African Journal of
Microbiology Research

  • Abbreviation: Afr. J. Microbiol. Res.
  • Language: English
  • ISSN: 1996-0808
  • DOI: 10.5897/AJMR
  • Start Year: 2007
  • Published Articles: 5128

Full Length Research Paper

Chemical analysis of the biomass of a native strain of Spirulina subsalsa Oersted ex Gomont 1892 (Spirulinaceae) cultivated in low-cost saline medium

Lolymar Romero Maza
  • Lolymar Romero Maza
  • Universidad Politécnica Territorial del Oeste de Sucre "Clodosbaldo Russián", Carretera Cumaná-Cumanacoa, km 4 Sucre, Venezuela.
  • Google Scholar
Miguel Angel Guevara Acosta
  • Miguel Angel Guevara Acosta
  • Instituto Superior de Formación Docente, Salome Ureña, ISFODOSU-FEM, Santo Domingo, Dominican Republic.
  • Google Scholar
Roraysi José Cortez Mago
  • Roraysi José Cortez Mago
  • Instituto Oceanográfico de Venezuela, Universidad de Oriente, Cerro Colorado, Cumaná, Sucre, Venezuela.
  • Google Scholar

  •  Received: 23 May 2019
  •  Accepted: 19 July 2019
  •  Published: 31 August 2019


Spirulina subsalsa, a filamentous cyanobacterium, was first described by Gomont in 1892. This microorganism has been subject to biotechnological evaluations, due to their high content of proteins and pigments. The objective of this study was to analyze the biochemical composition of the biomass of a native strain of S. subsalsa cultivated in low-cost saline medium and harvested in the exponential and stationary phases of growth. The highest protein contents (58.5%) were obtained in the exponential phase; while the highest amounts of carbohydrates (20%), lipids (19.7%), chlorophyll (51.6 μg/ml), total carotenoids (218,215 μg/ml), exopolysaccharides (7.30 ± 0.7 mg/ml) and phycocyanin (25.8 μg/ml) were accumulated in the stationary phase. Additionally, in the biomass of S. subsalsa, the presence of saponins and polyphenols was detected in both growth phases, whereas basic alkaloids and flavonoids were detected only in the stationary phase. This article concludes information on the potential future biotechnological applications of the cyanobacterium strain, S. subsalsa.


Key words: Cyanobacterium, biotechnology, Spirulina subsalsa.


Spirulina subsalsa Oersted ex Gomont is a filamentous cyanobacteria originally described by Gomont (1892, 1893). This microorganism inhabits saline and fresh waters all over the world (Szulbert et al., 2018). In Venezuela, Spirulina has been reported by Rodriguez (2001), Bernal (2002), González et al. (2003) and Petrash et al. (2012).
This  cyanobacterium  forms  mantles  on the substrate,
usually blue-green in color and has sometimes been observed to be part of the cyanobacteria blooms that cause poisoning in flamingos (Ballot et al., 2004) and shrimp (Lightner, 1978); however, there is no evidence that this cyanobacterium produces any cyanotoxin.
The biotechnological potential of S. subsalsa has been little studied, being used as a bioremediator agent of residual  contaminants  (Jiang  et al., 2015), biosensor for the evaluation of the toxicity of estuarine waters (Campanella et al., 2001), and producing bioactive metabolites (Mazur-Marzec et al., 2015). In addition, S. subsalsa is a source of polyhydroxyalkanoates (PHA), which are biopolymers for construction of implants and artificial tissues (Shrivastav et al., 2010).
Spirulina cultures are usually carried out in fresh water and need expensive culture media, due to the inclusion of a large number of analytical grade salts. Between these means, Zarrouk medium was emphasized (Zarrouk, 1966), Spirulina (Aiba and Ogawa, 1977), BG-11 (Rippka, 1988), and some modified media (Amala and Ramanathan, 2013; Kumari et al., 2014a, b). This situation has led to the search for alternative sources of culture media that allow obtaining high yields of biomass at low cost. Furthermore, it is necessary to evaluate new strains of this cyanobacterium, since it has been demonstrated that the responses of microalgae to changes in abiotic factors vary considerably from one species to another, between strains of the same species and even between clones originating from the same unialgal culture, which would be due to morphological and physiological differences, attributable to intraspecific genetic variations (Gómez and González, 2005; Guevara et al., 2016).
The objective of this investigation was to analyze the biomass of a native strain of S. subsalsa cultivated in low-cost saline medium and harvested in exponential and stationary phases of growth.


