases in cell size and volume when cultivated in seawater medium (SWM) compared to the standard BG11 medium. After 10 days of growth in SWM, the cell volume of C. thermophila increased dramatically from 5.83 μm³ to 54.76 μm³, while A. variabilis showed an increase from 43.8 μm³ to 50.36 μm³ (Table 2). The observed increases in cell size (Fig. 1), volume (Fig. 2), and biomass percentage (Fig. 3) in both C. thermophila and A. variabilis under artificial seawater stress likely arose from coordinated physiological and biochemical responses. This substantial enlargement is an adaptive response to the osmotic stress imposed by higher salinity in the SWM, and the expansion of cell volume can be attributed to the accumulation of osmoprotectants, such as proline and glycine betaine, which maintain cellular turgor and function in hypertonic environments (Kaewkannetra et al. 2012). Elevated salinity not only drives osmotic adjustment through compatible solute synthesis but also induces oxidative stress, prompting the production of antioxidant enzymes, such as superoxide dismutase and catalase, to mitigate reactive oxygen species and preserve cellular integrity (Sudhir and Murthy 2004). Concurrently, these conditions stimulate shifts in key metabolic pathways related to lipid and carbohydrate metabolism, further supporting osmotic balance and enhancing biomass accumulation (Banu et al. 2009;Zhila et al. 2011;Borovkov et al. 2019). The observed increase in cell size under saline conditions aligns with reports for other microalgae, such as Chlorella sorokiniana and Chlorella sorokiniana HS1, which also respond to salinity stress with increased cell size (Jiang et al. 2012;Kim et al. 2016). However, this response is not universal; some species, including Chlamydomonas sp., Chlorella vulgaris, C. salina, C. emersonii, and Scenedesmus opoliensis, often exhibit a reduction in cell size under similar conditions (Demetriou et al. 2007;Neelam and Subramanyam 2013;Talebi et al. 2013;Khona et al. 2016). The reduction in cell size in these species may serve as an energy-saving strategy that helps maintain osmotic balance and minimize water loss under saline stress.

In addition to changes in cell size, A. variabilis filaments cultivated in SWM displayed notable aggregation (Fig. 1d), a phenomenon that was absent in the BG11 control group (Fig. 1a). This aggregation could be the result of charge neutralization between the negatively charged algal cell surfaces and positively charged ions present in the seawater medium, which is consistent with the findings that aggregation in cyanobacteria and other microalgae is often induced by the screening or neutralization of surface electrostatic charges by divalent or monovalent cations in saline environments (Jiang et al. 2012;Lesniewska et al. 2024). Similar aggregation has been observed in other microalgae, such as Nannochloropsis sp. and Chlorella sorokiniana, when grown in media with high NaCl concentrations (Jiang et al. 2012).

Fig. 3 compares the biomass percentages of C. thermophila and A. variabilis after 10 days of growth in BG11 and SWM. For both species, exposure to artificial seawater resulted in a notable increase in biomass percentage compared to the control and surpassed the biomass achieved in BG11 medium over 20 days. This enhancement indicates an adaptive physiological response to saline conditions, likely mediated by osmotic adjustment and metabolic shifts that promote cell growth and biomass accumulation in the cells. These findings underscore the potential of using artificial seawater to cultivate freshwater microalgae and cyanobacteria to achieve higher biomass yields, which is advantageous for biofuel and bioproduct applications (Chokshi et al. 2017;Borovkov et al. 2019).

Marine algae have evolved genetic mechanisms to cope with salinity, which are embedded in their genomes through natural selection. In contrast, freshwater species such as C. thermophila and A. variabilis must rapidly adapt to high salinity, requiring both short-term physiological adjustments and, over time, the accumulation of advantageous mutations (Shetty et al. 2019). Although seawater can supplement but not fully replace freshwater media for these species, developing cultivation methods that leverage seawater is increasingly important, given the abundance of freshwater algal strains available for biofuel production (Mata et al. 2010;Jiang et al. 2012). Furthermore, the observed increases in the cell size, volume, and biomass of C. thermophila and A. variabilis under seawater highlight their adaptive capacity and potential for sustainable biomass and biofuel production in saline environments.

3.3. Photosynthetic pigment content

Fig. 4 presents the changes in chlorophyll a and carotenoid content for both C. thermophila and A. variabilis after 10 days of cultivation in BG11 and artificial seawater medium (SWM). For both species, a distinct increase in chlorophyll a content was observed when cells were grown in SWM relative to the BG11 control, with C. thermophila showing a greater magnitude of response than A. variabilis. The carotenoid content also increased under saline conditions for both strains. These results suggest that moderate salinity stimulates the synthesis or stabilization of photosynthetic pigments, potentially as part of an adaptive strategy to maintain or enhance the photosynthetic apparatus under osmotic stress. Notably, the more pronounced increase in pigment in C. thermophila supports its higher physiological plasticity and greater tolerance to saline environments compared to A. variabilis. The observed changes in pigment content are consistent with previous studies, indicating that exposure to salinity can upregulate pigment and antioxidant production in microalgae, providing increased protection against oxidative stress associated with saline conditions (Zhila et al. 2011;Chokshi et al. 2017;Borovkov et al. 2019).