A native strain of S. subsalsa, isolated from the Clavellino Reservoir, Sucre State, Venezuela (coordinates: between 10° 19 'to 10° 23' Lat. N and between 63° 35 'to 63° 40' Long. O) and deposited in the Algae Germplasm Bank of the Oceanographic Institute of Venezuela, Universidad de Oriente, with the code BGAUDO 161, was cultivated in seawater (9‰) previously treated, according to the methodology of Faucher et al. (1979).
The cultures were carried out in quadruplicate, discontinuously, for 30 days, in 45 cm diameter plastic bags, placed in cylindrical metal frames (Figure 1), containing 100 L of culture medium each with a nitrate concentration of 14 mM, 0.036 mM phosphate, 95.23 mM sodium bicarbonate, 0.0013 mM Fe and 0.0009 mM Mn. The bags were located in a controlled laboratory environment (T: 32 ± 1°C, continuous irradiance of 39 μmol/m2/s provided by 3 white light lamps of 40 W and photoperiod 12:12) and aerated with plastic hoses and diffuser stones. The salinity and nitrate concentration used were selected according to results in previous experiments (Romero et al., 2018).
The cultures were started with inocula previously acclimated to the mentioned environmental conditions. From the beginning of the test and every 48 h, samples were taken from each of the replicas to determine the pH and population growth according to the criteria of Pelizer and Oliveira (2014).
When the culture reached the exponential phase (2 replicas) and stationary (2 replicas), the entire culture was harvested, filtering it in permaline sleeve. The filtrate was used to quantify the exopolysaccharide content according to the methodology of Vicente et al. (2004). The harvested biomass, after several washes with acidulated water (pH 4), was maintained at low temperatures (-20°C) until the moment of realization, in triplicate, the protein analysis, according to Lowry et al. (1951); total lipids, according to Bligh and Dyer (1959) and Pande et al. (1963); carbohydrates, according to Dubois et al. (1956); secondary metabolites according to Domínguez (1973) and Marcano and Hasegawa (2002), and pigments according to Sharma et al. (2014) and Murugan and Rajesh (2014).
Analysis of the results
The data of the values ​​of exopolysaccharides, proteins, lipids, carbohydrates, and Spirulina pigments obtained in the exponential and stationary growth phases were contrasted by a one-way analysis of variance (phases), following recommendations of Sokal and Rolhf (1995).



Growth and pH
Figure 2 shows the population growth of the microalgae S. subsalsa in the low-cost culture medium during the 30 days of the trial. It is observed that during the first 6 days, this microalga was in adaptation phase; after which, the culture entered the exponential growth phase till day 12. Followed by and until the end of the trial, the culture remained in the stationary phase, and no signs of a descent phase were observed. The pH of the cultures was between 9 and 10.2.
The exopolysaccharide content obtained in the S. subsalsa cultures presented significant differences (p <0.05) between the growth phases (Figure 3). The concentration of these exocompounds was 7.30 ± 0.7 and 5.4 ± 0.4 mg/ml on stationary and exponential phase.
Proteins, carbohydrates, lipids and pigments
The contents of proteins, carbohydrates, lipids and pigments of S. subsalsa cultivated in a low-cost saline medium are shown in Table 1. Total proteins showed significant differences (p <0.05) between the phases, reaching their highest contents in the exponential phase (58.5 ± 0.58%). The rest of the analyzed compounds, like the proteins, showed significant differences between the phases (p <0.05), but with the difference that their highest values ​​were obtained in the stationary phase. In this way, carbohydrates, lipids, chlorophyll, phycocyanin and total carotenoids had percentages of 20.0 ± 2.71%, 19.7 ± 1.41%, 51.6 ± 0.64 µg/ml, 25.8 ± 0.40 µg/ml and 218.215 ± 2.27 µg/ml, respectively.
Secondary metabolites
As shown in Table 2, the presence of saponins and polyphenols in the fresh biomass of S. subsalsa was positive in both phases of growth; however, basic alkaloids and flavonoids were only evidenced in the stationary phase (Table 2).



The population growth observed in S. subsalsa in this study is related to the results presented by Rodríguez and Triana (2006), who indicated that in the Spirulina species, the adaptation phase usually lasts between zero and four days, because the microalga is coupled to the culture conditions and has a low specific growth rate. From there, the growth of the microalga gradually increases, entering the phase of exponential growth, where cell multiplication is at its maximum. This phase continues until it reaches its maximum value (days 12-16), where  depletion  of  nutrients  has  been  observed, hence a decrease in growth. The stationary phase begins, due to the decrease in the rate of growth, increased cellular respiration and accumulation of enhancement of toxic wastes. At this point, it is important to take care of the cultivation conditions to extend the phase and avoid unfavorable conditions that might cause the death of the cells (death phase). In the present study, death of the cells did not occur during the present test.
The amount of maximum biomass obtained in this study was 3.1 mg/ml. This biomass value is higher than those reported by Oliveira et al. (1999), where they determined an amount of 2.4 mg/ml at 30°C, in Spirulina platensis and Spirulina maxima. This difference may be due to the temperatures used for the culture, in this work the maximum temperature recorded was 32 ± 1°C.
Volkmann et al. (2008) and Licet et al. (2014) obtained higher biomass than those achieved in this research when cultivating Arthrospira platensis viz. 4.95 and 3.5 mg/ml, respectively.  This difference may be due to the fact that the previous authors used different culture conditions, among these are the irradiance (140 and 390 μmol/m2/s, respectively),  which  were  greater than those
implemented in this research (39 μmol/m2/s).
The pH of the cultures remained between 9 and 10.2, which is within the values ​​reported for this cyanobacterium, according to the criteria of Rincón et al. (2013).
Several studies have reviewed the ability of cyanobacteria to adapt to variations in salinity (Thajuddin and Subramanian, 2005; Nagle et al., 2010; Joset et al., 1996), but not all cyanobacteria are halotolerant (Blumwald   and    Tel-Or,     1982).      The      ability     of cyanobacteria to grow at high concentrations of Na+ may be related to their ability to regulate respiration (Gabbay- Azaria et al., 1992), the flow of Na+ (Molitor et al., 1986) and the production of osmolytic compounds (Reed et al., 1986), which help the cells to withstand the pressure caused by the large amount of sodium ions present in the medium. One of these compounds are the exopolysaccharides, which are exuded into the environment, as an osmoprotective effect.
The   higher   content   of   exopolysaccharides   in   the
stationary phase may be due to the deficiency of nitrogen that occurs in this phase, as indicated by De Philippis et al. (1993) and Otero and Vincenzini (2003). This situation probably contributes to the increase in the C: N ratio, which promotes the incorporation of carbon in polymers (Otero and Vin-cenzini, 2003; Kumar et al., 2007).
The higher contents of exopolysaccharides together with the growth of the cyanobacterium and increase in the pH of the medium, limit the availability of light, which leads to an increase in the content of accessory pigments and phycobiliproteins, thus reducing the phosphorus and nitrogen content, and subsequently the redirection of the cellular metabolism towards the synthesis of carbohydrates (Laloknam et al., 2010; Magro et al., 2018).
Although the characterization of the obtained exopolysaccharides was not satisfied in the development of this work, some authors have managed to isolate and identify some sulphated type of Spirulina polysaccharides, called spirulan calcium Ca-SP, in which antiviral (in vitro and ex vitro) microbiological tests has inhibited the replication of HIV, Herpes simplex, human cytomegalovirus, influenza A virus, mumps and measles (Chamorro et al., 2002). In vitro studies suggest that the polysaccharides, unique to Spirulina, improve the enzymatic activity of the cell nucleus and the synthesis and repair of DNA (Premkumar et al., 2004).