3.4. Determination of protein

Fig. 5 shows the protein (mg mL^-1) content of both selected algal species grown in different media types. The protein content increased in Chlorella thermophila when grown in SWM and BG11 media for 10 days. Moreover, Anabaena variabilis also showed an increase in protein content in SWM compared to that in BG11 media. Villaro et al. (2023) conducted production of the microalga Arthrospira platensis BEA 005B in 11.4 m³ raceway photobioreactors using a culture medium based on commercial fertilizers, with options of either freshwater or seawater. The biomass productivity of the reactors, operated at a fixed dilution rate of 0.3 d ^-1, decreased from 22.9 gm^-2 day^-1 when using freshwater to 16.3 gm^-2 day^-1 when using seawater for biomass production. Although the protein content of the seawaterproduced biomass was lower, it exhibited higher levels of essential amino acids such as valine, leucine, and isoleucine. Seawater also stimulated carotenoid production and induced changes in fatty acid synthesis and accumulation. Notably, the biomass produced using seawater showed a 319% increase in oleic acid and a 210% increase in eicosenoic acid content compared to that produced using freshwater.

The observed increase in protein content under saline conditions is consistent with previous findings in other microalgae, where salt stress acts as a metabolic trigger for the enhanced production of stress-response proteins and compatible solutes, aiding cellular adaptation and maintenance under osmotic challenge (Chokshi et al. 2017;Borovkov et al. 2019). Villaro et al. (2023) similarly highlighted the impact of seawater-based media on Arthrospira platensis, noting that although total protein percentages can fluctuate in saline conditions, seawatergrown microalgae often display altered amino acid profiles, sometimes with increased proportions of essential amino acids reflecting a shift in nitrogen metabolism and adaptation to ionic stress. These results suggest that cultivating freshwater algal species in artificial seawater can not only sustain but also stimulate cellular protein production, with potential implications for both nutritional quality and stress resilience. This adaptation likely involves the upregulation of stress-protective proteins, changes in amino acid biosynthetic pathways, and modulation of the proteome to optimize survival and productivity in saline environments.

3.5. Lipid productivity

The lipid productivity results summarized in Table 3 indicate that both C. thermophila and A. variabilis generally exhibited higher lipid productivity when cultivated in artificial seawater medium (SWM) than in BG11 medium during the initial 10-day growth period. Notably, Anabaena variabilis demonstrated a substantial increase in lipid productivity in SWM, whereas C. thermophila showed a moderate improvement under the same conditions. This enhanced lipid production under salinity stress highlights the metabolic flexibility of freshwater microalgae, which can redirect carbon flux from protein and carbohydrate synthesis towards lipid accumulation as an adaptive strategy (Singh and Gu 2010;Bellou et al. 2014;Arora et al. 2017). Mechanistically, exposure to elevated salinity can induce osmotic and oxidative stress, triggering pathways that increase the synthesis of storage lipids, particularly triacylglycerols, and upregulate reactive oxygen species-scavenging enzymes and accumulate compatible solutes such as proline and glycine betaine. Such adjustments not only help maintain cellular homeostasis but also support energy storage under stressful environments (Banu et al. 2009;Arora et al. 2017). The observed differences between the two species may reflect variations in their baseline metabolic profiles and specific adaptive responses to high salinity. However, when the cultivation period was extended to 20 days, the lipid productivity in the BG11 medium for both species was comparable to or slightly exceeded that observed in SWM at 10 days. This suggests that while salinity accelerates lipid accumulation, prolonged cultivation in standard medium can also yield substantial lipid quantities as cells transition to the stationary phase and experience nutrient depletion, another well-known trigger for lipid biosynthesis (Zhila et al. 2011;Bellou et al. 2014).

3.6. FTIR analysis

Fourier Transform Infrared (FTIR) spectroscopy was employed to characterize the biochemical composition of C. thermophila and A. variabilis grown under different media conditions. The FTIR spectra revealed distinct absorption bands corresponding to various functional groups, providing insights into the molecular changes associated with growth in artificial seawater medium (SWM). For C. thermophila, the major absorption bands were observed at 3,279.45 cm^-1 (water and protein), 2,924.86 cm^-1 (lipids), 1,637.99 cm^-1 (alkane), 1,541.97 cm^-1 (amide), 1,241.95 cm^-1 (nucleic acid and phosphoryl group), and 1,026.52 cm^-1 (polysaccharide) (Fig. 6). These bands indicate the presence of proteins, lipids, carbohydrates, and nucleic acids, reflecting the overall biochemical profile of the algal cells. Similarly, the FTIR spectrum of Anabaena variabilis showed characteristic peaks at 3,276.58 cm^-1 (water and protein), 2,923.14 cm^-1 (lipids), 1,538.76 cm^-1 (amide), 1,284.43 cm^-1 (alcohol ester), and 525 cm^-1 (alkyl stretch) (Fig. 7). The presence and intensity of these bands suggest that both protein and lipid content are prominent in the biomass, with additional contributions from polysaccharides and nucleic acids. The observed bands are consistent with standard biochemical assignments: broad bands near 3,270-3,280 cm^-1 correspond to O-H and N-H stretching vibrations (indicative of water and proteins), while peaks around 2,920- 2,925 cm^-1 are attributed to C-H stretching in lipids. The amide I and II bands (around 1,638 and 1,542 cm^-1) reflect protein content, and the peaks at 1,242 cm^-1 and 1,027 cm^-1 are associated with nucleic acids/phosphoryl groups and polysaccharides, respectively. In Anabaena variabilis, the 1,284 cm^-1 band is characteristic of alcohol esters, and the 525 cm^-1 band is related to alkyl stretching vibrations. These FTIR results confirmed that both algal species underwent significant biochemical changes when exposed to saline conditions, with notable alterations in lipid- and protein-associated bands. This analysis supports the findings from lipid productivity and protein content measurements, indicating that adaptation to saline media involves the modulation of key cellular macromolecules (Figs. 6 and 7).