The highest total protein contents of S. subsalsa, cultivated in low-cost saline medium, were obtained in the exponential phase (58.5%). These results may be due to the fact that in this phase, the culture medium did not present nutrient limitations, which favors protein synthesis. In addition, the salinity used in crops  does  not represent extreme stress levels that can interfere with protein accumulation.
Andrade et al. (2018) have observed protein content in Spirulina between 50 and 70%. These differences in biochemical composition, including proteins, are attributed to the variation between genera and species, and in the culture conditions (availability of nutrients, pH, light, temperature) of a particular species (Colla et al., 2007).
The highest contents of lipids (19%) and carbohydrates (20%) were observed in the stationary phase; this could be due to the fact that in this phase, the supply of nutrients usually decreases and the irradiance received by the culture becomes less, motivated by the overshadowing caused by the massive growth of this cyanobacterium, which have been referred to as stimulants of the accumulation of lipids and carbohydrates (Möllers et al., 2014).
Similar to carbohydrates and lipids, the pigment contents showed their highest values ​​in the stationary phase. Chlorophyll a reached contents of 51.6 μg/ml and total carotenoids of 218.215 μg/ml. These results differ from that reported by Marrez et al. (2013), who obtained values of chlorophyll a and total carotenoids ​​of 147.43 and 139.88 μg/ml, respectively for S. platensis. The discrepancies may be due to the dissimilarity of the salinities, since 9‰ was used in the present investigation and the mentioned authors cultivated salinities of 4.83‰.
Senthilkumar and Jeyachandran (2006) reported that the cultivation of cyanobacteria with high salt concentrations significantly affects the chlorophyll content. The  results  of  Ayachi  et  al.  (2007)  supports  this, who observed that the inhibition of chlorophyll synthesis under salt stress is due to a decrease in the energy level caused by the pumping of sodium ions entering the cell, and that also causes a significant inhibition of the chain of electron transport and transport of electrons in the photosystem (PS-II), due to damage in the PS-II reaction center and alterations in the water oxidation complex (Pulz and Gross, 2004).
The highest values ​​of phycocyanin were 25.8 μg/ml, which is lower than those reported (55.37 μg ml-1) by Marrez et al. (2013) obtained in S. platensis and cultivated in SHU medium. It is evident here that the composition of the culture medium exerts influence on the chemical composition of cyanobacteria (Marrez et al., 2014). The optimization of the culture conditions to maximize the accumulation of phycocyanin is due to the fact that this compound is indicated as being responsible for the antioxidant activity of this cyanobacterium (Ahmed et al., 2014).
The presence of saponins and flavonoids in both phases of cultivation, and basic alkaloids and flavonoids in the stationary phase, coincides with that reported by Borowitzka (1995), who proposes that almost all biologically active compounds of interest are secondary metabolites, thereby tending to be more abundant in the stationary phase or in slow-growing crops.
Some reports show that microalgae and cyanobacteria can contain many kinds of phenolic compounds, such as flavonoids (Klejdus et al., 2010). Hamouda and Doumandji (2017) performed the phytochemical analysis of S. platensis, testing with some solvents: acetone, methanol, ether, dichloromethane and hexane, and found the presence of flavonoids, phenolic compounds, alkaloids and cardiac glycosides.
Although no calculations were made to estimate the production costs of S. subsalsa with the culture medium used in this research, it can be inferred that this medium is less expensive, since it only has 5 commercial grade salts, while the zarrouk medium, the most widely used in the cultivation of Spirulina, has 21 analytical grade salts, with which 1000 L of medium can be prepared at a price of US$ 79.5 (Raoof et al., 2006).
The results obtained on the growth, as well as the contents of proteins, lipids, carbohydrates and pigments in the native strain of S. subsalsa when cultivated in low-cost saline medium, permit us to suggest the use of this cyanobacterium in the biotechnological industries with a view to their use as food in aquaculture and in humans, making it necessary to specify the degree of toxicity, since some strains can be toxic in certain culture conditions.


The authors have not declared any conflict of interests.


Ahmed H, Metwally S, Mohamed M, Ahmed E, Nour S, Azmuddin M (2014). Evaluation of antioxidants, pigments and secondary metabolites contents in Spirulina platensis. Applied Mechanics and Materials 625:160-163.


Aiba S, Ogawa T (1977). Assessment of growth yield of a blue-green alga: Spirulina platensis, in axenic and continuous culture. Journal of General Microbiology 102:179-182.


Amala K, Ramanathan N (2013). Comparative studies on production of Spirulina platensis on the standard and newly formulated alternative medium. Science Park 1(1):1-10.


Andrade L, Andrade C, Días M, Nascimento C, Mendes M (2018). Chlorella and Spirulina microalgae as sources of functional foods, nutraceuticals, and food supplements; an overview. Food Processing and Technology 6(1):45-58.


Ayachi S, El Abed A, Dhifi W, Marzouk B (2007). Chlorophylls, proteins and fatty acids amounts of Arthrospira platensis growing under saline conditions. Pakistan Journal of Biological Sciences 10:2286-2291.


Ballot A, Krienitz L, Kotut K, Wiegand C, Metcalf J, Codd G, Pflugmacher S (2004). Cyanobacteria and cyanobacterial toxins in three alkaline lakes of Kenya - Lakes Bogoria. Nakuru and Elmenteita. Journal Plankton Research 26:925-935.


Bernal J (2002). Taxonomy of microalgae on the banks of the Clavellinos Reservoir, Ribero Municipality, Sucre State, Venezuela. Thesis of Degree. Department of Biology, Universidad de Oriente, Cumaná, Venezuela.


Bligh E, Dyer W (1959). A rapid method of total lipid extraction and purification. The National Research Council of Canada. Canadian Journal of Biochemistry and Physiology 37:911-917.


Blumwald E, Tel-Or E (1982). Osmoregulation and cell composition in salt-adaptation of Nostoc muscorum. Archive of Microbiology 132:168-172.


Borowitzka M (1995). Microalgae as a source of pharmaceuticals and other biologically active compounds. Journal of Applied Phycology 7:13-15.


Campanella L, Cubadda F, Sammartino D, Saoncella A (2001). An algal biosensor for the monitoring of water toxicity in estuarine environments. Water Research 35(1):69-76.


Chamorro G, Salazar M, Gomes K, Pereira C, Ceballos G, Fabila L (2002). Update on the pharmacology of Spirulina (Arthrospira), an unconventional food. Archivo Latinoamericano de Nutrition 52:232-240.


Colla L, Reinehr C, Reichert C, Costa A (2007). Production of biomass and nutraceutical compounds by Spirulina platensis under different temperature and nitrogen regimes. Bioresource Technology 98:1489-1493.


De Philippis R, Margheri M, Pelosi E, Ventura S (1993). Exopolysaccharide production by a unicellular cyanobacterium isolated from a hypersaline habitat. Journal of Applied Phycology 5:387-394.


Domínguez X (1973). Methods of photochemical research. Mexico. Limusa pp. 81-226.


Dubois M, Gilles K, Hamilton J, Rebers P, Smith F (1956). Colorimetric method for determination of sugars and related substances. Analytical Chemistry 28(3):350-356.


Faucher O, Coupal B, Leduy A (1979). Utilization of seawater and urea as a culture medium for Spirulina maxima. Canadian Journal of Microbiology 25:752.


Gabbay-Azaria R, Schonfeld M, Tel-Or S, Messinger R, Tel-Or E (1992). Respiratory activity in the marine cyanobacterium Spirulina subsalsa and its role in salt tolerance. Archive of Microbiology 157:183-190.


Gómez P, González M (2005). The effect of temperature and irradiance on the growth and carotenogenic capacity of seven strains of Dunaliella salina (Chlorophyta) cultivated under laboratory conditions. Biological Research 38(2-3):151-162.


Gomont M (1892). Monographie des Oscillariées (Nostocacées homocystées) of Annales des Sciences Naturelles, Botanique Series. Fortin 7(15):91-264.


González E, Ortaz M, Peñaherrera C, Montes E, Matos M, Mendoza J (2003). Phytoplankton from five reservoirs of Venezuela with different trophic states. Limnetica 22(1-2):15-35.


Guevara M, Arredondo-Vega B, Palacios Y, Saéz K, Gómez P (2016). Comparison of growth and biochemical parameters of two strains of Rhodomonas salina (Cryptophyceae) cultivated under different combinations of irradiance, temperature, and nutrients. Journal of Applied Phycology 28(5):2651-2660.


Hamouda I, Doumandji A (2017). Comparative phytochemical analysis and in vitro antimicrobial activities of the cyanobacterium Spirulina platensis and the green alga Chlorella pyrenoidosa: potential application of bioactive components as an alternative to infectious diseases. Bulletin de l'Institut Scientifique 39:41-49.


Jiang L, Pei H, Hu W, Ji Y, Han L, Ma G (2015). The feasibility of using complex wastewater from a monosodium glutamate factory to cultivate Spirulina subsalsa and accumulate biochemical composition. Bioresource Technology 180:304-310.


Joset F, Jeanjean R, Hagemann M (1996). Dynamics of the response of cyanobacteria to salt stress: deciphering the molecular events. Physiology Plant 96:738-744.


Klejdus B, Lojkovo L, Plaza M, Snoblovo M, Stěrbovo D (2010). Hyphenated technique for the extraction and determination of isoflavones in algae: ultrasound-assisted supercritical fluid extraction followed by fast chromatography with tandem mass spectrometry. Journal of Chromatography A 1217:7956-7965.