3.7. Fatty acid composition of the algal lipids

The lipid profiles of the two algal species (Table 4) exhibited distinct peaks corresponding to specific fatty acids: C14:0 (myristic acid), C16:0 (palmitic acid), C18:0 (stearic acid), C18:1 (oleic acid), and C18:2 (linoleic acid) acids. Notably, C. thermophila grown in SWM displayed a peak for palmitoleic acid, unlike A. variabilis and C. thermophila grown in BG11 medium. Pantetheic acid (C15:0) was exclusively detected in C. thermophila grown in SWM when comparing the lipid profiles of these two species across different nutrient media. C. thermophila and A. variabilis cultivated in BG11 medium contained 3.38% and 2.88% total unsaturated fatty acids and 4.01% and 1.93% polyunsaturated fatty acids, respectively. In contrast, when grown in SWM, C. thermophila and A. variabilis had 2.95% and 3.38% of total unsaturated fatty acids and 0.80% and 1.09% of polyunsaturated fatty acids, respectively. Furthermore, there are notable variations in the percentages of nutritionally important omega-3, omega-6, and omega-7 fatty acids among these species. Although Chlorella cultivated in SWM shows higher lipid productivity than that in BG11, its nutritional quality appears inferior to that of the other species. These findings underscore the need for biotechnological interventions aimed at developing ideal strains for industrial applications or utilizing combinations of diverse algal species in cultivation to optimize biomass production.

The percentage of saturated fatty acids ranged from 80.89% in C. thermophila to 95.33% in A. variabilis, with no monounsaturated fatty acids detected in either species grown in the BG11 medium. In contrast, when grown in SWM, C. thermophila and A. variabilis had 84.43% and 95.51% of the total saturated fatty acids, respectively. Additionally, 4.66% of the total monounsaturated fatty acids were present only in C. thermophila grown in SWM, whereas none were detected in A. variabilis. Typically, saturated and polyunsaturated fatty acids containing 14-18 carbons are utilized as feedstocks for producing high-quality biodiesel (Schenk et al. 2008;Stansell et al. 2012). Thus, the C14 to C18 fatty acid content in lipid profiles serves as a preliminary criterion for assessing oil suitability for biodiesel production. The chemical composition of these algae oils includes palmitic, stearic, oleic, and linoleic acids, all of which are favourable for biodiesel production (Kumar et al. 2016;Santhakumaran et al. 2019).

4. CONCLUSION

This study demonstrates that freshwater microalgae, specifically Chlorella thermophila and Anabaena variabilis, possess remarkable adaptability to artificial seawater, as evidenced by their significant physiological and biochemical changes in response to treatment. Both species not only survived but also showed enhanced lipid productivity under saline conditions, with C. thermophila exhibiting particularly high lipid yields. The shift in fatty acid profiles, featuring valuable omega-3, omega-6, and omega-7 fatty acids, underscores the potential of these microalgae not only for sustainable energy and nutritional applications but also for broader biotechnological uses such as wastewater treatment and industrial effluent remediation. Environmental manipulation, such as cultivation in saline media, induces the synthesis of unique and valuable metabolites. For instance, C. thermophila produces palmitoleic and pantetheic acids exclusively in seawater, demonstrating that growth conditions can be optimized to yield target compounds of industrial relevance. FTIR analysis further confirmed substantial biochemical remodeling under saline stress, as evidenced by intensified lipid-associated absorption bands around 2,926 cm^-1 and 2,854 cm^-1 (C-H stretching) and changes in the amide I region (~1,650 cm^-1), indicative of protein structural modifications. These results highlight the versatility and adaptive capacity of freshwater microalgae for diverse sustainability- focused bioprocesses. These findings not only deepen our understanding of microalgal stress tolerance but also support the feasibility of using saline or brackish water for large-scale microalgal cultivation in the future. This approach can help reduce reliance on freshwater resources, aligning with sustainability goals. The metabolic flexibility of C. thermophila and A. variabilis opens up new opportunities for biofuel production, nutraceutical development, and environmental remediation. Future research should focus on optimizing the cultivation conditions and unraveling the genetic mechanisms underlying these adaptations.