Kumar A, Mody K, Jha B (2007). Bacterial exopolysaccharides- a perception. Journal of Basic Microbiology 47:103-117.


Kumari A, Kumar A, Pathak A, Guria C (2014a). Carbon dioxide assisted Spirulina platensis cultivation using NPK-10:26:26 complex fertilizers in sintered disk chromatographic glass bubble column. Journal of CO2 Utilization 8:49-59.


Kumari A, Sharma V, Pathak A, Guria C (2014b). Cultivation of Spirulina platensis using NPK-10:26:26 complex fertilizer and simulated flue gas in sintered disk chromatographic glass bubble column. Journal of Environmental Chemical Engineering 2:1859-1869.


Laloknam S, Bualuang A, Boonburapong B, Rai V, Takabe T, Incharoensakdi A (2010). Salt stress induced glycine-betaine accumulation with amino and fatty acid changes in cyanobacterium Aphanothece halophytica. Asian Journal of Food and Agro-industry 3:25-34.


Licet B, Guevara M, Lemus N, Freites L, Romero L, Lodeiros C, Arredondo - Vega B (2014). Growth and biochemical composition of Arthrospira platensis (Cyanophyta Division) cultivated at different salinities and nitrogen sources. Boletín del Instituto Oceanográfico de Venezuela 53(1):3-13.


Lightner D (1978). Possible toxic effects of the marine blue-green alga, Spirulina subsalsa, on the blue shrimp, Penaeus stylirostris. Journal of Invertebrate Pathology 32(2):139-150.


Lowry O, Rosebrough N, Farr J, Randall, R (1951). Protein measurement with the Folin phenol reagent. US National Library of Medicine. National Institutes of Health. The Journal of Biological Chemistry 193(1):265-275.


Magro F, Margarites C, Reinehr O, Gonçalves C, Rodigheri G, Costa J, Colla L (2018). Spirulina platensis biomass composition is influenced by the light availability and harvest phase in raceway ponds. Environmental Technology 39(14):1868-1877.


Marcano D, Hasegawa M (2002). Organic phytochemistry Council of Scientific and Humanistic Development, Central University of Venezuela P 588.


Marrez D, Naguib M, Sultan Y, Daw Z, Higazy A (2014). Evaluation of chemical composition for Spirulina platensis in different cultures media. Research Journal of Pharmaceutical, Biological and Chemical Sciences 5(4):1161-1171.


Marrez D, Naguib M, Sultan Y, Daw Z, Higazy A (2013). Impact of culturing media on biomass production and pigments content of Spirulina platensis. International Journal of Advanced Research 1:951-961.


Mazur-Marzec H, BÅ‚aszczyk A, Felczykowska A, Hohlfeld N, Kobos J, ToruÅ„ska-Sitarz A, Devi P, Montalvão P, D'souza L, Tammela P,


Mikosik A, Bloch S, Nejman-Faleńczyk B, Węgrzyn G (2015). Baltic cyanobacteria - a source of biologically active compounds. European Journal of Phycology 50:343-360.


Molitor V, Erber W, Peschek G (1986). Increased levels of cytochrome oxidase and sodium-proton antiporter in the plasma membrane of Anacystis nidulans after growth in sodium enriched media. FEBS Letters 204:251-256.


Möllers K, Cannella D, Jørgensen H, Frigaard N (2014). Cyanobacterial biomass as carbohydrate and nutrient feedstock for bioethanol production by yeast fermentation. Biotechnology for Biofuels 7:64. doi:10.1186/1754-6834-7-64.


Murugan T, Rajesh R (2014). Cultivation of two species of Spirulina (Spirulina platensis and Spirulina platensis var lonar) on sea water medium and extraction of C-phycocyanin. European Journal of Experimental Biology 4(2):93-97.


Nagle V, Mhalsekar N, Jagtap T (2010). Isolation, optimization and characterization of selected cyanophycean members. Indian Journal of Marine Science 39:212-218.


Oliveira M, Monteiro M, Robbs P, Leite S (1999). Growth and chemical composition of Spirulina maxima and Spirulina platensis biomass at different temperatures. Aquaculture International 7:261-275.


Otero A, Vincenzini M (2003). Extracellular polysaccharide synthesis by Nostoc strains as affected by N source and light intensity. Journal of Biotechnology 102:143-152.


Pande S, Parvin K, Venkitasubramanian T (1963). Microdetermination of lipids and serum total fatty acids. Analytical Biochemistry 6:415-423.


Pelizer L, Oliveira I (2014). A method to estimate the biomass of Spirulina platensis cultivated on a solid medium. Brazilian Journal of Microbiology 45(3):933-936.


Petrash D, Gingras M, Lalonde S, Orange F, Pecoits E, Konhauser K (2012). Dynamic controls on accretion and lithification of modern gypsum-dominated thrombolites, Los Roques, Venezuela. Sedimentary Geology 245-246:29-47.


Premkumar K, Abraham S, Santhiya S, Ramesh A (2004). Protective effect of Spirulina fusiformis on chemical-induced genotoxicity in mice. Fitoterapia 75(1):24-31.


Pulz O, Gross W (2004). Valuable products from biotechnology of microalgae. Applied Microbiology Biotechnology 65:635-648.


Raoof B, Kaushik B, Prasanna R. (2006). Formulation of a low-cost medium for mass production of Spirulina. Biomass and Bioenergy 30:537-542


Reed R, Borowitzka L, Mackay M, Chudek J, Foster R, Warr S, Moore D, Stewart W (1986). Organic solute accumulation in osmotically stressed cyanobacteria. FEMS Microbiology Reviews 39:51-56.


Rincón D, Semprun A, Dávila M, Velásquez H, Morales E, Hernández J (2013). Production of Spirulina maxima flour to be used as an ingredient in the elaboration of fish diets. Zootecnia Tropical 31(3):187-191.


Rippka R (1988). Isolation and purification of cyanobacteria. Methods Enzymology 167:3-27.


Rodríguez A, Triana F (2006). Evaluation of the pH in the culture of Spirulina spp. (= Arthospira) under laboratory conditions. Science Faculty. Pontifical Javeriana University. Bogota Colombia. [Online document] 



Rodríguez G (2001). The Maracaibo System, Venezuela. En Seeliger, U. & Kjerfve, B (Eds.), Coastal Marine Ecosystems of Latin America. Editorial Springer, Alemania.


Romero L, Guevara M, Bernal J (2018). Crecimiento y pigmentos de Spirulina subsalsa cultivada a diferentes salinidades y concentraciones de nitrógeno. Revista Mutis 8(2):25-36.


Senthilkumar T, Jeyachandran S (2006). Effect of salinity stress on the marine cyanobacterium Oscillatoria acuminata Gomont with reference to proline accumulation. Seaweed Research and Utilization 28:99-104.


Sharma G, Kumar M, Irfan M, Dut N (2014). Effect of carbon content, salinity and pH on Spirulina platensis for phycocyanin, allophycocyanin and phycoerythrin accumulation. Journal of Microbial and Biochemical Technology 6(4):202-206.


Shrivastav A, Mishra S, Mishra S (2010). Polyhydroxyalkanoate (PHA) synthesis by Spirulina subsalsa from Gujarat coast of India. International Journal of Biology Macromolecular 46(2):255-260.


Sokal R, Rohlf F (1995). Biometry. The Principles and Practice of Statistics in Biological Research. 3rd Edition, W.H. Freeman and Co., New York.


Szulbert K, Wiglusz M, Mazur-Marzec H (2018). Bioactive metabolites produced by Spirulina subsalsa from the Baltic Sea. Oceanologia 60(3):245-255.


Thajuddin N, Subramanian G (2005). Cyanobacterial biodiversity and potential applications in biotechnology. Current Science 89:47-57.


Vicente V, Ríos-Leal E, Calderón G, Cañizares R, Olvera R (2004). Detection, isolation, and characterization of exopolysaccharide produced by a strain of Phormidium 94a isolated from an arid zone of Mexico. Biotechnology Bioengineering 85(3):306-310.


Volkmann H, Imianovsky U, Oliveira J, Sant'anna E (2008). Cultivation of Arthrospira (spirulina) platensis in desalinator wastewater and salinated synthetic medium: protein content and amino-acid profile. Brazilian Journal of Microbiology 39:98-101.


Zarrouk C (1966). Contribution of the study of a cyanophycea. Influence of various physical and chemical factors on the growth and photosynthesis of Spirulina maxima (Setch and Gardner) Geitler. Trab. Doct. University of Paris, Paris, Francia P 41.